H.B. is supported by startup funding from the Medical Faculty and SFB829 of the University of Cologne. C.X. is supported by DFG grant BA 5810/1-1. We would like to thank the Imaging Facilities at the CECAD research center and Memorial Sloan Kettering Cancer Center (New York, USA). We thank Joaqu&#237;n Grego-Bessa (Spanish National Center for Cardiovascular Research, Madrid, Spain) for his insight on mounting the embryos for WMIF.

All experiments involving animal experiments were approved by the responsible authorities in North Rhein Westphalia (LANUV-NRW).
1. Scanning Electron Microscopy of the Mouse Embryonic Node
<ol>
	<li>Sacrifice the pregnant female mouse at ~ E7.5 (2-4 somite stage) by cervical dislocation. A detailed explanation with diagrams of Steps 1.1 - 1.7 is available in mouse embryo laboratory manuals13.</li>
	<li>Open the abdomen through the skin and mesenteries and remove the uterus using scissors and fine forceps.</li>
	<li>Rinse the uterus briefly in distilled water and place it in a small clean Petri dish (6 cm) containing 1x phosphate-buffered saline (PBS).</li>
	<li>Under a dissecting microscope and using fine forceps, remove the uterine muscles to free the individual deciduae or implantation sites.</li>
	<li>Hold each decidua with one pair of forceps and use the other pair to make a longitudinal full-thickness incision between the red part (future placenta) and white part (where the embryo is located). Make superficial perforations vertically along the white part of the decidua contiguous with the incision. Pull the decidua apart horizontally into two halves and carefully scoop out the embryo in the white part of the decidua.</li>
	<li>Transfer the embryo to a new Petri dish (35 mm) with fresh sterile-filtered PBS. Repeat for all deciduae/embryos.</li>
	<li>Remove Reichert&#39;s membrane, a relatively opaque membrane engulfing the embryo, from each embryo by teasing it away like a sock starting at the ectoplacental cone (reddish implantation site). For genotyping, take a little piece (~ 0.1 mm2) of the yolk sac at this stage.</li>
	<li>Under a chemical hood and wearing appropriate protection (gloves), transfer the embryos to EM grade fixative composed of 2.5% glutaraldehyde in sterile-filtered PBS in a microcentrifuge (1.5 mL) tube at room temperature. Fix the embryos overnight at 4 &#176;C.</li>
	<li>Carefully remove the glutaraldehyde fixative from the tube without touching the embryos and discard in a proper waste container. Wash the embryos three times in sterile-filtered PBS, for 15 min each at room temperature.</li>
	<li>Dehydrate the embryos in an ethanol series for 5 min each: 50%, 70%, 85%, and three times in 100% or absolute ethanol. Store the embryos at &#8722;20 &#176;C in ethanol or proceed directly to the next step.</li>
	<li>Transfer the embryos in ethanol to baskets for critical point drying (CPD) in a critical point dryer machine. Fill the chamber with ethanol to cover the baskets completely.</li>
	<li>Exchange the ethanol by carefully flushing with liquid CO2 for ten times at 10 &#176;C. Drain off liquid CO2 after the last step until the chamber is half full. Heat up to 40 &#176;C until the pressure reaches 80 bars (the critical point) and the liquid CO2 changes to gas. Wait for 10 min then slowly blow off the gas over approximately 45 min.</li>
	<li>As an easier alternative to CPD for drying, add hexamethyldisilazane (HMDS) at a ratio of 1:1 to the embryos in ethanol for 30 min. Then transfer the embryos to pure HMDS for 30 min. Remove the embryos out of the liquid using a pipette and leave them to dry for 30 min. 
	NOTE: Both drying methods worked equally well in our hands.</li>
	<li>Use a fine brush to mount the dried embryos with the ventral side (node) up on an SEM stub with double sided tape.</li>
	<li>Insert the stubs with the embryos into a sputter coating machine for gold particle coating, which is preferred to charge the long thin cilia. Apply a layer of 120-150 &#197;; the time is dependent on the current, which will vary with each sample.</li>
