We thank Prof. Michael Klymkowsky in the University of Colorado at Boulder and Prof. Nancy Papalopulu in the University of Manchester for kindly providing anti-Sox3 and anti-Myt1 antibodies, respectively. We thank the members of the Papalopulu lab for their insightful suggestions in developing this protocol. This work was supported by two Healing Foundation Studentships to SZ and JL, two project grants from the Healing Foundation to SZ/EA and JL/EA respectively, one program grant from the Wellcome Trust to EA (WT082450MA), and one Institutional Strategic Support grant from the Wellcome Trust [097820/Z/11/Z].

All animal experiments were approved from the University of Manchester Animal Welfare Centre and were covered by a UK Home Office Project License.
1. Collection and Fixation of Xenopus Embryos
<ol>
	<li>Prepare Reagents and Materials for Experiments.
	<ol>
		<li>Prepare 10x Marc&#39;s Modified Ringers (MMR) by dissolving 56.5 g of NaCl in approximately 800 ml ultrapure water and adding stock solutions of 1 M KCl, 1 M MgSO4, 1 M CaCl2, and 1 M HEPES pH 7.4 to achieve a final concentration of 20 mM KCl, 10 mM MgSO4, 20 mM CaCl2, 50 mM HEPES. Adjust pH to 7.4 by 10 M NaOH and then adjust the final volume to 1 L.</li>
		<li>Sterilize the 10x MMR solution by autoclaving at 121 &#176;C for 20 min on a liquid cycle. Upon using, dilute with ddH2O to 0.1x final concentration and add 20 mg/L gentamycin to inhibit microbial growth.</li>
		<li>Make 10x TBS solution by mixing 24 g Tris-HCl, 5.6 g Tris-base, 88 g NaCl and dissolving in approximately 900 ml ultrapure water. The final solution will have a pH value around 7.6. Adjust with either 10 M NaOH or concentrated HCl to achieve a final pH of 7.6 and final volume to 1 L.</li>
		<li>Upon using, make 1x TBS by diluting 1 part of 10x TBS solution with 9 parts of ultrapure water.</li>
		<li>Make 10x MEM Salt by dissolving 209.2 g MOPS in approximately 800 ml ultrapure water and adding stock solutions of 0.5 M EGTA and 1M MgSO4 to achieve a final concentration of 20 mM EGTA, 10 mM MgSO4. Adjust pH to 7.4 by 10 M NaOH and then adjust the final volume to 1 L.</li>
		<li>Sterilize the MEM salt solution by autoclaving at 121 &#176;C for 20 min on a liquid cycle (the solution may turn yellow by a few months of storage at room temperature or after it has been autoclaved, but this change in color does not affect its use). However, do not use the solution after prolonged storage (more than 6 month).</li>
		<li>Make 1x MEMFA solution by diluting 1 part of MEM Salts, 1 part of 37% formaldehyde with 8 parts of Ultrapure water (v/v, stable at 4 &#176;C for at least 1-2 weeks).</li>
		<li>Prepare 4% paraformaldehyde in TBS (for subsequent staining involving phalloidin) by dissolving 4 g of paraformaldehyde powder in 100 ml of 1x TBS solution Heat the solution to 60 &#176;C and add a few drops of 10 M NaOH to assist dissolving. Aliquot in 5-10 ml volume and freeze in -20 &#176;C. Do not re-freeze once thawed. 
		CAUTION: Paraformaldehyde powder is an irritant and is toxic if inhaled, thus the weighing step should be performed in a fume hood.</li>
		<li>Label as many 4 ml glass vials with screw caps prior to sample collection.</li>
		<li>Prepare 15% gelatin/15% sucrose by pouring 20 ml of 40% fish gelatin (pre-heat in a 50 &#176;C water bath) into a 50 ml centrifuge tube. Add 8 g of sucrose and fill the tube to the 50 ml line with 1x TBS.</li>
		<li>Place the gelatin tube on a rotary mixer or rolling bed to mix overnight at room temperature. This gelatin solution is stable at 4 &#176;C for 1 week. Do not use expired solution and do not freeze-thaw.</li>
	</ol>
	</li>
	<li>Prepare and Fix X. laevis or X. tropicalis Embryos Cultured to Desired Stages.
	<ol>
		<li>Culture the fertilized X. laevis or X. tropicalis embryos in 0.1x MMR with gentamycin until the desired stages. 
		NOTE: Generally, collect embryos between stages 23 and 40. Later collection of embryos after stage 40 is possible, especially when observing axonal growth from the spinal cord, but keep in mind that additional gelatin penetration time may be required. The embryos may be wild-type, transgenic, mutant, inhibitor-treated, Morpholino (MO)-injected, or electroporated 23,24.</li>
		<li>Collect 20-50 embryos in each 4 ml glass vial, remove as much medium as possible and replace with MEMFA. 
		NOTE: If subsequent staining involves phalloidin, use 4% paraformaldehyde instead of MEMFA since commercial supplied formaldehyde solutions usually contain up to 10% methanol as a stabilizer that will interfere with phalloidin staining.</li>
		<li>Fix overnight at 4 &#176;C or, if in an urgency, at room temperature for 2 hr on a rotary mixer. The embryos will be stable in fixation solution for at least 1 week at 4 &#176;C.</li>
		<li>After fixation, wash the embryos 3 times for 20 min using 1x TBS with 0.05% Triton X-100. After the final wash, remove as much TBS-Triton as possible and add 3 ml of 15% gelatin/15% sucrose into each vial.</li>
		<li>Place the vials on a roller bed overnight at room temperature. For embryos older than stage 40, use at least 24 hr of penetration time. After penetration, proceed immediately to section 2.2 on the next day.</li>
	</ol>
	</li>
</ol>
2. Mounting and Cryosectioning of Xenopus Embryos
<ol>
	<li>Prepare Reagents and Materials for Experiments.
	<ol>
		<li>Take one box of positively charged slides, ideally unopened. 
		NOTE: If opened, keep the slides in a dry condition (such as a dry box) and use within 1 month to ensure the static charge on the slides is maintained. Do not use expired slides since samples will fall off during immunostaining.</li>
		<li>Pre-chill the cryostat chamber to -30 &#176;C. Set the instrument parameters as -35 &#176;C for microtome and 12 &#181;m section thickness. Install the thick cover glass plate onto the stage and let the cryostat equilibrate for at least 30 min before section starts.</li>
		<li>Prepare painting brushes for moving section strips, keep inside the cryostat chamber.</li>
		<li>Prepare pencils for writing on slides, put them at room temperature.</li>
		<li>Wear gloves during cryosection. Do not use bare hands.</li>
	</ol>
	</li>
	<li>Mounting Xenopus Embryos
	<ol>
		<li>Carefully aspirate 5-10 embryos out of the glass vial using a plastic or glass pipette without introducing air bubbles. Transfer the embryos into the mounting chamber and observe under a stereoscope. Fill up the mounting chamber with gelatin solution to ensure the rigidity of the section block.</li>
		<li>Arrange the embryos as per Figure 1 using a pair of fine-tip forceps. Mark the orientation of heads by drawing an arrow on the rim of the chamber using a cryogenic-compatible marker. For multiple groups of embryos, write down the description of each group on the rim of the corresponding chamber as well.</li>
		<li>Carefully place the chamber horizontally in a foam box half-filled with dry ice and close the lid. Observe the mounting chamber freeze in 5-10 min. Process each chamber serially (i.e. place the previous chamber onto dry ice before proceeding to the next one) as this will leave sufficient time for each chamber to freeze and prevent the dry ice box to become overcrowded. 
		NOTE: Frozen mounting chambers do not need to stand horizontally and can be stacked up inside the box.</li>
		<li>Proceed with cryosectioning or, if required, keep frozen samples at -80 &#176;C for at least 1-2 week without losing immunogenicity.</li>
	</ol>
