Name: right ventricular systolic pressure measurements in combination with harvest of lung and immune tissue samples in mice
Uploaded: 2013-01-16T12:00:00.000+00:00
Duration: 10 min 20 s
Description: Watch this Scientific Journal Video about Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice at JoVE.com

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 can be studied simultaneously. Therefore, the procedure is suitable for small, targeted experiments 12 and large scale screens performed in a wide range of settings, from a small laboratory to a large pharmaceutical study.
The combination of heart physiology measurements and tissue harvest opens the possibility to perform analysis designed to detect biological networks that control or synchronize heart, lung and immune function, utilizing transgenic and KO mouse resources. An important advantage of the simultaneous procedure flow is the reduction in the number of animals needed per study. Another application of this procedural workflow is the possibility to study other or additional organs from the same animal as needed.
The procedure outlined here generates right heart pressure data very rapidly, within 5-10 min of anaesthesia induction. This makes it possible to study the animals while they are breathing room air spontaneously. A limitation of the approach is that it is invasive: the animals are anaesthetized, they are placed onto their backs, and a catheter is passed via the jugular vein through the atrium into the right ventricle. The limitation is also the advantage of this procedure because the data represent direct measurements of pressure changes over time in the right ventricle. Further technical improvements, particularly relying on miniaturizing the pressure catheters, may allow to place permanent, indwelling catheters in mice that allow for the direct recording of the pulmonary artery pressure over several weeks time, as is being performed in rats 22-24 and other larger animals. If insertion of a heart-catheter through the jugular vein in close-chested, spontaneously breathing animals is not possible, an alternative recording of right ventricular systolic pressure changes can be made in ventilated animals by opening of the thorax and placing the pressure catheter through an incision directly into the right ventricle 25. Non-invasive measurements of right heart function can be made using echocardiography. This procedure can either be performed prior to right heart catheterization, or mice can be examined by echocardiography for multiple times to study disease progression prior to the invasive measurements5,14-17.
Another application of this procedure is to obtain additional data. As an example, using a catheter that can measure flows and pressure, right heart output could be simultaneously recorded together with right heart pressure changes. The procedure outlined here can also be used to further understand the physiologic, cellular and molecular changes that are triggers of pulmonary hypertension other than the immune response, for example, genetic mutations 8,10, cigarette smoke exposure 26, or microbial infections 27-29.

