This work was funded by NIH/NHLBI R56 HL128646 and the Mindich Child Health and Development Institute (MCHDI) at ISMMS (to N.D.). E.B. is supported by an NIH/NIDCR Traineeship T32 HD075735.Microscopy and image analysis was performed at the Microscopy Core at the Icahn School of Medicine at Mount Sinai, which is supported in part by the Tisch Cancer Institute at Mount Sinai P30 CA196521 &#8211; Cancer Center Support Grant.

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
<ol>
	<li>Mate a fertile female mouse with a fertile stud male.</li>
	<li>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.</li>
	<li>On the morning of the 8th 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Under a dissection microscope, use fine forceps (#5) to remove the uterine tissue away from the decidual tissue.</li>
	<li>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.</li>
	<li>Dissect away extraembryonic tissues as much as possible without damaging the morphology of the embryo (Figure 1B).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Fix embryos with 4% paraformaldehyde (PFA) in PBS for 1 h at RT. 
	NOTE: Embryos can be fixed overnight (O/N) at 4 &#176;C.</li>
	<li>Rinse three times with PBS. Keep embryos at 4 &#176;C until ready to proceed with immunofluorescence. 
	NOTE: Pause point. Embryos can be safely stored in PBS at 4 &#176;C for several weeks.</li>
</ol>
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.
<ol>
	<li>Remove PBS and add 1 mL of blocking buffer (0.5% saponin, 1% bovine serum albumin (BSA) in PBS).</li>
	<li>Incubate at least 4 h at RT. This can also be done O/N at 4 &#176;C.</li>
	<li>Remove blocking buffer and add primary antibody mixture diluted in blocking buffer. Incubate O/N at 4 &#176;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.</li>
	<li>Remove primary antibodies by aspiration.</li>
	<li>Wash 3 times for 1 h each with 0.1% Triton in PBS.</li>
	<li>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 &#176;C.</li>
	<li>Wash 3 times for 1 h each with 0.1% Triton in PBS. This can also be done O/N at 4 &#176;C.</li>
	<li>Counterstain with 4&#39;,6-diamidino-2-phenylindole (DAPI) in PBS for 10 min. 
	NOTE: This counterstain can be performed simultaneously with secondary antibody.</li>
	<li>Wash 2 times for 5 min each with 0.1% Triton in PBS.</li>
	<li>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.</li>
</ol>
3. Mounting Embryos for Microscopy
<ol>
	<li>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).</li>
	<li>Place a 15 &#181;L 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Use a pipette to add additional anti-fade between the slide and coverslip to keep the sample from dying out during storage/imaging.</li>
</ol>
4. Confocal Imaging
<ol>
	<li>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 &#34;optimize&#34; 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.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Create a new surface for the reference channel using the Surface algorithm dialogue.
	<ol>
		<li>Select the &#34;Segment only a Region of Interest&#34; checkbox.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#34;split touching objects&#34; algorithm, to further separate your signal from background.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Using the newly created surface, create a masked channel for the experimental channel.
	<ol>
		<li>With the surface(s) selected, choose the &#34;Mask Sel ...&#863;&#34; function from the Edit window.</li>
		<li>Select the experimental channel of interest (i.e. YFP). Make sure that the &#34;Duplicate channel before applying mask&#34; checkbox is selected.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>To obtain the volumetric data, with each surface selected select the Statistics tab. In the &#34;Detailed&#34; sub-tab, choose &#34;Average Values&#34; 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 &#34;All Values&#34; from the drop-down menu. Selecting each value will make the corresponding region appear yellow, which aids in determining values to exclude.</li>
</ol>

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The authors have nothing to disclose.

