Images were acquired using microscopes maintained by the Cell and Molecular Biology Imaging Core at St. Jude Children&#39;s Research Hospital. Mouse strains were obtained from JAX. This work was supported by National Institutes of Health grant R35GM122546 (SKO) and by ALSAC of St. Jude Children&#39;s Research Hospital.

This protocol follows the approved animal care guidelines of Institutional Animal Care and Use Committee (IACUC) and St. Jude Children&#39;s Research Hospital. All strains were backcrossed five generations to the C57BL/6J strain.
1. Embryo isolation and whole mount staining
<ol>
	<li>Breed 6-week-old females and monitor for the presence of vaginal plug.</li>
	<li>Euthanize the pregnant dam by CO2 inhalation in a CO2 chamber followed by cervical dislocation according to the AVMA guidelines18. Make a Y-incision to the peritoneal cavity using dissecting scissors and forceps. Excise the uterus containing the E9.5 embryos as per approved institutional guidelines.</li>
	<li>Dissect the embryos in complete growth medium (Dulbecco&#39;s Modified Eagle Medium, supplemented with nonessential amino-acids, Na-pyruvate, L-glutamine, and 10% fetal bovine serum). Use forceps to remove the yolk sac, placenta, and surrounding membranes. 
	NOTE: When required, save each yolk sac for embryo genotyping.</li>
	<li>Rinse the isolated embryos in Hank&#39;s balanced salt solution (HBSS) to remove any residual amniotic tissue and blood.</li>
	<li>Prepare the fixative. Add paraformaldehyde (PFA) to HBSS for a final working concentration of 4% PFA. 
	CAUTION: PFA is a toxic chemical, so inhalation or direct skin exposure must be avoided. Preparation of the fixative solution is performed under a fume hood with the proper personal protective equipment of gloves and a lab coat.</li>
	<li>Add 1 mL of fixative to each well of a 24-well plate and place each embryo in an individual well. Incubate the embryos in fixative for 45 min with gentle agitation on a rocker. 
	NOTE: Critical: All washes and incubations must be done with gentle agitation (maximum 20 RPM) on a rocker or circular shaker, as abrupt handling or movement of the embryos will destroy the fixed cytonemes. Use a pipette to gently remove all solutions.</li>
	<li>Remove the fixative and wash the embryos 3 x 30 min in Phosphate-Buffered Saline (PBS) with Ca2+ and Mg2+ with added 0.1% Triton.</li>
	<li>After washing, incubate the embryos in blocking solution (PBS with Ca2+ and Mg2+, 0.1% Triton, and 5% goat serum) with gentle agitation. Block 2 x 1 h. After the second blocking incubation, perform one quick rinse of the embryos using fresh blocking solution.</li>
	<li>During the second blocking step, prepare the primary antibody solution11. Dilute the antibodies to the optimized concentration in PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum. 
	NOTE: For enhanced visualization of membrane GFP, chicken anti-GFP (1:250) can be used.</li>
	<li>Remove the blocking solution and add 1 mL of primary antibody solution to each well. Incubate at 4 &#176;C with gentle rotation for 3 days.</li>
	<li>Following the primary antibody incubation, wash the embryos 5 x 1 h at 20 RPM on a rocker in PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum.</li>
	<li>Prepare the secondary antibody solution using F(ab&#39;)2 fragment secondaries at a 1:1,000 dilution in PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum. 
	NOTE: Use of F(ab&#39;)2 fragments greatly enhances antibody penetration into sample.</li>
	<li>Add 1 mL of secondary antibody solution to each well. Incubate with gentle rocking at 4 &#176;C in the dark for 3 days. 
	NOTE: Important: From this point on, minimize embryo exposure to direct light. Optional: To prevent bacterial growth, add 0.2% sodium azide to the secondary antibody solution.</li>
	<li>Remove the secondary antibody solution and wash the embryos 3 x 30 min in PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum. If not performing actin staining with phalloidin or other actin dyes, after the first wash, add 4&#8242;,6-diamidino-2-phenylindole (DAPI) to PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum and incubate for 1 h, followed by 3 x 30 min washes as described above. 
	&#8203;NOTE: At this point, the embryos can be stored overnight at 4 &#176;C in PBS or HBSS in the dark until the embedding step the following day. However, it is recommended that embryos are embedded and sectioned immediately.</li>
</ol>
2. Embryo embedding, sectioning, and mounting
<ol>
	<li>Prepare a 4% w/v Low Melting Point (LMP) agarose solution dissolved in HBSS or PBS with Ca2+ and Mg2+ to a final volume of ~3 mL per embryo. Add the appropriate weight of LMP agarose to HBSS or PBS with Ca2+ and Mg2+ and microwave until the LMP agarose is dissolved. 
	NOTE: Once the LMP agarose is dissolved, store the solution in a bead bath, incubator, or water bath set to 55 &#176;C to prevent the LMP agarose solution from solidifying.</li>
	<li>Use a 12-well plate as the embedding mold for embryos. Place the 12-well plate in the 55 &#176;C bead bath. Add 2.5-3 mL of 4% LMP agarose to each well that will hold an embryo. 
	NOTE: Although other molds may be used, 12-well plate volumes are optimal for preparing the agarose block at later stages for sectioning.</li>
	<li>Using a perforated spoon, transfer the embryos into individual wells containing 4% LMP agarose solution (Figure 1A).
	<ol>
		<li>Transfer the 12-well plate from the bead bath to a benchtop. Use pipette tips to gently embed and orient the embryo so it is centered within the solution. Once the embryos are oriented, place the plate at -20 &#176;C for 10 min to allow quick solidification of the block. 
		CRITICAL: Ensure the embryo is positioned in the center of the block. Embryos that sink to the bottom or are too close to the edge of the mold will likely be dislodged from the agarose block while sectioning.</li>
	</ol>
	</li>
	<li>Remove the entire agarose block from the well using a scalpel and cut a rectangular block around the embryo leaving ~0.3 cm of block on each side. Allow extra length along the caudal end of the embryo. When mounted on the vibratome, orient the embryo in an upright position in the upper section of the block (Figure 1B).</li>
	<li>Apply a strip of tape to the specimen holder of the vibratome and superglue the agarose block to the tape, oriented so the blade will generate axial sections of the embryo in an anterior (cranial) to posterior sequence.</li>
	<li>Fill the vibratome chamber with cold HBSS to ensure the sample is fully immersed, and then surround the chamber with ice.</li>
	<li>Set the vibratome speed to 0.2 mm/s and frequency between 5 and 7 (50-70 Hz) with cut thickness set to 100 &#181;m. Perform serial axial sectioning of the embryo. 
	NOTE: Use a slow speed for sectioning. If the cut speed is increased above 0.25 mm/s, sectioning may tear the embryo or dislodge the embryo from the block.
	<ol>
		<li>During the sectioning series, use forceps to gently transfer individual sections to a separate 60 mm dish filled with HBSS. Use forceps to grab the block, not the tissue, to avoid tissue damage and destruction of cytonemes. 
		NOTE: Tissue sections should remain within the agarose block. If the tissue falls out of the block into the vibratome chamber, gently transfer by lifting the section out. Do not grab or pinch the tissue. CRITICAL: Any folding of tissue sections or abrupt handling will destroy fixed cytonemes in the tissue sections.</li>
	</ol>
	</li>
	<li>If not performing F-actin staining, proceed to step 2.10.
	<ol>
		<li>To perform F-actin staining, remove HBSS and incubate sections for 40 min with Actin-Red and DAPI solution diluted in PBS with Ca2+ and Mg2+, 0.1% Tween-20, and 5% goat serum at room temperature.</li>
	</ol>
	</li>
	<li>Wash sections 3 x 20 min in PBS with Ca2+ and Mg2+ and 0.1% Tween-20.</li>
	<li>Using a hydrophobic marker pen, draw a hydrophobic barrier around the edges of a charged microscope slide and add a small volume of HBSS to fill the area.</li>
	<li>Use forceps or a perforated spoon to transfer sections to the slide. 
	NOTE: Any tissue sections that are not encased in agarose can be transferred via a transfer pipette.</li>
	<li>Remove excess agarose block using forceps.</li>
	<li>Once all sections have been transferred to the slide, remove excess liquid via pipetting and the corner of an absorbent towelette, and then add several drops of mounting medium to the slide. Use enough mounting medium to cover the entire coverslip area when cured. Mount the coverslip by gently placing it on the slide. 
	&#8203;NOTE: Avoid applying pressure or disturbing coverslip until the mounting medium has cured.</li>
</ol>
3. Imaging
<ol>
	<li>Perform imaging of tissue sections on any confocal or higher resolution microscope11. Analyze a minimum of three embryos per genotype. 
	NOTE: Images of tissue sections were acquired using a TCS SP8 STED 3x confocal microscope, followed by LIGHTNING deconvolution.</li>
</ol>

