I thank Alex L. Kolodkin, as I learned this filleted preparation protocol in his laboratory. I also thank Young Gi Hong for technical assistance. &#160;This study was supported by NRF-2013R1A1A4A01011329 (S.J.).

1. Preparation
<ol>
	<li>Prepare 500 mL of phosphate-buffered saline (PBS) with t-Octylphenoxypolyethoxyethanol (PBT) solution by adding 0.5 g of bovine serum albumin (BSA) and 0.5 mL of t-Octylphenoxypolyethoxyethanol (see the Table of Materials) to 500 mL of 1X PBS and stirring for at least 30 min. Store at 4 &#176;C. Use when relatively fresh and store the solution in a clean bottle.</li>
	<li>Make 10 mL of 4% paraformaldehyde by adding 2.5 mL of 16% stock paraformaldehyde solution and 1 mL of 10x PBS to 6.5 mL of deionized water. Store at 4 &#176;C and use within one week. 
	CAUTION: Avoid skin contact with paraformaldehyde solution because it is highly toxic.</li>
	<li>Make X-Gal substrate (10% w/v X-Gal in dimethyl sulfoxide (DMSO)) by dissolving 0.1 g of X-Gal substrate per 1 mL of DMSO. Store at -20 &#176;C in 100 &#181;L aliquots.</li>
	<li>Make X-Gal staining solution using 10 mM phosphate buffer (pH 7.2), 150 mM NaCl, 1 mM MgCl2, 3 mM K4[FeII(CN)6], 3 mM K3[FeIII(CN)6], and 0.3% t-Octylphenoxypolyethoxyethanol. Store at 4 &#176;C in the dark.</li>
	<li>Make 3% hydrogen peroxide solution by diluting 30% stock hydrogen peroxide 1:10 with deionized water.</li>
	<li>Make 3,3&#8242;-diaminobenzidine (DAB) solution by completely dissolving 1 tablet (10 mg) of DAB in 35 mL of fresh PBT solution. Filter through a 0.2-&#181;m syringe filter and store at -20 &#176;C in 910 &#181;L aliquots. 
	CAUTION: Dispose of all DAB waste in bleach to destroy the DAB.</li>
	<li>Make egg plates with apple juice. Add 35 g of agar and 1 L of deionized water to a 2 L flask. Add 33 g of sucrose, 2 g of tegosept, and 375 mL of apple juice to a 1- L flask. Melt them by autoclaving for 30 min. Allow both solutions to cool to around 60 &#176;C. Combine and mix these solutions well by stirring. Pour ~11 mL of combined solution per 60 mm culture dish.</li>
	<li>Make yeast paste by blending baker yeast into deionized water (1:0.8 ratio) and store at 4 &#176;C.</li>
</ol>
2. Embryo Collection (Day 1)
<ol>
	<li>Collect young female and male flies (younger than 5 days), and for 1-2 days, maintain them in a plastic beaker cage with an egg plate (60 mm x 15 mm) that contains a small amount of yeast paste in the center.</li>
	<li>For the optimal collection of late stage-16 embryos, set up a bottle cage with flies (&#62; 50) at around 5 PM and let them lay eggs at 25 &#176;C for 3 h.</li>
	<li>Transfer the flies to a fresh food bottle.</li>
	<li>Close the egg plate with the lid and incubate it upside down at 25 &#176;C in a humidified incubator (~70% humidity) overnight.</li>
</ol>
3. Preparation of Embryos for Immunostaining (Day 2)
<ol>
	<li>Add 1.8 mL of PBT solution to the egg plate around 10:20 AM and use a cotton swab to loosen the embryos from the plate while tilting it. 
	Note: By gently mashing the yeast paste using a cotton swab, dissolve it in case a large number of embryos are found on it. Begin collecting the embryos ~40 min before fixation (around 11:00 AM).</li>
	<li>Transfer the embryos from the egg plate to a 1.5-mL microtube using a 1-mL pipette tip.</li>
	<li>Allow the embryos to settle down and aspirate the PBT solution as much as possible. Rinse the embryos twice with 1 mL of PBT solution. Aspirate the PBT solution as much as possible.</li>
	<li>Fill the tube with 1 mL of 50% bleach (i.e., diluted in deionized water) and dechorionate the embryos by incubating them on a nutator for 3 min at room temperature (RT). 