The protocol above describes a strategy for obtaining quantitative data from high quality whole mount immunofluorescence images of post-implantation mouse embryos. When performed correctly, the 3D volumetric data generated through these steps can be used for comparative and intersectional analysis of multiple domains within the embryo. The surface signal masking approach described is of particular use when investigating novel cell populations in comparison with well-established reference structures.
We believe that this approach offers a distinct advantage over existing approaches. Whole mount immunofluorescence with wide-field imaging can offer tissue-level information but lacks cellular resolution. Conversely, immunofluorescence of serial sections can give detailed cellular-level resolution, though these data lack the three-dimensional advantage of whole mount imaging. While three-dimensional confocal imaging addresses these shortcomings, few users take advantage of the quantitative potential of this data, and we hope that our protocol will allow more researchers to take full advantage of the data they collect.
Here, this approach has been exemplified using the marker Nkx2-5 to delineate the cardiac crescent and examine the presence of Foxa2Cre:YFP lineage-traced cells within that domain. However, due to the ability to image at least three lineages concurrently and the high resolution three-dimension data generated, this protocol will likely be useful to researchers in many fields. We propose that this approach can be applied to multiple embryonic stages, organ systems, and lineages through minor adjustments to the protocol based on the specific needs of the system. In our experience, this protocol is directly applicable to a range of embryonic ages (E6.5-E9.5 at minimum) with limited adaptations.19 We have not encountered issues with permeability or antibody penetration in embryos as large as E9.5 with the incubation times listed, though we expect that with thicker or denser tissues these would have to be elongated to achieve full penetration.
The analysis described here was performed using a proprietary software package, but can be adapted to be performed in alternative proprietary packages optimized for big-data handling. Alternatively, similar pipelines can easily be implemented using computational languages or open-source software packages, which will lead to the rapid development of a shareable protocols database. The growth of a global network of scientists, with access to freely available tools and protocols for image acquisition and analysis, will rapidly advance the field and increase the reproducibility of studies.17
In our experience, the major limitation to this approach was related to imaging depth, especially for later embryonic stages, and will depend heavily on the specifics of the microscopy setup being used. Similarly, how samples are mounted for imaging will depend on the specific needs of the user. We have found that placing the region of interest closest to the lens improves imaging depth and decreases background signal, and we suggest using layered double-sided tape for mounting, as it enables users to create a chamber that fits their specific needs. Use of a glycerol-based mounting media with an anti-fade reagent, and equilibrating samples in this media before mounting, is highly recommended to suppress photobleaching. Finally, the samples used in this analysis are nearly transparent; users may need to incorporate clearing steps for more advanced embryonic stages, tissue fragments, or organoids, and determining the best clearing protocol for a given application will require empirical testing.
In the case of cardiac morphogenesis, various imaging methods can be used. While confocal microscopy is used here, light-sheet microscopy is extremely well suited for these studies because of the speed, resolution, and signal-to-noise ratio of the acquired image.20 We expect the increased availability of this new technology to further advance the utility and richness of similar imaging approaches. Similarly, the continuous advancement of fluorescent reporters,21 chromophores, dyes,22 and live-embryo culture23,24 will lead to these studies moving to four dimensions, adding the element of time, and yielding yet more information about how embryonic structures form during development.

The study of organogenesis has long relied on the observation of morphogenetic events in the developing embryo. These studies frequently rely on the use of fluorescent dyes or lineage tracing reporters in combination with labeling of defined reference populations.1 By comparing relative positions of these labels, information can be gleaned on the origin, movement, or ultimate contribution of a population of interest. Transplantation and fate mapping experiments use either morphological landmarks or injection of dyes into non-motile lineages to define the starting point of the cells of interest, which are then examined for contribution to the developed embryo.2,3,4,5 Genetic lineage-tracing experiments use the same concept with well-defined reporter alleles that are used to label cell populations without experimental manipulation. Key to these approaches is the ability to determine, with high spatial resolution, the locations of the experimental and reference labels. These approaches have yielded outstanding progress in pre-implantation development and explant organogenesis studies.6,7,8,9
The developmental events that underlie heart morphogenesis have been increasingly well described in recent years.10 One of the major discoveries in this area of research is the description of a number of progenitor populations that can be distinguished by expression of unique markers.11 These populations include the First and Second Heart Fields (FHF and SHF), which are present within the cardiac crescent at the anterior side of the embryo at embryonic day (E) 8.25 of mouse development.12 These populations are frequently examined through a combination of wide-field microscopy, which provides tissue-level information, and serial sectioning with immunofluorescence assays, which offers high cellular resolution but only two-dimensional spatial information.13 Thus, while these studies have greatly advanced our understanding of heart development, the available methods have limited in depth quantitative analysis of morphogenesis during these stages, creating the need for approaches to examine the organization of these populations on a whole-organism level.
The recent advances in both confocal microscopy and 3D image analysis allow for high-resolution and high-throughput algorithmic reconstructions of cells and structures in situ with relative ease, thus paving the way for detailed studies of complex cellular structures.14 With the increase of computational power and the development of big-data managing algorithms, both necessary to handle the exponential increase of the size of imaging data-sets, analyses can now be fully automated.15 Automated analysis of imaging data-sets has the benefit of being unbiased, but it is only as reliable as the quality of the input dataset; it is imperative, then, that best-practices are used during acquisition and image pre-processing to ensure the highest quality, unbiased analysis.16 Protocols can be completely automated and shared for reproducibility, and the algorithms used by proprietary software are readily available through libraries to be used by scientists who have familiarity with modern proprietary or open-source developer tools.17
The following protocol explains the necessary steps to perform such analysis on one well defined model of organogenesis, the formation of the cardiac crescent during heart development. Specifically, this protocol describes how to (1) harvest and dissect cardiac crescent stage embryos, (2) perform whole mount immunofluorescence for reference (Nkx2-5) and experimental (Foxa2Cre:YFP18,19) markers, (3) prepare and image the embryos using confocal microscopy, and finally (4) analyze and quantify the resulting images using advanced three-dimensional approaches. While the cardiac crescent is used as an example here, with appropriate modification, this protocol may be used for analysis of multiple lineages in gastrula to early somite stage embryos.

