This work is supported, in part, by R01HL139796 and R01HL128847 grants to CKB and RO1DE028297 and CBET1608058 for DWM.

The use of animals in this study was in accordance with Nationwide Children&#39;s Hospital institutional animal care and use committee under protocol AR13-00030.
1. Pulmonary valve excision
<ol>
	<li>Autoclave the necessary tools needed for the mouse dissection. This includes fine scissors, micro forceps, micro vascular clamps, clamp applying forceps, microneedle holders, spring scissors, and retractors.</li>
	<li>Acclimate all mice for a minimum of 2 weeks before the operation. Remove C57BL/6 mice, approximately 1 year of age, from their cages and weigh, then euthanize with a ketamine/xylazine cocktail (3:1 ketamine:xylazine, 0.01 mL per g) overdose.</li>
	<li>Place the mouse in a dorsal recumbence position on a tray and secure its limbs with tape. Once secured, perform the thoracotomy.
	<ol>
		<li>Expose the heart by removing any excess adipose tissue and fascia.</li>
		<li>Remove the right atrium and perfuse through the left ventricle with room temperature saline solution (0.9% NaCl). The perfusion should take approximately 20 mL over 30 s. This results in the exsanguination of the mouse.</li>
	</ol>
	</li>
	<li>Remove the entire heart by severing the superior vena cava, inferior vena cava, pulmonary artery, and aorta. Special care should be taken for the pulmonary artery. Cut approximately 2 mm above the ventriculo-arterial junction, as this will serve as the conduit for pressurization.</li>
	<li>Remove the left and right ventricles to expose chambers to atmospheric pressure. Ensure that the structure of the pulmonary trunk should be unaffected by the removal of the ventricles.</li>
</ol>
2. Pressure fixation of pulmonary valve
<ol>
	<li>Anastomose pressurization tubing with pulmonary artery, leaving approximately 1 mm distance between the sino-tubular junction and the end of the tubing to accommodate for large movements of the leaflets and pulmonary trunk.</li>
	<li>Elevate the reservoir to an analogous physiological pressure and fill it with the saline solution. Test the flow-through system to ensure there are no blockages or air bubbles.</li>
	<li>Attach a stopcock to the anastomosed pulmonary valve and ensure adequate flow through the tubing (i.e., no air bubbles) by switching the outflow tract. Once the flow is adequate, switch the outflow to the anastomosed pulmonary valve and ensure pressurization of the pulmonary trunk. This is identified by pulmonary trunk distention.</li>
	<li>Once the pressurization of the pulmonary trunk is confirmed, gradually incorporate primary fixative solution (1.25% glutaraldehyde, 1.0% paraformaldehyde in 0.15 M cacodylate) until the saline solution is purged. This is done by removing a portion, approximately 25% of the reservoir capacity, of the saline solution and replacing it with the primary fixative. 
	CAUTION: The fixatives used (paraformaldehyde and glutaraldehyde) are highly toxic and appropriate personal protective equipment (PPE) should be worn to ensure safety.</li>
	<li>Place a fixative-soaked gauze over the tissue sample to prevent drying.</li>
	<li>Perfuse the fixative for 3 h, refilling the reservoir as needed to maintain a constant pressure. Throughout fixation, it is not uncommon for the pulmonary valve to shrink because of the chemical fixation. If this is the case, constantly replenish the reservoir with primary fixative to maintain physiological pressure.</li>
	<li>Store the heart valve in the fixative solution at 4 &#176;C until use. Samples were stored for up to 1 week without any discernable difference.</li>
</ol>
3. En bloc sample staining and embedment15,16
CAUTION: The staining reagents used in this section (potassium ferrocyanide, osmium tetroxide, thiocarbohydrazide, lead aspartate, and uranyl acetate) are highly toxic and should be handled with extreme care. Use of a fume hood and proper PPE is advised.
<ol>
	<li>Staining
	<ol>
		<li>Wash the fixed heart valve sample for 5 min with cold 0.15 M cacodylate buffer. Repeat the wash two more times.</li>
		<li>Completely submerge the heart valve in a solution of 1.5% potassium ferrocyanide, 0.15 M cacodylate, 2 mM calcium chloride, and 2% osmium tetroxide, on ice for 1 h.</li>
