The use of mice for these studies is necessary as mice have four-chambered hearts that can mimic human CHDs. Mice were provided veterinary care and housed in the institution&#39;s Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC)-accredited animal care facility. Strict protocols were followed to minimize the mice&#39;s discomfort, stress, pain, and injury. Mice were euthanized using CO2 gas, which is acceptable for small rodents according to the American Veterinary Medical Association Guidelines on Euthanasia. The studies on mice in this manuscript were carried out with an approved IACUC protocol at the University of Pittsburgh.
1. Fetal echocardiogram
NOTE: An echocardiogram is a powerful tool for identifying cardiovascular malformation and extracardiac defects in mice. Due to the small size of the mouse embryos (about 1-2 mm at midgestation, 3.5 mm at birth), ultrahigh-frequency echocardiographic equipment with ultrasound biomicroscopy (UBM) is required. UBM provides different high-frequency (30-50 MHz) probes with a small imaging window (15 mm x 14 mm) that provides the resolution (30 &#181;m axial x 68 &#181;m lateral) to visualize one mouse fetus at a time. A 40 MHz transducer provides high-resolution images to identify cardiovascular phenotypes7.
<ol>
	<li>Turn on the echocardiogram machine and select the program Cardiology. 
	NOTE: The following protocol can be used for any mouse background from embryonic day (E)14.5 to 19.5.</li>
	<li>Anesthetize the desired mouse in an anesthetic induction chamber. Induce anesthesia using a concentration of 4% isoflurane and medical oxygen at a flow rate of 1 L/min and reduce it to 2%-3% for maintenance.</li>
	<li>Place the mouse on the imaging platform quickly. The imaging platform has heated steel to keep the mouse warm during the procedure. Put the mouth and nose of the mouse into the anesthetic nose cone. Secure the limbs with tape to avoid movement. Monitor the heart rate to ensure that it stays between 400-450 bpm.</li>
	<li>Monitor the temperature using a rectal thermometer probe and make sure it rests at 37 &#176;C &#177; 0.5 &#176;C. Monitor the breathing to avoid hypoxia. Keep a gentle force on the probe to prevent harm. 
	NOTE: A heat lamp can be set above the mouse to prevent hypothermia while under and to recover from anesthesia. Petrolatum ophthalmic ointment can be used as a lubricant to avoid dry eyes.</li>
	<li>Remove the fur from the thorax and abdomen using depilatory cream. Apply the cream and wait 3 min before removing. Clean the area with 70% ethanol. Ethanol works better than water as a shaving lubricant.</li>
	<li>Warm up ultrasound gel to a normal body temperature. Apply the ultrasound gel generously and place the transducer on the abdomen to orient it in a horizontal plane and identify the bladder on the screen. Once the bladder is identified, scan cranially from the bladder and look for the fetus. Measure the crown-to-rump length to determine the gestational age10 (Figure 1 and Table 1). 
	NOTE: Change transducer positions to visualize different planes, including the transverse four-chamber, sagittal, and frontal/coronal imaging planes7 (Figure 1).</li>
	<li>Use the color Doppler to analyze blood flow from the heart.</li>
	<li>Put the mouse back in the cage if the embryos have not reached the intended stage. Otherwise, prepare the mouse for harvest. 
	NOTE: Ensure that the mouse is already awake and recovering well from anesthesia before putting it back into the cage.</li>
	<li>Turn off isoflurane and oxygen, clean the work area, and turn off the machine. 
	NOTE: It is important to remove the gel from the transducer.</li>
</ol>
2. Necropsy
NOTE: Once abnormal cardiac phenotypes are suspected using fetal echocardiography, fetuses are collected and fixed via full-body submersion in the fixative solution: either 10% buffered formalin phosphate or 4% paraformaldehyde (PFA). Inspect the sample&#39;s external and internal morphology, looking for macroscopic anatomical abnormalities or malformations.
<ol>
	<li>Prepare the mouse.
	<ol>
		<li>If the mouse is an adult, euthanize the mouse using a standard CO2 protocol. Use forceps or dissecting scissors to make incisions (about 3 cm) in the lateral thorax and abdomen to allow penetration of the fixative into the internal organs. 
		NOTE: The sample should be fixed for at least 24 h before necropsy if the embryo is older than E14.5.</li>
	</ol>
	</li>
	<li>Analyze the exterior of the body.
	<ol>
		<li>Set up the software to save the pictures with a name, including the sample&#39;s identification, the microscope magnification, and the content of the picture. 
		NOTE: Magnification of 1.0x to 3.2x should be adequate for imaging most structures in E14.5 mice or older.</li>
		<li>Place the mouse on the plate under the stereomicroscope lens. Fill the plate with phosphate-buffered saline (PBS) to completely cover the sample to prevent dehydration and reflection in the pictures. Adjust the magnification so that the screen includes the whole embryo, and then take a picture of both the left and right sides of the embryo. 