	</li>
	<li>Cryosection of Mounted Xenopus Embryos
	<ol>
		<li>Remove a frozen sample block from the chamber by pressing the bottom of the chamber using a blunt stick (such as a pencil).</li>
		<li>Add several drops of tissue-freezing medium, (a polyethylene glycol and polyvinyl alcohol-containing medium) onto the sample holding disc and mount the sample block with the anterior end of the embryos points up (Figure 2). Allow the mounted- block stand inside the cryostat chamber for approximately 1 min or until the tissue-freezing medium becomes opaque.</li>
		<li>Immediately install the sample holding disc onto the microtome with the bottom side of the sample block facing up. Trim off a portion of the sample block using a blade when the sample block is still relatively &#34;soft&#34; to reduce the length of each section (if desired). Leave the sample holding disc on the microtome for at least 5 min to allow its temperature to reach equilibrium.</li>
		<li>Gradually trim the sample block down until the heads of tadpoles are visible through the translucent gelatin. To increase trimming speed, apply a higher (thicker) setting of section thickness (e.g. 20-25 &#181;M) at this stage (such as enabling the &#34;trim&#34; option) but not too thick because sample block can fall off from the sample holding disc. Observe the performance of cryostat, such as the sharpness and angle of the blade at this stage to ensure the generation of a long strip of subsequent sections.</li>
		<li>Once the tadpole heads become visible, adjust the settings of the cryostat back to normal (e.g. 10-12 &#181;M) and clean both the microtome and stage using a paint brush. Gently make 2-3 sections as a trial using the hand wheel setting (do not use the motorized setting) to make sure that finished slices can form strips, not overlapping or sticking to the blade.</li>
		<li>Continue trimming until the heads of tadpoles are almost exposed (this may need some experience but achievable). Brush off any remnant sections on the stage and the collection of sample sections will begin.</li>
		<li>Make approximately 10-15 sections and let them form a long strip.</li>
		<li>Flip over the thick cover glass plate to the side and gently remove the strip from the blade using a fine-tip paint brush and arrange it on the stage with long axis parallel to the blade.</li>
		<li>Pick one positively charged slide at room temperature, label it with a pencil (which will not be washed off during subsequent staining treatments), press it quickly but firmly onto the strip with the label side facing down and remove the slide from the cryostat chamber. If done correctly, the strip should immediately stick to the positively charged slide (Figure 3A).</li>
		<li>Repeat this step to have 20-30 slices on each slide and arranged in parallel (Figure 3B). This is sufficient to cover the entire brain region of X. tropicalis embryos and most of forebrain and midbrain regions of X. laevis embryos.</li>
		<li>Air-dry the slides for 10 min, proceed immediately or, store the slides in a slide box at -80 &#176;C for at least 3-6 months without losing immunogenicity.</li>
	</ol>
	</li>
</ol>
3. Immunostaining of Sectioned Xenopus Embryos
<ol>
	<li>Prepare Reagents and Materials for Experiments.
	<ol>
		<li>Prepare 0.05% TBS-Triton X-100 by adding Triton X-100 into 1x TBS to achieve a final concentration of 0.05%. This solution is stable at room temp for 3-4 days. Do not use expired solution.</li>
		<li>Prepare 5% heat-inactivated goat serum or 5% BSA in TBST as blocking buffer. Firstly, heat-inactivate the goat/lamb serum by placing approximately 20-30 ml of serum in a 65 &#176;C water bath for 30-60 min. Aliquot into 1.5 ml centrifuge tubes and freeze in -20 &#176;C. No re-inactivation is required upon using.</li>
		<li>Dilute the heat-inactivated serum or BSA in TBST to achieve a final concentration of 5% before use.</li>
		<li>Prepare appropriate primary antibodies according to dilutions in blocking buffer (e.g. 1:500 anti-Sox3 10,25/1:250 anti-Myt1 26/anti-acetylated tubulin). Use approximately 100 &#181;l of antibody solution on each slide.</li>
		<li>Prepare appropriate fluorescent antibodies (e.g. anti-mouse/rabbit red fluorescent dye-conjugated antibodies) in blocking buffer (usually 1:500).</li>
		<li>(optional) Add red fluorescent dye-conjugated Phalloidin (1:500) into secondary antibody mix to reveal actin network.</li>
		<li>(optional) Add DAPI (0.5 &#181;g/ml final concentration) into secondary antibody mix to visualize nuclear localization.</li>
		<li>Take the anti-fade mounting medium out of the freezer and thaw in room-temperature water bath.</li>
	</ol>
	</li>
	<li>Immunostaining
	<ol>
		<li>Remove the frozen slides from -80 &#176;C and place them on a piece of towel paper inside a ventilation hood for at least 1 hr to eliminate any condensed water droplets, then bake the dried slides on a 85-90 &#176;C heat block with the slices facing up for 15 min to activate the adhesion mechanism. Finally, allow the slides cool down to room temperature (at least 10 min).</li>
		<li>Fill the staining jar with pure acetone and incubate the slides in acetone for 10 min to remove fish gelatin. If multiple slides are to be treated, arrange them on a staining rack. Air-dry the treated-slides for 15 min in a ventilation hood. Do not re-use the acetone.</li>
		<li>Carefully draw a ring around the samples on the slide using a PAP pen without touching the samples. Ensure that the ring is self-enclosed, otherwise antibody solution will flow out during staining. Fully dry the PAP pen ring.</li>
		<li>Fill another staining jar with 1x TBS without Triton X-100. Insert the dried-slides into the staining jar and allow rehydration for at least 1 hr. In the meantime, prepare the blocking buffer by diluting either heat-inactivated goat serum or BSA to 5% final concentration using 1x TBS with 0.05% Triton X-100.</li>
		<li>Make a wet box by placing 1-2 6-well plates inside a click-lock food box and fill the wells halfway with ultrapure water. Remove rehydrated slides from the staining jar and place them horizontally on the 6-well plates (without the plate lid).</li>
		<li>Carefully add 300-600 &#181;l of blocking buffer inside the PAP ring. Seal the wet box by locking the lid into position and incubate at room temperature for at least 1 hr.</li>
		<li>Dilute the appropriate primary antibodies in blocking buffer. Generally, use 100-150 &#181;l diluted antibody solution per slide. If multiple slides are processed, scale up the volume proportionally.</li>
		<li>After blocking, carefully remove the blocking buffer from slides by aspiration and quickly add primary antibody solution to prevent dry-out, then seal the wet box and incubate at 4 &#176;C overnight.</li>
		<li>On the next day, remove the primary antibody solution from the slides by aspiration. Wash the slides by inserting into staining jars filled with 1x TBS with 0.05% Triton X-100, 3 times, 15 min each. Meanwhile, dilute the appropriate fluorescent secondary antibodies with blocking buffer (with or without DAPI or phalloidin).</li>
		<li>Carefully add 100-150 &#181;l of secondary antibody solution onto the slides. Place the slides inside the wet box and seal the lid. Incubate the slides for 1-2 hr at room temperature.</li>
		<li>Wash the slides in staining jars filled with 1x TBS with 0.05% Triton X-100, 3 times, 15 min each. At the final wash, thaw the anti-fade mounting medium in a 50 &#176;C water bath for 10 min if stored at -20 &#176;C. Cool down the mounting medium to room temperature before use.</li>
