Name: cardiac pressure-volume loop analysis using conductance catheters in mice
Uploaded: 2015-09-17T14:00:00.000+00:00
Duration: 8 min 15 s
Description: Watch this Scientific Journal Video about Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice at JoVE.com

This work is supported by the American Heart Association 14FTF20370058 (DMA) and NIH T32 HL007101-35 (DMA).

1. Conductance Catheter Preparations and Pressure Calibration
<ol>
	<li>Connect conductance catheter to the hemodynamic catheter module. Electronically calibrate pressure and volume measurements by recording preset pressure and volume set on the catheter module. Record a tracing of 0 mm Hg and 25 mm Hg (Figure 1A) and assign voltages to both pressure tracings (Figure&#160;1B and 1C). Similarly, record a volume tracing of 5 RVU and 25 RVU (Figure 1D) and assign voltages to both volume tracings (1E and 1F).</li>
	<li>Confirm the electronic pressure calibration with a manual pressure calibration, using a mercury column sphygmomanometer. Fit the sphygmomanometer cuff port with a 3-way stopcock. Fill a 3-port hemostasis valve system, often used for coronary angioplasty, with RT water using the side port.
	<ol>
		<li>Place the tip of a conductance catheter into the fluid filled hemostasis valve and gently secure without kinking catheter. Connect the hemostasis valve to the sphygmomanometer and inflate to 200 mmHg and lock 3-way stopcock. Examine whether the measured pressure corresponds with the pressure inflated on the sphygmomanometer.</li>
	</ol>
	</li>
	<li>Place catheter in saline heated to 37 &#176;C that is at the level of the operating field and measure pressure. Adjust pressure control until the recorded pressures is at zero.</li>
</ol>
2. Anesthesia/Intubation
<ol>
	<li>Administer ketamine/xylazine (80-100/10 mg kg-1) as an intraperitoneal injection. 
	Note: Alternative anesthetic agents can be used. An extensive list of anesthetics are provided in previous reviews of this technique 11,12. Proper anesthetization can be confirmed by gentle tail pinch.&#160;</li>
	<li>Once anesthetized, shave the neck and chest with hair clippers and place on a heated pad. Maintain the mouse rectal temperature at 36.5-37.5 &#186;C. Low body temperatures will result in depressed heart rates. Apply ointment to the eyes to prevent dryness</li>
	<li>Make a midline incision in the neck and dissect the tracheal muscles away to expose the trachea. Place an endotracheal tube through the mouth, while visualizing the trachea to ensure intubation, and connect to the respirator.</li>
	<li>Maintain mouse on the ventilator during the procedure. Set ventilator settings based on animal weight as previously described 11. Tidal Volume (ml)= 6.2 x (animal weight in kilograms) 1.01 and Respiratory rate= 53.5 x (animal weight in kilograms)-0.26.</li>
</ol>
3. Placement of Conductance Catheter in LV chamber
<ol>
	<li>Carotid approach
	<ol>
		<li>To ensure sterility, two sets of sterile surgical instruments are used- one for initial skin incision and one to operate in the thoracotomy. Instruments should be decontaminated using a dry sterilizer between animals during an individual surgical session and autoclaved at the end of each surgical day.</li>
		<li>&#160;After the skin has been decontaminated with three cycles of a Chlorhexidine + alcohol skin scrub (0.5% chlorhexidine/70% alcohol solution), make an incision over the right carotid from mandible to sternum. Dissect surrounding tissue to expose the right carotid, and cut the vagus nerve which runs adjacent to the carotid.</li>
		<li>Place a sterile 6-0 silk suture around the distal end (away from the chest) of the carotid artery, tie and secure. Place two additional sutures beneath the carotid artery proximal (closer to the chest) to the first suture, loosely tying the middle suture. Gently pull the proximal suture and secure by clamping it to the skin. Ensure that the carotid artery has been clamped both proximally and distally before proceeding.</li>
		<li>Make a small incision in the R. carotid artery, proximal to the first suture, and extend longitudinally toward the chest.</li>
		<li>Insert the conductance catheter tip, previously soaked in warm saline for 30 min, into the vessel through the incision and secure the catheter using the middle suture.</li>
		<li>Gently advance the catheter in the left ventricle through the carotid, while watching the pressure-volume loop tracing to ensure correct placement. 
		Note: Optimal placement of the catheter should yield pressure-volume loops that appear rectangular (See Figure 2). If any resistance to the passage of the catheter is encountered, gently pull back and advance again with gentle pressure. Gentle rotation of the catheter may help with placement into the left ventricle. Forcing the conductance catheter can lead to serious cardiovascular complications or damage to the catheter.</li>
		<li>Record baseline pressure-volume loops ~10 min&#160;after catheter placement and achieving a steady state (Figure 2).</li>
	</ol>
	</li>
	<li>Apical Approach
	<ol>
		<li>In an anesthetized and ventilated mouse, make an incision from the xiphoid process and cut through the chest wall laterally until the diaphragm is visible.</li>
		<li>Cut though the diaphragm and visualize the apex of the heart.</li>
		<li>Insert the conductance catheter into the apex of the left ventricle through a needle stab wound (using a 25-30 G needle), until the proximal electrode is just inside the ventricle.</li>
		<li>Record baseline pressure-volume loops ~10 min&#160;after catheter placement and achieving a steady state.</li>
	</ol>
	</li>
</ol>
4. Varying Afterload Using Transient Aortic Occlusion
<ol>
	<li>To perform transient aortic occlusion, make a small horizontal incision in the upper chest and dissect surrounding tissue to expose the transverse aorta.</li>
	<li>Place a 6-0 silk ligature underneath the transverse aorta. After pressure-volume loops have returned to baseline, clasp both ends of the suture with a needle clamp, gently and slowly raise the suture over 1-2 sec, and slowly release tension.</li>
	<li>Repeat this procedure, until three separate optimal recordings are made from the same animal. 
	Note: Optimal recordings should have at least 5 pressure volume loop cycles and a steady rise in end systolic pressures during the application of tension on the suture (Figure 3A and 3C).</li>