	<li>Place the coated stubs with embryos into an SEM microscope, apply vacuum and observe the embryonic node and notochordal plate cells with cilia at magnifications ranging from 1000X to 15,000X.</li>
</ol>
2. Whole Mount Immunofluorescence of the Mouse Node and Notochordal Plate
<ol>
	<li>Using ice-cold PBS with 0.05% Tween 20 (PBSTw), follow Steps 1.1-1.7 above to remove the embryos at E7.75 and place them in PBSTw in a 35 mm Petri dish on ice.</li>
	<li>Under a chemical hood and wearing appropriate protection (gloves), transfer the embryos to a fixative solution of 4% paraformaldehyde in PBS in a microcentrifuge tube. Fix the embryos overnight at 4 &#176;C.</li>
	<li>Carefully remove the paraformaldehyde fixative from the tube without touching the embryos and discard in a proper waste container. Wash the embryos three times in PBS containing 0.2% Triton X-100 (PBSTr) for 5 min each at room temperature. Perform all the wash and next incubation steps on a nutating shaker.</li>
	<li>Remove the last wash and add blocking solution containing PBSTr with 10% heat-inactivated serum (from the host species of the secondary antibody). Block from 2 h to overnight (or longer) on a nutator at 4 &#176;C.</li>
	<li>Remove the blocking and add ~ 1 mL of the primary antibody diluted in blocking solution, for example an anti-T antibody at 1:500 dilution. Incubate overnight (or longer) on a nutator at 4 &#176;C.</li>
	<li>Remove the primary antibody and save it for later use by adding sodium azide to a final concentration of 0.02% (1 &#181;L of 20% stock to 1 mL of antibody solution). The antibody can be reused ~ 10 times. Rinse the embryos twice with PBSTr and then wash them three times for 30 min each on a nutator at 4 &#176;C.</li>
	<li>Replace the wash with a fluorescence-conjugated secondary antibody, against the primary antibody host species, diluted at ~ 1:1000 overnight (or longer) on a nutator at 4 &#176;C.</li>
	<li>Remove the secondary antibody and rinse twice with PBSTr, then wash three times for 30 min with PBSTr.</li>
	<li>Replace the last wash with PBSTr containing 1:500 fluorescence-conjugated phalloidin, to stain F-Actin, and 1:1000 DAPI, to stain nuclei, for 1 h at room temperature.</li>
	<li>Rinse twice in PBSTr and wash once with PBSTr for 30 min at room temperature.</li>
	<li>Replace PBSTr with PBS and leave the embryos on ice. Prepare clean positively-charged slides (60 x 24 mm) and coverslips (24 x 24 mm) and aqueous glycerol-based mounting media, for example, 90% glycerol in 1xPBS and an antifade reagent, to mount the embryos.</li>
	<li>Put two pieces of clear tape at a distance of ~ 15 mm from each other on the clear part of the slide. This will create enough 3D space (in the Z dimension) that would allow flattening the embryos but not completely squishing them.</li>
	<li>Under a dissecting microscope, carefully move the embryos using a cut P200 pipette (to allow enough space for the embryo to be transferred and not damaged) to the slide.</li>
	<li>Using fine forceps, make two full cuts on the lateral sides of the yolk sac to unfold the embryo. Place the embryo with the ventral side (node and notochordal plate) up (dorsal of neural tube down on the slide).</li>
	<li>Add 50 &#181;L of mounting media on the embryo. Place 4-5 embryos per slide. Add a dab of mounting media to the side of the coverslip that will first touch the slide (top or bottom side), then place it straddling the two pieces of tape and lower it slowly onto the embryos using fine forceps or a bent fine needle while avoiding creating air bubbles.</li>
	<li>Clean the excess mounting media by using an absorbent wipe. Be careful not to move the coverslip in the process.</li>
	<li>Using a generous amount of nail polish, seal the sides of the coverslip without moving it.</li>
	<li>Observe under a scanning confocal microscope.</li>
</ol>