		NOTE: The bottom of the plate should be coated in paraffin, silicon, or another such substrate to facilitate pinning.</li>
		<li>Pin the sample through its throat, facing up, and take another picture. 
		NOTE: Orient the pin slightly upward to ensure no important structures of the thoracic cavity&#160;are pierced.</li>
	</ol>
	</li>
	<li>Analyze the chest.
	<ol>
		<li>While lifting the skin in the middle of the neck with forceps, cut the skin toward both the armpits with the scissors before cutting the skin along the median axis toward the tail. Then, cut the skin from the umbilicus to the legs. Pin the sample through its wrists and ankles. 
		NOTE: Be careful to cut only the skin. It is recommended to hold the scissor&#39;s blades horizontally or angled upward.</li>
		<li>To break away the connective tissue, lift the skin with one pair of forceps while holding the underlying tissue in place with the other pair. Pin the sample through the skin to help expose the chest and the abdomen (Figure 2). 
		NOTE: Too much stretching while pinning, cutting, or scraping may result in tissue damage.</li>
		<li>Take a photo of the exposed muscles, and then gently scrape away the muscles to expose the ribs.</li>
		<li>Take a picture of the exposed ribs. Separate the ribs from the diaphragm and cut the ribs on both sides as far as possible on the lateral axis toward the neck.</li>
		<li>Expose the heart by removing the cut ribs. Take a picture of the heart.</li>
		<li>Remove the thymus by peeling it up with one pair of forceps. Use the other pair of forceps to stabilize the base of the thymus to avoid tearing any underlying vessels. Take pictures of the heart and the great vessels. 
		NOTE: Using pins to stretch some adjacent structures may help get better imaging views. Additionally, take separate photos focused on the great arteries, heart, and other structures of interest as they may not be in focus together.</li>
	</ol>
	</li>
	<li>Analyze the abdomen.
	<ol>
		<li>Pull the diaphragm to remove it and expose the liver. Take a picture.</li>
		<li>Pin back the liver to expose the stomach and pancreas. Take a picture.</li>
		<li>Cut the esophagus. Remove the colon and the intestines by pulling them out with forceps. Cut just above the liver and remove it to reveal the kidneys and adrenal glands. Take a picture. 
		NOTE: The removed organs should be kept in the fixative solution.</li>
	</ol>
	</li>
	<li>Isolate the thorax for the ECM analysis.
	<ol>
		<li>Cut the lower thorax along a straight line between the liver and the lungs. Cut the head off high enough to not cut the branchings of the carotid arteries.</li>
		<li>Gently remove the lateral ribs while maintaining the dorsal ribs and the spine. Peel up and scrape the dorsal fat away.</li>
		<li>Detach the thorax from the rest of the body and place it in a 10% buffered formalin phosphate solution.</li>
	</ol>
	</li>
</ol>
3. Embedding
<ol>
	<li>Decant the fixative into an appropriate hazardous waste bottle. Wash the samples with 1x PBS for 15 min three times.</li>
	<li>Use increasing concentrations of ethanol, and xylene to dehydrate the samples. The duration of all the following steps depends on the stage of the embryos. Please refer to Table 2 for details. 
	NOTE: Caution should be taken when changing solutions to avoid damaging the samples. The optimal parameters for sample processing can be empirically adjusted. Xylene will dissolve certain plastics; glass instruments and containers should be used.</li>
	<li>Replace xylene with paraffin for the desired duration. Leave the bottles in a 65 &#176;C incubator for the appropriate duration (Table 2).</li>
	<li>Use fresh paraffin to embed the samples in the desired position. 
	&#8203;NOTE: It is recommended to orient the sample into the middle of the paraffin block, with its dorsal side facing the top and the posterior side facing the front of the block. When orienting the sample, remember that the sample is embedded with the block facing upside down.</li>
</ol>
4. Episcopic confocal microscopy (ECM)
NOTE: After appropriate embedding, embryos undergo image collection serially via ECM for histopathology analysis. Individual slides can be recovered from the microtome for further studies.
<ol>
	<li>Take out the paraffin block from the -20 &#176;C freezer and remove the metal molds.</li>
	<li>Use a razor blade to trim the wax on the edges and the back of the cassette. Cut the wax surrounding the sample until it is encased in a small square of wax attached to the cassette. 
	NOTE: Use extreme caution while handling blades.</li>
	<li>Use the metal lever to clamp the paraffin block against the slicing stage of the microtome. Select the MAN (manual) function in Run Mode, raise the surface of the paraffin close to the blade, and run a few slides to ensure that the blade contacts the paraffin block.</li>
	<li>Open the LAS AF application and select MatrixScreener. Select Single Regular Matrix and load the appropriate previously saved template. Click on the Quick LUT button to switch to white and orange ombre.</li>