<table><tbody><tr><td>Blunt probe</td><td>Roboz</td><td>RS-9580</td><td></td></tr><tr><td>Forceps</td><td>Roboz</td><td>RS-8100</td><td></td></tr><tr><td>Fine forceps</td><td>Roboz</td><td>RS-5015</td><td></td></tr><tr><td>Dissection scissors</td><td>Roboz</td><td>RS-5912</td><td></td></tr><tr><td>Saponin</td><td>Sigma Aldrich</td><td>84510</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma Aldrich</td><td>A8022</td><td></td></tr><tr><td>Triton</td><td>RPI</td><td>A4490</td><td></td></tr><tr><td>Goat anti-Nkx2-5</td><td>Santa Cruz Biotech</td><td>sc-8697</td><td>Used at 1:100-1:500</td></tr><tr><td>Chicken anti-GFP</td><td>abcam</td><td>ab13970</td><td>Used at 1:500</td></tr><tr><td>Rabbit anti-Islet1</td><td>abcam</td><td>ab109517</td><td>Used at 1:100</td></tr><tr><td>Rabbit anti-Hcn4</td><td>Millipore</td><td>AB5808</td><td>Used at 1:100</td></tr><tr><td>488 anti-chicken</td><td>Jackson Immunoresearch</td><td>703-546-155</td><td>Reconstituted in water and stored at -20 &amp;deg;C in final concentration of 50% glycerol. Used at 1:500.</td></tr><tr><td>555 anti-goat</td><td>Thermo Fisher Scientific</td><td>A21432</td><td>Used at 1:500</td></tr><tr><td>647 anti-rabbit</td><td>Jackson Immunoresearch</td><td>711-606-152</td><td>Reconstituted in water and stored at -20 &amp;deg;C in final concentration of 50% glycerol. Used at 1:500.</td></tr><tr><td>DAPI</td><td>Sigma Aldrich</td><td>D9542</td><td></td></tr><tr><td>n-Propyl gallate</td><td>Sigma Aldrich</td><td>2370</td><td>Stock solution is 20% w/v in DMSO. Working solution prepared by mixing 1 part 10x PBS with 9 parts 100% glycerol and slowly adding 0.1 part stock solution.</td></tr><tr><td>Superfrost Plus microscopy slides</td><td>VWR Scientific</td><td>48311-703</td><td></td></tr><tr><td>22x22 mm coverslips</td><td>VWR Scientific</td><td>48366-227</td><td></td></tr><tr><td>Imaris 8.4.1</td><td>Bitplane</td><td></td><td></td></tr></tbody></table>

quantitative whole-mount immunofluorescence analysis of cardiac progenitor populations in mouse embryos

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development and organogenesis are two areas of research that have benefitted greatly from these advances, due to the high quality of data that can be obtained directly from imaging pre-implantation embryos or ex vivo organs. While pre-implantation embryos have yielded data with especially high spatial resolution, later stages have been less amenable to three-dimensional reconstruction. Obtaining high-quality 3D or volumetric data for known embryonic structures in combination with fate mapping or genetic lineage tracing will allow for a more comprehensive analysis of the morphogenetic events taking place during embryogenesis.
This protocol describes a whole-mount immunofluorescence approach that allows for the labeling, visualization, and quantification of progenitor cell populations within the developing cardiac crescent, a key structure formed during heart development. The approach is designed in such a way that both cell- and tissue-level information can be obtained. Using confocal microscopy and image processing, this protocol allows for three-dimensional spatial reconstruction of the cardiac crescent, thereby providing the ability to analyze the localization and organization of specific progenitor populations during this critical phase of heart development. Importantly, the use of reference antibodies allows for successive masking of the cardiac crescent and subsequent quantitative measurements of areas within the crescent. This protocol will not only enable a detailed examination of early heart development, but with adaptations should be applicable to most organ systems in the gastrula to early somite stage mouse embryo.