		<li>While the sample is incubating, prepare thiocarbohydrazide (TCH) solution by dissolving 0.1 g of TCH in 10 mL of ddH2O. Place the solution in a 60 &#176;C oven for 1 h. Gently agitate periodically to ensure TCH is dissolved completely. Filter the solution through a 0.22 &#181;m syringe filter immediately before use.</li>
		<li>Wash the samples with room temperature ddH2O by placing them in a tube of ddH2O for 5 min and slightly agitate it by shaking the container. Repeat this process three times.</li>
		<li>Place it in filtered TCH solution for 20 min at room temperature. Perform the washing step three times (5 min each) with room temperature ddH2O as described in step 3.1.4.</li>
		<li>Once done, place the sample in 2% osmium tetroxide for 30 min at room temperature. Then wash it again for a total of three times for 5 min each with room temperature ddH2O.</li>
		<li>Incubate the sample in 1% uranyl acetate overnight at 4 &#176;C.</li>
		<li>During this time, make a solution of 0.066 g of lead nitrate in 10 mL of aspartic acid stock solution. Adjust the pH to 5.5 with 1 N KOH. Place the solution in a 60 &#176;C oven to dissolve lead nitrate.</li>
		<li>After the overnight incubation, perform the washing step as described in step 3.1.4. Repeat three times. Then, incubate the heart valve tissue in lead aspartate solution from step 3.1.8 in a 60 &#176;C oven for 30 min.</li>
	</ol>
	</li>
	<li>Dehydration
	<ol>
		<li>Wash the tissues for 5 min at room temperature with ddH2O. Repeat three times.</li>
		<li>Make fresh solutions of 20%, 50%, 70%, 90%, and 100% ethanol in ddH2O. Keep on ice until use.</li>
		<li>To dehydrate the heart valve tissue. Perform subsequent treatments of 20%, 50%, 70%, and 90% ethanol on ice for 5 min each. Then perform two subsequent treatments of 100% ethanol on ice for 5 min.</li>
		<li>Move the tissue to ice-cold acetone for 10 min. Then place in fresh acetone at room temperature for 10 min.</li>
	</ol>
	</li>
	<li>Embedding
	<ol>
		<li>Make the resin mixture (see Table of Materials) according to the manufacturer&#39;s specifications: 11.4 g of component A, 10 g of component B, 0.3 g of component C, and 0.05-0.1 g of component D. Initially mix components A and B by heating each of the components to 60 &#176;C before adding components C and D. The mixture will turn into an amber color upon adding components C and D. It will be uniform when properly mixed.</li>
		<li>Make mixtures with volume ratios of 25:75, 50:50, and 75:25 resin:acetone. Mix thoroughly.</li>
		<li>Place tissues in subsequent treatments of the 25:75, 50:50, and 75:25 resin:acetone mixture for 2 h each at room temperature.</li>
		<li>Place tissues in 100% resin overnight at room temperature.</li>
		<li>The following day, place tissues in fresh 100% resin for 2 h at room temperature.</li>
		<li>Transfer the heart valve tissues to an embedding capsule, substitute with fresh 100% resin, and place in a 60 &#176;C oven for 48 h to cure.</li>
	</ol>
	</li>
	<li>Perform correlative imaging as shown below.</li>
</ol>
4. Micro-computed tomography imaging
<ol>
	<li>Mount the resin sample block onto a &#181;CT sample holder with an adhesive (glue or double-sided tape works well).</li>
	<li>Place the holder into &#181;CT chamber and fix it onto the stage by screwing the collet such that there is no motion of the holder. Sample movement during the scan will decrease the image quality.</li>
	<li>Open the &#181;CT user interface by double-clicking on the icon. Add a project by selecting the + sign next to Projects and fill in the necessary fields.</li>
	<li>Once this is created, find the created project, and select it by clicking on it. This will open up a second column Acquisitions. Select the + next to Acquisitions and fill out the necessary fields.</li>
	<li>Close the chamber doors and arm the system by hitting the arming button on the front panel or the &#181;CT. Warmup x-rays from the user interface by selecting the button indicating Warmup. 
	NOTE: The system will automatically turn off the x-rays following a successful warm-up procedure. Turn on x-rays by selecting the button.</li>
	<li>Adjust the stage rotation such that the center of the rotation of the sample does not deviate from the center of the monitor (i.e., the sample is within the field of view for the entire scan). For the system used for this experiment, this involves adjusting the region of interest (ROI) stage under the drop-down tab as detailed below.
