Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Animal use and procedures were approved by the Institutional Animal Care and Use Committee at Rutgers University.

1. Preparation of solutions

1. Prepare 1 L of phosphate buffered saline with 0.1% Triton-X-100 (PBST) and filter sterilize. This solution can be stored at room temperature (RT) for at least a year.
2. Prepare 600 µL of blocking buffer consisting of 10% of normal donkey serum in PBST. Make this solution fresh each time.
3. Prepare 50 mL of the following methanol (MeOH) dilutions in a flow hood: 25% MeOH in deionized water (dH₂O), 50% MeOH in dH₂O, and 75% MeOH in dH₂O. Vortex to mix. Store at RT.
4. Prepare 50 mL of the following benzyl alcohol-benzyl benzoate (BABB) solutions in 50 mL conical tubes.
   1. For 100% BABB, add 32 mL of benzyl benzoate to 16 mL of benzyl alcohol (2:1 volume per volume ratio).
   2. For 50% BABB, add 16 mL of benzyl benzoate and 8 mL of benzyl alcohol to 24 mL of MeOH.
   3. Cover conical tubes in aluminum foil to protect from light. These solutions can be stored at RT for up to a year.
      CAUTION: BABB is toxic and corrosive. It should be handled and disposed according to MSDS.

2. Embryo dissection and fixation

NOTE: This protocol is suitable for E9.5 and E10.5 mouse embryos (male or female) isolated from any mouse strain. For younger and older embryos, incubations times should be experimentally determined to maximize signal to noise ratio of fluorescence signal.

1. Fill one 35 mm and one 60 mm Petri dishes with 1x PBS and place on ice until needed.
2. Euthanize a pregnant mouse via CO₂ inhalation. Perform cervical dislocation as a secondary measure of euthanasia.
3. Clean the abdominal area of the dam with 70% ethanol. Pinch the abdominal area using forceps and make a V-like incision using surgical scissors starting from the base of the abdominal wall at the midline; continue to open up the thoracic cavity. Lift the abdominal tissue and move the intestines to the side to expose the uterine horns.
4. Make a cut at the base of the vaginal canal, and with forceps, pull the uterus away from the dam. Make an additional cut at each ovary to free the uterus. Transfer the uterus into one of the 60 mm Petri dishes containing cold 1x PBS.
5. Using straight scissors, cut the uterine wall between each implantation site. Pick up a decidua with a glass pipet and transfer into the 35 mm Petri dish with 1x PBS. Under a dissection microscope, insert straight scissors into the space between the decidua and the uterine wall. Cut and remove the uterine wall.
6. With fine forceps, remove the decidua and Reichert's membranes from the embryo by carefully making transverse incisions along the tissue and pulling the tissue away from the yolk sac. Remove yolk sac and amniotic sac by carefully pulling the tissue away from the embryo and making cuts at the allantois and umbilical vein.
   NOTE: Yolk sacs can be used for genotyping embryos.
7. Transfer each embryo with a glass pipet into individual 2 mL tubes filled with 1 mL of 1x PBS. Label each tube with a unique identifier.
8. To fix embryos, carefully remove the 1x PBS and add 4% paraformaldehyde (PFA) solution in 1x PBS. Incubate at 4 °C with gentle agitation overnight.
   NOTE: 4% PFA fixation is suitable for the antibodies mentioned in this protocol. However, fixation procedures should be optimized for additional antibodies.

3. Embryo staining

NOTE: In this section, embryos are permeabilized and stained with primary and secondary antibodies. Because PAA development proceeds rapidly, differences in embryonic stage will greatly affect the analysis downstream. Therefore, embryos must be age-matched by carefully counting somites to match control and mutant pairs prior to further manipulations.