</ol>
5. Varying Preload Using Transient Inferior Vena Cava Occlusion (IVC)
<ol>
	<li>To perform transient inferior vena cava occlusion, make a horizontal incision below the xiphoid process, underneath the diaphragm to expose the IVC.</li>
	<li>Place a 6-0 silk ligature underneath the IVC. After pressure-volume loops have returned to baseline, clasp both ends of the suture with a needle clamp; gently and slowly raise the suture over 1-2 sec, and slowly release tension. IVC occlusion can also be performed by the gentle application of pressure using a cotton tipped swab.</li>
	<li>Repeat this procedure, until three separate optimal recordings are made from the same animal. 
	Note: Optimal recordings should have at least 5 pressure volume loop cycles and a steady drop in left ventricular end-diastolic pressures during the application of tension on the suture (Figure 4A and 4B).</li>
</ol>
6. Saline Calibration
<ol>
	<li>At the conclusion of the study, a parallel conductance (Vp) value can be obtain by injecting a 10 &#956;l bolus of hypertonic saline (15%) into the animal through the jugular vein (Figure 5A and 5B). 
	Note: This bolus will cause an apparent increase in volume with no change in pressure. This apparent change in volume is the result of a change in blood pool conductance rather than due to an actual increase in volume. A transient decline in dP/dtmax can be observed, as hypertonic saline has a negative inotropic effect 13. The calculated Vp can be entered into pressure volume loop analysis software along with the cuvette calibration parameters to and convert from RVUs in microliters.</li>
</ol>
7. Cuvette Calibration
<ol>
	<li>To perform a cuvette calibration, place a cuvette with wells of known diameters provided by manufacturer on a heating pad or a water bath warmed to 37 &#186;C. Fill the first 4&#8211;5 holes with fresh warm heparinized blood from mice undergoing hemodynamic assessments. 
	Note: A cuvette calibration allows for accurate assessment of left ventricular blood pool using the mouse blood and allows for conversion of volume data from RVUs to microliters.</li>
	<li>Insert the conductance catheter into the first well, until all the electrodes are submerged. Gently move the catheter in the well, which will generate varying RVUs.</li>
	<li>Record the conductance changes in the volume channel in RVUs. Select the highest RVU for the calibration. 
	Note: The volume of the wells can either be calculated by 1) using the equation for the volume of a cylinder, where the radius is that of the cuvette well and the length is based on the length between the inner two sensing electrodes or 2) checking in the manufacturer&#39;s instructions. The conductance output can be correlated with the known volumes to develop a calibration equation that converts the data from RVUs into microliters 11.</li>
</ol>
8. Euthanasia
<ol>
	<li>&#160;At the conclusion of the protocol, mice are euthanized using cervical dislocation while under anesthesia.</li>
	<li>&#160;In order to ensure death, mice having undergone cervical dislocation must undergo a secondary method of euthanasia. We use exsanguination under anesthesia, with harvesting of cardiac tissue for experimentation, or bilateral thoracotomy under anesthesia.</li>
</ol>
9.&#160;Data Analysis using Pressure Volume Loop Analysis Software
<ol>
	<li>Calculate Vp from Saline Calibration
	<ol>
		<li>Select pressure and volume loops obtained during the ejection of hypertonic saline (Figure 5A and 5B)</li>
		<li>Export loops to pressure volume analysis software. Choose option for Saline calibration (Figure 5C)</li>
		<li>Record the calculated Vp value (Figure 5D). 
		Note: The Vp value is calculated by identifying the intersection of the 1) End-diastolic volume vs. End-systolic volume from the saline calibration and 2) End-diastolic volume = End-systolic volume line. The intersection of these lines provides the Vp, which is calculated by the pressure volume loop analysis software.</li>
	</ol>
	</li>
	<li>Enter conductance to volume (RVU) relation in volume channel options (Figure 1F)</li>
	<li>Measure Baseline Pressure-Volume loop relation
	<ol>
		<li>Select 8-10 cardiac cycles from pressure and volume channels once steady state has been achieved (Figure 2A) and export to analysis software. Identify 5-6 end-expiratory loops (Figure 2B)</li>
		<li>Use the Vp value to correct for parallel conductance. Choose the &#8220;Steady State&#8221; option and generate a hemodynamic summary table (Figure 2C)</li>
	</ol>
	</li>
	<li>Measure Pressure-Volume loop Relation during Aortic Constriction
	<ol>
		<li>Select 8-10 cardiac cycles from pressure and volume channels that correspond to aortic constriction prior to the increase of end-diastolic pressures (Figure 3A) and export to analysis software. Identify 5-6 end-expiratory loops (Figure 3C)</li>
		<li>Select &#8220;Contractility&#8221; analysis (Figure 3B), which will calculate the End-Systolic Pressure Volume Relation (ESPVR)</li>
	</ol>
	</li>
	<li>Measure Pressure-Volume loop Relation during IVC Constriction
	<ol>
		<li>Select 8-10 cardiac cycles from pressure and volume channels that correspond to IVC constriction (Figure 4A) and export to analysis software. Identify 5-6 end-expiratory loops (Figure 4B)</li>
		<li>Select &#8220;Contractility&#8221; analysis (Figure 3B), which will calculate Preload Recruitable Stroke Work (PRSW) (Figure 4C), Maximum dP/dt vs EDV (Figure 4D), as well as the ESPVR and End-diastolic pressure volume relation (Figure 4E)</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We describe a method for perfoming pressure-volume loop analysis using a conductance catheter in mice, to derive comprehensive analyses of both cardiac contractility and relaxation. Suga, Sagawa and colleagues utilized pressure-volume loops to define measures of cardiac contractility, specifically the slope of the ESPVR, or the end-systolic elastance (Ees), and Emax. Elastance, defined by the ratio of pressure to volume (P/V), varies over the duration of systole. During each systole, the instantaneous elastance is dependent on heart rate and cardiac contractility, but is largely independent of preload or afterload 3,17. Thus, the peak elastance or Emax