This work was funded by the National Institutes of Health 1R21HL092370-01 (GG), 1R01 HL095764-01 (GG); R01HL082694 (JW); American Heart Association, Founders affiliate (0855943D, GG); Stony Wold - Herbert Fund, New York (SHP).

1. Preparation
<ol>
 <li>Prepare the following solutions and tubes (Table 1) as follows:
 <ol>
 <li>Hanks solution, no calcium, magnesium or indicator with Penicillin (100 U/ml) / Streptomycin (100 mg/ml).</li>
 <li>Phosphate buffered saline (PBS), 1x, no calcium, no magnesium.</li>
 <li>Ethanol, 70 %, make 500 ml.</li>
 <li>Buffered formaldehyde, 7-10 % with PBS, make 500 ml.</li>
 <li>Anaesthesia solutions:
 <ol>
 <li>Avertin. Carefully add 5 ml of 2-Methyl-2-butanol to 5 g of 2,2,2-Tribromoethanol. Once dissolved, keep stock solution at room temperature in the dark. Take 0.25 ml of the stock solution and dilute it with 10 ml of 1xPBS in glass bottom bottle. Wrap the bottle with aluminium foil, place at 37 &deg;C until dissolved and then aliquot into 5 ml polypropylene tubes and keep in refrigerator. Warm one aliquot on the morning of the experiment.</li>
 <li>Barbiturate. Dilute stock solution with PBS to give 2.6% barbiturate solution. Place 3-4 ml aliquots in polypropylene tubes.</li>
 </ol>
 </li>
 </ol>
 </li>
 <li>Organize three work-areas (Figure 1).</li>
 <li>
 <ol>
 <li>Area A: Arrange the following items: study form to record study identification number, date, body weight, weights of right heart, left heart and septum; precision scale (0-50 g at 0.01 g precision) to determine body weight; micro scale to determine heart weights at 0.001 g precision; weigh boats; anaesthesia solutions (clearly marked); small dewar with liquid nitrogen; insulated container with ice; tubes and 24-well plate to collect blood and tissue specimens (Table 1); surgical instruments (Table 2, Figure 2); and solutions to clean the instruments (Table 1).</li>
 <li>Area B: Arrange the following items: dissecting microscope; right heart catheter connected to a pressure control unit that is connected to an amplifier and laptop computer. The catheter is placed into a beaker containing water. Arrange surgical instruments (Table 2), pieces of suture cut to optimal length and placed into a weigh boat; tape (autoclave tape is optimal) cut to optimal length and width and lined up on microscope or lab bench for easy access; cotton swabs and hair remover solution. Calibrate the catheter at the beginning of the study.</li>
 <li>Area C: Arrange the following items: magnifier lens; tracheal cannula; 1 ml syringes; suture cut to optimal length and placed into a weigh boat; small dewar with liquid nitrogen; insulated container with ice; tubes and 24-well plate to collect BAL and tissue specimens (Table 1); surgical instruments (Table 2); solutions to perform BAL and to clean the tracheal cannula and the instruments (Table 1).</li>
 </ol>
 </li>
</ol>
2. Right Heart Catheterization
<ol>
 <li>Weigh mouse using a precision scale by gently placing mouse into a weigh boat. Make sure to handle the animal gently and quietly. Record weight. The described protocol works best for mice that are 20 g or more because the diameter of the jugular vein has to accommodate the pressure catheter; it is possible to catheterize smaller mice (17 g or more) as long as the vein is large enough.</li>
 <li>Anaesthetize the mouse by injection with Avertin depending on the body weight (10 &mu;l/g), e.g. for 20 g give 200 &mu;l, for 25 g give 250 &mu;l, for 30 g give 300 &mu;l. The exact injection volume needs to be adjusted depending on the mouse strain. Make sure to handle the animal gently and quietly because stress can alter the response to the anaesthetic.</li>
 <li>Once the mouse is sedated, place on double-pleated tissue paper. Note the ID number on the paper. Gently rub hair removal lotion into the ventral aspect of the neck to clear the surgical field. Place the mouse back onto the paper and wait for 1-2 min.</li>