<ol>
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R., 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=Fgf8+morphogen+gradient+forms+by+a+source-sink+mechanism+with+freely+diffusing+molecules.">Fgf8 morphogen gradient forms by a source-sink mechanism with freely diffusing molecules.</a> Nature. 461 (7263), 533-536 (2009).</li><li>Zhou, 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=Free+extracellular+diffusion+creates+the+Dpp+morphogen+gradient+of+the+Drosophila+wing+disc.">Free extracellular diffusion creates the Dpp morphogen gradient of the Drosophila wing disc.</a> Current Biology. 22 (8), 668-675 (2012).</li><li>Ribes, V., Briscoe, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishing+and+interpreting+graded+Sonic+Hedgehog+signaling+during+vertebrate+neural+tube+patterning:+the+role+of+negative+feedback.">Establishing and interpreting graded Sonic Hedgehog signaling during vertebrate neural tube patterning: the role of negative feedback.</a> Cold Spring Harbor Perspectives in Biology. 1 (2), 002014 (2009).</li><li>Le Dr&#233;au, G., Mart&#237;, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dorsal-ventral+patterning+of+the+neural+tube:+a+tale+of+three+signals.">Dorsal-ventral patterning of the neural tube: a tale of three signals.</a> Developmental Neurobiology. 72 (12), 1471-1481 (2012).</li><li>Kornberg, T. B., Guha, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Understanding+morphogen+gradients:+a+problem+of+dispersion+and+containment.">Understanding morphogen gradients: a problem of dispersion and containment.</a> Current Opinion in Genetics &amp; Development. 17 (4), 264-271 (2007).</li><li>Ramirez-Weber, F. A., Kornberg, T. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytonemes:+cellular+processes+that+project+to+the+principal+signaling+center+in+Drosophila+imaginal+discs.">Cytonemes: cellular processes that project to the principal signaling center in Drosophila imaginal discs.</a> Cell. 97 (5), 599-607 (1999).</li><li>Sanders, T. 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D., 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=Evidence+for+an+expansion-based+temporal+Shh+gradient+in+specifying+vertebrate+digit+identities.">Evidence for an expansion-based temporal Shh gradient in specifying vertebrate digit identities.</a> Cell. 118 (4), 517-528 (2004).</li><li>Chamberlain, C. E., Jeong, J., Guo, C., Allen, B. L., McMahon, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notochord-derived+Shh+concentrates+in+close+association+with+the+apically+positioned+basal+body+in+neural+target+cells+and+forms+a+dynamic+gradient+during+neural+patterning.">Notochord-derived Shh concentrates in close association with the apically positioned basal body in neural target cells and forms a dynamic gradient during neural patterning.</a> Development. 135 (6), 1097-1106 (2008).</li><li>Muzumdar, M. D., Tasic, B., Miyamichi, K., Li, L., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+global+double-fluorescent+Cre+reporter+mouse.">A global double-fluorescent Cre reporter mouse.</a> Genesis. 45 (9), 593-605 (2007).</li><li>Bacallao, R., Sohrab, S., Phillips, C., Pawley, J. 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The authors have no competing interests to declare.

