Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos

All methods described here have been approved by the Institutional Animal Care and Use Committee at the Icahn School of Medicine at Mount Sinai.

1. Harvesting and Processing Cardiac Crescent Stage Embryos

1. Mate a fertile female mouse with a fertile stud male.
2. Check for the presence of a vaginal copulation plug using a blunt probe or forceps. Noon on the day of plug detection is considered embryonic day (E) 0.5 (See Figure 1A for full timeline).
   NOTE: Plugs should be checked for in the morning, as they are lost throughout the day.
3. On the morning of the 8^th day (E8.25), sacrifice the pregnant dam by CO₂ inhalation, or according to local and institutional regulations.
   NOTE: Exact timing can be strain dependent and should be determined empirically by morphology.
4. Spray the abdomen of the mouse with 70% ethanol to clean the area and minimize shedding. Expose the viscera by performing an abdominal incision through both the skin and body wall.
5. Locate the uterus and carefully remove the entire uterine horn from the animal. Start by cutting above one oviduct, trimming fat away from the uterus while proceeding to the cervical end. Cut through the cervix and continue to the upper end of the other oviduct, cutting to release the entire uterus.
6. Place the uterus in a 10 cm dish with phosphate buffered saline (PBS, pH 7.4) to wash away excess blood. Sub-dissect the uterus by cutting the mesometrium between each deciduum, which contains the embryos. Transfer the embryos to a 6 cm dish with fresh PBS.
7. Under a dissection microscope, use fine forceps (#5) to remove the uterine tissue away from the decidual tissue.
8. Using forceps, carefully slice the tip of the embryonic half of the deciduum to reveal the embryo. Pinch the deciduum to push the embryo out and carefully pull the embryo out.
9. Dissect away extraembryonic tissues as much as possible without damaging the morphology of the embryo (Figure 1B).
10. Use a transfer pipette to place the embryos in a 1.5 mL tube with fresh PBS and place on ice. Repeat 1.7-1.9 for the remaining embryos before continuing.
11. Aspirate PBS and rinse once with PBS. Aspirate PBS.
    NOTE: Try to remove as much PBS as possible without damaging or drying the embryos. Manual removal of solutions is recommended at all steps to avoid loss of embryos.
12. Fix embryos with 4% paraformaldehyde (PFA) in PBS for 1 h at RT.
    NOTE: Embryos can be fixed overnight (O/N) at 4 °C.
13. Rinse three times with PBS. Keep embryos at 4 °C until ready to proceed with immunofluorescence.
    NOTE: Pause point. Embryos can be safely stored in PBS at 4 °C for several weeks.

2. Immunofluorescence Staining

NOTE: The incubation conditions below can be adjusted to accommodate different schedules. Use of gentle shaking or rocking for all long incubation steps is recommended.

1. Remove PBS and add 1 mL of blocking buffer (0.5% saponin, 1% bovine serum albumin (BSA) in PBS).
2. Incubate at least 4 h at RT. This can also be done O/N at 4 °C.
3. Remove blocking buffer and add primary antibody mixture diluted in blocking buffer. Incubate O/N at 4 °C.
   NOTE: See Materials section for description of antibodies used and suggested dilutions. Antibody dilutions should be determined empirically. The use of Nkx2-5 as a reference stain for the cardiac crescent is recommended, and is key to downstream image segmentation and analysis steps.
4. Remove primary antibodies by aspiration.
5. Wash 3 times for 1 h each with 0.1% Triton in PBS.
6. Remove wash and add secondary antibody mixture diluted in Blocking buffer. Incubate for 3 h at RT. This can also be done O/N at 4 °C.
7. Wash 3 times for 1 h each with 0.1% Triton in PBS. This can also be done O/N at 4 °C.
8. Counterstain with 4',6-diamidino-2-phenylindole (DAPI) in PBS for 10 min.
   NOTE: This counterstain can be performed simultaneously with secondary antibody.
9. Wash 2 times for 5 min each with 0.1% Triton in PBS.
10. Slowly suspend embryos in anti-fade mounting media (2% w/v n-Propyl gallate (nPG), 90% glycerol, 1x PBS; See Materials section for more details). Allow to equilibrate at least 1 h before mounting. Gently flick tube periodically to help embryos drop into anti-fade solution.
    NOTE: Pause point. Embryos can be stored for several days in anti-fade solution until ready to mount and image.