	<ol>
		<li>Set the ROI stage adjustment by setting the rotation angle to 0&#176; and marking the edge of the sample. The sample is then rotated to 180&#176; and the edge of the sample is marked again.</li>
		<li>Adjust the ROI stage x-axis position such that the edge of the sample is between these two extremes. Repeat this process for rotation angles of 90&#176; and 270&#176; for the y-position.</li>
		<li>In the case where the rotation of the sample causes the edges to move out of the field of view, decrease the magnification of the &#181;CT needs until the edges of the sample are visible at the above set angles and a coarse ROI stage adjustment can be made.</li>
		<li>Once adjusted at the lower magnification, move the sample or the detector to increase the magnification and the ROI positions can be fine adjusted. 
		NOTE: This process may need to be repeated to ensure alignment. Proper alignment results in a sample that shows little to no horizontal movement through all sample rotation angles.</li>
	</ol>
	</li>
	<li>Adjust the &#181;CT to desired scanning parameters using the manufacturer&#39;s software. These parameters include tube potential, tube current, detector distance, sample distance, exposure time, trajectory, and the number of projections (see Table S1 for the &#181;CT acquisition parameters used in this study). 
	NOTE: Calibration parameters, such as clear field and dark field scans, should be kept at the manufacturer&#39;s specifications unless otherwise stated. For example, if a sample is too large and the system cannot move the sample out of the field view, then the sample needs to be removed to perform clear field calibration scans.</li>
	<li>Once the parameters are set, an approximation of the duration of the scan can be seen by hitting the Time Estimate button. Select the Start button to begin the scan.</li>
	<li>Following the scan completion, ensure that the x-rays are turned off, unscrew the collet, and carefully remove the sample from the &#181;CT chamber.</li>
</ol>
5. Sample processing and image correlation
<ol>
	<li>Reconstruct &#181;CT projections using a filtered back projection algorithm with the software provided by the manufacturer (see Table of Materials).
	<ol>
		<li>Select the Recon tab at the top of the screen. Select the Project and Acquisition tab.</li>
		<li>Select the recon template associate with filtered back projection. Hit the Start Recon button.</li>
	</ol>
	</li>
	<li>Identify and segment the pulmonary valve using image processing software. This requires prior knowledge of the anatomy of the pulmonary valve17. At elevated arterial pressures, the leaflets coapt and block the lumen of the pulmonary valve.</li>
	<li>Identify the slicing orientation with respect to the scanning direction and sample. Reorient the sample such that the slicing direction is aligned with the desired axis.</li>
	<li>Identify regions of interest for high-resolution imaging. In this experiment, the belly (middle) of the leaflet and the arterio-valvular junction was chosen.</li>
	<li>Remove the excess resin and sample by either razor blade or sanding for large pieces of material, or by microtome for finer pieces.</li>
	<li>Once at a location of interest, compare the physical specimen with virtual &#181;CT slices to confirm location. This was done by comparing anatomical features at the cross section.</li>
</ol>
6. Serial block face scanning electron microscopy18
<ol>
	<li>Reduce the cross-section of the specimen to accommodate SBF-SEM, approximately 2.0 x 1.5 x 1.8 mm.</li>
	<li>Mount the trimmed sample onto the SBF-SEM aluminum stub using epoxy.</li>
	<li>Coat the specimen block with 35 nm gold. Spin the sample on the platform to apply an even layer of coating.</li>
	<li>Vent the SBF-SEM chamber and adjust the height of the knife blade to microscope eucentric height.</li>
	<li>Insert the sample and level the sample stub.</li>
	<li>Image under low vacuum conditions to prevent charging and with backscatter detector (see Table S2 for SBF-SEM acquisition parameters).</li>
	<li>Image and stitch multiple regions of interest at different magnifications using automated software (see Table of Materials).</li>
</ol>