is used to define cardiac contractility that is mostly independent of loading conditions from individual cardiac cycles18. A closely related contractility term, Ees is defined by the slope of the ESPVR over a series of cardiac cycles in a stable condition. While Ees appears linear over a limited range of loads, Ees can be curvilinear and curvilinearity is correlated with contractile state19. &#160;An increase in Ees or Emax indicates increased contractility and a decrease denotes diminished contractility. In addition to Ees or Emax, pressure-volume loop data can be used to derive alternative contractility indices such as: dP/dtmax-EDV relation 20, preload-recruitable stroke work (PRSW) 21 or maximal power-EDV relations 22. These alternative parameters examine the cardiac response over a range of preloads and can be obtained with IVC constriction. It is worth noting that while ESPVR is relatively load-independent, this is not absolute. There are differences in ESPVRs derived from aortic or IVC constriction 23, with aortic constriction having a larger impact on the duration of systole and the extent of shortening 8.
Similar to the ESPVR, the end-diastolic pressure-volume relation (EDPVR) provides a load independent measure of cardiac compliance. This relation is derived by identifying end-diastolic pressures over a range of loading conditions, which is then fit to either an exponential model defined as P=&#945;(e&#223;V-1) +P0 (&#945; is a stiffness and scaling coefficient,&#223;=chamber stiffness coefficient and P0 = pressure at a 0 volume)8 or a linear model (shown in Figure 4C). Pressure-volume loop analysis can provide additional information about diastolic function. A measure of active relaxation is derived from the decline of ventricular pressure during isovolumic relaxation. The monoexponential decay from the peak rate of relaxation to the onset of LV filling is expressed as time constant t8.
The measurement of parallel conductance is critical to the assessment of cardiac volume. While we have described the use of saline calibration to assess parallel conductance, a growing body of literature has identified alternative methods to assess parallel conductance. Saline calibration utilizes Baan&#8217;s equation 9, in which&#160;&#945; is uniform field correction factor. However, the moving heart wall changes the electrical field thus raising the issue of a dynamic &#945; constant 24. Moreover, the contribution of the ventricular wall to conductance varies during systole and diastole, however the saline calibration uses a fixed value for parallel conductance 24. To address this, the concepts of time varying field correction, termed Wei&#8217;s equation, and instantaneous parallel conductance, termed &#8220;admittance&#8221;, have been devised 25. Newer micro-catheters that measure admittance have been created and successfully utilized in cardiac injury models 1. These technologies represent a significant advance in the assessment of pressure-volume loop analysis.
The systolic and diastolic parameters obtained from pressure-volume loop analysis provides a comprehensive evaluation of cardiac function. The accuracy and precision of these analyses depend on attention to experimental detail. To obtain good quality cardiac pressure-volume loops in mice, a skillful operator is crucial. Additionally, care must be taken in the selection of anesthesia, proper ventilation, body temperature, and positioning of the catheter into the LV. The correct analysis of obtained data will depend on consistent instrument, saline and cuvette calibration. These aspects are highlighted in these written methods and the accompanying video should provide a framework upon which to embark on these cardiac physiology studies.
Trouble Shooting
1. Hypotension or bradycardia in a basal state: Normal mouse blood pressure and heart rate have been summarized by in previous reviews of this topic 11.
a) Ensure that body temperature is above 36 &#186;C using a rectal thermometer. If below 36&#160;&#186;C, a heating pad or heating lamps can be used to raise the mouse body temperature.
b) Assess for bleeding during the surgical procedure. Hemostasis can be achieved with manual pressure or with cautery. Significant volume loss can be treated with saline fluid boluses.
c) Assess whether the mouse is over-anesthetized. If this is suspected, saline fluid boluses can be used to treat hypotension. Of note, ketamine/xylazine, which is used for the demonstrated experiments, can be cardiodepressive. Alternatively, inhaled anesthesia using isoflurane (Induction 3-4%, maintenance 1.5% mixed with 95% oxygen &#38; 5% CO2) can be substituted. Alternative anesthetic agents such as urethane (800 mg kg-1)/etomidate (5-10 mg kg-1)/ morphine (2 mg kg-1) or pentobarbital sodium (40-80 mg kg-1) can be administered intraperitoneally11.
2. Noise in the pressure or volume channels: This may result from electromagnetic interference or a broken/dirty conductance catheter. Make a careful assessment of electronic devices that may contribute to interference. If noise persists, examine the conductance catheter under a microscope to assess for adherent material or damage to the catheter tip. If available, try a fresh conductance catheter to see if it remedies the problem.
3. Volume recording drifts over time: This may be due to an inadequate warm-up time for the hemodynamic module. Allow the module to warm up for 30 min before calibration steps and analysis.
5. Volume reading is larger than expected: This can occur if the measured volume has not been corrected for parallel conductance. Perform the saline calibration outlined in protocol step 6 to obtain the parallel conductance.
6. Poor quality pressure volume loops: The ideal pressure-volume loop has a square or rectangular appearance (see Figure 2 for example). If the shape of the loop is irregular, gently manipulate or twist the conductance catheter in the LV cavity to see if it changes the shape of the loop. Our preferred approach to accessing the LV is through the carotid artery (Protocol step 3.1) as this does not require opening the chest, which can affect hemodynamics. However, the carotid approach can be prone to catheter artifact an irregular loops. Thus, an apical approach (Protocol step 3.2) can be used to address this issue.