<ol>
	<li>Rivera-P&#233;rez, J. A., Hadjantonakis, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dynamics+of+morphogenesis+in+the+early+mouse+embryo.">The dynamics of morphogenesis in the early mouse embryo.</a> Cold Spring Harbor Perspectives in Biology. , (2015).</li><li>Lee, J. D., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphogenesis+of+the+node+and+notochord:+The+cellular+basis+for+the+establishment+and+maintenance+of+left-right+asymmetry+in+the+mouse.">Morphogenesis of the node and notochord: The cellular basis for the establishment and maintenance of left-right asymmetry in the mouse.</a> Developmental Dynamics. 237 (12), 3464-3476 (2008).</li><li>Balmer, S., Nowotschin, S., Hadjantonakis, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notochord+morphogenesis+in+mice:+Current+understanding+and+open+questions.">Notochord morphogenesis in mice: Current understanding and open questions.</a> Developmental Dynamics. 245 (5), 547-557 (2016).</li><li>Yoshiba, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cilia+at+the+node+of+mouse+embryos+sense+fluid+flow+for+left-right+determination+via+Pkd2.">Cilia at the node of mouse embryos sense fluid flow for left-right determination via Pkd2.</a> Science. , (2012).</li><li>Jurand, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Some+aspects+of+the+development+of+the+notochord+in+mouse+embryos.">Some aspects of the development of the notochord in mouse embryos.</a> Journal of Embryology and Experimental Morpholog. 32 (1), 1-33 (1974).</li><li>Poelmann, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+head-process+and+the+formation+of+the+definitive+endoderm+in+the+mouse+embryo.">The head-process and the formation of the definitive endoderm in the mouse embryo.</a> Anatomy and Embryology (Berl). , 41-49 (1981).</li><li>Sulik, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphogenesis+of+the+murine+node+and+notochordal+plate.">Morphogenesis of the murine node and notochordal plate.</a> Developmental Dynamics. 201 (3), 260-278 (1994).</li><li>Bazzi, H., Soroka, E., Alcorn, H. L., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=STRIP1,+a+core+component+of+STRIPAK+complexes,+is+essential+for+normal+mesoderm+migration+in+the+mouse+embryo.">STRIP1, a core component of STRIPAK complexes, is essential for normal mesoderm migration in the mouse embryo.</a> Proceedings of the National Academy of Sciences of the United States of America. 114 (51), 10928-10936 (2017).</li><li>Bazzi, H., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acentriolar+mitosis+activates+a+p53-dependent+apoptosis+pathway+in+the+mouse+embryo.">Acentriolar mitosis activates a p53-dependent apoptosis pathway in the mouse embryo.</a> Proceedings of the National Academy of Sciences of the United States of America. 111 (15), 1491-1500 (2014).</li><li>Huangfu, D., Liu, A., Rakeman, A. S., Murcia, N. S., Niswander, L., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hedgehog+signalling+in+the+mouse+requires+intraflagellar+transport+proteins.">Hedgehog signalling in the mouse requires intraflagellar transport proteins.</a> Nature. 426 (6962), 83-87 (2003).</li><li>McMullan, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+electron+microscopy+1928-1965.">Scanning electron microscopy 1928-1965.</a> Scanning. 17 (3), 175-185 (2006).</li><li>Bai, S. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+characterization+of+a+set+of+conserved+and+new+regulators+of+cytoskeletal+organization,+cell+morphology+and+migration.">Identification and characterization of a set of conserved and new regulators of cytoskeletal organization, cell morphology and migration.</a> BMC Biology. 9, (2011).</li><li>Behringer, R., Gertsenstein, M., Vintersen Nagy, K., Nagy, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manipulating+the+Mouse+Embryo:+A+Laboratory+Manual,+Fourth+Edition.">Manipulating the Mouse Embryo: A Laboratory Manual, Fourth Edition.</a> Cold Harb Lab Press. , (2014).</li><li>Braet, F., De Zanger, R., Wisse, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drying+cells+for+SEM,+AFM+and+TEM+by+hexamethyldisilazane:+a+study+on+hepatic+endothelial+cells.">Drying cells for SEM, AFM and TEM by hexamethyldisilazane: a study on hepatic endothelial cells.</a> Journal of Microscopy. 186, 84-87 (1997).</li></ol>