	Note: This step does not kill the dechorionated embryos.</li>
	<li>Allow the dechorionated embryos to settle down, aspirate the 50% bleach as much as possible, and rinse the embryos 3 times with 1 mL of PBT solution.</li>
	<li>Add 0.5 mL of heptane and subsequently add 0.5 mL of 4% paraformaldehyde solution to the tube at RT at around 11:00 AM.</li>
	<li>Incubate the embryos on a nutator for 15 min at RT.</li>
	<li>Remove the 4% paraformaldehyde solution (the bottom layer).</li>
	<li>Add 0.5 mL of 100% methanol and devitellinize the embryos by shaking the tube vigorously for 30 s.</li>
	<li>Remove the heptane (i.e., the top layer).</li>
	<li>Add 0.5 mL of 100% methanol and shake the tube vigorously for 10 s.</li>
	<li>Allow the embryos to settle down by tapping the tube and aspirate the methanol as much as possible. Rinse the embryos with 1 mL of 100% methanol, shaking for less than 10 s. 
	Note: Extended exposure to methanol destroys &#946;-Gal activity.</li>
	<li>Remove the methanol and rinse the devitellinized embryos 3 times with 1 mL of PBT solution.</li>
</ol>
4. Genotyping the Embryos Using LacZ Staining (Day 2)
<ol>
	<li>Add 1 mL of PBT solution to the tube and incubate the tube in a 37 &#176;C heat block or water bath for 15 min.</li>
	<li>During incubation, place X-Gal staining solution (1 mL per tube) at 65 &#176;C in a water bath until it gets cloudy. Incubate it in a 37 &#176;C water bath.</li>
	<li>Aspirate the PBT solution and add 1 mL of X-Gal staining solution and 20 &#181;L of X-Gal substrate to the tube.</li>
	<li>Incubate the tube at 37 &#176;C with nutation until a blue precipitate is evident. 
	Note: The incubation time for the 2nd and 3rd blue balancers in general ranges from 2-4 h.</li>
	<li>Remove the X-Gal staining solution and rinse the embryos 3 times with 1 mL of RT PBT solution.</li>
	<li>Using a needle probe or forceps, hand-sort the embryos of the desired genotype (i.e., non-stained white embryos) with a light dissecting microscope (1.6X-2.5X objectives). 
	Note: Since some blue balancers, such as CyO, actin-LacZ and TM3, actin-LacZ, carry a transgenic construct expressing bacterial &#946;-Gal (LacZ) under the control of the actin promoter, embryos stained blue in a ubiquitous manner contain one or two copies of blue balancer chromosomes. Therefore, non-stained white embryos are homozygous for the desired lethal allele.</li>
</ol>
5. Immunostaining of Embryos with Anti-Fasciclin II Antibodies (Day 2-3)
<ol>
	<li>Collect and transfer the embryos of the desired genotype (i.e., non-stained white embryos) to a 0.5 mL microtube using a 1 mL pipette tip. 
	Note: In this step, the embryos can be roughly staged according to morphological criteria, including the segmentation patterns of the cuticle and the structures of the head and tail9. During head involution, between 10 and 16 h after the eggs are laid, the embryo undergoes a pronounced reduction of the most anterior segment9.</li>
	<li>Wash the embryos once with 0.4 mL of PBT solution and add 0.3 mL of blocking solution (5% normal goat serum and 5% DMSO in PBT solution).</li>
	<li>Block the embryos with nutation at RT for 15 min.</li>
	<li>Add 75 &#181;L of anti-Fasciclin II antibody and incubate the tube with nutation at RT overnight.</li>
	<li>Wash the embryos 4 times with 0.4 mL of PBT solution for at least 20 min per wash.</li>
	<li>Add 0.3 mL of blocking solution (5% normal goat serum in PBT solution) containing goat anti-mouse-HRP antibody (2 &#181;g/mL) and incubate the tube with nutation at RT overnight.</li>
	<li>Wash the embryos 6 times with 0.4 mL of PBT solution for at least 20 min per wash.</li>
	<li>Add 0.3 mL of DAB solution to the tube. 
	CAUTION: Wear gloves and place all DAB wastes/tubes/tips into bleach.</li>
	<li>Add 2 &#181;L of 3% hydrogen peroxide and incubate the tube in the dark with nutation at RT until the desired amount of precipitate is produced. 
	Note: The incubation time for 1D4 immunohistochemistry in general ranges from 0.5-1 h.</li>
	<li>Wash the embryos 4 times with 0.4 mL of PBT solution. 
	CAUTION: Remove the DAB solution and the first two washes with a pipette and discard them in bleach.</li>
	<li>Add 0.2 mL of mounting 70% glycerol/PBS solution to the tube and store at RT or 4 &#176;C. 