This protocol was developed for the preservation of delicate cytonemes in embryonic tissue sections for high-resolution fluorescence microscopy. To date, visualization of cytonemes in tissue across various model organisms has relied largely upon ectopic expression of fluorescently labeled proteins using live-tissue imaging. Unfortunately, the use of fluorescently labeled live-imaging is rarely conducive to the analysis of endogenously expressed proteins, can introduce time constraints, and requires specialized imaging stages and microscopes. The staining and sectioning method described here preserves cytonemes and other cellular extensions to permit immunohistochemistry for the eventual detection of endogenously expressed proteins. This technology will facilitate new insights into how morphogens are transported across different tissues in vivo and expands the scope of model systems where cytonemes can be studied.
This protocol uses whole-mount staining and vibratome thick-sectioning, which avoids the use of equilibration solution such as glycerol or cryoprotectant solutions such as sucrose. It aims to minimize sectioned tissue handling to allow for the maximal preservation of cellular extensions. Reduced handling of the tissue after sectioning provided consistently better results in the overall preservation of cytonemes. Thus, sectioning of the embryo is one of the final steps prior to imaging.
MEM-fix, a fixation technique developed for the preservation of cytonemes of cultured cells, consists of a combination of glutaraldehyde and PFA that allows for improved 3D preservation of the innate cell architecture comparable to their live dimensions16,17. Unfortunately, MEM-fix was not useful for embryo fixation because glutaraldehyde can auto-fluoresce and severely limits antibody penetration and epitope binding in cells due to high protein cross-linking activity22,23. Thus, sectioning protocols post PFA fixation conditions were optimized to allow for the preservation of cellular extensions in embryonic tissue. Although PFA can preserve cytoneme-like extensions in embryonic tissue, image analysis should be performed with caution. PFA can cause partial dehydration of individual cells and tissues, leading to minor volume reduction and the formation of small extracellular spaces within the fixed tissue.
This protocol avoids the extra steps of sucrose resaturation for cryoprotection and tissue clearing because sucrose or glycerol solutions can cause a minor resaturation of the tissue24. The resulting minor swelling can destroy fixed cellular extensions and cytonemes migrating between cells and across tissues. This was evident when comparing thick-sectioned vibratome to cryostat sections. Although increasing tissue thickness improved the preservation of intact cytonemes, the sucrose-resaturated cryostat sections consistently had a lower incidence of detectable cytonemes.
Optimization of this protocol revealed that 100 &#181;m thick sections were the best for ensuring preservation of intact cytonemes. Thinner sections are typically used for imaging because most confocal microscopes are only capable of imaging at sufficient resolution to visualize cellular extensions to a depth of ~20-40 &#181;m without clearing25. However, the mechanical forces and distortion applied to embryonic cells from the blade while cutting thin sections destroys cytonemes. Thicker sections allow for a larger area of depth within the section while preserving intact extensions within the tissue. Additionally, the use of thicker sections allows for ease of handling to prevent excessive folding or buckling of the tissue sections.
This protocol was optimized for the maximum preservation of cytonemes in tissue of E8-10.5 mouse embryos. Minimal manipulation of the tissue after sectioning provided consistently improved overall preservation of cytonemes. It is likely that further optimization will be required to adapt the technique for later developmental stages and adult tissue sections due to increased size and tissue complexity. This will require sectioning followed by immunofluorescence staining. Such tissue may require additional clearing steps and adaptation of tissue handling during sectioning to preserve cytoneme-like extensions. These additional steps will need to be considered and addressed for future applications of this protocol.