	<li>Select Set Up Jobs and drag the 405 nm visible laser to the maximum. Match the left edge of the spectrum block to the 405 nm line and drag the right edge to the 800 nm line. Mark the Pinhole option and start the Live view.</li>
	<li>Adjust the position of the laser projection to center the specimen on the screen and adjust the Zoom knob to ~20x. To optimize the resolution, set the gain to 1,250 V and maximize the blue area using the focus knob; then, reset the gain to approximately 750 V for imaging. 
	NOTE: Specific values may vary depending on the condition of the embryo.</li>
	<li>Switch the cutting method to Auto, set the thickness to around 50 &#181;m, and run the slides. Stop cutting when the lungs and airway are in view.</li>
	<li>Choose a cutting thickness between 8-10 &#181;m and stop the Live view. Open Microtome Communicator to start imaging. Ensure the temporary storage folder is empty before collecting images.</li>
	<li>Stop cutting when no additional heart structure is visualized. Close the Microtome Communicator application, and export the temporary file into one .tiff image series via the image processing software for 3D reconstructions later.</li>
</ol>
5. Three-dimensional (3D) reconstruction
NOTE: The purpose of 3D reconstruction is to process a 2D image stack from ECM imaging into 3D videos in the coronal, sagittal, and transverse orientations and to use the 3D videos for diagnosis of the structural and anatomical abnormalities in the samples.
<ol>
	<li>Open the ECM image stack in the image processing software.
	<ol>
		<li>Drag and drop image the files into the image processing software. Flip the ECM images horizontally by selecting Image &#62; Transform &#62; Flip Horizontally in the menu bar.</li>
		<li>Save the flipped image and close the image processing software.</li>
	</ol>
	</li>
	<li>Import the ECM image stack in the DICOM viewing software.
	<ol>
		<li>Drag and drop the horizontally flipped ECM images into the DICOM viewing software. Make sure there is a light blue border around the sample list, or images may be added to an existing sample folder.</li>
		<li>Select the links or the files to copy into the database when the pop-up window appears. A new sample will appear in the sample list with the same name as the file copied into the DICOM viewing software.</li>
		<li>Click on the newly added file to open it.</li>
	</ol>
	</li>
	<li>Perform 3D reconstruction.
	<ol>
		<li>Once the file is opened, click on the 2D/3D Reconstruction Tools menu from the toolbar and select 3D MPR.</li>
		<li>For Pixel X Resolution and Pixel Y Resolution, input the image resolution by giving the zoom used during ECM imaging. For the Slice Interval, input the slice thickness used to cut during ECM imaging. 
		NOTE: The camera resolution changes depending on the camera&#39;s zoom and may differ from camera to camera.</li>
	</ol>
	</li>
	<li>Use the different tools from the Tools tab on the left side to adjust image stacks as desired.
	<ol>
		<li>Use the WW/WL tool to adjust Window Width and Window Level. Click and drag the tool on the image upward to decrease image brightness and downward to increase it. Click and drag the tool on the image rightward to decrease image contrast and leftward to increase it. 
		NOTE: The optimal WW/WL settings for one structure may be suboptimal for another. For this reason, creating separate videos to view different structures is recommended.</li>
		<li>Use the Pan tool to drag the images into the desired positions. Use the Zoom tool to enlarge or shrink the image as desired and the Rotate tool to rotate the image as desired. 
		NOTE: Enlarging images may cause decreased image quality. Be cautious when rotating images, as doing so may cause the axes to flip.</li>
	</ol>
	</li>
	<li>Click and drag the colored axis of the first panel. Notice how rotating this axis changes the orientation of the other two panels. Rotate the three panels&#39; axes until the three panels represent coronal, sagittal, and transverse views of the sample. 
	NOTE: While reorienting the samples, maintain correct anterior/posterior orientation.</li>
	<li>Generate videos.
	<ol>
		<li>Once all three panels are properly positioned, oriented, and brightened, click on the panel representing the coronal view.</li>
		<li>Click on Movie Export on the right side of the menu bar. Click on Batch and drag the From and To sliders to encompass the entire region of interest. For Interval, select the Same as Thickness option. Save the video indicating the view&#39;s orientation.</li>
		<li>Review the video to see whether the structures of interest can be adequately identified. If not, use the tools (step 5.4) to readjust the videos as needed and re-save the video.</li>
	</ol>
	</li>
	<li>Repeat step 5.6 for transverse and sagittal views. Diagnose the samples using reconstructed videos. 
	NOTE: Carefully watch the videos in each orientation to make a complete assessment of whether the sample exhibits any anatomical abnormalities or disease phenotypes (Figure 3).</li>
</ol>