		<li>Add approximately 20 &#181;l (one drop) of mounting medium onto the slide and apply a large cover slip (at least 22 mm x 64 mm) over the samples. Observe using fluorescent or confocal microscope within 6 hr post mounting.
		<ol>
			<li>If imaging cannot be done on the same day, seal slides using nail polish and place them inside the wet box at 4 &#176;C overnight. 
			NOTE: The anti-fade mounting medium will gradually get oxidized after a few days (and turn brownish) so it is recommended to take the images as soon as possible.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we demonstrate a convenient and efficient method for visualizing primary neurogenesis in Xenopus embryos. This method permits the assessment of different types of neural cells, including neuronal stem cells and differentiated primary neurons using cell type-specific markers.
The protocol is generally robust with very high level of reproducibility. For embryos that are at pre-hatching stages (i.e. up to stage 28), we recommend manually removing the vitelline membrane prior to fixation as this allows embryos to fully uncurl before fixation. This is especially important when collecting embryos at relatively early stages (before hatching), since bent embryos are extremely difficult to arrange at the desired orientation inside the mounting chamber. We have not experienced a loss of immunogenicity after MEMFA or PFA fixation hence additional antigen retrieval step is not necessary for this protocol. In addition, this immunostaining procedure can be carried out in samples from chromogenic/fluorescent in situ hybridization. In such scenario, it is recommended to skip the protease K treatment step during the in situ hybridization protocol. After chromogenic/fluorescent substrate reaction, samples can be washed with TBS and embedded in fish gelatin solution in a similar way to MEMFA/PFA fixed samples. Sample vials can be protected from light by wrapping the vials in aluminum foil if fluorescent in situ hybridization will be imaged together with immunofluorescent staining later.
In some cases, embryos may shrink excessively after gelatin penetration. This is most likely caused by either insufficient fixation or insufficient TBS-Triton extraction. Ensure that embryos have been fixed in PFA/MEMFA for at least 2-3 hr or preferably overnight at 4 &#176;C. If the problem persists, increase the washing time of TBS-Triton to 3 for 1 hr.
During cryo-sectioning, the rigidity of sample block may vary as different batches of fish gelatin have different properties. This problem can be partially compensated by adjusting the temperature of the microtome. A lower temperature will result in a &#34;harder&#34; sample block however under excessive low temperature the sample block becomes crisp and difficult to section. Some fine-tuning or practice (using mock sample blocks without embryos) before each sectioning may be beneficial prior to use on particularly valuable samples.
A commonly encountered problem during sectioning is the thin sections sometimes tend to stick onto the thick glass plate rather than stay flat on the steel stage. This can be particularly disrupting since it constantly interrupts the continuous sectioning process. Such cases are usually caused by one of two reasons: the section chamber and (especially) the thick glass plate not being cold enough, or excessive static charge on the machine and the operator, especially in dry weather. The former can be solved by turning down the temperature of the section chamber and leaving the glass plate within for extra time (possibly overnight); and the latter by properly grounding the cryostat. Connecting the metal surface of the cryostat to a metal tap or similar water tubing system made of metal can be an alternative way to release the static charges. After each use, the thick glass plate should be washed well with non-corrosive detergent (such as dishwashing liquid), rinsed by ultrapure water, sprayed with pure ethanol, and wrapped in towel paper to protect the surface and edges from damaging.
The antibodies listed in the protocol are generally specific and we rarely encounter nonspecific adsorption or high background signal. It should be noted that, if using heat-inactivated goat/lamb serum as blocking agent, excessive heat-inactivation (as visualized by the formation of fluffy precipitant in the serum) should be avoided as this precipitant is highly attractive to secondary antibodies and may contribute to a source of high background.
It is possible, and sometimes desired, to perform double-staining to visualize different populations of neuronal cells simultaneously. Such double-staining is possible and generally gives satisfactory results with the following combinations: Sox3/N-tub or Myt1/N-tub. However, double-staining of Sox3 and Myt1 is complicated by the fact that the two antibodies are both from rabbit origin. We have tested several antibody direct labeling kits, however no satisfactory results were observed (low signal-to-noise level), possibly due to the lack of signal amplification from secondary antibodies. One possible approach to circumvent this problem would be to raise transgenic lines in Xenopus, as discussed below.
One of the main restrictions of this protocol is that, while this protocol could sufficiently distinguish two main pools of neuronal cells, namely, the Sox3-expressing neural stem cell pool and the Myt1-expressing differentiated neurons, it lacks the ability to reveal different sub-populations of differentiated neurons. Such sub-populations, which including, but are not limited to, primary motor neurons, interneurons, and sensory neurons, are usually characterized by their differentially-expressed marker genes 28-30. As mentioned above, recent advances in the development of multicolor in situ hybridization combined with antibody-based immunofluorescence detection method in Xenopus embryos may fill in the gap to reveal these sub-populations of differentiated neurons for potential investigation 31. Alternatively, as has been demonstrated in mice model, it would also be desired to raise a more comprehensive set of cell type-specific antibodies to distinguish such different cell populations in Xenopus.
It is also worth mentioning that, as an addition to this method, recent advances in optical imaging and image analysis, such as multiphoton microscopy, 3D reconstruction, and segmentation, can also be applied after initial assessments to achieve more comprehensive observations of Xenopus oocytes as well as early embryos, particularly under a live setting 32,33. Therefore, to track proliferation, differentiation, and movement of neuronal cells in live animals, it would be desired to establish one or more transgenic lines that harbor fluorescent proteins driven by cell type-specific promoter to allow live observation of such cell populations in early Xenopus embryos. The establishment of a X. laevis line with neuro-specific &#946;-tubulin promoter driving tauGFP and its applications have provided a nice example 23,34. With the full promoter sequences of both sox3 and myt1 characterized in vertebrates 35-37, it is should be relatively easy to establish additional transgenic lines in Xenopus which should contribute extensively to both the Xenopus community and a more general field of primary neurogenesis research.