1. To wash embryo(s), carefully remove 4% PFA and add 1x PBS. Gently invert the tube(s) several times. Place tube(s) right-side up and allow the embryo(s) to sink. Repeat wash 3 times. Place the tube(s) with embryo(s) on ice.
   NOTE: (Optional stopping point) Following the washes, embryos can be dehydrated in graded series of MeOH for 30 min per dilution, as in section 1.3, and stored at -20 °C in 100% MeOH for later use for up to 6 months.
2. For E10.5 embryos, use a glass pipet to transfer one embryo to a 35 mm Petri dish filled with chilled 1x PBS. Carefully pinch the embryo just above the hind limb with fine forceps and make a transverse cut to remove the posterior half of the embryo. This allows the embryo to lay flat in a sagittal position for step 4.2. Place the embryo back into the 2 mL tube with fresh 1x PBS.
   NOTE: A control and mutant embryo can be paired and stained with the same antibody solution in one tube, for steps 3.3 through 3.8.
   1. If staining two embryos together, cut the head off of one embryo above the first pharyngeal arch by pinching with fine forceps to make a transverse cut. This will distinguish embryos of two different genotypes within each tube.
3. To permeabilize the embryo(s), pipet out 1x PBS from the tube, being careful not to touch the embryo(s). Add 1 mL of PBST. Place the tube at 4 °C with gentle agitation overnight.
   NOTE: (Optional stopping point) Embryos can be kept in PBST solution at 4 °C for several days.
4. To prevent non-specific binding of antibodies, first remove PBST from the tube, being careful not to touch the embryo(s). Add 600 µL of blocking buffer solution to the embryo(s). Block the embryo(s) at 4 °C with gentle agitation overnight.
   NOTE: Blocking solution needs to be spun at top speed on a benchtop centrifuge immediately before use to remove debris.
5. To stain and quantify ECs, use antibodies against VEGFR2 and ERG. Antibody solutions are made in the blocking buffer. Anti-VEGFR2 antibody is diluted 1:200 and ERG antibody is diluted 1:1000.
   NOTE: Antibody solutions need to be spun at top speed on a bench-top centrifuge immediately before use to remove particulates.
   1. To incubate embryo(s) with primary antibodies, remove blocking buffer solution from the tube, being careful not to touch the embryo(s). Add 600 µL of primary antibody solution to each tube. Incubate embryo(s) at 4 °C with gentle agitation for 4-5 days.
6. To wash the embryo(s) of the antibody solution, first remove the primary antibody solution from the tube. Wash embryo(s) every hour with 1 mL of PBST at room temperature (RT) with gentle agitation. Wash embryo(s) 4-5 times during the day and then incubate at 4 °C with gentle agitation overnight. Repeat washes the following day.
7. Make secondary antibody solutions by diluting anti-goat Alexa Fluor 488 and anti-mouse Alexa Fluor 555 1:300 in blocking buffer. Dilute the stock DAPI 1:1000 in blocking buffer.
   NOTE: Antibody solutions must be spun at top speed on a bench-top centrifuge immediately before use to remove particulates. In addition, other Alexa Fluor dyes can be used in lieu of 488 or 555.
   1. To incubate the embryo(s) with secondary antibodies, remove PBST from the tube. Add 600 µL of secondary antibody solution to each tube. Incubate embryo(s) at 4 °C with gentle agitation for 4-5 days.
8. To wash the embryo(s) of the antibody solution, first remove the secondary antibody solution from the tube. Wash the embryo(s) every hour with 1 mL of PBST at RT with gentle agitation. Wash embryo(s) 4-5 times during the day and then incubate at 4 °C with gentle agitation overnight. Repeat washes the following day.

4. Embedding embryos in agarose

NOTE: In section 4, the embryo(s) will be embedded in agarose. This embedding process serves two purposes: to properly orient the embryo prior to imaging, and to aid in locating the embryo after it has been cleared in BABB (steps 5.2.2 – 5.3.2).

1. Prepare 200 mL of 1% agarose solution by adding 2 g of agarose to 200 mL of dH₂O. Microwave until all agarose is dissolved.
   NOTE: Remaining agarose can be stored at 4 °C and reheated for later uses.
2. Using a plastic paraffin mold and glass pipet, gently transfer one embryo to the mold. Carefully remove PBST from embryo. Place the embryo in a sagittal position. Quickly, add about 0.5 mL of hot agarose to the mold – just enough to cover the embryo and fill the mold. Ensure that no air bubbles surround the embryo.
3. Place the mold on ice and cover with aluminum foil until the agarose has solidified.
   NOTE: Do not allow the embryo to dry following the removal of PBST. Agarose solution must be warm enough to remain liquid when it is being added to the embryo. Add just enough agarose to cover the embryo, but not too much, otherwise it will be difficult to image. Image depth is determined in part by the working distance of the objective.