<ol>
	<li>Wu, W., He, J., Shao, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incidence+and+mortality+trend+of+congenital+heart+disease+at+the+global,+regional,+and+national+level,+1990-2017.">Incidence and mortality trend of congenital heart disease at the global, regional, and national level, 1990-2017.</a> Medicine. 99 (23), e20593 (2020).</li><li>vander Linde, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Birth+prevalence+of+congenital+heart+disease+worldwide:+a+systematic+review+and+meta-analysis).">Birth prevalence of congenital heart disease worldwide: a systematic review and meta-analysis).</a> Journal of the American College of Cardiology. 58 (21), 2241-2247 (2011).</li><li>Yang, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Racial+differences+in+infant+mortality+attributable+to+birth+defects+in+the+United+States.">Racial differences in infant mortality attributable to birth defects in the United States.</a> Birth Defects Research. Part A, Clinical and Molecular Teratology. 76 (10), 706-713 (1989).</li><li>Patel, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prevalence+of+noncardiac+and+genetic+abnormalities+in+neonates+undergoing+cardiac+operations:+Analysis+of+the+society+of+thoracic+surgeons+congenital+heart+surgery+database.">Prevalence of noncardiac and genetic abnormalities in neonates undergoing cardiac operations: Analysis of the society of thoracic surgeons congenital heart surgery database.</a> The Annals of Thoracic Surgery. 102 (5), 1607-1614 (2016).</li><li>Pierpont, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+basis+for+congenital+heart+disease:+Revisited:+A+scientific+statement+from+the+American+Heart+Association.">Genetic basis for congenital heart disease: Revisited: A scientific statement from the American Heart Association.</a> Circulation. 138 (21), e653-e711 (2018).</li><li>Krishnan, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+detailed+comparison+of+mouse+and+human+cardiac+development.">A detailed comparison of mouse and human cardiac development.</a> Pediatric Research. 76 (6), 500-507 (2014).</li><li>Liu, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interrogating+congenital+heart+defects+with+noninvasive+fetal+echocardiography+in+a+mouse+forward+genetic+screen.">Interrogating congenital heart defects with noninvasive fetal echocardiography in a mouse forward genetic screen.</a> Circulation. Cardiovascular Imaging. 7 (1), 31-42 (2014).</li><li>Liu, X., Tobita, K., Francis, R. J., Lo, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+techniques+for+visualizing+and+phenotyping+congenital+heart+defects+in+murine+models.">Imaging techniques for visualizing and phenotyping congenital heart defects in murine models.</a> Birth Defects Research. Part C, Embryo Today: Review. 99 (2), 93-105 (2013).</li><li>Tsuchiya, M., Yamada, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+histological+3D-imaging:+episcopic+fluorescence+image+capture+is+widely+applied+for+experimental+animals.">High-resolution histological 3D-imaging: episcopic fluorescence image capture is widely applied for experimental animals.</a> Congenital Anomalies (Kyoto. 54 (4), 250-251 (2014).</li><li>Yu, Q., Tian Leatherbury, ., Lo, X., W, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiovascular+assessment+of+fetal+mice+by+in+utero+echocardiography).">Cardiovascular assessment of fetal mice by in utero echocardiography).</a> Ultrasound in Medicine and Biology. 34, 741-752 (2008).</li><li>Rosenthal, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+high+resolution+three-dimensional+reconstruction+of+embryos+with+episcopic+fluorescence+image+capture.">Rapid high resolution three-dimensional reconstruction of embryos with episcopic fluorescence image capture.</a> Birth Defects Research. 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The authors declare no conflicts of interest in this manuscript.