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The authors have nothing to disclose.

Removal of the ventricles serves two purposes. First, exposing the ventricle side to the atmospheric pressure, thereby only needing to apply a transvalvular pressure from the arterial side of the pulmonary valve to close, and second, providing a stable base to prevent twisting of the pulmonary trunk. During pressurization, the pulmonary trunk distends radially and inferiorly, making it prone to twisting, causing the collapse of the pulmonary trunk. Preloading the pulmonary valve with a saline solution offers an additiona...

The pulmonary valve (PV) serves to ensure unidirectional blood flow between the right ventricle and the pulmonary artery. Pulmonary valve malformations are associated with several forms of congenital heart disease. The current treatment for congenital heart valve disease (HVD) is valvular repair or valve replacement, which can necessitate multiple invasive surgeries throughout a patient&#39;s lifetime1. It has been widely accepted that the function of the heart valve is derived from its structure, often referred to as the structure-function correlate. More specifically, the geometric and biomechanical properties of the heart dictate its function. The mechanical properties, in turn, are determined by the composition and organization of the ECM. By developing a method for determining the biomechanical properties of murine heart valves, transgenic animal models can be used to interrogate the role of the ECM on heart valve function and dysfunction2,3,4,5.
The murine animal model has long been regarded as the standard for molecular studies because transgenic models are more readily available in mice compared to other species. Murine transgenic models provide a versatile platform for researching heart valve-related diseases6. However, the surgical expertise and instrumentation requirements to characterize both the geometry and ECM organization have been a major hurdle in progressing HVD research. Hstological data in literature provides a picture into murine heart valve extracellular matrix content, but only in the form of 2D images, and are unable to describe its 3D architecture7,8. Additionally, the heart valve is both spatially and temporally heterogeneous, making it difficult to draw conclusions across experiments regarding ECM organization if the sampling and conformation are not fixed. Conventional 3D characterization methods, such as MRI or 3D echocardiography, do not provide the resolution necessary to resolve ECM components9,10.
This work details a fully correlative workflow where the temporal heterogeneity due to the cardiac cycle was addressed by fixing the conformation of the murine PV with hydrostatic transvalvular pressure. The spatial heterogeneity was controlled precisely by sampling regions of interest and registering data sets from different imaging modalities, specifically &#181;CT and serial block face scanning electron microscopy, across different length scales. This method of scouting with &#181;CT for guiding downstream sampling has been proposed previously, but because the pulmonary valve exhibits temporal variation, an additional level of control was needed on the surgical level11.
In vivo studies describing murine heart valve biomechanics are sparse and, instead, rely on computational models when describing the deformation behavior. It is of critical importance that local extracellular data on the nanometer length scale be related to the geometry and location of the heart valve. This, in turn, provides quantifiable, spatially mapped distributions of mechanically contributing ECM proteins, which can be used to reinforce existing biomechanical heart valve models12,13,14.