Embryonic development is orchestrated through coordinated activation of morphogen signaling pathways. Morphogens are small, secreted proteins that are categorized into Sonic Hedgehog (SHH), transforming growth factor &#946; (TGF-&#946;)/bone morphogenic protein (BMP), Wingless-related integration site (WNT), and fibroblast and epidermal growth factor (FGF/EGF) families. Morphogens are produced and released from cellular organizing centers during tissue development and establish signaling gradients across organizing fields of cells to inform tissue morphogenesis1,2,3,4,5. One representation of morphogen gradients is found in the developing nervous system, where the presumptive central nervous system is patterned through morphogen pathway activation. This tissue, referred to as the neural tube, is comprised of opposing gradients of SHH secreted by the ventral-most notochord and floor plate, and WNTs/BMPs secreted from the dorsal roofplate, to pattern distinct neural progenitor regions6. The neural tube is commonly used to interrogate morphogen gradient integrity in developmental research.
Morphogen gradient formation relies on tight regulation of signal dispersion7. One cellular mechanism by which this occurs is through the formation of long signaling filopodia called cytonemes that facilitate direct delivery of morphogens from signal-producing cells to specific target cell populations. Cytonemes have been observed to extend hundreds of micrometers to deposit morphogens on signal-receiving cell membranes8,9. Disruption of cytoneme-mediated morphogen transport leads to developmental anomalies in both flies and vertebrates, highlighting their importance during tissue patterning10,11,12,13,14.
To date, cytonemes have been documented in Drosophila, chick, and zebrafish models, but imaging of the structures in developing mammalian embryos remains challenging8,9,15. A hurdle to effectively imaging cytonemes in complex mammalian tissues in situ is their thin and fragile nature, which makes them susceptible to damage by conventional fixation methods8. We had previously developed and optimized protocols for a modified electron microscopy fixative (MEM-fix) to preserve cytonemes in cultured cells and enable their study using confocal microscopy16,17.
Use of the MEM-fix technique has permitted identification of some of the molecules involved in SHH-induced cytoneme formation and function11,16,17. However, confirmation of these findings in the physiologically relevant context of neural tube patterning necessitated the development of novel techniques to fix and image mouse embryonic tissue. A protocol to fix mouse embryos in a manner that maintains cytoneme integrity and permits immunostaining and subsequent sectioning of embryonic tissue for confocal analysis is outlined here. This protocol was developed using a membrane-tethered green fluorescent protein (GFP) to label membrane extensions from the SHH-producing cells in the developing neural tube. Implementation of this protocol will address unanswered questions pertaining to the prevalence and significance of cytonemes in developing mammalian systems.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>12-well plate; Nunc Cell-Culture Treated&#160;</td>
			<td>Thermo Scientific&#160;</td>
			<td>150628 (Fisher Scientific 12-565-321)</td>
			<td></td>
		</tr>
		<tr>
			<td>24-Well Plate, Round Nunc</td>
			<td>Thermo Fisher Scientific</td>
			<td>142475</td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm dish&#160;</td>
			<td>Corning, Falcon</td>
			<td>353004 (Fisher Scientific 08772F)</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorbant towels (Professional Kimtech Science Kimwipes)</td>
			<td>Kimberly-Clark KIMTECH</td>
			<td>34155 (Fisher Scientific 06666A)</td>
			<td></td>
		</tr>
		<tr>
			<td>Actin Red 555 ReadyProbes Reagent&#160;</td>
			<td>Invitrogen&#160;</td>
			<td>R37112</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 AffiniPure F(ab&#39;)2 Fragment Donkey Anti-Chicken IgY (IgG) (H+L)</td>
			<td>Jackson Immunoresearch Lab Inc</td>
			<td>703-546-155</td>
			<td>1:1,000 working concentration</td>
		</tr>
		<tr>
			<td>Bead Bath 6L 230V</td>
			<td>Lab Armor</td>
			<td>Item #12L048 (Mfr. Model #74220-706)</td>
			<td>set to 55 &#176;C</td>
		</tr>
		<tr>
			<td>chicken anti-GFP</td>
			<td>Aves Labs</td>
			<td>SKU GFP-1020</td>
			<td>1:250 working concentration</td>
		</tr>
		<tr>
			<td>CO2 chamber</td>
			<td></td>
			<td></td>
			<td>plexiglass chamber connected to a CO2 emitting line.&#160;</td>
		</tr>
		<tr>
			<td>Confocal laser-scanning microscope</td>
			<td>Leica Microsystems</td>
			<td>model: Leica TCS SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI Solution (1 mg/mL)</td>
			<td>Thermo Fisher Scientific</td>
			<td>62248</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors (Vannas Scissors)</td>
			<td>World Precision Instruments</td>