Genetically modified mice have been used to understand the pathomechanisms of congenital heart defects. The protocols we provide in this study attempt to streamline and standardize the process of assessing murine fetal heart defects. However, there are critical steps to note during the protocol. Mouse embryos grow significantly during each day of gestation, and the correct time to harvest a mouse can be determined by performing a fetal echocardiogram accurately. The fetal echocardiogram can be used to screen the fetal ca...

Congenital heart diseases (CHDs) are the most common neonatal birth defect1,2, affecting about 0.8%-1.7% of neonates and resulting in significant neonatal mortality and morbidity3. A genetic etiology is strongly indicated with CHDs4,5. Genetically modified mouse models have been used widely to understand the complexity of CHDs and the mechanisms that cause them due to the mice having four-chamber hearts and comparable cardiac developmental DNA sequences in mouse and human fetuses6. Identifying the phenotype of the mouse mutants is the fundamental first step in characterizing the function of the targeted gene. Mouse models expressing gene dosage effects, in which a single genetic mutation can result in a spectrum of cardiac defects that mimic human CHDs, are important for understanding the complexity of CHDs and the mechanisms that cause them.
This article outlines a pipeline to characterize cardiac phenotypes in mouse models. The applied methods utilize fetal echocardiogram7, followed by necropsy and ECM histopathology7,8, which can display the detailed anatomy of developing murine cardiac phenotypes. A fetal echocardiogram is a noninvasive modality that allows direct visualization of multiple embryos with reasonable imaging resolution. In addition, a fetal echocardiogram provides a quick determination of the total number of embryos in a litter, their developing stages, and the relative orientation and location in the uterine horn. Using a spectral Doppler/color flow, abnormal embryos can be identified based on the structure, the hemodynamic disturbance, the growth restriction, or the development of&#160;hydrops. Since a fetal echocardiogram study is a noninvasive technique, it can be used to scan on multiple days and to observe the changes in hemodynamics or cardiac morphology. Obtaining high-quality imaging of fetal echocardiograms requires practice and skill, as specific heart defects may be missed due to a lack of experience and knowledge. Because of this, a more definitive analysis of cardiac morphology may be obtained through a combination of necropsy and ECM histopathology. Necropsy provides direct visualization of the arch structure, the relative relationships of the aorta and pulmonary artery, the size of the ventricles and atria, the position of the heart relative to the chest, and the bronchopulmonary structures. However, interior features such as the heart valves and wall thickness may be difficult to assess through necropsy alone. Thus, ECM histopathology is recommended for a conclusive diagnosis. ECM histopathology is a high-resolution visualization technique that allows for both 2D and 3D reconstruction of the image stack9. These images are obtained through serial Episcopic fluorescent imaging of a paraffin-embedded sample as it is thinly sectioned at a consistent interval by an automatic microtome. Unlike classical histology, images are captured as a section before it is cut from the block such that all images are captured within the same reference frame. Because of this, the 2D image stack produced by ECM histopathology may easily and reliably be reconstructed in three dimensions. This is done using a DICOM viewer, which allows 3D visualization of the images in the three anatomical planes: coronal, sagittal, and transverse. From these high-resolution 3D reconstructions, a definitive cardiac diagnosis may be made. The application of these three different visualization modalities, either individually or in combination, can provide accurate characterizations of structural heart defects in mouse embryos.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1x phosphate-buffered saline solution (PBS), PH7.4</td>
			<td>Sigma Aldrich</td>
			<td>P3813</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL Eppendorf tubes (or preferred vial for tissue storage)</td>
			<td>SealRite</td>
			<td>1615-5599</td>
			<td></td>
		</tr>
		<tr>
			<td>10% buffered formalin phosphate solution</td>
			<td>Fisher Chemical</td>
			<td>SF100-4</td>
			<td></td>
		</tr>
		<tr>
			<td>100% Ethanol</td>
			<td>Decon Laboratories</td>
			<td>2701</td>
			<td></td>
		</tr>
		<tr>
			<td>16% paraformaldehyde (PFA) fixative&#160;</td>
			<td>ThermoScientific</td>
			<td>28908</td>
			<td>4% working concentration freshly prepared in 1x PBS at 4 &#176;C</td>
		</tr>
		<tr>
			<td>50 mL tubes</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>6&#8211;12 Well plate or 20 mL vial&#160; for embryo storage</td>
			<td>Falcon</td>
			<td>353046</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope&#160;</td>
			<td>Leica</td>
			<td>MDG36</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting Pins (A1 or A2 grade)</td>
			<td>F.S.T</td>
			<td>26002-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting Plate&#160;</td>
			<td>F.S.T</td>
			<td>FB0875713</td>
			<td>Petri dish with paraffin base</td>
		</tr>
		<tr>
			<td>Embedding molds</td>
			<td>Sakura</td>
			<td>4133</td>
			<td></td>
		</tr>
		<tr>
			<td>Extra narrow scissors (10.5 cm)</td>
			<td>F.S.T</td>
			<td>14088-10</td>
			<td>1&#8211;2 pairs&#160;</td>
		</tr>
		<tr>
			<td>Fiji application/Image J</td>
			<td>NIH</td>
			<td></td>
			<td>Fiji.sc</td>
		</tr>
		<tr>
			<td>Fine tip (0.05 mm x 0.01 mm) Dissecting Forceps (11 cm)</td>
			<td>F.S.T</td>
			<td>11252-00</td>
			<td>2 Pairs</td>
		</tr>
		<tr>
			<td>Hot forceps&#160;</td>
			<td>F.S.T</td>
			<td>11252-00</td>
			<td>For orientation of embryos</td>
		</tr>
		<tr>
			<td>Industrial Marker for Wax Blocks&#160;</td>
			<td>Sharpie</td>
			<td>2003898</td>
			<td>Formatted for labratory use</td>
		</tr>
		<tr>
			<td>Jenoptik ProgRes C14plus Microscope Camera&#160;</td>
			<td>Jenoptik</td>
			<td>017953-650-26</td>
			<td></td>
		</tr>
		<tr>
			<td>Jenoptik ProgRess CapturePro acquisition software</td>
			<td>Jenoptik</td>
			<td></td>
			<td>jenoptik.com</td>
		</tr>
		<tr>
			<td>Large glass beaker&#160;</td>
			<td>Fisher Scientific</td>
			<td>S111053</td>
			<td>For melting paraffin</td>
		</tr>
		<tr>
			<td>Leica M165 FC binocular microscope (16.5:1 zoom optics)</td>
			<td>Leica</td>
			<td>M165 FC</td>
			<td></td>
		</tr>
		<tr>
			<td>OsiriX MD Version 12.0</td>
			<td>OsiriX</td>
			<td></td>
			<td>osirix-viewer.com&#160;</td>
		</tr>
		<tr>
			<td>Paraplast embedding paraffin wax</td>
			<td>Millipore Sigma</td>
			<td>1003230215</td>
			<td></td>
		</tr>
		<tr>
			<td>Small glass beaker</td>
			<td>Fisher Scientific</td>
			<td>S111045</td>
			<td></td>
		</tr>
		<tr>
			<td>Small, perforated spoon (14.5 cm)</td>
			<td>F.S.T</td>
			<td>10370-17</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight Vannas Scissors (4&#8211;8 mm)</td>
			<td>F.S.T</td>
			<td>15018-10</td>
			<td>A pair</td>
		</tr>
		<tr>
			<td>Vevo2100 ultrahigh-frequency ultrasound biomicroscope</td>
			<td>FUJIFILM VisualSonics Inc.</td>
			<td>Vevo2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylene</td>
			<td>Fisher Chemical</td>
			<td>UN1307</td>
			<td></td>
		</tr>
	</tbody>
</table>