In vertebrates, the development of the central nervous system comprises several distinctive yet consecutive stages. The first step is neural induction, when naive ectodermal cells are specified toward a neural fate rather than an epidermal fate. Several interconnected regulatory mechanisms are involved during this stage in Xenopus and other model systems 1,2. This process is primarily coordinated by secreted factors produced by the underlying mesoderm, such as chordin, noggin, and follistatin 3-7. After neural induction, a subset of neural progenitors exit the cell cycle and begin to differentiate in a process referred to as primary neurogenesis. Not all neuronal precursors differentiate at this time. The remaining neural precursor cells continue to proliferate, thereby maintaining the stem cell pool needed for continued growth of central nervous system throughout development and into adulthood.
These proliferating neural precursor cells are characterized by their expression of the SRY (sex determining region Y)-box 3 (sox3) gene 8-11. The other population of cells, which exit the cell cycle and commit to a differentiated fate, are identified by the expression of the differentiation gene markers, tubulin, beta 2B class IIb (tubb2b,N-tub) and myelin transcription factor 1 (myt1) 12-14. Such differentiated neuronal cells eventually give rise to different categories of neurons including, but not limited to, motor, inter-, and sensory neurons positioned in distinct areas within the neural tube 15-17.
While significant efforts have been devoted to uncovering the regulatory mechanisms that govern the patterning and fate determining events in the anterior neuroectoderm, less attention has been made on investigating the neurogenic events that occur following the initial patterning stage. Indeed, signal transduction, transcriptional regulations, as well as post-translational modifications are all involved in this later stage, controlling both timing and lineage specification during neurogenesis 18-20. Further investigations into these mechanisms require a reliable method to easily visualize and distinguish different populations of neuronal cells. The above-mentioned neural markers, including, Sox3, Myt1, and N-tub, can provide a means for identifying these different cell populations, thus providing the necessary foundations for revealing the underlying mechanisms of neuronal differentiation 21-23.
Although differential labeling of neuronal cell populations have been demonstrated in other model organisms, relatively few studies have exploited the Xenopus system to its fullest in this regard. This is mainly due to a paucity of compatible antibodies that reliably identify the various neuronal cell populations in the neural tube. Here, we describe a method for visualizing neuronal differentiation in early Xenopus embryos via immunostaining, which provides a robust and convenient approach for investigating primary neurogenesis in Xenopus. This protocol should give sufficient guidance for researchers interested in the early development of Xenopus central nervous system between stage 26 and stage 45.

