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>

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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><tbody><tr><td>12-well plate; Nunc Cell-Culture Treated&amp;nbsp;</td><td>Thermo Scientific&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</td><td>Invitrogen&amp;nbsp;</td><td>R37112</td><td></td></tr><tr><td>Alexa Fluor 488 AffiniPure F(ab&#039;)&lt;sub&gt;2&lt;/sub&gt; 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 &amp;deg;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>CO&lt;sub&gt;2&lt;/sub&gt; chamber</td><td></td><td></td><td>plexiglass chamber connected to a CO&lt;sub&gt;2&lt;/sub&gt; emitting line.&amp;nbsp;</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&#039;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 &amp;micro;L</td><td>Eppendorf</td><td>3123000012</td><td></td></tr><tr><td>Eppendorf Research Plus single channel pipette, 0.5-10 &amp;micro;L</td><td>Eppendorf</td><td>3123000020</td><td></td></tr><tr><td>Eppendorf Research Plus single channel pipette, 100-1,000 &amp;micro;L</td><td>Eppendorf</td><td>3123000063</td><td></td></tr><tr><td>Eppendorf Research Plus single channel pipette, 20-200 &amp;micro;L</td><td>Eppendorf</td><td>3123000055</td><td></td></tr><tr><td>Feather Disposable Scalpel #10</td><td>Fisher Scientific&amp;nbsp;</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&amp;nbsp;</td><td>22-327379</td><td></td></tr><tr><td>Fisherbrand Labeling Tape</td><td>Fisher Scientific&amp;nbsp;</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&#039;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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</td><td>1037018</td><td></td></tr><tr><td>Mounting media (ProLong Diamond Antifade Mountant)&amp;nbsp;</td><td>Thermo Fisher Scientific, Invitrogen</td><td>P36961&amp;nbsp;</td><td></td></tr><tr><td>Mouse: B6.129X1(Cg)-Shh&lt;sup&gt;tm6Amc&lt;/sup&gt;/J (Shh&lt;sup&gt;GFP/+&lt;/sup&gt;)</td><td>The Jackson Laboratory</td><td>Strain #:008466</td><td></td></tr><tr><td>Mouse: B6.Cg-Shh&lt;sup&gt;tm1(EGFP/cre)Cjt&lt;/sup&gt;/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)26Sor&lt;sup&gt;tm4(ACTB-tdTomato,-EGFP)Luo&lt;/sup&gt;/J (Rosa26 mTmG)</td><td>The Jackson Laboratory</td><td>Strain #:007576</td><td></td></tr><tr><td>Normal Goat Serum&amp;nbsp;</td><td>Jackson ImmunoResearch</td><td>005-000-121 (Fisher Scientific NC9660079)</td><td></td></tr><tr><td>PBS + Ca&lt;sup&gt;2+&lt;/sup&gt; + Mg&lt;sup&gt;2+&lt;/sup&gt;&amp;nbsp;</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&amp;nbsp;</td><td>05-408-129</td><td></td></tr><tr><td>SHARP Precision Barrier Tips, Extra Long for P-10 and Eppendorf 10 &amp;micro;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 &amp;micro;L</td><td>Denville Scientific</td><td>P1126</td><td></td></tr><tr><td>SHARP Precision Barrier Tips, For P-20 and Eppendorf 20 &amp;micro;L</td><td>Denville Scientific</td><td>P1121</td><td></td></tr><tr><td>SHARP Precision Barrier Tips, For P-200 and Eppendorf 200 &amp;micro;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&amp;nbsp;</td><td>Fisher Scientific&amp;nbsp;</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&amp;nbsp;</td><td>Sigma-Aldrich</td><td>T9284</td><td></td></tr><tr><td>Tween-20&amp;nbsp;</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://www.jove.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://www.jove.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://www.jove.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

Research

JoVE Journal

Developmental Biology

2.4K 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

A Seminiferous Tubule Squash Technique for the Cytological Analysis of Spermatogenesis Using the Mouse Model

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during meiosis can lead to infertility, pregnancy loss or genetic defects. Commencing at the onset of puberty and continuing throughout adulthood, continuous semi-synchronous waves of spermatocytes undergo spermatogenesis and ultimately form haploid sperm. The first wave of mouse spermatocytes undergoing meiotic initiation appear at day 10 post-partum (10 dpp) and are released into the lumen of seminiferous tubules as mature sperm at 35 dpp. Therefore, it is advantageous to utilize mice within this developmental time-window in order to obtain highly enriched populations of interest. Analysis of rare cell stages is more difficult in older mice due to the contribution of successive spermatogenic waves, which increase the diversity of the cellular populations within the tubules. The method described here is an easily implemented technique for the cytological evaluation of the cells found within the seminiferous tubules of mice, including spermatogonia, spermatocytes, and spermatids. The tubule squash technique maintains the integrity of isolated male germ cells and allows examination of cellular structures that are not easily visualized with other techniques. To demonstrate the possible applications of this tubule squash technique, spindle assembly was monitored in spermatocytes progressing through the prophase to metaphase I transition (G2/MI transition). In addition, centrosome duplication, meiotic sex chromosome inactivation (MSCI), and chromosome bouquet formation were assessed as examples of the cytological structures that can be observed using this tubule squash method. This technique can be used to pinpoint specific defects during spermatogenesis that are caused by mutation or exogenous perturbation, and thus, contributes to our molecular understanding of spermatogenesis.