Cardiac pressure volume loop analysis provides detailed information of cardiac function and are the gold standard for functional assessment 1. While imaging techniques such as echocardiography or cardiac MRI provide functional measures, these measures are highly dependent on loading conditions. Load-independent measures of cardiac contractility and relaxation require dynamic measurements of the ventricular pressure and volume relation over a range of preload and afterload. This understanding of the pressure-volume relation arises from the groundbreaking work of Sagawa and colleagues 2,3. They demonstrated in ex vivo perfused canine hearts that the pressure-volume loop derived contractility measures were independent of loading conditions 4.
In vivo application of these analyses became possible with the development of conductance catheters in the 1980s. This technical advance allowed Kass and colleagues to perform pressure-volume loop analysis in humans 5,6. Miniaturization of conductance catheters and improvements in surgical techniques in the late 1990&#8217;s 7 made analysis of rodent cardiac function feasible, allowing for genetic and pharmacologic studies to be performed. This advance has since lead to the widespread use of pressure-volume loop analysis and has generated a great deal of insight into mammalian cardiac physiology.
A key concept in the use of conductance catheters and the interpretation of data obtained from it is the relationship between volume and conductance. Conductance is inversely related to voltage, which is measured using a catheter with electrodes placed proximally, usually placed below the aortic valve, and distally, at the LV apex 8. Changes in voltage or conductance are measured by changes in current flowing from proximal to distal electrode. Although the blood pool contributes significantly to conductance, the contribution of the ventricular wall, termed parallel conductance (Vp), to measured conductance must be subtracted to obtain absolute LV volume measurements.
The methods to perform this correction, called a saline calibration, are discussed in the protocol below. The mathematical relationship between conductance and volume, described by Baan and colleagues, is that volume=1/&#945;; (&#961;L2)(G-Gp), where &#945;=uniform field correction factor, &#961;=blood resistivity, L=distance between the electrodes, G=conductance and Gp=non-blood conductance 9. Of note, the uniform field correction factor in mice approaches 1.0 due to small chamber volumes10. Coupled with pressure transducers, the conductance catheter provides real time simultaneous pressure and volume data.
Cardiac pressure-volume analysis presents particular advantages over other measures of cardiac function, as they allow for measurement of ventricular function independent of loading conditions and of heart rate. Specific load-independent cardiac indices of contractility include: end-systolic pressure volume relation (ESPVR), dP/dtmax&#8211;end-diastolic volume relation, maximal elastance (Emax) and preload recruitable stroke work (PRSW). A load-independent measure of diastolic function is the end-diastolic pressure volume relationship (EDPVR) 11. The following protocol describes the conduct of cardiac pressure volume loop analysis, using both a carotid and an apical approach. While the methodology to perform these studies have been described in detail previously 8,11, we will review key steps to obtain precise pressure-volume measurements, including both saline and cuvette calibration correction, and provide a visual demonstration of these procedures. &#160;&#160;Research with animals carried out for this study was handled according to approved protocols and animal welfare regulations of Duke University Medical Center&#8217;s Institutional Animal Care and Use Committee.