5. Dehydration and tissue clearing

NOTE: In this section, embryo(s) are dehydrated using methanol series, then cleared in the organic solvent, BABB, and mounted between two coverslips separated by a rubber spacer; in this protocol Fast Well rubber spacers are used. The Fast Well bumper has a double-sided adhesive surface. The spacer is needed to create a well, in which the embryo will be placed and held between two coverslips.

1. Methanol dehydration
   1. Label new 2 mL tubes, one per embryo. Add 1 mL of 25% MeOH per tube.
   2. Using a clean scalpel, gently cut the agarose around the embryo, leaving enough around the embryo so that it can be picked up by forceps. Use fine forceps to gently grab the agarose with the embedded embryo and place it into the labeled tube with 25% MeOH. Do not allow the forceps to touch the embryo.
   3. Incubate embryo(s) at RT with gentle agitation for 1 hour in the dark.
   4. Remove 25% MeOH from the tube, being careful not to touch the embryo. Add 1 mL of 50% MeOH per tube. Incubate at RT with gentle agitation for 1 hour in the dark.
   5. Remove 50% MeOH from the tube, being careful not to touch the embryo. Add 1 mL of 75% MeOH per tube. Incubate at RT with gentle agitation for 1 hour in the dark.
   6. Remove 75% MeOH from the tube, being careful not to touch the embryo(s). Add 1 mL of 100% MeOH per tube. Incubate at RT with gentle agitation for 1 hour in the dark. Repeat 100% MeOH wash twice.
2. Clearing with BABB
   1. Remove 100% MeOH from the tube, being careful not to touch the embryo. Add 1 mL of 50% BABB per tube. Incubate at RT with gentle agitation for 1 hour in the dark.
   2. Remove 50% BABB from the tube, being careful not to touch the embryo. Add 1 mL of 100% BABB per tube. Incubate at RT with gentle agitation for 1 hour in the dark. Repeat 100% BABB wash twice.
      NOTE: (Optional stopping point) Embryos can remain in 100% BABB in tubes for about a week. Longer storage will cause BABB to dissolve the plastic of tubes.
3. Mounting embryos for imaging
   1. Place a Fast Well bumper on a 24 mm x 60 mm #1.5 glass cover slip, by peeling off the plastic adhesive from one side. Make sure there are no air bubbles between the coverslip and bumper by applying gentle pressure on the plastic adhesive atop the rubber bumper. Label the coverslip according to embryo number, genotype, and antibodies used for staining.
      NOTE: Any spacer can be placed between the coverslips as long as it is thick enough to prevent crushing or squishing an embryo. We use Fast Well spacers due to their thickness and convenience, which includes adhesive surfaces on either side of the spacer for securing it to coverslips.
   2. Carefully pipet out and discard the 100% BABB from the tube. After visualizing the agarose-embedded embryo in the tube, use fine forceps to pick up the agarose and carefully transfer the embryo onto the coverslip inside the Fast Well – do not allow the forceps to touch the embryo.
   3. Remove the second plastic adhesive from the bumper and place the second coverslip on top. Remove air bubbles by gently pressing on the coverslip. Be careful to not break the glass.
      NOTE: Samples can be stored flat in a slide holder in the dark at RT for up to a year if the seal is tight.

6. Acquisition of data

NOTE: In the following steps, the endothelium of pharyngeal arches 3, 4, and 6 will be imaged using confocal microscopy.

1. Positioning of slides on microscope stage
   1. To image embryos, use a confocal microscope equipped with a 20x water immersion objective, numerical aperture 0.95, working distance 0.95 mm, and the NIS-Elements AR 5.11.01 64-bit software.
   2. Using wide-field fluorescence, visually locate the pharyngeal arches. Center the field view around the 4^th PAA.
   3. If the objective's field of view does not capture the entire pharyngeal arch area, take and stitch a large panel of images with 1% overlap. To prevent movement of the sample during the acquisition of the large image, gently secure the cover slip assembly to the stage using molding clay.
2. Setting up acquisition parameters
   1. Set the pinhole size to 1.0.
   2. Under the ND Acquisition tab, set the top and bottom limits of imaging using the coarse adjustment. Set Z step size according to software specifications. Determine the thickness that can be imaged by the working distance of the objective and the clarity of the sample.
   3. Due to the thickness of the embryo, adjust the gain throughout the Z-stack. Set the laser intensity and gain at the middle of the Z-stack for each channel (405, 488, and 555) and assign values under the Z Intensity Correction tab. Set the same values for the bottom slice.
   4. Scroll through the embryo until fluorescence signals begin to appear dimmer. Increase the gain of each channel until signal intensity appears similar to the previous segment. Assign the new value under the Z Intensity Correction tab. Repeat until z-stack is complete. Import settings back to ND Acquisition.
   5. Run scan using Run Z Correction option.