			<td>14003</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium, high glucose, no glutamine</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>11960044</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus single channel pipette, 0.1-2.5 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000012</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus single channel pipette, 0.5-10 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000020</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus single channel pipette, 100-1,000 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000063</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research Plus single channel pipette, 20-200 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000055</td>
			<td></td>
		</tr>
		<tr>
			<td>Feather Disposable Scalpel #10</td>
			<td>Fisher Scientific&#160;</td>
			<td>Catalog No.NC9999403</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum, certified, United States</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>16000044</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Fine Point High Precision Forceps</td>
			<td>Fisher Scientific&#160;</td>
			<td>22-327379</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Labeling Tape</td>
			<td>Fisher Scientific&#160;</td>
			<td>15-954</td>
			<td></td>
		</tr>
		<tr>
			<td>Formaldehyde, 16%, methanol free, Ultra Pure EM Grade</td>
			<td>Polysciences</td>
			<td>18814-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s Balanced Salt Solution (HBSS)</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>14025</td>
			<td></td>
		</tr>
		<tr>
			<td>ImmEdge Hydrophobic Barrier Pap Pen&#160;</td>
			<td>Vector laboratories</td>
			<td>H-4000</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Application Suite X (LAS X) with LIGHTNING</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td>Microscope software</td>
		</tr>
		<tr>
			<td>L-Glutamine (200 mM)</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>25030081</td>
			<td></td>
		</tr>
		<tr>
			<td>Low Melting Point Agarose- UltraPure</td>
			<td>Invitrogen&#160;</td>
			<td>16520</td>
			<td></td>
		</tr>
		<tr>
			<td>MEM Non-Essential Amino Acids Solution (100x)</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>11140050</td>
			<td></td>
		</tr>
		<tr>
			<td>Moria MC 17 BIS Perforated Spoon</td>
			<td>Fine Science Tools&#160;</td>
			<td>1037018</td>
			<td></td>
		</tr>
		<tr>
			<td>Mounting media (ProLong Diamond Antifade Mountant)&#160;</td>
			<td>Thermo Fisher Scientific, Invitrogen</td>
			<td>P36961&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse: B6.129X1(Cg)-Shhtm6Amc/J (ShhGFP/+)</td>
			<td>The Jackson Laboratory</td>
			<td>Strain #:008466</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse: B6.Cg-Shhtm1(EGFP/cre)Cjt/J (Shh-Cre)</td>
			<td>The Jackson Laboratory</td>
			<td>Strain #:005622</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse: C57BL/6J (B6)</td>
			<td>The Jackson Laboratory</td>
			<td>Strain #:000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse: Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J (Rosa26 mTmG)</td>
			<td>The Jackson Laboratory</td>
			<td>Strain #:007576</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Goat Serum&#160;</td>
			<td>Jackson ImmunoResearch</td>
			<td>005-000-121 (Fisher Scientific NC9660079)</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS + Ca2+ + Mg2+&#160;</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>14040133</td>
			<td></td>
		</tr>
		<tr>
			<td>Premium Microcentrifuge Tubes: 1.5 mL</td>
			<td>Fisher Scientific&#160;</td>
			<td>05-408-129</td>
			<td></td>
		</tr>
		<tr>
			<td>SHARP Precision Barrier Tips, Extra Long for P-10 and Eppendorf 10 &#181;L</td>
			<td>Denville Scientific</td>
			<td>P1096-FR</td>
			<td></td>
		</tr>
		<tr>
			<td>SHARP Precision Barrier Tips, For P-1000 and Eppendorf 1,000, 1,250 &#181;L</td>
			<td>Denville Scientific</td>
			<td>P1126</td>
			<td></td>
		</tr>
		<tr>
			<td>SHARP Precision Barrier Tips, For P-20 and Eppendorf 20 &#181;L</td>
			<td>Denville Scientific</td>
			<td>P1121</td>
			<td></td>
		</tr>
		<tr>
			<td>SHARP Precision Barrier Tips, For P-200 and Eppendorf 200 &#181;L</td>
			<td>Denville Scientific</td>
			<td>P1122</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td>Sigma-Aldrich</td>
			<td>S8032</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Pyruvate (100 mM) (100x)</td>
			<td>Thermo Fisher Scientific, Gibco</td>
			<td>11360070</td>
			<td></td>
		</tr>
		<tr>
			<td>Super Glue</td>
			<td>Gorilla</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SuperFrost Plus microscope slides&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>TPP centrifuge tubes, volume 15 mL, polypropylene</td>
			<td>TPP Techno Plastic Products</td>
			<td>91015</td>
			<td></td>
		</tr>
		<tr>
			<td>TPP centrifuge tubes, volume 50 mL, polypropylene</td>
			<td>TPP Techno Plastic Products</td>
			<td>91050</td>
			<td></td>
		</tr>
		<tr>
			<td>Transfer pipette, polyethylene</td>
			<td>Millipore Sigma</td>
			<td>Z135003</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>T9284</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>P1379</td>
			<td></td>
		</tr>
		<tr>
			<td>Vari-Mix Platform Rocker</td>
			<td>Thermo Fisher Scientific</td>
			<td>09-047-113Q</td>
			<td>set to 20 RPM</td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica Biosytems</td>