The authors have nothing to disclose.

In this work, we demonstrate how to perform SEM and WMIF to visualize the mouse embryonic node and notochordal plate. The small size of gastrulating mouse embryos ~ E7.5 and the presence of these structures on the surface make them ideal to study using the techniques described2,7,8. The availability of good antibodies, such as T and cilia markers, gives excellent 3D information using WMIF on the structure, organization and formation of these essential embryonic organizers8.
Because mouse embryonic development proceeds at a very rapid pace and the node and notochordal plate are only transiently present on the surface of the embryo, timing is essential for the success of these experiments2,3. For example, 2-4 somite embryos are good for SEM analysis of a mature node pit with long cilia. In much earlier or later embryos (for example, 12 h before or after), the node might not be present on the surface. WMIF is a little more flexible in this regard but the structures themselves are also transient during development and the timing in this case depends on the researchers&#39; interests.
The purity of the reagents is also essential for the success of these techniques, especially in probing the ultrastructure by SEM. Tiny impurities that stick to the embryos usually result in huge artifacts.
We have tested two different methods of embryo fixation for SEM one using half Karnovsky&#39;s fixative (2.5% glutaraldehyde, 2% paraformaldehyde and 0.1 M cacodylate buffer) and a simpler 2.5% glutaraldehyde in 1x PBS. We prefer to use the glutaraldehyde and PBS fixative as described in Step 1, however, we and others have also used the half Karnovsky&#39;s fixative successfully for SEM.
We have also compared two methods of drying the embryos for SEM and found no difference in the quality of the sample by using either a critical point dryer or HMDS as described in Step 1 and reported elsewhere14.
For Step 2, we tested embedding the embryos after the final washing steps in 1% low melting agarose mounted on a 35 mm glass-bottom dish and then topping it with ~ 10 &#181;L of mounting medium. This embedding method works and preserves the original 3D structure of the embryo and associated structures; however, a multiphoton microscope is required to image the specimen because a regular confocal microscope cannot reach as deep into the intact embryos (~ 1 mm).
We believe that using these two techniques gives complementary information on the structure of the node and the notochordal plate during normal development and in mutants which show defects in the formation of these structures.

Gastrulation and the accompanying morphogenetic movements are crucial for shaping the mouse embryo1. The changes in cellular shape and organization during morphogenesis dictate positional information to regulate cell fate and also allow the ensuing signaling pathways to precisely perform their functions to diversify the newly formed germ layers1. The formation of transient organizing structures and signaling centers such as the node and notochord is essential for the execution of the developmental program2. Developmental biologists have used a variety of techniques to study the morphogenesis of these structures, most notable of which is the use of cellular reporters and live ex vivo imaging to follow the dynamics in cellular and subcellular behavior2,3,4. In this report, we focus on describing the details of our optimized protocols for two of these techniques: scanning electron microscopy (SEM) and whole mount immunofluorescence (WMIF), which were and are still instrumental in studying the morphogenesis of the node and the notochordal plate, the precursor of the notochord.
The mouse embryonic node is a teardrop-shaped cup of cells that is located on the ventral surface of the mouse embryo around the early to late head fold stages during gastrulation and morphogenesis (embryonic day, E7.5-E8)2,5,6,7. The notochordal plate morphologically emanates anteriorly from the node3. Each cell in the node and notochordal plate is characterized by a single cilium that protrudes to the outside, which is longer in node cells but whose length varies with the developmental stage2. The rotation of cilia in the node pit has been shown to be important for signaling that determines left-right asymmetry4. The notochordal plate is the precursor of the notochord, the signaling center that is important for the patterning of the adjacent somites and the overlying neural tube3.
Because of the attributes of location (surface), shape (cup) and possessing distinct outer cellular structures (cilia), SEM has traditionally been used to visualize the node and notochordal plate and study their formation and structure2,7. SEM is also used to study the changes in the structure of the node itself or the cilia on its cells in mutations that affect gastrulation, morphogenesis, as well as cilia formation8,9,10. SEM is a technique that utilizes a focused beam of electrons to interrogate the topological ultrastructure of the outer surface of materials such as biological specimens11. The sample is usually fixed, dried and then sputter-coated with metals for observation under a scanning electron microscope as we describe in Step 1.
WMIF is a staining technique to visualize gene products, such as proteins, in three-dimensions (3D). WMIF of tissues, organs or even whole organisms provides spatial information about the distribution of the signal and the shape of the resulting structure in 3D. The technique is based on fixing the sample then staining it with fluorescent conjugates. Mouse embryos ~ E7.5 are small and transparent and therefore ideal for WMIF protocols to visualize the node and notochordal plate. For example, the transcription factor Barchyury (T) is expressed in the nuclei of the node and notochordal plate, and to a lesser extent in the primitive streak, around E7.5-E8 of embryonic development and good working antibodies against T by WMIF are commercially available and make the staining procedure possible. The cells of the node and notochordal plate are also characterized by constricted apical surfaces, which face the outside and thus can be stained with fluorescence-conjugated Phalloidin to mark F-Actin at the apical constrictions. Using these reagents as examples, the combination of T and F-Actin staining by WMIF provides a representation of the node and notochordal plate in 3D in gastrulating mouse embryos as we demonstrate in Step 2 8. However, markers of cilia, such as ARL13B or acetylated tubulin, as well as other markers of the node and notochordal plate, such as FOXA2, can also be used to perform WMIF on developing mouse embryos3,4.
We have shown that striatin-interacting protein 1 (STRIP1) is essential for normal gastrulation and morphogenesis in the mouse embryo8. STRIP1 is a core component of the striatin-interacting phosphatases and kinases complexes (STRIPAK), which we and others have implicated in the actin cytoskeleton organization8,12. A major defect in Strip1 mutant embryos is in the formation of the axial mesoderm (node and notochordal plate) and extension of the antero-posterior body axis. We have used SEM and WMIF to analyze the node and notochordal plate in wild-type (WT) and Strip1 mutant embryos as we show in the Representative Results and corresponding figures.