<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 animals, the standard deviation of the &Delta;P, and the heart rate needs to be calculated. Both values are expected to be not significantly different between treatment groups. Furthermore, the expectation is that the &Delta;P values and the heart rate do not show a drift across the recording time. For example, slowly increasing &Delta;P values would indicate hypoxic pulmonary vasoconstriction 18-20. Decreasing heart rate would indicate that the level of anaesthesia is too high leading to the death of the animal. In our laboratory different people perform the procedure. Across all groups of mice we routinely achieve 90-95 % success rates in obtaining right ventricular pressure data.
While learning the procedure, it is best to start with older female mice (25-30 g and more) that have a larger jugular vein and tend to have less subcutaneous fat deposits than male mice. The best way to ascertain the placement of the catheter is to euthanize the animal as soon as the catheter is secured inside of the vein. Dissect the animal starting from the heart to directly visualize the placement of the catheter. This will greatly facilitate problem solving.
The first outcome for obtaining BAL fluid is the correct placement of the tracheal cannula. This is indicated by the inflation of the lungs when Hanks buffer is instilled, and deflation of the lungs when the buffer is removed. The expected recovery is 70-80 % of the instilled volume in control animals, less (down to 50 %) in animals that have lung inflammation. Foam on top of the recovered BAL fluid indicates the presence of surfactant. Blood contamination of the recovered BAL fluid either occurs because of the presence of significant lung inflammation, or because instillation and recovery of the wash fluid was too fast. It is of note that some acute measures of injury within the lungs can be induced by the BAL procedure. If the experimental design is to understand these acute injury parameters and examination of BAL is also important, then one lung lobe needs to be tied off and removed prior to conducting BAL.
In the procedure shown here, we harvest lung tissues for histology without standardized inflation and perfusion of the lungs. In cases where exact histological morphometry is the major read-out, the lungs should be inflated via the trachea, and perfusion should be performed via the pulmonary artery, all under standardized pressure.
In the protocol shown, the heart is dissected to provide the Fulton index as measure of right heart hypertrophy (weight of right ventricle / weight of left ventricle and septum). In addition, new methods are being developed that are based on histological evaluation of the right heart. Emerging histological measures of right heart hypertrophy are a) the number of nuclei within a defined area of randomly selected right heart, and b) counts of vessels, and fibrotic index per area of heart.
In our laboratory, the correct recovery of lung draining mediastinal and tracheobronchial lymph nodes (Figure 5) is routinely verified by flow cytometry because these tissues are very small in control mice. However, depending on the microbiome in the housing facility and the age of the animals, the size of the lymph nodes in unchallenged, control mice may be sufficiently large for only visual confirmation. When verifying by flow cytometry, lymph node cells need to be distinguished from thymocytes. By accident, the thymus can easily be sampled together with the lymph node because the thymus and the lymph nodes are located close to each other and are contained within the rib cage. The distinction is that the thymus is located close to the breast-bone and the lymph node close to the spine. Further challenging is that the thymus is very big, containing many more cells than the small lymph nodes. Thymocytes can easily be distinguished from lymph node cells by using flow cytometry and CD3, CD4, and CD8 markers. Lymph nodes have characteristically CD3+ cells, the majority of which are either positive for CD4 or CD8. In contrast, the majority of thymocytes are positive for all three markers (CD3+, CD4+, CD8+). Further in depth information on the localization of mouse lymph nodes can be found in the manuscript by van den Broeck et al.21.
Consistent timing of each main procedural step (heart catheterization, recovery of blood and spleen, harvest of BAL, lung and heart tissues) is critical for the generation of data that allows for robust comparison between treatment groups. Therefore, it is recommended that at least two, and best three, investigators collaborate to accomplish the experiment. Furthermore, it is critical that the investigators perform the procedure without knowledge of the group identity of the animals. Therefore, the identification of the data has to be made in a manner that obscures the group identity. Finally, it is also important that the order in which the animals are analyzed is randomized. For example, a simple identifier is the date of the experiment and consecutive numbering of the animals. If there are for example 4 groups of animals (A-D), assign identification and study the animals in the following order: date_1: Aa, date_2: Ba, date_3: Ca, date_4: Da, date_5: Ab, date_6: Bb, date_7: Cb, and so on.
This experimental flow allows for an in-depth, simultaneous study of the molecular mechanisms that control lung inflammation and right heart function. The simultaneous data and sample collection achieves better insights into molecular networks, and a reduction in the number of animals needed to obtain novel biological knowledge.
<table border="1">
 <tbody>
 <tr>
 <td>Type of tube / beaker</td>
 <td>Solution (volume)</td>
 <td>Utilization</td>
 </tr>
 <tr>
 <td colspan="3">Processing of blood</td>
 </tr>
 <tr>
 <td>Eppendorf, 1.5 ml</td>
 <td></td>
 <td>Blood for serum</td>
 </tr>
 <tr>
 <td>EDTA coated 2.0 ml</td>