<table><tbody><tr><td>25% glutaraldehyde (aq)</td><td>EMS</td><td>16210</td><td>Primary fixative component</td></tr><tr><td>0.9% sodium chloride injection</td><td>Hospira Inc.</td><td>NDC 0409-4888-10</td><td></td></tr><tr><td>1 mL syringe</td><td>BD</td><td>309659</td><td></td></tr><tr><td>10 mL syringe</td><td>BD</td><td>309604</td><td></td></tr><tr><td>200 proof ethanol</td><td>EMS</td><td>15055</td><td></td></tr><tr><td>22G needle</td><td>BD</td><td>305156</td><td></td></tr><tr><td>3 mL syringe</td><td>BD</td><td>309657</td><td></td></tr><tr><td>3-way stopcock</td><td>Smiths Medical ASD, Inc.</td><td>MX5311L</td><td></td></tr><tr><td>4% osmium tetroxide</td><td>EMS</td><td>19150</td><td>Staining component</td></tr><tr><td>4% paraformaldehyde (aq)</td><td>EMS</td><td>157-4-100</td><td>Primary fixative component</td></tr><tr><td>Absorbable hemostat</td><td>Ethicon</td><td>1961</td><td></td></tr><tr><td>Acetone</td><td>EMS</td><td>10012</td><td></td></tr><tr><td>Black polyamide monofilament suture, 10-0</td><td>AROSurgical instruments Corporation</td><td>TI38402</td><td></td></tr><tr><td>Black polyamide monofilament suture, 6-0</td><td>AROSurgical instruments Corporation</td><td>SN-1956</td><td></td></tr><tr><td>C57BL/6 mice</td><td>Jackson Laboratories</td><td>664</td><td>Approximately 1 yo</td></tr><tr><td>Calcium chloride</td><td>Sigma-Aldrich</td><td>10043-52-4</td><td></td></tr><tr><td>Clamp applying forcep</td><td>FST</td><td>00072-14</td><td></td></tr><tr><td>Cotton tip applicators</td><td>Fisher Scientific</td><td>23-400-118</td><td></td></tr><tr><td>DPBS</td><td>Gibco</td><td>14190-144</td><td></td></tr><tr><td>Dumont #5 forcep</td><td>FST</td><td>11251-20</td><td></td></tr><tr><td>Dumont #5/45 forceps</td><td>FST</td><td>11251-35</td><td></td></tr><tr><td>Dumont #7 fine forcep</td><td>FST</td><td>11274-20</td><td></td></tr><tr><td>Durcupan ACM resin</td><td>EMS</td><td>14040</td><td>For embedding</td></tr><tr><td>Fine scissor</td><td>FST</td><td>14028-10</td><td></td></tr><tr><td>Heliscan microCT</td><td>Thermo Fisher Scientific</td><td></td><td>Micro-CT</td></tr><tr><td>Ketamine hydrochloride injection</td><td>Hospira Inc.</td><td>NDC 0409-2053</td><td></td></tr><tr><td>L-aspartic acid</td><td>Sigma-Aldrich</td><td>56-84-8</td><td>Staining component</td></tr><tr><td>Lead nitrate</td><td>EMS</td><td>17900</td><td>Staining component</td></tr><tr><td>low-vacuum backscatter detector</td><td>Thermo Fisher Scientific</td><td>VSDBS</td><td>SEM backscatter detector</td></tr><tr><td>Micro-adson forcep</td><td>FST</td><td>11018-12</td><td></td></tr><tr><td>Millex-GP filter, 0.22 um, PES 33mm, non-sterile</td><td>EMD Millipore</td><td>SLGP033NS</td><td></td></tr><tr><td>Non-woven songes</td><td>McKesson Corp.</td><td>94442000</td><td></td></tr><tr><td>Potassium hexacyanoferrate(II) trihydrate</td><td>Sigma-Aldrich</td><td>14459-95-1</td><td>Staining component</td></tr><tr><td>Potassium hydroxide</td><td>Sigma-Aldrich</td><td>1310-58-3</td><td></td></tr><tr><td>Pressure monitor line</td><td>Smiths Medical ASD, Inc.</td><td>MX562</td><td></td></tr><tr><td>Saline solution (sterile 0.9% sodium chloride)</td><td>Hospira Inc.</td><td>NDC 0409-0138-22</td><td></td></tr><tr><td>Size 3 BEEM capsule</td><td>EMS</td><td>69910-01</td><td>Embedding container</td></tr><tr><td>Sodium cacodylate trihydrate</td><td>Sigma-Aldrich</td><td>6131-99-3</td><td>Buffer</td></tr><tr><td>Solibri retractors</td><td>FST</td><td>17000-04</td><td></td></tr><tr><td>Sputter, carbon and e-beam coater</td><td>Leica</td><td>EM ACE600</td><td>Gold coater</td></tr><tr><td>Surgical microscope</td><td>Leica</td><td>M80</td><td></td></tr><tr><td>Thiocarbohydrazide (TCH)</td><td>EMS</td><td>21900</td><td>Staining component</td></tr><tr><td>Tish needle holder/forcep</td><td>Micrins</td><td>MI1540</td><td></td></tr><tr><td>Trimmer</td><td>Wahl</td><td>9854-500</td><td></td></tr><tr><td>Uranyl acetate</td><td>EMS</td><td>22400</td><td>Staining component</td></tr><tr><td>Volumescope scanning electron microscope</td><td>Thermo Fisher Scientific</td><td>VOLUMESCOPESEM</td><td>Serial Block Face Scanning Electron Microscope</td></tr><tr><td>Xylazine sterile solution</td><td>Akorn Inc.</td><td>NADA# 139-236</td><td></td></tr></tbody></table>

surgery and sample processing for correlative imaging of the murine pulmonary valve

