A Pipeline to Characterize Structural Heart Defects in the Fetal Mouse

Summary

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.

Abstract

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.

Introduction

Congenital heart diseases (CHDs) are the most common neonatal birth defect^1,2, affecting about 0.8%-1.7% of neonates and resulting in significant neonatal mortality and morbidity³. A genetic etiology is strongly indicated with CHDs^4,5. 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 histopathology^7,8, 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 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.

Protocol

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's Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC)-accredited animal care facility. Strict protocols were followed to minimize the mice'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 µm axial x 68 µm lateral) to visualize one mouse fetus at a time. A 40 MHz transducer provides high-resolution images to identify cardiovascular phenotypes⁷.

1. 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.
2. 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.
3. 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.
4. Monitor the temperature using a rectal thermometer probe and make sure it rests at 37 °C ± 0.5 °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.
5. 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.
6. 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).
7. Use the color Doppler to analyze blood flow from the heart.
8. 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.
9. 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.

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's external and internal morphology, looking for macroscopic anatomical abnormalities or malformations.

1. Prepare the mouse.
   1. 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.
2. Analyze the exterior of the body.
   1. Set up the software to save the pictures with a name, including the sample'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.
   2. 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.
   3. 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 are pierced.
3. Analyze the chest.
   1. 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's blades horizontally or angled upward.
   2. 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.
   3. Take a photo of the exposed muscles, and then gently scrape away the muscles to expose the ribs.
   4. 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.
   5. Expose the heart by removing the cut ribs. Take a picture of the heart.
   6. 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.
4. Analyze the abdomen.
   1. Pull the diaphragm to remove it and expose the liver. Take a picture.
   2. Pin back the liver to expose the stomach and pancreas. Take a picture.
   3. 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.
5. Isolate the thorax for the ECM analysis.
   1. 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.
   2. Gently remove the lateral ribs while maintaining the dorsal ribs and the spine. Peel up and scrape the dorsal fat away.
   3. Detach the thorax from the rest of the body and place it in a 10% buffered formalin phosphate solution.

3. Embedding

1. Decant the fixative into an appropriate hazardous waste bottle. Wash the samples with 1x PBS for 15 min three times.
2. 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.
3. Replace xylene with paraffin for the desired duration. Leave the bottles in a 65 °C incubator for the appropriate duration (Table 2).
4. Use fresh paraffin to embed the samples in the desired position.
   ​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.

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.

1. Take out the paraffin block from the -20 °C freezer and remove the metal molds.
2. 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.
3. 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.
4. 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.
5. 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.
6. 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.
7. Switch the cutting method to Auto, set the thickness to around 50 µm, and run the slides. Stop cutting when the lungs and airway are in view.
8. Choose a cutting thickness between 8-10 µm and stop the Live view. Open Microtome Communicator to start imaging. Ensure the temporary storage folder is empty before collecting images.
9. 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.

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.

1. Open the ECM image stack in the image processing software.
   1. Drag and drop image the files into the image processing software. Flip the ECM images horizontally by selecting Image > Transform > Flip Horizontally in the menu bar.
   2. Save the flipped image and close the image processing software.
2. Import the ECM image stack in the DICOM viewing software.
   1. 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.
   2. 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.
   3. Click on the newly added file to open it.
3. Perform 3D reconstruction.
   1. Once the file is opened, click on the 2D/3D Reconstruction Tools menu from the toolbar and select 3D MPR.
   2. 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's zoom and may differ from camera to camera.
4. Use the different tools from the Tools tab on the left side to adjust image stacks as desired.
   1. 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.
   2. 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.
5. 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' axes until the three panels represent coronal, sagittal, and transverse views of the sample.
   NOTE: While reorienting the samples, maintain correct anterior/posterior orientation.
6. Generate videos.
   1. Once all three panels are properly positioned, oriented, and brightened, click on the panel representing the coronal view.
   2. 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's orientation.
   3. 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.
7. 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).

Results

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...

Discussion

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...

Materials

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