 <td></td>
 <td>Blood for plasma</td>
 </tr>
 <tr>
 <td>Eppendorf, 0.5 ml</td>
 <td></td>
 <td>Freeze serum or plasma</td>
 </tr>
 <tr>
 <td>Processing of BAL</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Polypropylene, round bottom, 4.5 ml</td>
 <td></td>
 <td>Collect bronchoalveolar lavage</td>
 </tr>
 <tr>
 <td>Polypropylene, round bottom, 4.5 ml</td>
 <td></td>
 <td>Freeze BAL supernatant</td>
 </tr>
 <tr>
 <td>96-well plate, skirted</td>
 <td></td>
 <td>BAL cells</td>
 </tr>
 <tr>
 <td colspan="3">Processing of tissue</td>
 </tr>
 <tr>
 <td>Polypropylene, 50 ml</td>
 <td>Formaldehyde, 7.5 ml</td>
 <td>Fix lung tissue </td>
 </tr>
 <tr>
 <td>Eppendorf, 1.5 ml</td>
 <td></td>
 <td>Snap freeze lung lobe</td>
 </tr>
 <tr>
 <td>Eppendorf, 1.5 ml</td>
 <td></td>
 <td>Snap freeze right heart</td>
 </tr>
 <tr>
 <td>Eppendorf, 1.5 ml</td>
 <td></td>
 <td>Snap freeze left heart</td>
 </tr>
 <tr>
 <td>24-well plate</td>
 <td>Hanks buffer, 0.5-1 ml</td>
 <td>Quick-storage of lung lobes for later isolation of single cells</td>
 </tr>
 <tr>
 <td>24-well plate</td>
 <td>Hanks buffer, 0.5-1 ml</td>
 <td>Quick storage of spleens for later isolation of single cells</td>
 </tr>
 <tr>
 <td>24-well plate</td>
 <td>Hanks buffer, 0.5-1 ml</td>
 <td>Quick storage of lymph nodes for later isolation of single cells</td>
 </tr>
 <tr>
 <td colspan="3">For the procedures</td>
 </tr>
 <tr>
 <td>Glass beaker, 300 ml</td>
 <td>Autoclaved water</td>
 <td>Moisturizing and cleaning catheter</td>
 </tr>
 <tr>
 <td>Polypropylene tube, 50 ml</td>
 <td>Hanks buffer, 50 ml</td>
 <td>Perform BAL</td>
 </tr>
 <tr>
 <td>Polypropylene tube, 50 ml</td>
 <td>Hanks buffer, 50 ml</td>
 <td>Wash tracheal cannula</td>
 </tr>
 <tr>
 <td>Polypropylene tube, 50 ml</td>
 <td>Disinfectant solution</td>
 <td>Cleaning instruments</td>
 </tr>
 <tr>
 <td>Polypropylene tube, 50 ml</td>
 <td>70 % ethanol</td>
 <td>Cleaning instruments</td>
 </tr>
 <tr>
 <td>Polypropylene tube, 50 ml</td>
 <td>Tap water</td>
 <td>Cleaning instruments</td>
 </tr>
 </tbody>
</table>
Table 1. Preparation and organization of tubes and solutions.
<table border="1">
 <tbody>
 <tr>
 <td>Instrument</td>
 <td>Description</td>
 <td>Use</td>
 </tr>
 <tr>
 <td>Scissors</td>
 <td>rounded or straight 5.5 inch, round or sharp ends </td>
 <td rowspan="4">Area A</td>
 </tr>
 <tr>
 <td>Scissors</td>
 <td>rounded 4.0 inches, sharp ends </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.5 inch, bent, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.0 inch, bent, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Scissors</td>
 <td>Slim Blades/Angled- 9 cm </td>
 <td rowspan="7">Area B</td>
 </tr>
 <tr>
 <td>Micro-scissors </td>
 <td>Vannas Spring Scissors - 2 mm Blades </td>
 </tr>
 <tr>
 <td>Forceps </td>
 <td>forceps (Dumon #5 Fine) </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.0 inch, bent, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.0 inch, straight, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Suture</td>
 <td>Braided Silk Suture 6-0</td>
 </tr>
 <tr>
 <td>Catheter</td>
 <td>Pressure catheter</td>
 </tr>
 <tr>
 <td>Scissors</td>
 <td>rounded or straight 5.5 inch, round or sharp ends </td>
 <td rowspan="6">Area C</td>
 </tr>
 <tr>
 <td>Scissors</td>
 <td>rounded 4.0 inches, sharp ends </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.5 inch, bent, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td>4.0 inch, bent, with flat, grasping ends </td>
 </tr>
 <tr>
 <td>Cannula</td>
 <td>Tracheal cannula</td>
 </tr>
 <tr>
 <td>Suture</td>
 <td>Braided Silk Suture 4-0</td>
 </tr>
 </tbody>
</table>
Table 2. Organization of instruments, suture, catheter, tracheal cannula.
<img src="/files/ftp_upload/50023/50023fig1.jpg" alt="Figure 1"/> 
 Figure 1. Schematic sketch of the work-areas. A) Work-area A for recording, weighing, collection of blood and spleen tissue and temporary holding space for the animals. B) Work-area B for right heart catheterization. C) Work-area C for harvest of BAL, lungs, lung lymph node, and heart tissues.
<img src="/files/ftp_upload/50023/50023fig2.jpg" alt="Figure 2"/> 
 Figure 2. Instruments used for the procedure. A) Instruments for work-area A; B) instruments for right heart catheterization (work-area B); C) instruments for work-area C. D) Tracheal cannulas. The ruler shows the size of the instruments.
<img src="/files/ftp_upload/50023/50023fig3.jpg" alt="Figure 3"/> 
 Figure 3. Right heart catheter. Right heart catheter shown with marks and tape (A), or held by hand (B). The pencil points to the marks made on the catheter (A). Insertion of the catheter (C-E): cleaning of the right jugular vein (C); sutured vein held with forceps, catheter is being advanced towards a hole previously made into the vein (D); catheter following insertion into the vein (E). Arrows point to the catheter (D, E).
<img src="/files/ftp_upload/50023/50023fig4.jpg" alt="Figure 4" fo:content-width="4in" fo:src="/files/ftp_upload/50023/50023fig1highres.jpg"/> 
 Figure 4. Sample trace of pressure changes in the right heart. The traces show pressure changes (mmHg) over time (min : sec); the software displays each of the curves at two different speeds. The traces of a control mouse (A); and a mouse with inflammation induced pulmonary arterial remodeling and hypertension (B) are shown. Curves used for the measurement of the right systolic ventricular pressure are highlighted. Note the clear pressure plateaus in the highlighted curves. The arrow points to a curve that has a spike indicating a technical error. These spiky types of curves cannot be used for measurement of right ventricular systolic pressures. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50023/50023fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/50023/50023fig5.jpg" alt="Figure 5"/> 
 Figure 5. Location of a mediastinal / tracheobronchial lymph node relative to rib cage and spine. The arrow points to the lymph node.