7. Analysis using the Imaris software

NOTE: In these steps, confocal images will be analyzed using the microscopy image analysis software, Imaris version 9.2.0. During this analysis, we will first select regions of interest to be analyzed by creating surfaces. Next, we will use the Mask function to visually separate these regions. Finally, we will use the Spot function to quantify the number of ECs within each region of interest.

1. Depending on the imaging software used in step 6, convert images to .ims using Imaris File Converter.
2. Open the .ims files. Set image to Orthogonal under Camera/Labels | Camera Type panel.
3. Locate the PAAs and orient the image for surfacing.
   NOTE: When the files are first opened, they will appear as a 3D compilation of all slices imaged. In this step, the PAAs will be located by making the 3D image into a 2D image. The 2D image then allows for the PAAs to be properly oriented for analysis.
   1. Under the Properties panel, turn off Volume. Under the Properties panel, click on Add New Ortho Slicer. Set Slice Orientation to the XY Plane. Use the Slice Position to scroll through the image until finding the PAAs.
   2. If the PAAs are not parallel to the top and bottom of the image, freely rotate the image using the mouse cursor so that PAAs run from left to right across the screen. Under the Image Processing drop down menu, select Free Rotate and click OK.
4. Surfacing the 3 ^rd Pharyngeal Arch (Figure 2A, B – B")
   NOTE: In these steps, the pharyngeal arches and PAAs will be traced using the Surface tool to generate a 'surfaced' region of interest. This will allow for each region of interest to be visually isolated from the surrounding tissue. Herein we describe the steps to surface and analyze the endothelial components of the 3^rd pharyngeal arch. Pharyngeal arches 4 and 6 are analyzed similarly.
   1. To surface the endothelium in the entire 3^rd pharyngeal arch, click on the Add New Surface button located under the Properties panel. Double click on Surface 1 and rename the new surface to "3^rd Pharyngeal Arch".
   2. Select Skip automatic creation, edit manually. Set Surface Orientation to the YZ Plane (coronal orientation). Use the Slice Position to place the 3^rd pharyngeal arch surface plane to where the 3^rd PAA and Dorsal Aorta connect.
   3. Rotate the image so that the 3^rd pharyngeal arch surface plane is in view. Turn off Ortho Slicer 1.
   4. Under the Draw | Contour | Mode tab, select the Distance Drawing Mode function. Adjust parameter settings if needed. Maintain consistent surfacing parameters between samples. For this example, Vertex spacing is 10 μm.
   5. To begin surfacing, press the Esc key and then click on the Draw button. Trace the perimeter of the 3^rd pharyngeal arch with the mouse cursor. Use the Slice Position to move 10-25 slices. Trace the perimeter of the pharyngeal arch. Repeat until the pharyngeal arch is fully traced.
   6. To generate the surface of the traced region, select the Create Surface button in the Properties panel.
5. Surfacing the 3 ^rd PAA (Figure 2C – C")
   1. To surface the endothelium of the 3^rd PAA, first turn off the surfaced region from step 7.4, by deselecting the 3^rd Pharyngeal Arch surface box. Then, click on the Add New Surface button again. Double click on Surface 1 and rename the new surface to "3^rd PAA".
   2. Select Skip automatic creation, edit manually. Set Surface Orientation to the YZ Plane (coronal orientation). Use the Slice Position to place the 3^rd PAA surface plane to where the 3^rd PAA and Dorsal Aorta connect, then repeat steps 7.4.
6. Masking of surfaced structures
   NOTE: In the following steps, each surfaced region of interest will be Masked. Masking allows the region of interest to be visually distinct from the rest of the imaged tissue and allows for the quantification of these distinct structures of interest. Below, we describe the steps in Imaris to visualize and analyze the PAA endothelium, as well as the plexus – the smaller vasculature surrounding the PAAs within the pharyngeal arches. In these steps, the 3^rd PAA surface will be masked in order to visualize and perform analysis only on the PAAs using the Spot function described in Section 7.7.