 <li>Remove hair from the ventral aspect of the neck by gently stroking the hair against the direction of the hair growth with cotton swabs.</li>
 <li>Carefully determine the sufficient depth of anaesthesia by checking for the absence of the toe reflex: the mouse does not react to pinching of its toes.</li>
 <li>Working quickly and relaxed, the remainder of the right heart catheterization should not take more than 10 min, optimally 3-6 min. It is important that the tissue does not dry out because the catheter needs to glide into the vein and move inside of the vein on a layer of moisture. Consistent timing of the procedure is important for the generation of data that are comparable between groups.</li>
 <li>Place mouse back on tissue paper and transfer with tissue paper onto styrofoam board. Fix paws and head into place using autoclave tape.</li>
 <li>Make incision of skin from mandibles to sternum (breast-bone).</li>
 <li>Carefully dissect the thyroid grand upwards towards the mandibles to expose the right jugular vein and trachea.</li>
 <li>Place styrofoam board holding the mouse underneath the dissection microscope with the focus plane already in place and the lens at approximately 0.8x magnification (total magnification [lens x eye piece] is approximately 8x.</li>
 <li>Carefully microdissect the right jugular vein by removing surrounding connective tissue with surgical forceps.</li>
 <li>Place two pieces of suture on top of the right jugular vein. Tie the suture closest to the mandibles to close off blood flow in the vein to a tiny trickle, place the suture that is closest to the sternum / breast-bone at a loose knot around the vein. One can use the tiny trickle of blood to later find the hole in the vein, should this become necessary.</li>
 <li>Breathe and relax (loosen the muscles in the neck and jaw) to keep your eyes on the vein through the eyepiece. This will ensure that your hands and fingers will move securely, smoothly and relaxed. While holding the vein with forceps at the side closest to the mandibles, cut a hole into the vein between the two sutures using micro-scissors operated by the other hand. Put down the micro-scissors, and keeping that hand relaxed, feel for the catheter. Gently retrieve the catheter holding it between thumb and first finger (Figure 3). Insert the catheter into the hole of the vein.</li>
 <li>Gently advance the catheter into the right heart (Figure 3). Observe the marks at the catheter that give an approximate index of how far the catheter needs to be inserted and observe the monitor. Once the pressure curves are displayed, gently tighten the lower suture around the catheter. Because the catheter has tensile strength that gives it a coil, and because the breathing and heart beat of the mouse add motion, additionally affix the catheter using tape.</li>
 <li>Record pressure measurements for 2 min for later analysis (Figure 4).</li>
 <li>Open the suture holding the catheter, then gently retrieve catheter. Wipe the part behind the pressure transducer, place back into the beaker with water. Do not touch the pressure transducer.</li>
 <li>Transfer the mouse on top of the tissue paper to area A for euthanasia, collection of blood and spleen tissue. Clean surgical instruments.</li>
 <li>Start the procedure with the next animal. When the timing permits, inject the anaesthetic while waiting to record the pressure curve (2.15).</li>
</ol>
3. Collection of Blood and Spleen Samples
<ol>
 <li>Inject barbiturate solution, 400 &mu;l per mouse and wait for 1-2 min. This is routinely performed in our laboratory to avoid the possibility that the mice inadvertently recover from avertin-anaesthesia while being bled. Appropriate standardization of the experiment is achieved by injecting the same dose of barbiturate solution per mouse, and by waiting the same amount of time prior to harvest of blood. If permitted by the Institutional Animal Care and Use Committee and as long as the animals are carefully monitored for the depth of anaesthesia prior to exsanguination, this step can be omitted.</li>