The quality of the final data and analysis depends greatly on (1) the integrity and morphology of the dissected embryos, (2) the use of high specificity antibodies, and (3) the proper setup of imaging parameters. Damaged embryos will confound the surface generation process and hinder quantitative analysis. Examples of properly dissected and staged embryos are shown in Figure 2B. The embryos can be further dissected by removing the yolk sac, to which antibodies will frequently bind non-specifically. However, removal of the yolk sac will make the embryos less robust, so care should be taken during downstream handling.
The combination of high quality antibodies and proper imaging settings are crucial to achieve high signal-to-noise ratio images. Figure 2A shows an example image using antibodies against Nkx2-5 as a reference marker to label the cardiac crescent, and GFP to label a lineage traced population of interest (Foxa2Cre:YFP ventricular cardiovascular progenitor cells) (See supplemental data for raw image and data files).19 This strategy allows for the comparison of a given population with the entire cardiac crescent region. For some experiments, it may be of interest to examine the FHF and SHF subdomains of the cardiac crescent. In this case, the FHF and SHF can be labeled with Hcn4 and Islet1, respectively (see reference19 for images and quantitative analysis with these antibody combinations). With the appropriate setup and secondary antibody combinations, we have successfully imaged and analyzed three lineages concurrently (i.e. the cardiac crescent, YFP, and first or second HF).
Inability to achieve strong signal will also limit the ability to generate high confidence 3D models, as it will become difficult to remove background signal from the final surfaces. In these cases, care should be taken when adjusting image gamma (during pre-processing, Figure 2B, 2C, 2N, 2O) or surface thresholding (during surface generation algorithm, Figure 2G, 2H, 2L) and similar settings should be used for all images within an experiment. Failure to do so will result in inconsistent volumetric data that would be inappropriate to use for direct comparison. Often, erroneous surfaces resulting from background signal can be removed through size filtering (Figure 2I, 2J), though doing so from high-background images will be difficult without losing true surface fragments as well.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56446/56446fig1.jpg" /> 
Figure 1: Experimental timeline and schematics. (A) The entire experiment, from mating to data analysis, can be carried out in approximately two weeks. Embryos are collected in the morning on the 8th day post copulation (E8.25) for cardiac crescent stage analysis (B). Once fixed, embryos are blocked and stained with primary and secondary antibodies before mounting. Embryos are mounted with standard microscopy slides and coverslips, using double-sided tape as a spacer (C). Embryos are placed in a drop of anti-fade in the displayed orientation (C, upper) and a coverslip is slowly lowered onto the tape (C, lower). Confocal imaging and image analysis are performed to generate high resolution z-stack images (D) and 3D surface models (E). HF, head fold; CC, cardiac crescent; A, anterior; P, posterior; AF, anti-fade. Scale bars are 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56446/56446fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56446/56446fig2.jpg" /> 
Figure 2: Stepwise confocal image processing and 3D surface generation. (A) Confocal z-series loaded in the volume view. The reference channel (Nkx2-5) is selected (B) and the intensity and gamma adjusted (C) before starting the Create Surfaces algorithm. After the region of interest is selected (D), and the level of surface detail is chosen (E). The initial surface (F) is thresholded to include all true signal in the surface (G). Filtering is then performed to remove small background fragments (H) to yield a final reference surface (I). By selecting the reference surface (J), the comparison channel (YFP) can be duplicated and masked (K). The intensity and gamma for this channel are then adjusted (L) before generating a second surface through the same sequence of steps (M, N). The total volume for each surface is calculated automatically and can be compared both quantitatively and visually (O, note that surface colors do not merge when overlapped). Scale bars = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56446/56446fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos at JoVE.com

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

Here, we describe a protocol for whole mount immunofluorescence and image-based quantitative volumetric analysis of early stage mouse embryos. We present this technique as a powerful approach to qualitatively and quantitatively assess cardiac structures during development, and propose that it may be widely adaptable to other organ systems.