a pipeline to characterize structural heart defects in the fetal mouse

Congenital heart diseases (CHDs) are major causes of infant death in the United States. In the 1980s and earlier, most patients with moderate or severe CHD died before adulthood, with the maximum mortality during the first week of life. Remarkable advances in surgical techniques, diagnostic approaches, and medical management have led to marked improvements in outcomes. To address the critical research needs of understanding congenital heart defects, murine models have provided an ideal research platform, as they have very similar heart anatomy to humans and short gestation rates. The combination of genetic engineering with high-throughput phenotyping tools has allowed for the replication and diagnosis of structural heart defects to further elucidate the molecular pathways behind CHDs. The use of noninvasive fetal echocardiography to screen the cardiac phenotypes in mouse models coupled with the high fidelity of Episcopic fluorescence image capture (EFIC) using Episcopic confocal microscopy (ECM) histopathology with three-dimensional (3D) reconstructions enables a detailed view into the anatomy of various congenital heart defects. This protocol outlines a complete workflow of these methods to obtain an accurate diagnosis of murine congenital heart defects. Applying this phenotyping protocol to model organisms will allow for accurate CHD diagnosis, yielding insights into the mechanisms of CHD. Identifying the underlying mechanisms of CHD provide opportunities for potential therapies and interventions.

The mouse embryos with significant hemodynamic defects were noted to be embryonic lethal. A wide variety of CHDs can be identified through the high output, noninvasive fetal echocardiogram using different views (Figure 1).
Septal defects: The most common CHDs are septal defects such as a ventricular septal defect (VSD), an atrioventricular septal defect (AVSD), and an atrial septal defect (ASD)1. VSD or AVSD can be easily v...

Watch this Scientific Journal Video about A Pipeline to Characterize Structural Heart Defects in the Fetal Mouse at JoVE.com

A Pipeline to Characterize Structural Heart Defects in the Fetal Mouse

This article details murine congenital heart disease (CHD) diagnostic methods using fetal echocardiography, necropsy, and Episcopic fluorescence image capture (EFIC) using Episcopic confocal microscopy (ECM) followed by three-dimensional (3D) reconstruction.

Congenital heart diseases (CHDs) are major causes of infant death in the United States. In the 1980s and earlier, most patients with moderate or severe ...

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Research

JoVE Journal

Developmental Biology

1.7K Views.  University of Pittsburgh. This article details murine congenital heart disease (CHD) diagnostic methods using fetal echocardiography, necropsy, and Episcopic fluorescence image capture (EFIC) using Episcopic confocal microscopy (ECM) followed by three-dimensional (3D) reconstruction.

Video: A Pipeline to Characterize Structural Heart Defects in the Fetal Mouse

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.

Mutations that lead to congenital heart defects benefit from in vivo investigation of cardiac structure during development, but high-resolution structural studies in the mouse embryonic heart are technically challenging. Here we present a robust immunofluorescence and image analysis method to assess cardiomyocyte-specific structures in the developing mouse heart.

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated ...

Analysis of Coronary Vessels in Cleared Embryonic Hearts

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard microscopy techniques, due to the scattering of imaging light by the thick heart tissue, as these vessels are localized deep within the walls of the developing heart. As optical clearing of tissues using organic solvents such as BABB (1 part benzyl alcohol to 2 parts benzyl benzoate) has been shown to greatly improve the optical penetration depth that can be achieved, we combined clearance of whole, PECAM1-immunostained hearts, with laser-scanning confocal microscopy, in order to obtain high-resolution images of vessels throughout the entire heart. BABB clearance of embryonic hearts takes place rapidly and also acts to preserve the fluorescent signal for several weeks; in addition, samples can be imaged multiple times without loss of signal. This straightforward method is also applicable to imaging other types of blood vessels in whole embryos.

We present a protocol for the analysis of coronary vessels in whole embryonic murine hearts up to E15.5, using standard immunological staining methods followed by optical clearance and confocal microscopy. This technique enables visualization of blood vessels throughout the entire heart without the need for time-consuming analysis of serial sections.

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard ...

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is routinely used as an in vivo model to study embryo implantation and pregnancy progression. Unfortunately, such murine studies require pregnancy interruption to enable follow-up phenotypic analysis. To address this issue, we used three-dimensional (3-D) reconstruction of HFUS imaging data for early detection and characterization of murine embryo implantation sites and their individual developmental progression in utero. Combining HFUS imaging with 3-D reconstruction and modeling, we were able to accurately quantify embryo implantation site number as well as monitor developmental progression in pregnant C57BL6J/129S mice from 5.5 days post coitus (d.p.c.) through to 9.5 d.p.c. with the use of a transducer. Measurements included: number, location, and volume of implantation sites as well as inter-implantation site spacing; embryo viability was assessed by cardiac activity monitoring. In the immediate post-implantation period (5.5 to 8.5 d.p.c.), 3-D reconstruction of the gravid uterus in both mesh and solid overlay format enabled visual representation of the developing pregnancies within each uterine horn. As genetically engineered mice continue to be used to characterize female reproductive phenotypes derived from uterine dysfunction, this method offers a new approach to detect, quantify, and characterize early implantation events in vivo. This novel use of 3-D HFUS imaging demonstrates the ability to successfully detect, visualize, and characterize embryo-implantation sites during early murine pregnancy in a non-invasive manner. The technology offers a significant improvement over current methods, which rely on the interruption of pregnancies for gross tissue and histopathologic characterization. Here we use a video and text format to describe how to successfully perform ultrasounds of early murine pregnancy to generate reliable and reproducible data with reconstruction of the uterine form in mesh and solid 3-D images.