<table border="0" cellpadding="0" cellspacing="0" width="573">
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			<td>EMB-200-010K</td>
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			<td>ThermoFisher</td>
			<td>12-550-15</td>
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			<td height="21" style="height:21px;width:216px;">&#160;85-90&#176;C Heat block</td>
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			<td height="42" style="height:42px;">PAP pen</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td>Z377821</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Acetone, analytical grade</td>
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			<td height="21" style="height:21px;">Staining jar with slide racks</td>
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			<td colspan="4" height="21" style="height:21px;">Click-lock food box of appropriate size (has the capacity to hold 1-2 6-well plate inside)</td>
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			<td height="42" style="height:42px;">anti-acetylated tubulin</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td>T7451</td>
			<td></td>
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			<td height="63" style="height:63px;width:216px;">Anti-mouse IgG (H+L), F(ab&#39;)2&#160;Fragment (Alexa Fluor&#174;&#160;488 Conjugate)</td>
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			<td align="right" style="width:119px;">4408</td>
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		<tr height="63">
			<td height="63" style="height:63px;width:216px;">Anti-rabbit IgG (H+L), F(ab&#39;)2&#160;Fragment (Alexa Fluor&#174;&#160;555 Conjugate)</td>
			<td style="width:71px;">Cell Signaling</td>
			<td align="right" style="width:119px;">4413</td>
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			<td height="42" style="height:42px;width:216px;">Alexa Fluor&#174; 647 Phalloidin</td>
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			<td align="right" style="width:119px;">8940</td>
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			<td height="42" style="height:42px;width:216px;">DAPI</td>
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			<td height="63" style="height:63px;width:216px;">ProLong Gold Antifade Mountant</td>
			<td style="width:71px;">Life Technologies</td>
			<td>P36930</td>
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		</tr>
	</tbody>
</table>