The goal of this tubule squash technique is to rapidly assess cytological features of developing mouse spermatocytes while preserving cellular integrity. This method allows for the study of all stages of spermatogenesis, and can be easily implemented alongside other biochemical and molecular biological approaches for the study of mouse meiosis.

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during ...

Immunohistochemical Detection of 5-Methylcytosine and 5-Hydroxymethylcytosine in Developing and Postmitotic Mouse Retina

The epigenetics of retinal development is a well-studied research field, which promises to bring a new level of understanding about the mechanisms of a variety of human retinal degenerative diseases and pinpoint new treatment approaches. The nuclear architecture of mouse retina is organized in two different patterns: conventional and inverted. Conventional pattern is universal where heterochromatin is localized to the periphery of the nucleus, while active euchromatin resides in the nuclear interior. In contrast, inverted nuclear pattern is unique to the adult rod photoreceptor cell nuclei where heterochromatin localizes to the nuclear center, and euchromatin resides in the nuclear periphery. DNA methylation is predominantly observed in chromocenters. DNA methylation is a dynamic covalent modification on the cytosine residues (5-methylcytosine, 5mC) of CpG dinucleotides that are enriched in the promoter regions of many genes. Three DNA methyltransferases (DNMT1, DNMT3A and DNMT3B) participate in methylation of DNA during development. Detecting 5mC with immunohistochemical techniques is very challenging, contributing to variability in results, as all DNA bases including 5mC modified bases are hidden within the double-stranded DNA helix. However, detailed delineation of 5mC distribution during development is very informative. Here, we describe a reproducible technique for robust immunohistochemical detection of 5mC and another epigenetic DNA marker 5-hydroxymethylcytosine (5hmC), which colocalizes with the &#34;open&#34;, transcriptionally active chromatin in developing and postmitotic mouse retina.

The objective of this report is to describe the protocol for robust immunohistochemical detection of epigenetic markers, 5-methylcytosine (5mC) and 5-hydroxymethylcytosine (5hmC) in developing and postmitotic mouse retina.

The epigenetics of retinal development is a well-studied research field, which promises to bring a new level of understanding about the mechanisms of a ...

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.

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 ...

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and simple protocol to detect protein expression during craniofacial morphogenesis/pathogenesis using mouse craniofacial tissues as examples. The protocol consists of preparation and cryosectioning of tissues, indirect immunofluorescence, image acquisition, and quantification. In addition, a method for preparation and cryosectioning of undecalcified hard tissues for immunostaining is described, using craniofacial tissues and long bones as examples. Those methods are key to determine the protein expression and morphological/anatomical changes in various tissues during craniofacial morphogenesis/pathogenesis. They are also applicable to other tissues with appropriate modifications. Knowledge of the histology and high quality of sections are critical to draw scientific conclusions from experimental outcomes. Potential limitations of this methodology include but are not limited to specificity of antibodies and difficulties of quantification, which are also discussed here.

Here, we present a detailed protocol to detect and quantify protein levels during craniofacial morphogenesis/pathogenesis by immunostaining using mouse craniofacial tissues as examples. In addition, we describe a method for preparation and cryosectioning of undecalcified hard tissues from young mice for immunostaining.

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Neuroscience

Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits during embryonic maturation of the brain however is technically challenging because most transsynaptic tracing methods developed to date depend on stereotaxic tracer injection. To overcome this problem, we developed a binary genetic strategy for conditional genetic transsynaptic tracing in the mouse brain. Towards this end we generated two complementary knock-in mouse strains to selectively express the bidirectional transsynaptic tracer barley lectin (BL) and the retrograde transsynaptic tracer Tetanus Toxin fragment C from the ROSA26 locus after Cre-mediated recombination. Cell-specific tracer production in these mice is genetically encoded and does not depend on mechanical tracer injection. Therefore our experimental approach is suitable to study neural circuit formation in the embryonic murine brain. Furthermore, because tracer transfer across synapses depends on synaptic activity, these mouse strains can be used to analyze the communication between genetically defined neuronal populations during brain development at a single cell resolution. Here we provide a detailed protocol for transsynaptic tracing in mouse embryos using the novel recombinant ROSA26 alleles. We have utilized this experimental technique in order to delineate the neural circuitry underlying maturation of the reproductive axis in the developing female mouse brain.