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development ...

quantitative-whole-mount-immunofluorescence-analysis-cardiac

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

9.6K Views.  Icahn School of Medicine at Mount Sinai. Hier beschrijven we een protocol voor hele mount immunofluorescentie en beeld-gebaseerde kwantitatieve volumetric analysis van vroege fase muis embryo's. Wij presenteren deze techniek als een krachtige aanpak van kwalitatief en kwantitatief cardiale structuren beoordelen tijdens de ontwikkeling, en voor te stellen dat het wellicht grote schaal aan te passen aan andere orgaansystemen.

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

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 ...

Quantitative Analysis of Protein Expression to Study Lineage Specification in Mouse Preimplantation Embryos

This protocol presents a method to perform quantitative, single-cell in situ analyses of protein expression to study lineage specification in mouse preimplantation embryos. The procedures necessary for embryo collection, immunofluorescence, imaging on a confocal microscope, and image segmentation and analysis are described. This method allows quantitation of the expression of multiple nuclear markers and the spatial (XYZ) coordinates of all cells in the embryo. It takes advantage of MINS, an image segmentation software tool specifically developed for the analysis of confocal images of preimplantation embryos and embryonic stem cell (ESC) colonies. MINS carries out unsupervised nuclear segmentation across the X, Y and Z dimensions, and produces information on cell position in three-dimensional space, as well as nuclear fluorescence levels for all channels with minimal user input. While this protocol has been optimized for the analysis of images of preimplantation stage mouse embryos, it can easily be adapted to the analysis of any other samples exhibiting a good signal-to-noise ratio and where high nuclear density poses a hurdle to image segmentation (e.g., expression analysis of embryonic stem cell (ESC) colonies, differentiating cells in culture, embryos of other species or stages, etc.).

This protocol presents a method to perform quantitative, single-cell in situ analysis of protein expression to study lineage specification in mouse preimplantation embryos. The procedures necessary for collection of blastocysts, whole-mount immunofluorescent detection of proteins, imaging of samples on a confocal microscope, and nuclear segmentation and image analysis are described.

This protocol presents a method to perform quantitative, single-cell in situ analyses of protein expression to study lineage specification in mouse ...

Imaging Cleared Embryonic and Postnatal Hearts at Single-cell Resolution

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, and allow for the study of the cellular and molecular basis of normal and abnormal cardiac morphogenesis. Recent emerging technologies of retrospective single clonal analyses make the study of cardiac morphogenesis at single cell resolution feasible. However, tissue opacity and light scattering of the heart as imaging depth is increased hinder whole-heart imaging at single cell resolution. To overcome these obstacles, a whole-embryo clearing system that can render the heart highly transparent for both illumination and detection must be developed. Fortunately, in the last several years, many methodologies for whole-organism clearing systems such as CLARITY, Scale, SeeDB, ClearT, 3DISCO, CUBIC, DBE, BABB and PACT have been reported. This lab is interested in the cellular and molecular mechanisms of cardiac morphogenesis. Recently, we established single cell lineage tracing via the ROSA26-CreERT2; ROSA26-Confetti system to sparsely label cells during cardiac development. We adapted several whole embryo-clearing methodologies including Scale and CUBIC (clear, unobstructed brain imaging cocktails and computational analysis) to clear the embryo in combination with whole mount staining to image single clones inside the heart. The heart was successfully imaged at single cell resolution. We found that Scale can clear the embryonic heart, but cannot effectively clear the postnatal heart, while CUBIC can clear the postnatal heart, but damages the embryonic heart by dissolving the tissue. The methods described here will permit the study of gene function at a single clone resolution during cardiac morphogenesis, which, in turn, can reveal the cellular and molecular basis of congenital heart defects.

We describe a protocol to volumetrically image fluorescent protein labeled cells deep inside intact embryonic and postnatal hearts. Utilizing tissue-clearing methods in combination with whole mount staining, single fluorescent protein-labeled cells inside an embryonic or postnatal heart can be imaged clearly and accurately.

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, ...