assessing primary neurogenesis in xenopus embryos using immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

The representative results show cross-sections of stage 30 Xenopus embryos at different levels, namely the forebrain, midbrain, hindbrain, and spinal cord, stained with different antibodies (Figure 4). As mentioned, Sox3-labeled neuronal progenitor cell population locates in the proximity of the lumen of the neural tube, while MyT1-labeled differentiated primary neurons migrate outward and locate near the marginal layer (basal lamina) of the neural tube. Anti-acetyl-tubulin labels axons in differentiated neurons, which can be observed within the whole neural tube.
Side-specific gene knockdown or overexpression is a broadly used method in neurobiology to evaluate whether modulating the expression of specific gene(s) disrupts the growth and differentiation of neuronal cells. In such cases, either Morpholinos (MOs) or DNA construct(s) carrying promoter-driven gene(s) are injected into one of the two blastomeres at two-cell stage 23. Alternatively, they can be injected into the brain ventricle followed by electroporation, which will result in knockdown or over-expression of desired gene(s) 27. In both cases, the effects will be restricted to one side of the embryo, making the opposite side of the embryo as untreated internal control.
After fixation, sectioning, and immunostaining, the impacts of gene knockdown/over-expression are quantified by counting and comparing the cell numbers of different populations at both sides of embryo. By accumulating these data from several embryos, statistical analysis can be performed. In our representative results, no perturbation was made in the embryos. Examples of gene perturbation and their effects on different neuronal cell populations can be found in the references 20,23.
<img alt="Figure 1" src="/files/ftp_upload/53949/53949fig1.jpg" /> 
Figure 1: Embryo arrangement and orientation in the section mould. (A) A cartoon depiction showing the mounting assembly, note the tray has been labelled with the date, stage, and the orientation of embryos. (B) An image showing the natural appearance of the mounting assembly. <a href="https://www.jove.com/files/ftp_upload/53949/53949fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53949/53949fig2.jpg" /> 
Figure 2: The orientation and position of the gelatin block on the sample holding disc. (A) A cartoon depiction showing the section block assembly, note that embryos are in a &#34;heads up&#34; position and the gelatin block is firmly fixed onto the sample holding disc by the tissue-freezing medium at the base (see Figure 2B). (B) An image showing the natural appearance of the section block assembly, note the accumulation of the tissue-freezing medium between the gelatin block and the sample holding disc. <a href="https://www.jove.com/files/ftp_upload/53949/53949fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53949/53949fig3.jpg" /> 
Figure 3: Continuous sectioning. (A) Model of a section chamber. The sample holding disc should be rotated and fixed to the position that the embryo side is facing up. After a few (6-10) sections, the long strip of continuous section tiles are gently separated from the blade (blue) using a fine brush, and then turned 90&#176; on the holding plate. Then a room-temp positively charged slide is firmly pressed onto the section strip facing down and immediately lifted up to collected finished strips. (B) Normally, each slide can accommodate 2 parallel lines of strips, as shown. Remember to make detailed record on the slide label using pencil. <a href="https://www.jove.com/files/ftp_upload/53949/53949fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53949/53949fig4.jpg" /> 
Figure 4: Cross-section of stage 30 X. laevis tadpoles and staining in the neural tissue, 20X. (A) Schematics indicating the relative positions (forebrain, midbrain, hindbrain, and spinal cord) of section planes on Xenopus tadpoles. (B) Corresponding cross-sections stained with anti-Sox3 (Red), anti-acetylated-tubulin (Green), and DAPI (Blue), showing the relative locations of neural stem cell pools as well as neurofilaments within the neural tube. (C) Corresponding cross-sections stained with anti-MyT1 (Red) and DAPI (Blue), showing the relative locations of differentiated primary neurons within the neural tube. Scale bar = 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/53949/53949fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining at JoVE.com