Capitalizing on a binary genetic strategy we provide a detailed protocol for neural circuit tracing in mice that express complementary transsynaptic tracers after Cre-mediated recombination. Because cell-specific tracer production is genetically encoded, our experimental approach is suitable to study the formation and maturation of neural circuitry during murine embryonic brain development at a single cell resolution.

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits ...

Cortex-, Hippocampus-, Thalamus-, Hypothalamus-, Lateral Septal Nucleus- and Striatum-specific In Utero Electroporation in the C57BL/6 Mouse

In utero electroporation is a widely used technique for fast and efficient spatiotemporal manipulation of various genes in the rodent central nervous system. Overexpression of desired genes is just as possible as shRNA mediated loss-of-function studies. Therefore it offers a wide range of applications. The feasibility to target particular cells in a distinct area further increases the range of potential applications of this very useful method. For efficiently targeting specific regions knowledge about the subtleties, such as the embryonic stage, the voltage to apply and most importantly the position of the electrodes, is indispensable.
Here, we provide a detailed protocol that allows for specific and efficient in utero electroporation of several regions of the C57BL/6 mouse central nervous system. In particular it is shown how to transfect regions the develop into the retrosplenial cortex, the motor cortex, the somatosensory cortex, the piriform cortex, the cornu ammonis 1-3, the dentate gyrus, the striatum, the lateral septal nucleus, the thalamus and the hypothalamus. For this information about the appropriate embryonic stage, the appropriate voltage for the corresponding embryonic stage is provided. Most importantly an angle-map, which indicates the appropriate position of the positive pole, is depicted. This standardized protocol helps to facilitate efficient in utero electroporation, which might also lead to a reduced number of animals.

Cortex-, Hippocampus-, Thalamus-, Hypothalamus-, Lateral Septal Nucleus- and Striatum-specific In Utero Electroporation in the C57BL/6 Mouse

This protocol describes in detail how to specifically transfect different regions in the C57BL/6 central nervous system via in utero electroporation. Included in this protocol are detailed instructions for transfections of regions that develop into the cortex, hippocampus, thalamus, hypothalamus, lateral septal nucleus and striatum.

In utero electroporation is a widely used technique for fast and efficient spatiotemporal manipulation of various genes in the rodent central nervous ...

Chromatin Immunoprecipitation Assay for Tissue-specific Genes using Early-stage Mouse Embryos

Chromatin immunoprecipitation (ChIP) is a powerful tool to identify protein:chromatin interactions that occur in the context of living cells 1-3. This technique has been widely exploited in tissue culture cells, and to a lesser extent, in primary tissue. The application of ChIP to rodent embryonic tissue, especially at early times of development, is complicated by the limited amount of tissue and the heterogeneity of cell and tissue types in the embryo. Here we present a method to perform ChIP using a dissociated embryonic day 8.5 (E8.5) embryo. Sheared chromatin from a single E8.5 embryo can be divided into up to five aliquots, which allows the investigator sufficient material for controls and for investigation of specific protein:chromatin interactions.
We have utilized this technique to begin to document protein:chromatin interactions during the specification of tissue-specific gene expression programs. The heterogeneity of cell types in an embryo necessarily restricts the application of this technique because the result is the detection of protein:chromatin interactions without distinguishing whether the interactions occur in all, a subset of, or a single cell type(s). However, examination of tissue-specific genes during or following the onset of tissue-specific gene expression is feasible for two reasons. First, immunoprecipitation of tissue specific factors necessarily isolates chromatin from the cell type where the factor is expressed. Second, immunoprecipitation of coactivators and histones containing post-translational modifications that are associated with gene activation should only be found at genes and gene regulatory sequences in the cell type where the gene is being or has been activated. The technique should be applicable to the study of most tissue-specific gene activation events.
In the example described below, we utilized E8.5 and E9.5 mouse embryos to examine factor binding at a skeletal muscle specific gene promoter. Somites, which are the precursor tissues from which the skeletal muscles of the trunk and limbs will form, are present at E8.5-9.54,5. Myogenin is a regulatory factor required for skeletal muscle differentiation 6-9. The data demonstrate that myogenin is associated with its own promoter in E8.5 and E9.5 embryos. Because myogenin is only expressed in somites at this stage of development 6,10, the data indicate that myogenin interactions with its own promoter have already occurred in skeletal muscle precursor cells in E8.5 embryos.

We demonstrate a chromatin immunoprecipitation (ChIP) method to identify factor interactions at tissue-specific genes during or after the onset of tissue-specific gene expression in mouse embryonic tissue. This protocol should be widely applicable for the study of tissue-specific gene activation as it occurs during normal embryonic development.

Chromatin immunoprecipitation (ChIP) is a powerful tool to identify protein:chromatin interactions that occur in the context of living cells 1-3. This ...

Biology

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

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.

Fixation of Embryonic Mouse Tissue for Cytoneme Analysis

Summary

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Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

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