Watch this Scientific Journal Video about Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice at JoVE.com

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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Nanyang Technological University

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Circulatory and Pulmonary Systems

SCIENTISTS IN ACTION

25.1K Views.  New York University School of Medicine, Tuxedo. Een specifieke en snelle protocol tegelijkertijd onderzoeken juiste hartfunctie, longontsteking en de immuunrespons wordt beschreven als leermiddel. Video en cijfers beschrijven fysiologie en microdissectie technieken in een georganiseerd team-aanpak die aanpasbaar is om te worden gebruikt voor kleine tot grote 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 ...

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. ...

Quantification of Biventricular Function and Morphology by Cardiac Magnetic Resonance Imaging in Mice with Pulmonary Artery Banding

Right ventricular (RV) function and failure are major determinants of outcome in acquired and congenital heart diseases, including pulmonary hypertension. Assessment of RV function and morphology is complex, partly due to the complex shape of the RV. Currently, cardiac magnetic resonance (CMR) imaging is the golden standard for noninvasive assessment of RV function and morphology. The current protocol describes CMR imaging in a mouse model of RV pressure load induced by pulmonary artery banding (PAB). PAB is performed by placing a 6-0 suture around the pulmonary artery over a 23 G needle. The PAB gradient is determined using echocardiography at 2 and 6 weeks. At 6 weeks, the right and left ventricular morphology and function is assessed by measuring both end-systolic and end-diastolic volumes and mass by ten to eleven cine slices 1 mm thick using a 9.4 T magnetic resonance imaging scanner equipped with a 1,500 mT/m gradient. Representative results show that PAB induces a significant increase in RV pressure load, with significant effects on biventricular morphology and RV function. It is also shown that at 6 weeks of RV pressure load, cardiac output is maintained. Presented here is a reproducible protocol for the quantification of biventricular morphology and function in a mouse model of RV pressure load and may serve as a method for experiments exploring determinants of RV remodeling and dysfunction.

To understand the pathophysiology of right ventricular (RV) adaptation to abnormal loading, experimental models are crucial. However, assessment of RV dimensions and function is complex and challenging. This protocol provides a method to perform cardiac magnetic resonance imaging (CMR) as a noninvasive benchmark procedure in mice subjected to RV pressure load.

Right ventricular (RV) function and failure are major determinants of outcome in acquired and congenital heart diseases, including pulmonary hypertension. ...

Induction of Right Ventricular Failure by Pulmonary Artery Constriction and Evaluation of Right Ventricular Function in Mice

The mechanism of right ventricular failure (RVF) requires clarification due to the uniqueness, high morbidity, high mortality, and refractory nature of RVF. Previous rat models imitating RVF progression have been described. Compared with rats, mice are more accessible, economical, and widely used in animal experiments. We developed a pulmonary artery constriction (PAC) approach which is comprised of banding the pulmonary trunk in mice to induce right ventricular (RV) hypertrophy. A special surgical latch needle was designed that allows for easier separation of the aorta and the pulmonary trunk. In our experiments, the use of this fabricated latch needle reduced the risk of arteriorrhexis and improved the surgical success rate to 90%. We used different padding needle diameters to precisely create quantitative constriction, which can induce different degrees of RV hypertrophy. We quantified the degree of constriction by evaluating the blood flow velocity of the PA, which was measured by noninvasive transthoracic echocardiography. RV function was precisely evaluated by right heart catheterization at 8 weeks after surgery. The surgical instruments made inhouse were composed of common materials using a simple process that is easy to master. Therefore, the PAC approach described here is easy to imitate using instruments made in the lab and can be widely used in other labs. This study presents a modified PAC approach that has a higher success rate than other models and an 8-week postsurgery survival rate of 97.8%. This PAC approach provides a useful technique for studying the mechanism of RVF and will enable an increased understanding of RVF.

Here, we provide a useful approach for studying the mechanism of right ventricular failure. A more convenient and efficient approach to pulmonary artery constriction is established using surgical instruments made inhouse. In addition, methods to evaluate the quality of this approach by echocardiography and catheterization are provided.

The mechanism of right ventricular failure (RVF) requires clarification due to the uniqueness, high morbidity, high mortality, and refractory nature of ...

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

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