Assessing Cardiomyocyte Subtypes Following Transcription Factor-mediated Reprogramming of Mouse Embryonic Fibroblasts

Direct reprogramming of one cell type into another has recently emerged as a powerful paradigm for regenerative medicine, disease modeling, and lineage specification. In particular, the conversion of fibroblasts into induced cardiomyocyte-like myocytes (iCLMs) by Gata4, Hand2, Mef2c, and Tbx5 (GHMT) represents an important avenue for generating de novo cardiac myocytes in vitro and in vivo. Recent evidence suggests that GHMT generates a greater diversity of cardiac subtypes than previously appreciated, thus underscoring the need for a systematic approach to conducting additional studies. Before direct reprogramming can be used as a therapeutic strategy, however, the mechanistic underpinnings of lineage conversion must be understood in detail to generate specific cardiac subtypes. Here we present a detailed protocol for generating iCLMs by GHMT-mediated reprogramming of mouse embryonic fibroblasts (MEFs). 
We outline methods for MEF isolation, retroviral production, and MEF infection to accomplish efficient reprogramming. To determine the subtype identity of reprogrammed cells, we detail a step-by-step approach for performing immunocytochemistry on iCLMs using a defined set of compatible antibodies. Methods for confocal microscopy, identification, and quantification of iCLMs and individual atrial (iAM), ventricular (iVM), and pacemaker (iPM) subtypes are also presented. Finally, we discuss representative results of prototypical direct reprogramming experiments and highlight important technical aspects of our protocol to ensure efficient lineage conversion. Taken together, our optimized protocol should provide a stepwise approach for investigators to conduct meaningful cardiac reprogramming experiments that require identification of individual CM subtypes.

This manuscript describes a step-by-step protocol for the generation and quantification of diverse reprogrammed cardiac subtypes using a retrovirus-mediated delivery of Gata4, Hand2, Mef2c, and Tbx5.

Direct reprogramming of one cell type into another has recently emerged as a powerful paradigm for regenerative medicine, disease modeling, and lineage ...

Induction of Endothelial Differentiation in Cardiac Progenitor Cells Under Low Serum Conditions

Cardiac progenitor cells (CPCs) may have therapeutic potential for cardiac regeneration after injury. In the adult mammalian heart, intrinsic CPCs are extremely scarce, but expanded CPCs could be useful for cell therapy. A prerequisite for their use is their ability to differentiate in a controlled manner into the various cardiac lineages using defined and efficient protocols. In addition, upon in vitro expansion, CPCs isolated from patients or preclinical disease models may offer fruitful research tools for the investigation of disease mechanisms.
Current studies use different markers to identify CPCs. However, not all of them are expressed in humans, which limits the translational impact of some preclinical studies. Differentiation protocols that are applicable irrespective of the isolation technique and marker expression will allow for the standardized expansion and priming of CPCs for cell therapy purpose. Here we describe that the priming of CPCs under a low fetal bovine serum (FBS) concentration and low cell density conditions facilitates the endothelial differentiation of CPCs. Using two different subpopulations of mouse and rat CPCs, we show that laminin is a more suitable substrate than fibronectin for this purpose under the following protocol: after culturing for 2 - 3 days in medium including supplements that maintain multipotency and with 3.5% FBS, CPCs are seeded on laminin at &#60;60% confluence and cultured in supplement-free medium with low concentrations of FBS (0.1%) for 20 - 24 hours before differentiation in endothelial differentiation medium. Because CPCs are a heterogeneous population, serum concentrations and incubation times may need to be adjusted depending on the properties of the respective CPC subpopulation. Considering this, the technique can be applied to other types of CPCs as well and provides a useful method to investigate the potential and mechanisms of differentiation and how they are affected by disease when using CPCs isolated from respective disease models.

This protocol describes an endothelial differentiation technique for cardiac progenitor cells. It particularly focuses on how serum concentration and cell-seeding density affect the endothelial differentiation potential.

Cardiac progenitor cells (CPCs) may have therapeutic potential for cardiac regeneration after injury. In the adult mammalian heart, intrinsic CPCs are ...