			<td>VT1000 S</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Use of a larger mold of a 12-well plate with 2.5-3 mL of agarose solution per well was ideal for embedding and suspending several embryos over a short period of time (Figure 1A). The excess area allows for correct orientation when cutting the agarose block for sectioning. When cutting the agarose block, it is important to maintain excess agarose along the bottom of the block where it will be glued to the tape on the object holder. The embryo should be in the upper half of the block (Figure 1B). However, the bottom section should not be too large as excess block increases the chances of altering the cut angle as the blade pushes into the block while sectioning. Examples of correctly oriented sections are shown (Figure 1C,D).
While developing this protocol, vibratome sectioning was compared to cryostat sectioning. Cryostat sectioning of the tissue rarely preserved cellular extensions (Figure 2 cryostat&#160;and Figure 3A,B vibratome). Cryostat sections allowed for the detection of some GFP-positive membrane fragments between cells of the notochord and neural tube (Figure 2A arrowhead) and between adjacent neural tube cells (Figure 2B,B&#39; arrowheads). However, F-actin staining of cellular extensions in the highly filamentous mesenchymal cells surrounding the neural tube was impaired in cryostat sections (Figure 2C,C&#39; arrow, vs Figure 3C,D). These cryostat results indicate that only some traces of broken cellular extensions remain following this method. Thus, vibratome sectioning is preferred to efficiently preserve these delicate extensions for subsequent analysis.
Minimal disruption of whole embryo and individual tissue sections is essential for high-quality imaging of cellular extensions. Tissue sections that have undergone any folding or buckling will be evident by the absence of the notochord or a large separation (&#62;30 &#181;m) between the notochord and the ventral floor plate of the neural tube (Figure 3B). This is usually accompanied by loss of mesenchymal cells that normally surround the neural tube (Figure 3A, arrow). Damaged sections will also result in a loss of visible cellular membrane extensions migrating between epithelial cells (Figure 3B compared to Figure 3A, arrowheads). Even minor distortions to the sections may cause fragmentation of actin-based extensions and large gaps to form between cells (Figure 3D, arrowheads, compared to well-preserved Figure 3C), underscoring the need for delicate handling at all steps.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64100/64100fig01.jpg" /> 
Figure 1: Example of E9.5 Shh-Cre; Rosa26 mTmG stained, embedded, and sectioned embryo. (A) A single embryo in a 12-well plate, embedded in 4% LMP agarose. (B) Example of a correctly oriented embryo within the agarose block cut down to size for vibratome mounting. Excess agarose block is present along the bottom. (C) Bright field image of 100 &#181;m thick embryo section embedded in LMP agarose. (D) Immunofluorescence imaging of a section after removal of agarose. Anti-GFP-stained mGFP (green) represents Shh-expressing cells in the tissue, with other cell lineages expressing membrane Tomato (red), DAPI in blue. Neural tube (bracket) and mGFP-positive notochord (arrow) are clearly visible. Scale bars = 1 cm (A), 5 mm (B), and 100 &#181;m (C,D). Abbreviations: LMP = low melting point; mGFP = membrane green fluorescent protein; Shh = Sonic Hedgehog; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64100/64100fig01large.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/64100/64100fig02.jpg" /> 
Figure 2: Example of cryostat-sectioned neural tube floor plate and notochord with fragmented membrane extensions. (A) 20 &#181;m thick E9.5 Shh-Cre; Rosa26 mTmG19,20,21 section stained for GFP (green), F-actin (red), and DAPI (blue). mGFP puncta are visible between the notochord and floorplate (arrowhead) and (B,B&#39;) migrating between adjacent cells of the neural tube. (C,C&#39;) F-actin staining fails to detect any clear cytonemes on mesenchymal cells surrounding the neural tube (arrow). Scale bars = 20 &#181;m. Abbreviations: mGFP = membrane green fluorescent protein; Shh = Sonic Hedgehog; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64100/64100fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64100/64100fig03.jpg" /> 
Figure 3: Examples of optimal and suboptimal vibratome tissue sections and staining of the neural tube floor plate, notochord, and surrounding cells. Examples of delicately handled sections (A,C), compared to (B) a folded tissue section and (D) a poorly handled section from a Shh-Cre; Rosa26 mTmG and a ShhGFP/+ embryo19,20,21. Optimally handled sections allow detection of cytonemes between adjacently localized floor plate neural epithelial cells (A, arrowheads). Neural tube adjacent notochord (A, mGFP-expressing) and mesenchymal cells (A, arrow) should be visible. (C,D) F-actin- and DAPI-stained sections should have consistent spacing of mesenchymal cells and F-actin-based cytonemes (arrows) surrounding the neural tube and notochord (C). Any minor folding or disruption of the sections can cause gaps and broken F-actin fragments (D, arrowheads). Scale bars = 10 &#181;m. Abbreviations: mGFP = membrane green fluorescent protein; Shh = Sonic Hedgehog; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64100/64100fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fixation of Embryonic Mouse Tissue for Cytoneme Analysis at JoVE.com

