This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.
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.
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.
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.
2. Surgical Approach
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.
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 aortic valve. In a healthy wild type mouse left ventricular pressures of 90 – 110 mmHg are expected with maximal dP/dt of 8,000 – 12,000 mmHg/sec (see Table 1 for ranges of normal hemodynamic parameters). The normal ranges provided in the table are based on values obtained from wild type C57BL/10 and C57BL/6 mice; however, it is important to note that there are significant differences between strains 21. The procedure outlined here focuses on the use of an apical stab incision to position the catheter within the left ventricle. Another popular approach is to insert the catheter retrograde through the aortic valve following catheter insertion in the right carotid artery. The retrograde approach as the advantage in that it can be performed with a closed chest, resulting in the maintenance of normal intra-thoracic pressures; however, animals are frequently ventilated during this procedure which limits this advantage. The closed chest approach limits control over the catheter orientation within the ventricle, while the open chest approach provides greater ability to maneuver the catheter within the ventricle. A potential problem with retrograde insertion of the catheter is the potential for outflow track obstruction. The diameter of the mouse aorta varies from 0.8 to 1.2 mm 22,23, the diameter of commercially available pressure-volume catheters ranges from 0.33 (1.0 French) to 0.47 mm (1.4 French). The relative sizes of these catheters in the context of a large and small aorta are depicted in Figure 2. The fraction of the outflow tract blocked by the catheter crossing the aorta can become a significant issue in smaller hearts and should be taken into account when performing PV loop studies in small hearts. There are several artifacts of measurement that can complicate the analysis of PV data, one of the most common is catheter entrapment. This is evident as a spike in pressure at the end of systole likely resulting from direct compression of the pressure transducer by a papillary muscle or other dynamic structure within the ventricle (Figure 3). This is problematic, because most methods for determining systolic function use the maximum pressure. The systolic pressure and maximum derivative of pressure (dP/dt) from data sets with catheter entrapment need to be examined closely and the analysis may need to be altered to obtain meaningful data.
Pressure-Volume loops can be used in a wide variety of protocols to assess cardiac function. These include assessments of cardiac reserve via β-adrenergic stimulation 12,14,16,17. A wide variety of drugs may be infused to assess any acute effects on cardiac physiology. Control of the airway also allows for the administration of altered gas mixtures allowing the effects of hypoxia and/or acidosis on cardiac function to be addressed directly 24-27. Furthermore, analysis of this PV data can also provide a detailed assessment of the passive properties of the left ventricle, which can be altered significantly in various disease states 12.
Parameter | Normal Range |
Systolic Pressure | 90 – 110 mmHg |
Diastolic Pressure | 4 – 10 mmHg |
Maximum dP/dt | 8,000 – 12,000 mmHg/sec |
Minimum dP/dt | -8,500 – -12,000 mmHg/sec |
Tau | 5 – 6 sec |
Heart Rate | 550 – 600 bpm |
Cardiac Output | 10 – 13 ml/min |
Ejection Fraction | 40 – 60% |
Table 1. Normal Values for Selected Hemodynamic Parameters.
Figure 1: Representative Pressure-volume Loops from a Wild Type C57BL/10 Mouse. Representative data that was collected using the procedures described in this manuscript. Please click here to view a larger version of this figure.
Figure 2: Schematic Representation of the Importance of Aortic Diameter and Potential Outflow Track Obstruction. Schematic drawing of the relative sizes of the mouse aorta and the commercially available pressure-volume catheters for mice. These data underscore the importance of considering outflow track obstruction when assessing smaller mice using a retrograde catheter insertion approach. Please click here to view a larger version of this figure.
Figure 3: Pressure-volume Loops with Catheter Entrapment. These data represent an example of a pressure spike in late systole, likely resulting from direct compression of the catheter by an internal cardiac structure. Declines in end-diastolic volume (A) result in an increase in the pressure peak as the ventricle gets smaller, however the systolic pressure prior to the peak declines as expected, depicted as a dashed line in B. (C and D) represent the pressure-volume loops (C) and pressure-time (D) curves following abdominal compression. As the left ventricular volume increases, the pressure spike gets smaller and the underlying increases in systolic function becomes apparent; dashed line in D. Please click here to view a larger version of this figure.
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. This is important as too little ventilation will result in respiratory acidosis and too much ventilation will lead to respiratory alkalosis, both conditions that will alter cardiac function. A simple way of optimizing the respiratory rate is to take cues from the mouse's respiratory effort and use the lowest respiratory rate that eliminates respiratory effort from the anesthetized mouse.
A critical feature of PV loop analysis is to maintain a sufficient level of anesthesia without significant depression of cardiac function. There are many anesthetic regimes that can be successfully applied to perform PV loop analysis. These can be divided into two broad categories: inhalant and injectable anesthetics. Within this latter group are a variety of mixtures that can be used, however, many of these mixtures have potential cardio-depressive effects 19. Of the injectable cocktails, urethane based anesthetics have the least cardio-depressive effects 4,19. Take care to limit exposure of personnel to urethane as it is a suspected carcinogen 28. Isoflurane and sevoflurane are the currently available inhalant anesthetic agents. Both of these agents have cardio-depressive effects at doses sufficient for surgical manipulation. Importantly, inhaled anesthetic agents can be titrated to effect. This allows a higher analgesic dose during the surgical preparation and then a lower sedative dose for the measuring of cardiac function, thus minimizing the cardio-depressive effects of these compounds.