<table><tbody><tr><td>1,1,1,3,3,3 Hexamethyldisilazane (HMDS)</td><td>Carl Roth</td><td>3840</td><td></td></tr><tr><td>Anti-T antibody</td><td>R&amp;amp;D Systems</td><td>AF2058</td><td></td></tr><tr><td>Critical Point Dryer</td><td>Blazers Union</td><td>CPD 020</td><td></td></tr><tr><td>DAPI</td><td>AppliChem</td><td>A4099,0005</td><td></td></tr><tr><td>Glutardialdehyde solution 25%</td><td>Merck</td><td>1042390250</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>X100-100ML</td><td></td></tr><tr><td>Tween 20</td><td>AppliChem</td><td>A4974,0500</td><td></td></tr><tr><td>SEM coating unit PS3</td><td>Agar Aids for Electron Microscopy</td><td>PS3</td><td></td></tr><tr><td>SEM microscope Quantum FEG 250</td><td>ThermoFisher Scientific (FEI)</td><td>Quantum FEG 250</td><td></td></tr></tbody></table>

visualizing the node and notochordal plate in gastrulating mouse embryos using scanning electron microscopy and whole mount immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

In order to examine the formation of the node in WT and Strip1 mutant embryos at ~ E7.5, we used SEM as described in Step 1 and shown in Figure 18. The ultrastructural details of the outside topology using SEM were quite informative and it was immediately clear that unlike the pit-shaped node in WT embryos, the mutant embryos had a flattened and irregular node. Higher magnification of the embryos showed the characteristic cilia on node cells that identified them unambiguously. The apparent lower density of cilia in the mutant might be attributable to the loss of node pit structure and curvature or a lower number of node cells. The notochordal plate which appears emanating from the node was also irregular in the mutant embryos. They were identifiable with their shorter cilia. Therefore, SEM was important to reveal the node morphogenesis defects in Strip1 mutants8. We have also used SEM in previous studies to show the absence of cilia in the embryonic node of mutants that lacked centrioles, which provide the template for cilia9.
To study the axial mesoderm formation defects in Strip1 mutant embryos at the cellular level, we used WMIF as described in Step 2 and shown in Figure 2. Using this technique, the node and notochordal plate were easily identified by F-Actin and T staining. WT node and notochordal plate cells have constricted apical domains where F-Actin was enriched, and nuclear T staining was evident. The notochordal plate extended rostrally in the WT but was short and irregular in the mutant. The data showed that F-Actin organization is abnormal in the different germ layers of the mutant embryos including the axial mesoderm8. Thus, WMIF was instrumental to study the defects in node and notochordal plate formation in Strip1 mutant embryos.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58321/58321fig1.jpg" /> 
Figure 1. Scanning electron microscopy reveals the defects in node morphogenesis in Strip1 mutant mouse embryos. (Top) SEM analyses of WT and Strip1 mutant ventral embryonic nodes and notochordal plates (Noto)8. An example of a low magnification image of a WT embryo is shown on the left. (Bottom) Higher magnifications of the center of the nodes shown on top revealing the long monocilia projecting from the node cells. Anterior is up in all panels. Scale bars: 30 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58321/58321fig1large.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/58321/58321fig2.jpg" /> 
Figure 2. Whole mount immunofluorescence shows the abnormal node and notochordal plate at the cellular level in Strip1 mutant embryos. (Top) Ventral 3D rendering (Volocity software) of WMIF on WT and Strip1 mutant embryos using a combination of fluorescence-conjugated phalloidin (F-Actin, red) and T antibody (green) staining. (Bottom) More examples of the staining shown above focusing on the node with higher zoom and including DAPI. Anterior is up in all panels. Scale bars: 30 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58321/58321fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence at JoVE.com