 <li>Make incision into the abdomen. Open the caudal vein (vena cava) and collect blood using a transfer pipette.</li>
 <li>Remove the spleen, or a piece of the spleen into the Eppendorf tube and snap freeze in liquid nitrogen, or into the 24 well plate into a well containing Hanks buffer for later preparation of single cells.</li>
 <li>Transfer the mouse on top of the tissue paper to work-area C for collection of BAL, lung draining lymph node, lung, and heart tissues. Clean surgical instruments.</li>
</ol>
4. Collection of Lung and Heart Samples
<ol>
 <li>Dissect the trachea free of muscle and connective tissue. Placing forceps under the trachea, pull suture through. Gently generate sufficient stretch to cut a hole. Maintaining the gentle stretch, insert the tracheal cannula using a slightly turning motion, and suture cannula into place. For the insertion of the tracheal cannula, make sure that the trachea is moist, if necessary add a drop of Hanks solution. Throughout the procedure, breathe, relax neck and jaw muscles, to keep hand movements secure and smooth.</li>
 <li>Dissect open the rib cage by carefully removing the breast-bone starting from the abdominal end. Do not disturb the contents of the thorax because this will make it very hard to find the lymph node.</li>
 <li>Perform bronchoalveolar lavage (BAL) by gently inserting 1 ml of Hanks buffer, gently turn the syringe and retrieve BAL fluid. Repeat 3 times, close tube and place on ice. Throughout the procedure, keep eyes on the end of the tracheal cannula that connects to the syringe. Glance to thorax to observe the inflation and deflation of the lungs.</li>
 <li>Harvest lung lymph node by holding rib cage wall with one set of forceps, finding and retrieving the lymph node with another forceps. Place lymph node into 24 well plate. It is important to know where to search: starting from the neck, find the entrance of the rib cage from the right side of the mouse and look above the spine (Figure 5).</li>
 <li>Harvest heart and place on tissue paper. Isolate the right heart by carefully dissecting alongside the septum. Transfer to work-area A to weigh, record the weight and place into eppendorf tubes to snap freeze in liquid nitrogen.</li>
 <li>Place suture around the left lung lobe to close the main bronchus. Dissect the left lung lobe free, place into eppendorf tube and snap freeze in liquid nitrogen, or place into a well of a 24 well plate containing Hanks solution for later isolation of single cell suspensions.</li>
 <li>Insert approx. 0.5 ml buffered formaldehyde solution through the tracheal cannula into the remaining lung lobes. After inflation, dissect the lung lobes out of the thorax and place into 50 ml tube containing formaldehyde solution. For an alternative study design, the lungs can be inflated with optimal cutting media (OCT, diluted 1:4 with PBS) and frozen in a mold containing undiluted OCT for later preparation of frozen sections. Another study design might necessitate obtaining snap frozen lung tissue and single cell suspensions from the lungs. In that case, no inflation is necessary: dissect the remaining lung lobes; retrieve them from the thorax and place into the 24 well plate into a well containing Hanks buffer.</li>
 <li>Remove tracheal cannula, clean by rinsing with Hanks buffer, clean surgical instruments.</li>
</ol>
5. Analysis of Pressure Curves Generated from the Right Heart Catheter
The recorded right ventricular pressure data are analyzed using LabChart 7 software with no knowledge of the group identity of each recording. More than 20 curves are selected randomly and the difference between maximum and minimum ventricular pressure for each curve measured (&Delta;P). The average &Delta;P is calculated to give the right ventricular systolic pressure.