<table><tbody><tr><td>AnaSed (xylazine)&amp;nbsp;</td><td>Lloyd Laboratories</td><td>NADA no. 139-236</td><td>Anesthetic</td></tr><tr><td>Ketaset (ketamine)</td><td>Pfizer</td><td>440842</td><td>Anesthetic</td></tr><tr><td>VIP3000</td><td>Matrx Medical Inc.</td><td></td><td>Anesthesia machine</td></tr><tr><td>Ventilator</td><td>Harvard Apparatus</td><td>Model 683</td><td>Surgical Equipment</td></tr><tr><td>Tubing kit</td><td>Harvard Apparatus</td><td>72-1049</td><td>Surgical Equipment</td></tr><tr><td>Homeothermic Blanket&amp;nbsp;</td><td>Kaz Inc.</td><td>5628</td><td>Surgical Equipment</td></tr><tr><td>Stereo microscope</td><td>Carl Zeiss Optical Inc.</td><td>Stemi 2000</td><td>Surgical Equipment</td></tr><tr><td>Illuminator</td><td>Cole&amp;ndash;Parmer</td><td>41720</td><td>Surgical Equipment</td></tr><tr><td>Dumont no. 55 Dumostar Forceps&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>11295-51</td><td>Surgical Instruments</td></tr><tr><td>Graefe forceps, curved&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>11052-10</td><td>Surgical Instruments</td></tr><tr><td>Moria MC31 forceps&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>11370-31</td><td>Surgical Instruments</td></tr><tr><td>Mayo scissors&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>14512-15</td><td>Surgical Instruments</td></tr><tr><td>Iris scissors&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>14041-10</td><td>Surgical Instruments</td></tr><tr><td>Halsey needle holder&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>12501-13</td><td>Surgical Instruments</td></tr><tr><td>Olsen&amp;ndash;Hegar needle holder&amp;nbsp;</td><td>Fine Science Tools Inc</td><td>12002-12</td><td>Surgical Instruments</td></tr><tr><td>spring scissors</td><td>Fine Science Tools Inc</td><td>15610-08</td><td>Surgical Instruments</td></tr><tr><td>disposable underpads</td><td>Kendall/Tyco Healthcare</td><td>1038</td><td>Surgical Supplies</td></tr><tr><td>Sterile gauze sponges, sterile&amp;nbsp;</td><td>Dukal</td><td>62208</td><td>Surgical Supplies</td></tr><tr><td>Cotton-tipped applicators, sterile&amp;nbsp;</td><td>Solon</td><td>368</td><td>Surgical Supplies</td></tr><tr><td>Surgical suture,&amp;nbsp; silk, 6-0&amp;nbsp;</td><td>DemeTECH</td><td>FT-639-1</td><td>Surgical Supplies</td></tr><tr><td>1 cc Insulin syringes&amp;nbsp;</td><td>Becton Dickenson</td><td>329412</td><td>Surgical Supplies</td></tr><tr><td>Access 9 Hemostasis Valve</td><td>Merit Medical&amp;nbsp;</td><td>MAP111</td><td>Hemodynamic equipment</td></tr><tr><td>Sphygmomanometer</td><td>Baumanometer</td><td>320</td><td>Hemodynamic equipment</td></tr><tr><td>Millar PV system MPVS-300/400 or MPVS Ultra (includes calibration cuvette)</td><td>ADInstruments Inc</td><td></td><td>Hemodynamic equipment</td></tr><tr><td>1.4F conductance catheter&amp;nbsp;</td><td>ADInstruments Inc</td><td>SPR-839</td><td>Hemodynamic equipment</td></tr><tr><td>PowerLab 4/30 with Chart Pro</td><td>ADInstruments Inc.</td><td>ML866/P</td><td>Hemodynamic software</td></tr><tr><td>animal clipper</td><td>Wahl</td><td>8787-450A</td><td>Miscellaneous</td></tr><tr><td>Intradermic tubing PE-10</td><td>Becton Dickenson</td><td>427401</td><td>Miscellaneous</td></tr><tr><td>Intradermic tubing PE-50</td><td>Becton Dickenson</td><td>427411</td><td>Miscellaneous</td></tr><tr><td>Needle assortment (18, 25 and 30 gauge; Thomas Scientific)</td><td></td><td></td><td>Miscellaneous</td></tr><tr><td>0.9% (wt/vol) sodium chloride injection, USP)</td><td>Hospira</td><td>NDC no. 0409-4888-50</td><td>Miscellaneous</td></tr><tr><td>Surgical tape</td><td></td><td></td><td>Miscellaneous</td></tr><tr><td>Alconox (Alconox Inc.) for catheter cleaning</td><td>ADInstruments Inc.</td><td></td><td>Miscellaneous</td></tr></tbody></table>