If the hemodynamic parameters are below the normal levels, there are several common causes. First is low blood volume, secondary to blood loss or evaporation. This complication is generally addressed with the fluid administration outlined in section 2.7. As noted in section 1.5, maintenance of core body temperature is also very important for the assessment of hemodynamic function. Thus careful monitoring is essential, a digital feedback system can be useful for maintaining core body temperature during recording. Ensuring that the anesthetic has been properly reduced during the measurement phase is also an important aspect of improving cardiac performance.
Measuring load-independent parameters of contractility is one of the primary advantages of PV loop analysis. The real-time simultaneous collection of pressure and volume data provides the unique capacity to measure the hemodynamic changes in response to varying loading conditions on the heart. This analysis allows the contractile function of the heart to be isolated from the actions of the vessels. Reductions in preload through occlusion of venous return have been used to assess load-independent measures of contractility in many animal models and man 4,7,12-16,18,29-32. In small rodents, this procedure results in a curvilinear end-systolic pressure-volume relationship 4,31,33. This likely results from the reduction in coronary perfusion that dramatically accelerates the decline in systolic function 34. In the mouse, 2 – 3 sec of gentle abdominal compression results in a rightward shift of the PV loops. This results from both an increase in afterload and preload 12. Prolonged increases in afterload result in an increase in contractile function, a phenomena called the Anrep effect 35. However the short duration of the abdominal compression used in these studies indicates that the Anrep effect does not affect cardiac function with this procedure. In other studies, acute aortic constriction in dogs was demonstrated to increase contractile function, but this was hypothesized to result from the effects of reduced systemic perfusion 36. Again the duration and severity of the disruption of aortic flow resulting from abdominal compression, as described here, is not sufficient to significantly increase contractile function secondary to hypo-perfusion of systemic tissues. Analysis of PV loops obtained by abdominal compression align well with PV loops obtained shortly after caudal vena cava occlusion. Together, these observations demonstrate that the transient abdominal compression described here does not significantly alter the contractility of the heart. Furthermore, this procedure provides an important method to assess the passive properties of the heart over a broader range of end-diastolic volumes.
The analysis of load-independent measures requires the selection of specific PV loops to be included in the analysis. It is critically important that this be done consistently across an experimental data set. Positive pressure ventilation creates a hemodynamic artifact through respiratory dependent alterations of preload to the left ventricle 37. To avoid this artifact, PV loops should be collected during short periods of apnea (3 – 4 sec). Pauses in respiration are particularly useful as they provide better control of experimental changes in loading on the heart. It is important to keep these periods of apnea short to avoid hypoventilation. The procedures described in this manuscript collect load-independent data from two distinct procedures, caudal vena cava occlusion and abdominal compression, collected at roughly the same time. The loops isolated from these two procedures should be combined and analyzed together as they both measure the function of the same heart under, roughly, the same conditions. There are several principles to consider when selecting loops for analysis. Avoid arrhythmic beats. The beat following a dropped beat is always abnormally large and premature beats are abnormally small; both will disrupt the analysis of the data. Avoid beats where the pressure is declining but volume is constant. These are common in the mouse following the occlusion of the caudal vena cava and are likely secondary to poor perfusion of the myocardium. Finally, include only data from beats directly after the change in loading; the beats during the recovery period are likely affected by alterations in sympathetic nervous activity secondary to changes in systemic blood pressure resulting from manipulation of the loading conditions of the heart.
PV loop analysis provides an extremely detailed assessment of cardiac function. When applied in conjunction with the genetic flexibility and low housing costs of the mouse it can provide a practical means of assessing cardiac physiology on the molecular level. There are several important limitations that must be considered when deciding to perform these assays. First, this is an invasive procedure in which mice are anesthetized, which may impact important aspects of cardiac function. Furthermore, interpretation of PV loop data requires a detailed understanding of cardiac physiology both to identify patterns within the data and potential confounding variables. Additionally, because these assays are terminal they cannot be used to assess the same mouse repeatedly. The ventricular volumes derived from PV catheters tend to be less accurate than the anatomical ventricular volumes provided by MRI. This is not surprising as PV catheters model the ventricle as a cylinder, which is clearly an estimate of the overall volume of the ventricle. The real strength is the ability to collect this volume information at a high frequency, thus allowing beat-to-beat analysis of changes in ventricular volume.
The collection and analysis of PV data from the mouse can be challenging, but the method provides information regarding cardiac function that is not available through any other methodology. This procedure provides the most complete picture of cardiac function available and its utilization in the murine model will provide an important platform for the determination of the molecular underpinnings of complex cardiac disease states such as heart failure and inherited cardiomyopathies. This manuscript provides detailed information on the most critical aspects of performing this procedure. However, like all complicated procedures, this requires practice to build the microsurgical skills that are necessary to successfully perform these experiments.
The authors have nothing to disclose.
The author would like to acknowledge funding from NHLBI (K08 HL102066 and R01 HL114832).
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 |