3. Mounting Embryos for Microscopy

1. Prepare microscope slides for mounting using either double-stick tape or silicone spacers. If using double-stick tape, make two parallel stacks of 5-6 layers about 15-20 mm apart. This will leave enough space to place the embryos and secure the coverslip (Figure 1C).
2. Place a 15 µL drop of anti-fade on the slide, between the double-stick tape. Carefully transfer one embryo to the slide.
   NOTE: More than one embryo can be placed on a slide. However, the subsequent orientation steps are more difficult with multiple embryos, and long imaging durations may lead to photo-bleaching.
3. Under a dissection microscope using fine forceps, position the embryo such that the anterior side faces away from the slide with the body axis in line with the long axis of the slide. See Figure 1C for a schematic of the mounting setup.
4. Carefully place a coverslip over the sample by resting one side on one stack of double-stick tape and using forceps to gently lower the coverslip until it contacts the other tape.
   NOTE: Steps 3.3 and 3.4 are critical to ensure the correct orientation of the embryo for imaging. By lowering the coverslip along the posterior to anterior direction, the embryo should not roll and the cardiac crescent region will stay oriented away from the slide, which is ideal for imaging on an inverted microscope.
5. Use a pipette to add additional anti-fade between the slide and coverslip to keep the sample from dying out during storage/imaging.

4. Confocal Imaging

1. Acquire images using the highest magnification objective that allows for the whole cardiac crescent region to be captured in one field of view. Use Nyquist sampling rates to determine x-y-z voxel dimensions. Most microscope manufacturers will have an "optimize" button to ensure Nyquist sampling.
   NOTE: Staining for Nkx2-5 is beneficial here, as it will delineate the entire cardiac crescent region. Smaller Z-step sizes (oversampling) will yield better 3D modeling results but may lead to extended acquisition times and/or bleaching for some samples.
2. Set up imaging parameters using standard best-imaging practices. Namely, use the lowest possible laser intensity to achieve good signal-to-noise ratio for each fluorophore and choose gain and offset levels that yield a wide dynamic range without saturating the signal.
   NOTE: Be consistent with imaging parameters between samples that will be directly compared. This is especially important for downstream 3D analysis and quantification.

5. Image Analysis and Quantitative 3D Modeling

NOTE: For this protocol, images were analyzed using the Imaris software package. Similar analysis may be possible using alternative packages. The description below covers the analysis pipeline for a reference channel (i.e. Nkx2-5) and one experimental channel (i.e. YFP). Additional channels can be analyzed through repetition of these steps. An example dataset has been provided that can be used to replicate the analysis below.

1. Load raw image data sets into Imaris Arena view and open file for desired embryo in Surpass view. Data should load in 3D View in MIP (max) mode. Open the Display Adjustment window and select only the reference channel.
2. Adjust the threshold of the image if needed for better visualization. For this analysis, adjust the gamma if the signal-to-noise ratio is too low for segmentation.
   NOTE: If performing gamma corrections, which are non-linear adjustments, the adjusted images cannot be used for intensity measurements or comparisons. At this stage you can perform further image pre-processing (i.e. illumination corrections, Gaussian blur or other filtering), if your signal-to-noise ratio is not satisfactory. If you choose to do pre-processing, you must perform the same pre-processing steps for all images in your dataset.
3. Create a new surface for the reference channel using the Surface algorithm dialogue.
   1. Select the "Segment only a Region of Interest" checkbox.
   2. Adjust the bounding box region so that it contains the region of interest (ROI, i.e. the cardiac crescent area) as delineated by the reference stain.
   3. Select the reference channel from the Source Channel dropdown menu. The default setting for Smoothing (equal to the z voxel size) is typically sufficient but may require empirical optimization. For this analysis, use smoothing equal to half the z voxel-size.
   4. Perform thresholding by absolute intensity. To ensure that the surface does not extend beyond the channel signal, adjust the lower limit using the sliders.
      NOTE: For certain images where the cells can be delineated easily, you can include the "split touching objects" algorithm, to further separate your signal from background.
   5. Filter objects by voxel number or volume, and reject background (small or found in areas outside of the Nkx2-5 area) objects that the algorithm might have generated. Finish the algorithm.
4. Using the newly created surface, create a masked channel for the experimental channel.
   1. With the surface(s) selected, choose the "Mask Sel …͟" function from the Edit window.
   2. Select the experimental channel of interest (i.e. YFP). Make sure that the "Duplicate channel before applying mask" checkbox is selected.
5. In the Display Adjustment window select only the masked channel. Create a new surface for the masked channel by following steps 5.3-5.3.5.
   NOTE: At this point it may be helpful to adjust the appearance of each mask to facilitate the visualization of the 3D model: Select each surface and select the color tab. Adjust the Base color and transparency as needed. Surface colors do not merge when overlapped.
6. To obtain the volumetric data, with each surface selected select the Statistics tab. In the "Detailed" sub-tab, choose "Average Values" from the drop-down menu. The total volume of the surface can be found in the Sum column of the generated table.
   NOTE: If the generated surface contains erroneous fragments (i.e. fragments that are from background signal not excluded during segmentation or areas discontinuous from the main surface), it may be necessary to calculate the total volume manually, or filter the surfaces by volume or number of voxels and ensure that only signal segments are included in the calculation. The volume for each portion of the surface can be found by selecting "All Values" from the drop-down menu. Selecting each value will make the corresponding region appear yellow, which aids in determining values to exclude.