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No conflicts of interest declared.

The experimental flow outlined here allows for rapid and simultaneous measurement of right ventricular systolic pressure and harvest of samples for the analysis of the responses in the lungs, heart and the immune system in mice. The procedure combines heart physiology measurements, micro-dissection and subsequent tissue harvest for live cell studies, histological analysis, or omics-analysis of the tissues. The complete procedure takes less than 20 min per mouse. Because of the work-area-organized workflow, 2-3 animals ca...

<table><tbody><tr><td colspan="4">&lt;strong&gt;Reagents&lt;/strong&gt;</td></tr><tr><td>2-Methyl-2-butanol </td><td>Sigma-Aldrich</td><td>152463</td><td></td></tr><tr><td>2,2,2-Tribrom&amp;#8211;thanol </td><td>Sigma-Aldrich</td><td>T48402</td><td></td></tr><tr><td>disinfectant soap (Coverage Spray TB plus Steris)</td><td>Fisher Scientific</td><td>1629-08</td><td></td></tr><tr><td>Ethyl Alcohol, 200 Proof, Absolute, Anhydrous ACS/USP Grade </td><td>PHARMCO-AAPER</td><td>111000200</td><td>Dilute to 70 % with distilled water</td></tr><tr><td>Formaldehyde solution </td><td>Sigma-Aldrich</td><td>F1635-500ML</td><td>Dilute to a 7-10 % formaldehyde concentration at a PBS concentration of 1x using PBS stock solution and water</td></tr><tr><td>Hanks solution, no calcium, magnesium </td><td>Fisher Scientific</td><td>21-022-CV</td><td></td></tr><tr><td>O.C.T</td><td>Tissue-Tek</td><td>4583</td><td></td></tr><tr><td>Penicillin (10,000 U/ml) / Streptomycin (10,000 mg/ml) solution</td><td>Thermo Scientific</td><td>SV30010</td><td></td></tr><tr><td>Phosphate buffered saline (PBS), no calcium, no magnesium, 1x and 10x solutions </td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Sodium pentobarbital 26% </td><td>Fort Dodge Animal Health</td><td>NDC 0856-0471-01</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Labware&lt;/strong&gt; </td></tr><tr><td>Plates 12, 24, 96 well </td><td>Falcon</td><td></td><td></td></tr><tr><td>Transfer Pipet </td><td>Fisher Scientific</td><td>13-711-9BM</td><td></td></tr><tr><td>Tube, EDTA coated </td><td>Sarstedt</td><td>2013-08</td><td></td></tr><tr><td>Tubes 0.65 ml and 1.7 ml micro-centrifuge </td><td>VWR</td><td></td><td></td></tr><tr><td>Tubes 12 x 75 mm polypropylene </td><td>Fisher Scientific</td><td>14-956-1D</td><td></td></tr><tr><td>Tubes, various sizes, polypropylene</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Instruments&lt;/strong&gt; </td></tr><tr><td>Forceps, Dumon #5 Fine</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Forceps, extra fine graefe -0.5 mm tips curved</td><td>Fine Science Tools</td><td>11152-10</td><td></td></tr><tr><td>Forceps, extra fine graefe -0.5 mm tips straight</td><td>Fine Science Tools</td><td>11150-10</td><td></td></tr><tr><td>Cannula 18 ga, 19 ga</td><td>BD</td><td>Precision Glide Needles</td><td>Cut to optimal length, blunted and outside rasped to create a rough outside surface.</td></tr><tr><td>Scissors, Dissector scissors-slim blades 9 cm</td><td>Fine Science Tools</td><td>14081-09</td><td></td></tr><tr><td>Suture for BAL, braided silk suture, 4-0 </td><td>Fine Science Tools</td><td>SP116</td><td></td></tr><tr><td>Suture for right heart catheterization, braided silk suture, 6-0</td><td>Teleflex medical</td><td>18020-60</td><td></td></tr><tr><td>Syringe, 1 ml</td><td>BD</td><td>309659</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Equipment&lt;/strong&gt; </td></tr><tr><td>Amplifier, PowerLab 4/30</td><td>ADInstrument</td><td>Model ML866</td><td></td></tr><tr><td>Catheter, pressure F1.4</td><td>Millar Instruments, Inc</td><td>840-6719</td><td></td></tr><tr><td>Dissecting Microscope</td><td>Variscope</td><td></td><td></td></tr><tr><td>Forceps, Vannas spring scissors-2 mm blades </td><td>Fine Science Tools</td><td>15000-00</td><td></td></tr><tr><td>Halogen Illuminated Desk Magnifier</td><td>Fisher Scientific</td><td>11-990-56</td><td></td></tr><tr><td>Laptop computer</td><td>Asus</td><td>Model number A52F i5 processor; 15 inch</td><td></td></tr><tr><td>Light Source </td><td>Amscope</td><td>HL-250-A</td><td></td></tr><tr><td>Pressure Control Unit </td><td>Millar Instruments, Inc</td><td>PCU-2000</td><td></td></tr><tr><td>Software, Labchart-Pro V.7</td><td>AD Instruments</td><td></td><td></td></tr></tbody></table>