   1. Select Volume in order to visualize all masked channels. Press the Esc key in order to rotate the image and position the image into an XY position.
   2. Under the Edit tab, select Mask Selection for the 3^rd PAA. Select the DAPI channel and click OK. Repeat for the remaining channels.
   3. On the keyboard, press Ctrl + D to view the Display Adjustments panel. Select each new channel and rename them to make it clear what each channel shows. For example, this will lead to three new channels: "3^rd PAA DAPI", "3^rd PAA ERG", and "3^rd PAA VEGFR2".
      NOTE: In steps 7.6.4-7.6.7 we will create channels for pharyngeal arch plexus only.
   4. To visualize the endothelial plexus separately from the PAA, we will first select Mask Selection for the 3^rd PAA surface. Select the DAPI channel. Uncheck Select voxels outside surface to and check Select voxels inside surface to buttons, set Select voxels inside surface to zero. Click OK.
   5. Repeat for the remaining channels. This operation will exclude the region containing the PAA from the new masked channels. Rename channels to make it clear what each channel shows. For example, this will lead to three new channels: "Non-PAA DAPI", "Non-PAA ERG", and "Non-PAA VEGFR2".
   6. To visualize endothelial plexus within the 3^rd pharyngeal arch, select Mask Selection under the Edit tab, for the 3^rd Pharyngeal Arch surface. Select the Non-PAA DAPI channel. Click OK.
   7. Repeat for the remaining Non-PAA channels. Rename channels to make it clear what each channel shows. For example, this will lead to three new channels: "Plexus DAPI", "Plexus ERG", and "Plexus VEGFR2".
7. Quantification of EC numbers
   NOTE: The expression of ERG marks endothelial nuclei making it convenient to quantify EC numbers. In these steps, number of ECs will be quantified using the Spot function to generate a spot for each EC marked by ERG expression in the PAA and plexus. In section 7.7, spots will be generated for each ERG-positive cell in the masked PAA, followed by the deselection of spots in ERG-positive, VEGFR2-negative cells.
   1. On the keyboard, press Ctrl + D to view the Display Adjustments panel. Turn off all channels except for PAA ERG.
   2. Under the properties tab, click on the Add New Spots button. Click on Spots 1 and rename it to "PAA Total Number of ECs". Click on the blue arrow button. For Source Channel, select the PAA ERG channel. Adjust the Estimated XY Diameter to 4 μm. Proceed to the next panel by clicking on the blue arrow button.
   3. Adjust number of spots seen using the sliding scale, to ensure that each EC nucleus (marked by ERG expression) is represented by one spot. Click on the green double arrow button.
   4. Turn off the PAA ERG channel in the Display Adjustment. Turn on the PAA VEGFR2 channel to visualize PAA endothelium.
   5. To accurately quantify the number of ECs, we ensure that each spot expresses both EC markers, ERG and VEGFR2. To do this, select Surface of Object under the Edit tab | Add/Delete panel. Press the Esc key and delete any spots that are not VEGFR2 positive, by holding down shift and selecting the spot.
      NOTE: In the following step, spots will be generated for each ERG-positive cell in the masked plexus, followed by the deselection of spots in ERG-positive, VEGFR2-negative cells.
   6. Under the properties tab, click on the Add New Spots button. Select Spots 1 and rename to "Plexus Total Number of ECs". Click on the blue arrow button. For Source Channel, select the Plexus ERG channel. Adjust the Estimated XY Diameter to 4 μm. Repeat steps 7.7.1 – 7.7.5 for Plexus total number of ECs.
   7. Click on the Statistics tab of each spot function to determine the total number of ECs in the 3^rd PAA and pharyngeal arch plexus.
   8. Repeat steps 7.7.1 – 7.7.6 for the remaining PAAs and pharyngeal arch plexus.