The underlying causes of heart valve related-disease (HVD) are elusive. Murine animal models provide an excellent tool for studying HVD, however, the surgical and instrumental expertise required to accurately quantify the structure and organization across multiple length scales have stunted its advancement. This work provides a detailed description of the murine dissection, en bloc staining, sample processing, and correlative imaging procedures for depicting the heart valve at different length scales. Hydrostatic transvalvular pressure was used to control the temporal heterogeneity by chemically fixing the heart valve conformation. Micro-computed tomography (&#181;CT) was used to confirm the geometry of the heart valve and provide a reference for the downstream sample processing needed for the serial block face scanning electron microscopy (SBF-SEM). High-resolution serial SEM images of the extracellular matrix (ECM) were taken and reconstructed to provide a local 3D representation of its organization. &#181;CT and SBF-SEM imaging methods were then correlated to overcome the spatial variation across the pulmonary valve. Though the work presented is exclusively on the pulmonary valve, this methodology could be adopted for describing the hierarchical organization in biological systems and is pivotal for the structural characterization across multiple length scales.

Anastomosis of the pulmonary artery to the pressurization tubing is shown in Figure 1A. Following the application of hydrostatic pressure, the pulmonary trunk distends radially (Figure 1B) indicating that the pulmonary valve leaflets are in a closed configuration. Pulmonary valve conformation was confirmed by &#181;CT. In this case, the leaflets were coapt (closed) and the annulus was circular (Figure 2A). Figur...

Watch this Scientific Journal Video about Surgery and Sample Processing for Correlative Imaging of the Murine Pulmonary Valve at JoVE.com

Surgery and Sample Processing for Correlative Imaging of the Murine Pulmonary Valve

Here, we describe a correlative workflow for the excision, pressurization, fixation, and imaging of the murine pulmonary valve to determine the gross conformation and local extracellular matrix structures.

The underlying causes of heart valve related-disease (HVD) are elusive. Murine animal models provide an excellent tool for studying HVD, however, the ...

surgery-sample-processing-for-correlative-imaging-murine-pulmonary

Universidad de Cartagena UDEC

Research

JoVE Journal

Engineering

2.5K Views.  Ohio State University. Here, we describe a correlative workflow for the excision, pressurization, fixation, and imaging of the murine pulmonary valve to determine the gross conformation and local extracellular matrix structures.

Video: Surgery and Sample Processing for Correlative Imaging of the Murine Pulmonary Valve

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, ...

Medicine

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

Echocardiographic Approaches and Protocols for Comprehensive Phenotypic Characterization of Valvular Heart Disease in Mice

The aim of this manuscript and accompanying video is to provide an overview of the methods and approaches used for imaging heart valve function in rodents, with detailed descriptions of the appropriate methods for anesthesia, the echocardiographic windows used, the imaging planes and probe orientations for image acquisition, the methods for data analysis, and the limitations of emerging technologies for the evaluation of cardiac and valvular function. Importantly, we also highlight several future areas of research in cardiac and heart valve imaging that may be leveraged to gain insights into the pathogenesis of valve disease in preclinical animal models. We propose that using a systematic approach to evaluating cardiac and heart valve function in mice can result in more robust and reproducible data, as well as facilitate the discovery of previously underappreciated phenotypes in genetically-altered and/or physiologically-stressed mice.

This protocol provides a detailed description of the echocardiographic approach for comprehensive phenotyping of heart and heart valve function in mice.

The aim of this manuscript and accompanying video is to provide an overview of the methods and approaches used for imaging heart valve function in ...

Murine Echocardiography of Left Atrium, Aorta, and Pulmonary Artery

The present methodology teaches the investigator how to measure and use the LAV as a surrogate of chronic elevations in Left Ventricular diastolic pressure through echocardiography, as well as to obtain measurements of the Aorta and PA diameter in mice.
Mice older than 10 d of age can be analyzed using the present technique. The technique is composed of 3 main steps: set-up, image acquisition, and image analysis. The set-up step consists of getting the mouse anesthetized with 1% isoflurane, shaving it, and taping it in a supine position to a heated EKG board where the image acquisition will take place. The image acquisition step consists of learning to identify the cardiac structures and obtaining all the required images with its correspondent probe and axis in order to be able to calculate volumes and diameters. The image analysis step consists of measuring the previously acquired images with the aid of computer software.
Advantages of the proposed technique include a fast (15 min) procedure that would allow the researcher to evaluate interventions in a non-invasive, non-terminal approach and therefore follow the same mouse over time; each mouse can be used as its own control. This fact plus having the same operator perform all the acquisition and analysis for the entire experiment minimizes the limitation of operator-dependency. The present methodology is useful for mouse researchers in cardiovascular and pulmonary medicine.