right ventricular systolic pressure measurements in combination with harvest of lung and immune tissue samples in mice

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is a common modifier of heart and lung function, by elaborating cellular infiltration, production of cytokines and growth factors, and by initiating remodeling processes 1.
Compared to the left ventricle, the right ventricle is a low-pressure pump that operates in a relatively narrow zone of pressure changes. Increased pulmonary artery pressures are associated with increased pressure in the lung vascular bed and pulmonary hypertension 2. Pulmonary hypertension is often associated with inflammatory lung diseases, for example chronic obstructive pulmonary disease, or autoimmune diseases 3. Because pulmonary hypertension confers a bad prognosis for quality of life and life expectancy, much research is directed towards understanding the mechanisms that might be targets for pharmaceutical intervention 4. The main challenge for the development of effective management tools for pulmonary hypertension remains the complexity of the simultaneous understanding of molecular and cellular changes in the right heart, the lungs and the immune system.
Here, we present a procedural workflow for the rapid and precise measurement of pressure changes in the right heart of mice and the simultaneous harvest of samples from heart, lungs and immune tissues. The method is based on the direct catheterization of the right ventricle via the jugular vein in close-chested mice, first developed in the late 1990s as surrogate measure of pressures in the pulmonary artery5-13. The organized team-approach facilitates a very rapid right heart catheterization technique. This makes it possible to perform the measurements in mice that spontaneously breathe room air. The organization of the work-flow in distinct work-areas reduces time delay and opens the possibility to simultaneously perform physiology experiments and harvest immune, heart and lung tissues.
The procedural workflow outlined here can be adapted for a wide variety of laboratory settings and study designs, from small, targeted experiments, to large drug screening assays. The simultaneous acquisition of cardiac physiology data that can be expanded to include echocardiography5,14-17 and harvest of heart, lung and immune tissues reduces the number of animals needed to obtain data that move the scientific knowledge basis forward. The procedural workflow presented here also provides an ideal basis for gaining knowledge of the networks that link immune, lung and heart function. The same principles outlined here can be adapted to study other or additional organs as needed.

The primary outcome for obtaining right heart pressure curves is achieved by the correct position of the right heart catheter. The shape of the pressure time curves is critical because the correct placement of the catheter inside of the right ventricle will result in pressure plateaus (Figure 4). Spiky curves, instead, indicate a catheter that is moved by the breathing or heart beat motion against the wall of the right ventricle. To detect potential problems with the stage of survival of the anima...

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Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice | Immunology and Infection | Video

Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

A specific and rapid protocol to simultaneously investigate right heart function, lung inflammation, and the immune response is described as a learning tool. Video and figures describe physiology and microdissection techniques in an organized team-approach that is adaptable to be used for small to large sized studies.

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is ...

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25.1K Views.  New York University School of Medicine, Tuxedo. A specific and rapid protocol to simultaneously investigate right heart function, lung inflammation, and the immune response is described as a learning tool. Video and figures describe physiology and microdissection techniques in an organized team-approach that is adaptable to be used for small to large sized studies.