Fixation of Embryonic Mouse Tissue for Cytoneme Analysis

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 ...

fixation-of-embryonic-mouse-tissue-for-cytoneme-analysis

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

2.3K Views.  St. Jude Children’s Research Hospital. 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. 

Video: Fixation of Embryonic Mouse Tissue for Cytoneme Analysis

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

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 ...

Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel&#8217;s Cartilage

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. Patterning of the Meckel&#39;s cartilage, a first pharyngeal arch derivative, involves the migration of cranial neural crest (CNC) cells and the progressive partitioning, proliferation and organization of differentiated chondrocytes. Several studies have described CNC migration during lower jaw&#160;morphogenesis, but the details of how the chondrocytes achieve organization in the growth and extension of Meckel&#8217;s cartilage remains unclear. The sox10 restricted and chemically induced Cre recombinase-mediated recombination generates permutations of distinct fluorescent proteins (RFP, YFP and CFP), thereby creating a multi-spectral labeling of progenitor cells and their progeny, reflecting distinct clonal populations. Using confocal time-lapse photography, it is possible to observe the chondrocytes behavior during the development of the zebrafish Meckel&#8217;s cartilage.
Multispectral cell labeling enables scientists to demonstrate extension of the Meckel&#8217;s chondrocytes.&#160;During extension phase of the Meckel&#8217;s cartilage, which prefigures the mandible, chondrocytes intercalate to effect extension as they stack in an organized single-cell layered row. Failure of this organized intercalating process to mediate cell extension provides the cellular mechanistic explanation for hypoplastic mandible that we observe in mandibular malformations.

Cell organization of craniofacial bones has long been hypothesized but never directly visualized. Multi-spectral cell labeling and in vivo live imaging allows visualization of dynamic cell behavior in zebrafish lower jaw. Here, we detail the protocol to manipulate Zebrabow transgenic fish and directly observe cell intercalation and morphological changes of chondrocytes in the Meckel&#8217;s cartilage.

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. ...

Reprogramming Mouse Embryonic Fibroblasts with Transcription Factors to Induce a Hemogenic Program

This protocol details the induction of a hemogenic program in mouse embryonic fibroblasts (MEFs) via overexpression of transcription factors (TFs). We first designed a reporter screen using MEFs from human CD34-tTA/TetO-H2BGFP (34/H2BGFP) double transgenic mice. CD34+ cells from these mice label H2B histones with GFP, and cease labeling upon addition of doxycycline (DOX). MEFS were transduced with candidate TFs and then observed for the emergence of GFP+ cells that would indicate the acquisition of a hematopoietic or endothelial cell fate. Starting with 18 candidate TFs, and through a process of combinatorial elimination, we obtained a minimal set of factors that would induce the highest percentage of GFP+ cells. We found that Gata2, Gfi1b, and cFos were necessary and the addition of Etv6 provided the optimal induction. A series of gene expression analyses done at different time points during the reprogramming process revealed that these cells appeared to go through a precursor cell that underwent an endothelial to hematopoietic transition (EHT). As such, this reprogramming process mimics developmental hematopoiesis "in a dish," allowing study of hematopoiesis in vitro and a platform to identify the mechanisms that underlie this specification. This methodology also provides a framework for translation of this work to the human system in the hopes of generating an alternative source of patient-specific hematopoietic stem cells (HSCs) for a number of applications in the treatment and study of hematologic diseases.

The protocol described here details the induction of a hemogenic program in mouse embryonic fibroblasts via overexpression of a minimal set of transcription factors. This technology may be translated to the human system to provide platforms for future study of hematopoiesis, hematologic disease, and hematopoietic stem cell transplant.

This protocol details the induction of a hemogenic program in mouse embryonic fibroblasts (MEFs) via overexpression of transcription factors (TFs). We ...

Analysis of Coronary Vessels in Cleared Embryonic Hearts

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard microscopy techniques, due to the scattering of imaging light by the thick heart tissue, as these vessels are localized deep within the walls of the developing heart. As optical clearing of tissues using organic solvents such as BABB (1 part benzyl alcohol to 2 parts benzyl benzoate) has been shown to greatly improve the optical penetration depth that can be achieved, we combined clearance of whole, PECAM1-immunostained hearts, with laser-scanning confocal microscopy, in order to obtain high-resolution images of vessels throughout the entire heart. BABB clearance of embryonic hearts takes place rapidly and also acts to preserve the fluorescent signal for several weeks; in addition, samples can be imaged multiple times without loss of signal. This straightforward method is also applicable to imaging other types of blood vessels in whole embryos.

We present a protocol for the analysis of coronary vessels in whole embryonic murine hearts up to E15.5, using standard immunological staining methods followed by optical clearance and confocal microscopy. This technique enables visualization of blood vessels throughout the entire heart without the need for time-consuming analysis of serial sections.

Whole mount visualization of the embryonic coronary plexus from which the capillary and arterial networks will form is rendered problematic using standard ...