Mice are widely used to study gestational biology. However, pregnancy termination is required for such studies which precludes longitudinal investigations and necessitates the use of large numbers of animals. Therefore, we describe a non-invasive technique of high-frequency ultrasonography for early detection and monitoring of post-implantation events in the pregnant mouse.

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is ...

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound (30/45MHZ) System

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular mechanisms of CHDs is&#160;crucial for inventing new preventive and therapeutic strategies. Mutant mouse models are powerful tools to discover new&#160;mechanisms&#160;and environmental stress modifiers that drive cardiac development and their potential&#160;alteration in CHDs. However, efforts to establish the causality of these putative contributors have been limited to histological&#160;and molecular studies&#160;in non-survival animal experiments, in which&#160;monitoring the key physiological and hemodynamic parameters is&#160;often absent. Live imaging technology has become an essential tool to establish the&#160;etiology of CHDs. In particular,&#160;ultrasound imaging can be used prenatally without surgically exposing the fetuses,&#160;allowing maintaining&#160;their baseline physiology while monitoring the impact of environmental stress&#160;on the hemodynamic and structural aspects of cardiac chamber development. Herein, we use the High-Frequency Ultrasound (30/45)&#160;system to examine the cardiovascular system in fetal mice at E18.5 in utero at the baseline and in response to prenatal hypoxia exposure. We demonstrate the feasibility of the system to measure cardiac chamber size, morphology, ventricular function, fetal heart rate, and umbilical artery flow indices, and their alterations in fetal mice exposed to systemic chronic hypoxia in utero in real time.

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound 30/45MHZ System

High-frequency ultrasound imaging of the fetal mouse has improved imaging resolution and can provide precise non-invasive characterization of cardiac development and structural defects. The protocol outlined herein is designed to perform real-time fetal mice echocardiography in vivo.

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular ...

The Isolation and Culture of Primary Epicardial Cells Derived from Human Adult and Fetal Heart Specimens

The epicardium, an epithelial cell layer covering the myocardium, has an essential role during cardiac development, as well as in the repair response of the heart after ischemic injury. When activated, epicardial cells undergo a process known as epithelial to mesenchymal transition (EMT) to provide cells to the regenerating myocardium. Furthermore, the epicardium contributes via secretion of essential paracrine factors. To fully appreciate the regenerative potential of the epicardium, a human cell model is required. Here we outline a novel cell culture model to derive primary epicardial derived cells (EPDCs) from human adult and fetal cardiac tissue. To isolate EPDCs, the epicardium is dissected from the outside of the heart specimen and processed into a single cell suspension. Next, EPDCs are plated and cultured in EPDC medium containing the ALK 5-kinase inhibitor SB431542 to maintain their epithelial phenotype. EMT is induced by stimulation with TGF&#946;. This method enables, for the first time, the study of the process of human epicardial EMT in a controlled setting, and facilitates gaining more insight in the secretome of EPDCs that may aid heart regeneration. Furthermore, this uniform approach allows for direct comparison of human adult and fetal epicardial behavior.

The epicardium plays a crucial role in the development and repair of the heart by providing cells and growth factors to the myocardial wall. Here, we describe a method to culture human primary epicardial cells that enables the study and comparison of their developmental and adult characteristics.

The epicardium, an epithelial cell layer covering the myocardium, has an essential role during cardiac development, as well as in the repair response of ...

Intravenous and Intra-amniotic In Utero Transplantation in the Murine Model

In utero transplantation (IUT) is a unique and versatile mode of therapy that can be used to introduce stem cells, viral vectors, or any other substances early in the gestation. The rationale behind IUT for therapeutic purposes is based on the small size of the fetus, the fetal immunologic immaturity, the accessibility and proliferative nature of the fetal stem or progenitor cells, and the potential to treat a disease or the onset of symptoms prior to birth. Taking advantage of these normal developmental properties of the fetus, the delivery of hematopoietic stem cells (HSC) via an IUT has the potential to treat congenital hematologic disorders such as sickle cell disease, without the required myeloablative or immunosuppressive conditioning required for postnatal HSC transplants. Similarly, the accessibility of progenitor cells in multiple organs during development potentially allows for a more efficient targeting of stem/progenitor cells following an IUT of viral vectors for gene therapy or genome editing. Additionally, IUT can be used to study normal developmental processes including, but not limited to, the development of immunologic tolerance. The murine model provides a valuable and affordable means to understanding the potential and limitations of IUT prior to pre-clinical large animal studies and an eventual clinical application. Here, we describe a protocol for performing an IUT in the murine fetus through intravenous and intra-amniotic routes. This protocol has been used successfully to elucidate the necessary conditions and mechanisms behind in utero hematopoietic stem cell transplantation, tolerance induction, and in utero gene therapy.