Video: Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

Bioengineering

Echocardiographic Assessment of the Right Heart in Mice

Transgenic and toxic models of pulmonary arterial hypertension (PAH) are widely used to study the pathophysiology of PAH and to investigate potential therapies. Given the expense and time involved in creating animal models of disease, it is critical that researchers have tools to accurately assess phenotypic expression of disease. Right ventricular dysfunction is the major manifestation of pulmonary hypertension. Echocardiography is the mainstay of the noninvasive assessment of right ventricular function in rodent models and has the advantage of clear translation to humans in whom the same tool is used. Published echocardiography protocols in murine models of PAH are lacking.
In this article, we describe a protocol for assessing RV and pulmonary vascular function in a mouse model of PAH with a dominant negative BMPRII mutation; however, this protocol is applicable to any diseases affecting the pulmonary vasculature or right heart. We provide a detailed description of animal preparation, image acquisition and hemodynamic calculation of stroke volume, cardiac output and an estimate of pulmonary artery pressure.

This article provides a protocol for the echocardiographic assessment of right ventricular size and pulmonary hypertension in mice. Applications include phenotype determination and serial assessment in transgenic and toxin-induced mouse models of cardiomyopathy and pulmonary vascular disease.

Transgenic and toxic models of pulmonary arterial hypertension (PAH) are widely used to study the pathophysiology of PAH and to investigate potential ...

Medicine

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

Measurement of the Pressure-volume Curve in Mouse Lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

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 ...

Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured with independent control over pulmonary arterial flow rate, flow rate waveform, airway pressure and left atrial pressure. Pulmonary vascular resistance is calculated based on multi-point, steady pressure-flow curves; pulmonary vascular impedance is calculated from pulsatile pressure-flow curves obtained at a range of frequencies. As now recognized clinically, impedance is a superior measure of right ventricular afterload than resistance because it includes the effects of vascular compliance, which are not negligible, especially in the pulmonary circulation. Three important metrics of impedance - the zero hertz impedance Z0, the characteristic impedance ZC, and the index of wave reflection RW - provide insight into distal arterial cross-sectional area available for flow, proximal arterial stiffness and the upstream-downstream impedance mismatch, respectively. All results obtained in isolated, ventilated and perfused lungs are independent of sympathetic nervous system tone, volume status and the effects of anesthesia. We have used this technique to quantify the impact of pulmonary emboli and chronic hypoxia on resistance and impedance, and to differentiate between sites of action (i.e., proximal vs. distal) of vasoactive agents and disease using the pressure dependency of ZC. Furthermore, when these techniques are used with the lungs of genetically engineered strains of mice, the effects of molecular-level defects on pulmonary vascular structure and function can be determined.

The following protocol outlines the process of isolating, ventilating and instrumenting mouse lungs to measure steady or pulsatile pulmonary vascular pressure-flow relationships in order to quantify the effects of blood flow, airflow, airway changes and vascular changes on right ventricular afterload.

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured ...

Echocardiographic Measurement of Right Ventricular Diastolic Parameters in Mouse

Diastolic dysfunction is a prominent feature of right ventricular (RV) remodeling associated with conditions of pressure overload. However, the RV diastolic function is rarely quantified in experimental studies. This might be due to technical difficulties in the visualization of the RV in the apical four-chamber view in rodents. Here we describe two positions facilitating the visualization of the apical four-chamber view in mice to assess the RV diastolic function.
The apical four-chamber view is enabled by tilting the mouse fixation platform to the left and caudally (LeCa) or to the right and cranially (RiCr). Both positions provide images of comparable quality. The results of the RV diastolic function obtained from two positions are not significantly different. Both positions are comparably easy to perform. This protocol can be incorporated into published protocols and enables detailed investigations of the RV function.

Here we describe and compare two positions for obtaining the apical four-chamber view in mice. These positions enable the quantification of the right ventricular function, provide comparable results, and can be used interchangeably.

Diastolic dysfunction is a prominent feature of right ventricular (RV) remodeling associated with conditions of pressure overload. However, the RV ...