The following protocol describes the methodology for the acquisition and analysis of echocardiographic images used to obtain the Left Atrial Volume (LAV), Aorta (Ao) diameter, and Pulmonary Artery (PA) diameter in mice. This technique is a non-invasive, non-terminal procedure that allows assessment of the cardiopulmonary function.

The present methodology teaches the investigator how to measure and use the LAV as a surrogate of chronic elevations in Left Ventricular diastolic ...

Evaluation of Hydrodynamic Properties in Mounted Porcine Aortic Valves

An Ex Vivo Porcine Model for Hydrodynamic Testing of Experimental Aortic Valve Procedures and Novel Medical Devices

The options for testing new cardiac procedures and investigative medical devices prior to use in an animal model are limited. In this study, we present a method for mounting a porcine aortic valve in a pulse duplicator to evaluate its hydrodynamic properties. These properties can then be evaluated before and after the procedure under investigation is performed and/or the investigative medical device is applied. Securing the inflow segment presents some difficulty owing to the lack of circumferential myocardium in the left ventricular outflow tract. This method addresses that issue by securing the inflow segment using the anterior leaflet of the mitral valve and then suturing the left ventricular free wall around the inflow fixture. The outflow segment is secured simply by inserting the fixture into an incision in the superior aspect of the aortic arch. We found that specimens had significantly different hydrodynamic properties before and after tissue fixation. This finding induced us to use fresh specimens in our testing and should be considered when using this method. In our work, we used this method to test novel intracardiac patch materials for use in the valvular position by performing an aortic valve neocuspidization procedure (Ozaki procedure) on the mounted porcine aortic valves. These valves were tested before and after the procedure to assess the change in hydrodynamic properties in comparison to the native valve. Herein, we report a platform for hydrodynamic testing of experimental aortic valve procedures that enables comparison with the native valve and between different devices and techniques used for the procedure under investigation.

Author Spotlight: Advancing Cardiac Procedure Testing Prior to Embarking on Large Animal Studies 

We present a method for mounting a porcine aortic valve on a pulse duplicator to test its hydrodynamic properties. This method can be used to determine the change in hydrodynamics after the application of an experimental procedure or novel medical device prior to use in a large animal model.

The options for testing new cardiac procedures and investigative medical devices prior to use in an animal model are limited. In this study, we present a ...

Bioengineering

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. ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...

Biology

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, the preservation of pulmonary vessels that truly represents physiological conditions still presents challenges. In addition, the delicate configuration of murine lungs result in technical challenges preparing samples for high-quality images that preserve both cellular composition and architecture. Similarly, cellular contractility assays can be performed to study the potential of cells to respond to vasoconstrictors in vitro, but these assays do not reproduce the complex environment of the intact lung. In contrast to these technical issues, the precision-cut lung slice (PCLS) method can be applied as an efficient alternative to visualize lung tissue in three dimensions without regional bias and serve as a live surrogate contractility model for up to 10 days. Tissue prepared using PCLS has preserved structure and spatial orientation, making it ideal to study disease processes ex vivo. The location of endogenous tdTomato-labeled cells in PCLS harvested from an inducible tdTomato reporter murine model can be successfully visualized by confocal microscopy. After exposure to vasoconstrictors, PCLS demonstrates the preservation of both vessel contractility and lung structure, which can be captured by a time-lapse module. In combination with the other procedures, such as western blot and RNA analysis, PCLS can contribute to the comprehensive understanding of signaling cascades that underlie a wide variety of disorders and lead to a better understanding of the pathophysiology in pulmonary vascular diseases.

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

Presented here is a protocol for preserving the vascular contractility of PCLS murine lung tissue, resulting in a sophisticated three-dimensional image of the pulmonary vasculature and airway, which can be preserved for up to 10 days that is susceptible to numerous procedures.

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, ...