cardiac pressure-volume loop analysis using conductance catheters in mice

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular systolic and diastolic function. Measures of ventricular contractility and compliance are obtained through examination of cardiac response to changes in afterload and preload. These techniques were originally developed nearly three decades ago to measure cardiac function in large mammals and humans. The application of these analyses to small mammals, such as mice, has been accomplished through the optimization of microsurgical techniques and creation of conductance catheters. Conductance catheters allow for estimation of the blood pool by exploiting the relationship between electrical conductance and volume. When properly performed, these techniques allow for testing of cardiac function in genetic mutant mouse models or in drug treatment studies. The accuracy and precision of these studies are dependent on careful attention to the calibration of instruments, systematic conduct of hemodynamic measurements and data analyses. We will review the methods of conducting pressure-volume loop experiments using a conductance catheter in mice.

Pressure-volume loop analysis can be used to measure cardiac function in genetically modified mice 14,15 or mice undergoing drug studies16. Representative pressure volume loops are provided from previously published work 16 investigating the effect of &#223;-Arrestin biased AT1R ligand, TRV120023. To test whether TRV120023 affects cardiac function in vivo, pressure-volume loop analysis was performed on wild type mice receiving conventional and novel angiotensin receptor blockers. Intravenous infusion of TRV120023 increased cardiac contractility significantly (Figure 6 and Table 1, Figure modified from Kim et al.&#160;AJP 201216). Contractility measures were derived from aortic constriction.
<img alt="Figure 1" src="/files/ftp_upload/52942/52942fig1.jpg" /> 
Figure 1. Cardiac pressure and volume calibration. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In pressure volume loop software, assign channels for pressure and volume recordings.&#160;(A) Using hemodynamic catheter module, set volume setting to 5 RVU and 25 RVU and select both volumes, (B) Open option under the channel exhibiting volume recordings, (C) Select unit conversion and open &#8220;2 point calibration&#8221;, select &#8220;point 1&#8221; and assign as 5 RVU and select &#8220;point 2&#8221; and assign as 25 RVU , (D) Set pressure to 0 mm Hg and 25 mm Hg, (E) Open options under the channel exhibiting pressure recordings, (F) Select unit conversion and open &#8220;2 point calibration&#8221;, select &#8220;point 1&#8221; and assign as 0 mm Hg and select &#8220;point 2&#8221; and assign as 25 mm Hg. Accept assignments by selecting &#8220;OK&#8221;.
<img alt="Figure 2" src="/files/ftp_upload/52942/52942fig2.jpg" /> 
Figure 2. Baseline Hemodynamics <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(A) Screenshots from pressure and volume channels of representative cardiac cycles in a basal state, (B) Selected end expiratory baseline pressure-volume loops that have been corrected for parallel conductance for analysis. (C) Baseline hemodynamic summary table calculated from selected loops; Pes, End systolic pressure; Ped, End diastolic pressure; Ves, End systolic volume; Ved, End diastolic volume; SV= Stroke Volume
<img alt="Figure 3" src="/files/ftp_upload/52942/52942fig3.jpg" /> 
Figure 3. Aortic Constriction Hemodynamics <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(A) Screenshots from pressure and volume channels of representative cardiac cycles during aortic constriction, (B) Menu selection to perform contractility analysis, (C) Selected pressure-volume loops during aortic constriction for analysis. (D) ESPVR measured from aortic constriction loops.
<img alt="Figure 4" src="/files/ftp_upload/52942/52942fig4.jpg" /> 
Figure 4. IVC Constriction Hemodynamics <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(A) Screenshots from pressure and volume channels of representative cardiac cycles during IVC constriction, (B) Selected pressure-volume loops during IVC constriction for analysis. Using the PV loops from IVC constriction, Preload Recruitable Stroke Work (C), Maximal dP/dt vs EDV (D) as well as ESPVR and EDPVR can be measured.
<img alt="Figure 5" src="/files/ftp_upload/52942/52942fig5.jpg" /> 
Figure 5. Saline Calibration <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
(A) Screenshots from pressure and volume channels of representative cardiac cycles during hypertonic saline injection (B) Selected saline injection pressure-volume loops for analysis. Note that pressure will remain constant, while volume will increase significantly, (C) Menu selection to perform saline calibration, (D) Generated lines of measure end-systolic vs end-diastolic volume during saline injection and end-systolic volume=end-diastolic volume. The intersection of these lines provides the parallel conductance of Vp.
<img alt="Figure 6" src="/files/ftp_upload/52942/52942fig6.jpg" /> 
Figure 6. Change in Contractility with Drug Administration
Change in cardiac contractility, measured by end systolic elastance, assessed in wild type mice treated with saline, Losartan 5mg/kg/hr or TRV023 100&#956;g/kg/hr for 5 min. TRV023 treated mice developed a significant increase in end systolic elastance in comparison to Losartan treated mice. *p &#60; 0.05 vs Losartan by 1-way ANOVA.
<img alt="Table 1" src="/files/ftp_upload/52942/52942table1.jpg" /> 
Table 1. Wild type Mice; Hemodynamic Profile in Response of &#946;-arrestin 2 Biased AT1R Agonist
End-systolic pressure (ESP) was decreased significantly after TRV120023 (TRV) and losartan infusion. Cardiac contractility, Ees and Emax was increased significantly in TRV120023 100 &#956;g &#183; kg-1 &#183; min-1 infusion group. (*p &#60; 0.01; &#8224;p&#60;0.05; &#8225;p&#60; 0.001; n=5&#8211;6/group). p values reflect comparisons with basal condition within the same treatment group with the use of 1-way ANOVA. Cardiac contractility parameters were derived using an aortic constriction protocol. AT1R, ANG II type 1 receptor; HR, heart rate; EDP, end-diastolic pressure; ESV, end-systolic volume; EDV, end-diastolic volume; Ees, end-systolic elastance; EF, ejection fraction; Emax, maximal elastance; dP/dtmax and dP/dtmin, maximum and minimum rate of pressure change in the ventricle, respectively; &#964;, isovolumic relaxation constant. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52942/52942table1highres.jpg" target="_blank">Please click here to view a larger version of this table.</a>

Watch this Scientific Journal Video about Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice at JoVE.com

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters.

Cardiac pressure-volume loop analysis is the &#8220;gold-standard&#8221; in the assessment of load-dependent and load-independent measures of ventricular ...

cardiac-pressure-volume-loop-analysis-using-conductance-catheters

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19.4K Views.  Duke University Medical Center. Cardiac pressure-volume loop analysis is the most comprehensive way to measure cardiac function in the intact heart. We describe a technique to perform and analyze cardiac pressure volume loops, using conductance catheters. 