Induction and Characterization of Pulmonary Hypertension in Mice using the Hypoxia/SU5416 Model

Pulmonary Hypertension (PH) is a pathophysiological condition, defined by a mean pulmonary arterial pressure exceeding 25 mm Hg at rest, as assessed by right heart&#160;catheterization. A broad spectrum of diseases can lead to PH, differing in their etiology, histopathology, clinical presentation, prognosis, and response to&#160;treatment. Despite significant progress in the last years, PH remains an uncured disease. Understanding the underlying mechanisms can pave the way for the development of new therapies. Animal models are important research tools to achieve this goal. Currently, there are several models available for recapitulating PH. This protocol describes a two-hit mouse PH model. The stimuli for PH development are hypoxia and the injection of SU5416, a vascular endothelial growth factor (VEGF) receptor antagonist. Three weeks after initiation of Hypoxia/SU5416, animals develop pulmonary vascular remodeling imitating the histopathological changes observed in human PH (predominantly Group 1). Vascular remodeling in the pulmonary circulation results in the remodeling of the right&#160;ventricle (RV). The procedures for measuring RV pressures (using the open chest method), the morphometrical analyses of the RV (by dissecting and weighing both cardiac ventricles) and the histological assessments of the remodeling (both pulmonary by assessing vascular remodeling and cardiac by assessing RV cardiomyocyte hypertrophy and fibrosis) are described in detail. The advantages of this protocol are the possibility of the application both in wild type and in genetically modified mice, the relatively easy and low-cost implementation, and the quick development of the disease of interest (3 weeks). Limitations of this method are that mice do not develop a severe phenotype and PH is reversible upon return to normoxia. Prevention, as well as therapy studies, can easily be implemented in this model, without the necessity of advanced skills (as opposed to surgical rodent models).

This protocol describes the induction of pulmonary hypertension (PH) in mice based on the exposure to hypoxia and the injection of a VEGF receptor antagonist. The animals develop PH and right ventricular (RV) hypertrophy 3 weeks after the initiation of the protocol. The functional and morphometrical characterization of the model is also presented.

Pulmonary Hypertension (PH) is a pathophysiological condition, defined by a mean pulmonary arterial pressure exceeding 25 mm Hg at rest, as assessed by ...

Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. However, the evaluation of right ventricular pressure (RVP) and pulmonary artery pressure (PAP) remains technically challenging in mice. RVP and PAP are more difficult to measure than left ventricular pressure because of the anatomical differences between the left and right heart systems. In this paper, we describe a stable right heart hemodynamic measurement method and its validation using healthy and PAH mice. This method is based on open-chest surgery and mechanical ventilation support. It is a complicated procedure compared to closed chest procedures. While a well-trained surgeon is required for this surgery, the advantage of this procedure is that it can generate both RVP and PAP parameters at the same time, so it is a preferable procedure for the evaluation of PAH models.

Here, we present a protocol to perform an invasive hemodynamic assessment of the right ventricle and pulmonary artery in mice using an open-chest surgery approach.

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. ...

Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, there is no cure for PAH. It is important to discover new compounds that can be used to treat PAH. The mouse hypoxia-induced PAH model is a widely used model for PAH research. This model recapitulates human clinical manifestations of PAH Group 3 disease and is an important research tool to evaluate the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds in mice. For a compound that needs to be given directly into the bloodstream, optimizing intravenous (IV) administration is a key part of the experimental procedures. Ideally, the IV injection system should permit multiple injections over a set time course. Although the mouse hypoxia-induced PAH model is very popular in many laboratories, it is technically challenging to perform multiple IV bolus dosing and invasive hemodynamic assessment in this model. In this protocol, we present step-by-step instructions on how to carry out multiple IV bolus dosing via mouse jugular vein and perform arterial and right ventricle catheterization for hemodynamic assessment in mouse hypoxia-induced PAH model.

This protocol provides a step-by-step procedure for executing multiple intravenous bolus dose administration and invasive hemodynamic monitoring in mice. Investigators can use this protocol for future therapeutic compound screening for pulmonary artery hypertension.

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, ...