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

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9.5K Views.  University of Manchester. This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Video: Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages of the fly as an experimental system include its genetic tractability, short life span and the possibility to observe and quantitatively analyze complex behaviors. The expression of disease-linked genes in specific neuronal populations of the Drosophila brain, can be used to model human neurodegenerative diseases such as Parkinson's and Alzheimer's 5.
Dopaminergic (DA) neurons are among the most vulnerable neuronal populations in the aging human brain. In Parkinson's disease (PD), the most common neurodegenerative movement disorder, the accelerated loss of DA neurons leads to a progressive and irreversible decline in locomotor function. In addition to age and exposure to environmental toxins, loss of DA neurons is exacerbated by specific mutations in the coding or promoter regions of several genes. The identification of such PD-associated alleles provides the experimental basis for the use of Drosophila as a model to study neurodegeneration of DA neurons in vivo. For example, the expression of the PD-linked human &alpha;-synuclein gene in Drosophila DA neurons recapitulates some features of the human disease, e.g. progressive loss of DA neurons and declining locomotor function 2. Accordingly, this model has been successfully used to identify potential therapeutic targets in PD 8.
Here we describe two assays that have commonly been used to study age-dependent neurodegeneration of DA neurons in Drosophila: a climbing assay based on the startle-induced negative geotaxis response and tyrosine hydroxylase immunostaining of whole adult brain mounts to monitor the number of DA neurons at different ages. In both cases, in vivo expression of UAS transgenes specifically in DA neurons can be achieved by using a tyrosine hydroxylase (TH) promoter-Gal4 driver line 3, 10.

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Here we describe two assays that have been established to study age-dependent neurodegeneration of dopaminergic (DA) neurons in Drosophila: the climbing/startle-induced negative geotaxis assay which allows to study the functional effects of DA neurons degeneration and the tyrosine hydroxylase immunostaining which is used to identify and count DA neurons in whole brain mounts.

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages of the ...