Revealing Murine Pulmonary Veins Myocardial Architecture

Microdissection and Immunofluorescence Staining of Myocardial Sleeves in Murine Pulmonary Veins

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial fibrillation (AF). PVs contain myocardial sleeves (MS) composed of cardiomyocytes. MS are implicated in the initiation and maintenance of AF, as they preserve similarities to the cardiac working myocardium, including the ability to generate ectopic electrical impulses. Rodents are widely used and may represent excellent animal models to study the pulmonary vein myocardium since cardiomyocytes are widely present all over the vessel wall. However, precise microdissection and preparation of murine PVs is challenging due to the small organ size and intricate anatomy.
We demonstrate a microscopy-guided microdissection protocol for isolating the murine left atrium (LA) together with the PVs. Immunofluorescence staining using cardiac Troponin-T (cTNT) and connexin 43 (Cx43) antibodies is performed to visualize the LA and PVs in full length. Imaging at 10x and 40x magnification provides a comprehensive view of the PV structure as well as detailed insights into the myocardial architecture, particularly highlighting the presence of connexin 43 within the MS.

Author Spotlight: Developing a Translational Model for Atrial Fibrillation Research Across Species 

This protocol demonstrates microscopy-guided isolation and immunofluorescence staining of murine pulmonary veins. We prepare tissue samples containing the left atrium, pulmonary veins, and the corresponding lungs and stain them for cardiac Troponin T and Connexin 43.

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial ...

Creating a Murine Model of RV Hypertrophy and Failure by Pulmonary Trunk Banding

A Murine Model of Pressure Overload-Induced Right Ventricular Hypertrophy and Failure by Pulmonary Trunk Banding

Right ventricular (RV) failure caused by pressure overload is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary diseases. The pathogenesis of RV failure is complex and remains inadequately understood. To identify new therapeutic strategies for the treatment of RV failure, robust and reproducible animal models are essential. Models of pulmonary trunk banding (PTB) have gained popularity, as RV function can be assessed independently of changes in the pulmonary vasculature.
In this paper, we present a murine model of RV pressure overload induced by PTB in 5-week-old mice. The model can be used to induce different degrees of RV pathology, ranging from mild RV hypertrophy to decompensated RV failure. Detailed protocols for intubation, PTB surgery, and phenotyping by echocardiography are included in the paper. Furthermore, instructions for customizing instruments for intubation and PTB surgery are given, enabling fast and inexpensive reproduction of the PTB model.
Titanium ligating clips were used to constrict the pulmonary trunk, ensuring a highly reproducible and operator-independent degree of pulmonary trunk constriction. The severity of PTB was graded by using different inner ligating clip diameters (mild: 450 &#181;m and severe: 250 &#181;m). This resulted in RV pathology ranging from hypertrophy with preserved RV function to decompensated RV failure with reduced cardiac output and extracardiac manifestations. RV function was assessed by echocardiography at 1 week and 3 weeks after surgery. Examples of echocardiographic images and results are presented here. Furthermore, results from right heart catheterization and histological analyses of cardiac tissue are shown.

Author Spotlight: Investigating the Underlying Mechanisms of Right Ventricular Failure in Pulmonary Hypertension

We describe a murine model of right ventricular pressure overload-induced by pulmonary trunk banding. Detailed protocols for intubation, surgery, and phenotyping by echocardiography are included in the paper. Custom-made instruments are used for intubation and surgery, allowing for fast and inexpensive reproduction of the model.

Right ventricular (RV) failure caused by pressure overload is strongly associated with morbidity and mortality in a number of cardiovascular and pulmonary ...

Surgery and Sample Processing for Correlative Imaging of the Murine Pulmonary Valve

Summary

Explore More Videos

Chapters in this video

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

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Echocardiographic Approaches and Protocols for Comprehensive Phenotypic Characterization of Valvular Heart Disease in Mice

Murine Echocardiography of Left Atrium, Aorta, and Pulmonary Artery

An Ex Vivo Porcine Model for Hydrodynamic Testing of Experimental Aortic Valve Procedures and Novel Medical Devices

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

Isolation of Murine Valve Endothelial Cells

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

Microdissection and Immunofluorescence Staining of Myocardial Sleeves in Murine Pulmonary Veins

A Murine Model of Pressure Overload-Induced Right Ventricular Hypertrophy and Failure by Pulmonary Trunk Banding