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

visualizing-node-notochordal-plate-gastrulating-mouse-embryos-using

Nanyang Technological University

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Developmental Biology

8.8K Views.  University of Cologne. Het knooppunt en de notochordalplaat zijn essentiële organisatoren tijdens de embryonale ontwikkeling van de muis. Vandaag zullen we twee technieken beschrijven die door ontwikkelingsbiologen worden gebruikt om deze structuren te visualiseren en te bestuderen. In het eerste protocol laten we zien hoe we scanning electro microscopie, SEM en in het tweede, hele mount immunofluorescentie kunnen uitvoeren.Het knooppunt en de notochordalplaat zijn tijdelijk aanwezig op het oppervlak van he...

Video: Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

Labeling and Imaging Cells in the Zebrafish Hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during neurulation.

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

Neuroscience

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

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 ...

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a wide range of techniques encompassing ex vivo and in vitro approaches. However, most studies still rely on the analysis of two-dimensional (2D) imaging data, which limits proper evaluation of a developmental process like axial extension and somitogenesis involving highly dynamic interactions in a complex 3D space. Here we describe techniques that allow mouse live imaging acquisition, dataset processing, visualization and analysis in 3D and 4D to study the cells (e.g., neuromesodermal progenitors) involved in these developmental processes. We also provide a step-by-step protocol for optical projection tomography and whole-mount immunofluorescence microscopy in mouse embryos (from sample preparation to image acquisition) and show a pipeline that we developed to process and visualize 3D image data. We extend the use of some of these techniques and highlight specific features of different available software (e.g., Fiji/ImageJ, Drishti, Amira and Imaris) that can be used to improve our current understanding of axial extension and somite formation (e.g., 3D reconstructions). Altogether, the techniques here described emphasize the importance of 3D data visualization and analysis in developmental biology, and might help other researchers to better address 3D and 4D image data in the context of vertebrate axial extension and segmentation. Finally, the work also employs novel tools to facilitate teaching vertebrate embryonic development.

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a ...

Fixation of Embryonic Mouse Tissue for Cytoneme Analysis

Developmental tissue patterning and postdevelopmental tissue homeostasis depend upon controlled delivery of cellular signals called morphogens. Morphogens act in a concentration- and time-dependent manner to specify distinct transcriptional programs that instruct and reinforce cell fate. One mechanism by which appropriate morphogen signaling thresholds are ensured is through delivery of the signaling proteins by specialized filopodia called cytonemes. Cytonemes are very thin (&#8804;200 nm in diameter) and can grow to lengths of several hundred microns, which makes their preservation for fixed-image analysis challenging. This paper describes a refined method for delicate handling of mouse embryos for fixation, immunostaining, and thick sectioning to allow for visualization of cytonemes using standard confocal microscopy. This protocol has been successfully used to visualize cytonemes that connect distinct cellular signaling compartments during mouse neural tube development. The technique can also be adapted to detect cytonemes across tissue types to facilitate the interrogation of developmental signaling at unprecedented resolution.

Here, an optimized step-by-step protocol is provided for fixation, immunostaining, and sectioning of embryos to detect specialized signaling filopodia called cytonemes in developing mouse tissues.

Developmental tissue patterning and postdevelopmental tissue homeostasis depend upon controlled delivery of cellular signals called morphogens. Morphogens ...

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in vertebrates. Thus, it is critical to uncover the molecular mechanisms of how motor axons navigate to, arborize, and innervate their peripheral muscle targets during development and degeneration. Although transgenic Hb9::GFP mouse lines have long served to visualize motor axon trajectories during embryonic development, detailed descriptions of the full spectrum of axon terminal arborization remain incomplete due to the pattern complexity and limitations of current optical microscopy. Here, we describe an improved protocol that combines light sheet fluorescence microscopy (LSFM) and robust image analysis to qualitatively and quantitatively visualize developing motor axons. This system can be easily adopted to cross genetic mutants or MN disease models with Hb9::GFP lines, revealing novel molecular mechanisms that lead to defects in motor axon navigation and arborization.

Here, we describe a protocol for visualizing motor neuron projection and axon arborization in transgenic Hb9::GFP mouse embryos. After immunostaining for motor neurons, we used light sheet fluorescence microscopy to image embryos for subsequent quantitative analysis. This protocol is applicable to other neuron navigation processes in the central nervous system.

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

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