Video: Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

Cardiac Catheterization in Mice to Measure the Pressure Volume Relationship: Investigating the Bowditch Effect

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these strategies is to examine their effects on heart function. There are several techniques to measure in vivo cardiac mechanics (e.g., echocardiography, pressure/volume relations, etc.). Compared to echocardiography, real-time left ventricular (LV) pressure/volume analysis via catheterization is more precise and insightful in assessing LV function. Additionally, LV pressure/volume analysis provides the ability to instantaneously record changes during manipulations of contractility (e.g., &#946;-adrenergic stimulation) and pathological insults (e.g., ischemia/reperfusion injury). In addition to the maximum (+dP/dt) and minimum (-dP/dt) rate of pressure change in the LV, an accurate assessment of LV function via several load-independent indexes (e.g., end systolic pressure volume relationship and preload recruitable stroke work) can be attained. Heart rate has a significant effect on LV contractility such that an increase in the heart rate is the primary mechanism to increase cardiac output (i.e., Bowditch effect). Thus, when comparing hemodynamics between experimental groups, it is necessary to have similar heart rates. Furthermore, a hallmark of many cardiomyopathy models is a decrease in contractile reserve (i.e., decreased Bowditch effect). Consequently, vital information can be obtained by determining the effects of increasing heart rate on contractility. Our and others data has demonstrated that the neuronal nitric oxide synthase (NOS1) knockout mouse has decreased contractility. Here we describe the procedure of measuring LV pressure/volume with increasing heart rates using the NOS1 knockout mouse model.

This article describes the measurement of murine left ventricular function via pressure/volume analysis at different heart rates.

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these ...

Medicine

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Hemodynamic Characterization of Rodent Models of Pulmonary Arterial Hypertension

Pulmonary arterial hypertension (PAH) is a rare disease of the pulmonary vasculature characterized by endothelial cell apoptosis, smooth muscle proliferation and obliteration of pulmonary arterioles. This in turn results in right ventricular (RV) failure, with significant morbidity and mortality. Rodent models of PAH, in the mouse and the rat, are important for understanding the pathophysiology underlying this rare disease. Notably, different models of PAH may be associated with different degrees of pulmonary hypertension, RV hypertrophy and RV failure. Therefore, a complete hemodynamic characterization of mice and rats with PAH is critical in determining the effects of drugs or genetic modifications on the disease. 
Here we demonstrate standard procedures for assessment of right ventricular function and hemodynamics in both rat and mouse PAH models. Echocardiography is useful in determining RV function in rats, although obtaining standard views of the right ventricle is challenging in the awake mouse. Access for right heart catheterization is obtained by the internal jugular vein in closed-chest mice and rats. Pressures can be measured using polyethylene tubing with a fluid pressure transducer or a miniature micromanometer pressure catheter. Pressure-volume loop analysis can be performed in the open chest. After obtaining hemodynamics, the rodent is euthanized. The heart can be dissected to separate the RV free wall from the left ventricle (LV) and septum, allowing an assessment of RV hypertrophy using the Fulton index (RV/(LV+S)). Then samples can be harvested from the heart, lungs and other tissues as needed.

Pulmonary arterial hypertension (PAH) is a disease of pulmonary arterioles that leads to their obliteration and the development of right ventricular failure. Rodent models of PAH are critical in understanding the pathophysiology of PAH. Here we demonstrate hemodynamic characterization, with right heart catheterization and echocardiography, in the mouse and rat.

Pulmonary arterial hypertension (PAH) is a rare disease of the pulmonary vasculature characterized by endothelial cell apoptosis, smooth muscle ...

Bioengineering Human Microvascular Networks in Immunodeficient Mice

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks in vivo. In this regard, the search for experimental models to build blood vessel networks in vivo is of utmost importance 1. The feasibility of bioengineering microvascular networks in vivo was first shown using human tissue-derived mature endothelial cells (ECs) 2-4; however, such autologous endothelial cells present problems for wide clinical use, because they are difficult to obtain in sufficient quantities and require harvesting from existing vasculature. These limitations have instigated the search for other sources of ECs. The identification of endothelial colony-forming cells (ECFCs) in blood presented an opportunity to non-invasively obtain ECs 5-7. We and other authors have shown that adult and cord blood-derived ECFCs have the capacity to form functional vascular networks in vivo 7-11. Importantly, these studies have also shown that to obtain stable and durable vascular networks, ECFCs require co-implantation with perivascular cells. The assay we describe here illustrates this concept: we show how human cord blood-derived ECFCs can be combined with bone marrow-derived mesenchymal stem cells (MSCs) as a single cell suspension in a collagen/fibronectin/fibrinogen gel to form a functional human vascular network within 7 days after implantation into an immunodeficient mouse. The presence of human ECFC-lined lumens containing host erythrocytes can be seen throughout the implants indicating not only the formation (de novo) of a vascular network, but also the development of functional anastomoses with the host circulatory system. This murine model of bioengineered human vascular network is ideally suited for studies on the cellular and molecular mechanisms of human vascular network formation and for the development of strategies to vascularize engineered tissues.

Here, we describe a methodology to deliver human cord blood-derived endothelial colony-forming cells (ECFCs) and bone marrow-derived mesenchymal stem cells (MSCs), embedded in a collagen/fibronectin gel, subcutaneously into immunodeficient mice. This cell/gel combination generates a human vascular network that connects with the mouse vasculature.