Intravenous and Intra-amniotic In Utero Transplantation in the Murine Model

We describe a protocol for performing an in utero transplantation (IUT) through intravenous and intra-amniotic routes of injection in the murine model. This protocol can be used to introduce cells, viral vectors, and other substances into the unique immune-tolerant fetal environment.

In utero transplantation (IUT) is a unique and versatile mode of therapy that can be used to introduce stem cells, viral vectors, or any other substances ...

High Frequency Ultrasound for the Analysis of Fetal and Placental Development In Vivo

Ultrasound imaging is a widespread method used to detect organ anomalies and tumors in human and animal tissues. The method is non-invasive, harmless, and painless, and the application is easy, fast, and can be done anywhere, even with mobile devices. During pregnancy, ultrasound imaging is standardly used to closely monitor fetal development. The technique is important to assess intrauterine growth restriction (IUGR), a pregnancy complication with short- and long-term health consequences for both the mother and fetus. Understanding the process of IUGR is indispensable for developing effective therapeutic strategies. 
The ultrasound system used in this manuscript is an ultrasound device produced for the analysis of small animals and can be used in various research fields, including pregnancy research. Here we describe the usage of the system for in vivo analysis of fetuses from natural killer (NK) cell/mast cell (MC)-deficient mothers that give birth to growth-restricted pups. The protocol includes preparation of the system, handling of the mice before and during measurements, and the usage of the B-mode, color doppler mode, and pulse-wave doppler mode. Fetal size, placental size, and blood supply to the fetus were analyzed. We found reduced implantation sizes and smaller placentas in NK/MC-deficient mice from mid-gestation onwards. In addition, MC/NK-deficiency was associated with absent and reversed end diastolic flow in the fetal Arteria umbilicalis(UmA) and an elevated resistance index. The methods described in the protocol can easily be used for related and non-related research topics.

High Frequency Ultrasound for the Analysis of Fetal and Placental Development In Vivo

Here we describe the technique of high frequency ultrasound for in vivo analysis of fetuses in mice. This method allows the follow-up of fetuses and the analysis of placental parameters as well as maternal and fetal blood flow throughout pregnancy.

Ultrasound imaging is a widespread method used to detect organ anomalies and tumors in human and animal tissues. The method is non-invasive, harmless, and ...

An In Vitro 3D Model and Computational Pipeline to Quantify the Vasculogenic Potential of iPSC-Derived Endothelial Progenitors

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent endothelial progenitors (EPs), which can differentiate into the cell types necessary to assemble mature, functional vasculature, have been derived from both embryonic and induced pluripotent stem cells. However, these cells have not been rigorously evaluated in three-dimensional environments, and a quantitative measure of their vasculogenic potential remains elusive. Here, the generation and isolation of iPSC-EPs via fluorescent-activated cell sorting are first outlined, followed by a description of the encapsulation and culture of iPSC-EPs in collagen hydrogels. This extracellular matrix (ECM)-mimicking microenvironment encourages a robust vasculogenic response; vascular networks form after a week of culture. The creation of a computational pipeline that utilizes open-source software to quantify this vasculogenic response is delineated. This pipeline is specifically designed to preserve the 3D architecture of the capillary plexus to robustly identify the number of branches, branching points, and the total network length with minimal user input.

Endothelial progenitors derived from induced pluripotent stem cells (iPSC-EPs) have the potential to revolutionize cardiovascular disease treatments and to enable the creation of more faithful cardiovascular disease models. Herein, the encapsulation of iPSC-EPs in three-dimensional (3D) collagen microenvironments and a quantitative analysis of these cells&#8217; vasculogenic potential are described.

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...

Recapitulating Suckling-to-Weaning Transition In Vitro using Fetal Intestinal Organoids

At the end of the suckling period, many mammalian species undergo major changes in the intestinal epithelium that are associated with the capability to digest solid food. This process is termed suckling-to-weaning transition and results in the replacement of neonatal epithelium with adult epithelium which goes hand in hand with metabolic and morphological adjustments. These complex developmental changes are the result of a genetic program that is intrinsic to the intestinal epithelial cells but can, to some extent, be modulated by extrinsic factors. Prolonged culture of mouse primary intestinal epithelial cells from late fetal period, recapitulates suckling-to-weaning transition in vitro. Here, we describe a detailed protocol for mouse fetal intestinal organoid culture best suited to model this process in vitro. We describe several useful assays designed to monitor the change of intestinal functions associated with suckling-to-weaning transition over time. Additionally, we include an example of an extrinsic factor that is capable to affect suckling-to-weaning transition in vivo, as a representation of modulating the timing of suckling-to-weaning transition in vitro. This in vitro approach can be used to study molecular mechanisms of the suckling-to-weaning transition as well as modulators of this process. Importantly, with respect to animal ethics in research, replacing in vivo models by this in vitro model contributes to refinement of animal experiments and possibly to a reduction in the use of animals to study gut maturation processes.

This protocol describes how to mimic suckling-to-weaning transition in vitro using mouse late fetal intestinal organoids cultured for 30 days.

At the end of the suckling period, many mammalian species undergo major changes in the intestinal epithelium that are associated with the capability to ...