Structure-function Studies in Mouse Embryonic Stem Cells Using Recombinase-mediated Cassette Exchange

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of the cell and often consist of multiple functional domains, which can be influenced by posttranslational modifications. The depletion of the entire protein in conditional or constitutive knock-out (KO) mice does not take into account this functional diversity and regulation. An mESC line and a derived mouse model, in which a docking site for FLPe recombination-mediated cassette exchange (RMCE) was inserted within the ROSA26 (R26) locus, was previously reported. Here, we report on a structure-function approach that allows for molecular dissection of the different functionalities of a multidomain protein. To this end, RMCE-compatible mice must be crossed with KO mice and then RMCE-compatible KO mESCs must be isolated. Next, a panel of putative rescue constructs can be introduced into the R26 locus via RMCE targeting. The candidate rescue cDNAs can be easily inserted between RMCE sites of the targeting vector using recombination cloning. Next, KO mESCs are transfected with the targeting vector in combination with an FLPe recombinase expression plasmid. RMCE reactivates the promoter-less neomycin-resistance gene in the ROSA26 docking sites and allows for the selection of the correct targeting event. In this way, high targeting efficiencies close to 100% are obtained, allowing for insertion of multiple putative rescue constructs in a semi-high throughput manner. Finally, a multitude of R26-driven rescue constructs can be tested for their ability to rescue the phenotype that was observed in parental KO mESCs. We present a proof-of-principle structure-function study in p120 catenin (p120ctn) KO mESCs using endoderm differentiation in embryoid bodies (EBs) as the phenotypic readout. This approach enables the identification of important domains, putative downstream pathways, and disease-relevant point mutations that underlie KO phenotypes for a given protein.

Proteins often contain multiple domains that can exert different cellular functions. Gene knock-outs (KO) do not consider this functional diversity. Here, we report a recombination-mediated cassette exchange (RMCE)-based structure-function approach in KO embryonic stem cells that allows for the molecular dissection of various functional domains or variants of a protein.

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of ...

Analysis of Retinoic Acid-induced Neural Differentiation of Mouse Embryonic Stem Cells in Two and Three-dimensional Embryoid Bodies

Mouse embryonic stem cells (ESCs) isolated from the inner mass of the blastocyst (typically at day E3.5), can be used as in vitro model system for studying early embryonic development. In the absence of leukemia inhibitory factor (LIF), ESCs differentiate by default into neural precursor cells. They can be amassed into a three dimensional (3D) spherical aggregate termed embryoid body (EB) due to its similarity to the early stage embryo. EBs can be seeded on fibronectin-coated coverslips, where they expand by growing two dimensional (2D) extensions, or implanted in 3D collagen matrices where they continue growing as spheroids, and differentiate into the three germ layers: endodermal, mesodermal, and ectodermal. The 3D collagen culture mimics the in vivo environment more closely than the 2D EBs. The 2D EB culture facilitates analysis by immunofluorescence and immunoblotting to track differentiation. We have developed a two-step neural differentiation protocol. In the first step, EBs are generated by the hanging-drop technique, and, simultaneously, are induced to differentiate by exposure to retinoic acid (RA). In the second step, neural differentiation proceeds in a 2D or 3D format in the absence of RA.

We describe a technique for using murine embryonic stem cells for generating two or three dimensional embryoid bodies. We then explain how to induce neural differentiation of the embryoid body cells by retinoic acid, and how to analyze their state of differentiation by progenitor cell marker immunofluorescence and immunoblotting.

Mouse embryonic stem cells (ESCs) isolated from the inner mass of the blastocyst (typically at day E3.5), can be used as in vitro model system for ...

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

Dissection and Culture of Mouse Embryonic Kidney

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney development, the two progenitor tissues, the ureteric bud and the metanephric mesenchyme, communicate and reciprocally induce cellular mechanisms to eventually form the collecting system and the nephrons of the kidney. As mammalian embryos grow intrauterine and therefore are inaccessible to the observer, an organ culture has been developed. With this method, it is possible to study epithelial-mesenchymal interactions and cellular behavior during kidney organogenesis. Furthermore, the origin of congenital kidney and urogenital tract malformations can be investigated. After careful dissection, the metanephric rudiments are transferred onto a filter that floats on culture medium and can be kept in a cell culture incubator for several days. However, one must be aware that the conditions are artificial and could influence the metabolism in the tissue. Also, the penetration of test substances could be limited due to the extracellular matrix and basal membrane present in the explant.
One main advantage of organ culture is that the experimenter can gain direct access to the organ. This technology is cheap, simple, and allows a large number of modifications, such as the addition of biologically active substances, the study of genetic variants, and the application of advanced imaging techniques.

This protocol describes a method for isolating and culturing metanephric rudiments from mouse embryos.

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Fixation of Embryonic Mouse Tissue for Cytoneme Analysis

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Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel’s Cartilage

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Analysis of Coronary Vessels in Cleared Embryonic Hearts

Structure-function Studies in Mouse Embryonic Stem Cells Using Recombinase-mediated Cassette Exchange

Analysis of Retinoic Acid-induced Neural Differentiation of Mouse Embryonic Stem Cells in Two and Three-dimensional Embryoid Bodies

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Dissection and Culture of Mouse Embryonic Kidney

Live Imaging of Mouse Secondary Palate Fusion