Functional Interrogation of Adult Hypothalamic Neurogenesis with Focal Radiological Inhibition

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral delivery or transgenic animals have potential confounds that make interpretation of results from these studies difficult. New radiological tools are emerging, however, that allow one to noninvasively investigate the function of select groups of adult-born neurons through accurate and precise anatomical targeting in small animals. Focal ionizing radiation inhibits the birth and differentiation of new neurons, and allows targeting of specific neural progenitor regions. In order to illuminate the potential functional role that adult hypothalamic neurogenesis plays in the regulation of physiological processes, we developed a noninvasive focal irradiation technique to selectively inhibit the birth of adult-born neurons in the hypothalamic median eminence. We describe a method for Computer tomography-guided focal irradiation (CFIR) delivery to enable precise and accurate anatomical targeting in small animals. CFIR uses three-dimensional volumetric image guidance for localization and targeting of the radiation dose, minimizes radiation exposure to nontargeted brain regions, and allows for conformal dose distribution with sharp beam boundaries. This protocol allows one to ask questions regarding the function of adult-born neurons, but also opens areas to questions in areas of radiobiology, tumor biology, and immunology. These radiological tools will facilitate the translation of discoveries at the bench to the bedside.

The function of adult-born mammalian neurons remains an active area of investigation. Ionizing radiation inhibits the birth of new neurons. Using computer tomography-guided focal irradiation (CFIR), three-dimensional anatomical targeting of specific neural progenitor populations can now be used to assess the functional role of adult neurogenesis.

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral ...

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response is a vision-based, reflexive behavior that has been observed in all vertebrates tested. Tadpole eyes are small so the tail flip response was used as alternative measure, which requires a trained technician to record the subtle response. We developed an alternative behavior assay based on the fact that tadpoles prefer to swim on the white side of a tank when placed in a tank with both black and white sides. The assay presented here is an inexpensive, simple alternative that creates a response that is easily measured. The setup consists of a tripod, webcam and nested testing tanks, readily available in most Xenopus laboratories. This article includes a movie showing the behavior of tadpoles, before and after severing the optic nerve. In order to test the function of one eye, we also include representative results of a tadpole in which each eye underwent retinal axotomy on consecutive days. Future studies could develop an automated version of this assay for testing the vision of many tadpoles at once.

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Xenopus laevis tadpoles prefer swimming on the white side of a black/white tank. This behavior is guided by their vision. Based on this behavior, we present a simple assay to test the visual function of tadpoles.

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

The Mouse Hindbrain As a Model for Studying Embryonic Neurogenesis

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded forebrain neuroepithelium is not amenable to wholemount analysis to examine organ-wide neurogenesis patterns. Moreover, defining the mechanisms of forebrain neurogenesis is not necessarily predictive of neurogenesis in other parts of the brain; for example, due to the presence of forebrain-specific progenitor subtypes. The mouse hindbrain provides an alternative model for studying embryonic neurogenesis that is amenable to wholemount analysis, as well as tissue sections to observe the spatiotemporal distribution and behavior of neural progenitors. Moreover, it is easily dissected for other downstream applications, such as cell isolation or molecular biology analysis. As the mouse hindbrain can be readily analyzed in the vast number of cell lineage reporter and mutant mouse strains that have become available, it offers a powerful model for studying the cellular and molecular mechanisms of developmental neurogenesis in a mammalian organism. Here, we present a simple and quick method to use the mouse embryo hindbrain for analyzing mammalian neural progenitor cell (NPC) behavior in wholemount preparations and tissue sections.

This article demonstrates how the mouse embryonic hindbrain can be used as a model for studying developmental neurogenesis in both whole organ and tissue section preparations.

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Rapid and Specific Immunomagnetic Isolation of Mouse Primary Oligodendrocytes

The efficient and robust isolation and culture of primary oligodendrocytes (OLs) is a valuable tool for the in vitro study of the development of oligodendroglia as well as the biology of demyelinating diseases such as multiple sclerosis and Pelizaeus-Merzbacher-like disease (PMLD). Here, we present a simple and efficient selection method for the immunomagnetic isolation of stage three O4+ preoligodendrocytes cells from neonatal mice pups. Since immature OL constitute more than 80% of the rodent-brain white matter at postnatal day 7 (P7) this isolation method not only ensures high cellular yield, but also the specific isolation of OLs already committed to the oligodendroglial lineage, decreasing the possibility of isolating contaminating cells such as astrocytes and other cells from the mouse brain. This method is a modification of the techniques reported previously, and provides oligodendrocyte preparation purity above 80% in about 4 h.

We describe the immunomagnetic isolation of primary mouse oligodendrocytes, which allows the rapid and specific isolation of the cells for in vitro culture.