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

En Face Endocardial Cushion Preparation for Planar Morphogenesis Analysis in Mouse Embryos

The study of the cellular and molecular mechanisms underlying the development of the mammalian heart is essential to address human congenital heart disease. The development of the primitive cardiac valves involves the epithelial-to-mesenchymal transition (EMT) of endocardial cells from the atrioventricular canal (AVC) and outflow tract (OFT) regions of the heart in response to local inductive myocardial and endocardial signals. Once the cells delaminate and invade the extracellular matrix (cardiac jelly) located between the endocardium and the myocardium, the primitive endocardial cushions (EC) are formed. This process implies that the endocardium has to fill the gaps left by the delaminated cells and has to reorganize itself to converge (narrow) or extend (lengthen) along an axis. Current research has implicated the planar cell polarity (PCP) pathway in regulating the subcellular localization of the factors involved in this process. Classically, the initial phases of cardiac valve development have been studied in cross-sections of embryonic hearts or in ex vivo AVC or OFT explants cultured on collagen gels. These approaches allow the analysis of apico-basal polarity but do not allow the analysis of cell behavior within the plane of the epithelium or of the morphological changes of migrating cells. Here, we show an experimental approach that allows the visualization of the endocardium at valvulogenic regions as a planar field of cells. This experimental approach provides the opportunity to study PCP, planar topology, and intercellular communication within the endocardium of the OFT and AVC during valve development. Deciphering new cellular mechanisms involved in cardiac valve morphogenesis may contribute to understanding congenital heart disease associated with endocardial cushion defects.

En Face Endocardial Cushion Preparation for Planar Morphogenesis Analysis in Mouse Embryos

Classically, the endocardium of the mouse embryonic valve primordium has been analyzed using transversal, coronal, or sagittal sections. Our novel approach for en face, two-dimensional imaging of the endocardium at valvulogenic regions allows planar polarity and cell rearrangement analysis of the endocardium during valve development.

The study of the cellular and molecular mechanisms underlying the development of the mammalian heart is essential to address human congenital heart ...

Live Imaging of Early Cardiac Progenitors in the Mouse Embryo

The first steps of heart development imply drastic changes in cell behavior and differentiation. While analysis of fixed embryos allows studying in detail specific developmental stages in a still snapshot, live imaging captures dynamic morphogenetic events, such as cell migration, shape changes, and differentiation, by imaging the embryo as it develops. This complements fixed analysis and expands the understanding of how organs develop during embryogenesis. Despite its advantages, live imaging is rarely used in mouse models because of its technical challenges. Early mouse embryos are sensitive when cultured ex vivo and require efficient handling. To facilitate a broader use of live imaging in mouse developmental research, this paper presents a detailed protocol for two-photon live microscopy that allows long-term acquisition in mouse embryos. In addition to the protocol, tips are provided on embryo handling and culture optimization. This will help understand key events in early mouse organogenesis, enhancing the understanding of cardiovascular progenitor biology.

We present a detailed protocol for mouse embryo culture and imaging that enables 3D + time imaging of cardiac progenitor cells. This video-toolkit addresses the key skills required for successful live imaging otherwise hard to acquire from text-only publications.

The first steps of heart development imply drastic changes in cell behavior and differentiation. While analysis of fixed embryos allows studying in detail ...

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Biology

Protein Isolation from the Developing Embryonic Mouse Heart Valve Region

Western blot analysis is a commonly employed technique for detecting and quantifying protein levels. However, for small tissue samples, this analysis method may not be sufficiently sensitive to detect a protein of interest. To overcome these difficulties, we examined protocols for obtaining protein from adult human cardiac valves and modified these protocols for the developing early embryonic mouse counterparts. In brief, the mouse embryonic aortic valve regions, including the aortic valve and surrounding aortic wall, are collected in the minimal possible volume of a Tris-based lysis buffer with protease inhibitors. If required based on the breeding strategy, embryos are genotyped prior to pooling four embryonic aortic valve regions for homogenization. After homogenization, an SDS-based sample buffer is used to denature the sample for running on an SDS-PAGE gel and subsequent western blot analysis. Although the protein concentration remains too low to quantify using spectrophotometric protein quantification assays and have sample remaining for subsequent analyses, this technique can be used to successfully detect and semi-quantify phosphorylated proteins via western blot from pooled samples of four embryonic day 13.5 mouse aortic valve regions, each of which yields approximately 1 &#956;g of protein. This technique will be of benefit for studying cell signaling pathway activation and protein expression levels during early embryonic mouse valve development.

The analysis of protein expression in young embryonic mouse valves has been hampered by the limited tissue available. This manuscript provides a protocol for preparing protein from developing embryonic mouse valve regions for western blot analysis.

Western blot analysis is a commonly employed technique for detecting and quantifying protein levels. However, for small tissue samples, this analysis ...

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

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