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks ...

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) generated by recording both pressure and volume during cardiac catheterization are vital when assessing both systolic and diastolic cardiac function. Left and right heart function are closely related, reflected in ventricular interdependence. Thus, recording biventricular function in the same animal is important to get a complete assessment of cardiac function. In this protocol, a closed chest approach to cardiac catheterization consistent with the way catheterization is performed in patients is adopted in mice. While challenging, the closed chest strategy is a more physiological approach, because opening the chest results in major changes in preload and afterload that create artifacts, most notably a fall in systemic blood pressure. While high-resolution echocardiography is used to assess rodents, cardiac catheterization is invaluable, particularly when assessing diastolic pressures in both ventricles.
Described here is a procedure to perform invasive, closed chest, sequential left and right ventricular pressure-volume (PV) loops in the same animal. PV loops are acquired using&#160;admittance technology with a mouse pressure-volume catheter and pressure-volume system acquisition. The procedure is described, beginning with the neck dissection, which is required to access the right jugular vein and the right carotid artery, to the insertion and positioning of the catheter, and finally the data acquisition. Then, the criteria required to ensure the acquisition of high-quality PV loops are discussed. Finally, the analysis of the left and right ventricular PV loops and the different hemodynamic parameters available to quantify systolic and diastolic ventricular function are briefly described.

Presented here is a protocol to assess biventricular heart function in mice by generating pressure-volume (PV) loops from the right and left ventricle in the same animal using closed chest catheterization. The focus is on the technical aspect of surgery and data acquisition.

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) ...

Cardiac Response to &#946;-Adrenergic Stimulation Determined by Pressure-Volume Loop Analysis

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&#160;in vivo using pressure-volume loop measurements and analysis during basal conditions and under &#946;-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.

Here we describe a cardiac pressure-volume loop analysis under increasing doses of intravenously infused isoproterenol to determine the intrinsic cardiac function and the &#946;-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.

Determination of the cardiac function is a robust endpoint analysis in animal models of cardiovascular diseases in order to characterize effects of ...

Closed Chest Biventricular Pressure-Volume Loop Recordings with Admittance Catheters in a Porcine Model

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular evaluation is often performed in preclinical research. However, the right and left ventricles exert functional interdependence due to their parallel and serial connections, encouraging simultaneous evaluation of both ventricles. Furthermore, various pharmacological interventions may affect the ventricles and their preloads and afterloads differently. 
We describe our closed chest approach to admittance-based bi-ventricular PV loop recordings in a porcine model of acute right ventricular (RV) overload. We utilize minimally invasive techniques with all vascular accesses guided by ultrasound. PV catheters are positioned, under fluoroscopic guidance, to avoid thoracotomy in animals, as the closed chest approach maintains the relevant cardiopulmonary physiology. The admittance technology provides real-time PV loop recordings without the need for post-hoc processing. Furthermore, we explain some essential troubleshooting steps during critical timepoints of the presented procedure. 
The presented protocol is a reproducible and physiologically relevant approach to obtain a bi-ventricular cardiac PV loop recording in a large animal model. This can be applied to a large variety of cardiovascular animal research.

Here we present a closed chest approach to admittance-based bi-ventricular pressure-volume loop recordings in pigs with acute right ventricular dysfunction.

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular ...

Measuring Pressure Volume Loops in 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&#39;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.

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 ...

Monitoring Cardiac Function of Rats Using the Pressure-Volume Conductance Technique

Real-Time Detection of Ferulic Acid Effects on Rat Left Ventricle Using Pressure-Volume Conductivity Catheter

Decreased cardiac function can have a negative impact on other organs. The left ventricular pressure-volume relationship is considered to be a valid method for evaluating cardiac function. Real-time monitoring of cardiac function is important for drug evaluation. Under closed-chest conditions, the miniature transducer, which is an important component of the pressure-volume catheter, enters the left ventricle of the rat through the right carotid artery. The device visualizes the changes in cardiac function during the experiment in the form of a pressure-volume loop. The actual volume of the ventricle is calculated by altering the conductivity of the blood by injecting 50 &#181;L of a 20% sodium chloride solution into the rat's left jugular vein. The actual volume of the rat's ventricular cavity is calculated by measuring the conductivity of the blood in a known volume using a pressure-volume conductance catheter. This protocol allows for continuous observation of the effects of drugs on the heart and will promote the rationale for the use of specialty ethnic drugs in cardiovascular disease.

Author Spotlight: Advantages of Pressure-Volume Loop Measurement Method in Cardiovascular Research

This protocol describes a method for measuring left ventricular pressure and volume using the pressure-volume conductance technique. This method enables continuous real-time monitoring of the effects of drugs on the heart.

Decreased cardiac function can have a negative impact on other organs. The left ventricular pressure-volume relationship is considered to be a valid ...

Exploring HR-Dependent Cardiac Performance Using an Atrial-Pacing Protocol

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

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.

Author Spotlight: Investigating HR-Dependent Cardiac Function in Mouse Models Through a Novel Atrial-Pacing Approach 

The present protocol describes obtaining the pressure-volume relationship through transesophageal pacing, which serves as a valuable tool in evaluating diastolic function&#160;in mouse models of heart failure.

Cardiac Pressure-Volume Loop Analysis Using Conductance Catheters in Mice

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