	Note: Keep the tube away from light. Clearer preparations can be obtained by placing the embryos in a 90% glycerol/PBS solution10. However, it is more difficult to dissect embryos cleared in 90% glycerol/PBS solution than those cleared in 70% glycerol/PBS solution10.</li>
</ol>
6. Staging, Dissecting, Mounting, and Imaging the Embryos (Day 4)
<ol>
	<li>For staging, transfer the embryos equilibrated with 70% glycerol/PBS onto a glass slide.</li>
	<li>Collect late stage-16 embryos that have abdominal segment 7 (A7), A8, and A9 ISN nerves converging to touch each other or/and have midgut that is subdivided into three bands. 
	Note: Embryos stained with 1D4 antibody can be staged based on the projection patterns of ISN and ISNb motor axons. After exiting the CNS, A7, A8, and A9 ISN nerves of late stage-16 embryos converge to touch each other and subsequently diverge to extend far out to the dorsal muscles (arrowhead in Figure 1A). In mid-stage-16 embryos, however, A7 ISN nerves extend in parallel with A8/A9 nerves (square bracket in Figure 1B). In addition, the projection axes of A6 ISNb nerves (perpendicular to the ventrolateral muscles) in both hemisegments are roughly aligned with the posterior end of the CNS longitudinal axon fascicles (arrows and a line in Figure 1C). These morphological criteria somewhat vary between different genotypes. Alternatively, embryos can be staged based on gut morphology9,11. Prior to stage 17, the midgut has a heart-like shape, whereas the midgut is subdivided into three bands by stage 1712.</li>
	<li>Dissect the embryos.
	<ol>
		<li>Using a 1 mL syringe, move each embryo out of the glycerol drop and place the embryo ventral-face up. Cut off the anterior part at 1/4 of the body length and the most posterior region that is free of motor axons.</li>
		<li>Using a needle probe, move the end-cut embryo out of the glycerol drop, roll to orient it dorsal-side up, and place it horizontally in the field. 
		Note: When the embryo is moved out of the glycerol drop, a little bit of glycerol associated with the embryo makes it sticky.</li>
		<li>Cut the embryo along the dorsal midline using a single very fine tungsten needle13. Make a small cut at the posterior end of the embryo and then continue to cut down the dorsal midline toward the anterior; cutting in the opposite direction is also fine.</li>
		<li>Using a needle probe, move the midline-cut embryo into the glycerol drop, place it dorsal-side up, and detach the gut from the body wall by unfolding each body wall in a ventrolateral (diagonal) direction (2.5X objective). 
		Note: Make sure that the dorsal midline is fully cut, resulting in the complete separation between the left and right body walls.</li>
		<li>Carefully move the midline-cut embryo out of the glycerol drop and then lay two flaps of the body wall down on the glass slide using a fine needle probe (2.5X objective).</li>
		<li>Using a very fine tungsten needle, remove the internal organs from the dissected embryo by pushing them laterally (5X objective). 
		Note: A more detailed description of another similar dissection procedure can be found on the website5.</li>
	</ol>
	</li>
	<li>For mounting, add 2-3 &#181;L of 70% glycerol/PBS solution to clean, dissected embryos and, using a fine needle probe, transfer them to a new glass slide onto which 8 &#181;L of 70% glycerol/PBS solution has been spread in the center (2.5X objective).</li>
	<li>Put on a coverslip (18 mm x 18 mm). Seal the edges of the coverslip with regular nail polish (either clear or colored is fine).</li>
	<li>Capture images at high resolution (20X, 40X, and 63X oil-immersion objectives) under a differential interference contrast (DIC) light microscope, according to the manufacturer&#39;s instructions. 
	Note: Better visualization and phenotypic characterization, especially for ISNb motor axons, can be produced by using a 63X oil-immersion objective.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+analysis+of+growth+cone+guidance+in+Drosophila:+fasciclin+II+functions+as+a+neuronal+recognition+molecule.">Genetic analysis of growth cone guidance in Drosophila: fasciclin II functions as a neuronal recognition molecule.</a> Cell. 67 (1), 45-57 (1991).</li><li>Vactor, D. V., Sink, H., Fambrough, D., Tsoo, R., Goodman, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genes+that+control+neuromuscular+specificity+in+Drosophila.">Genes that control neuromuscular specificity in Drosophila.</a> Cell. 73 (6), 1137-1153 (1993).</li><li>. 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A., Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The embryonic development of Drosophila melanogaster. , (1985).</li><li>Patel, N. H., Goldstein, L. S. B., Fyrberg, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+neuronal+subsets+and+other+cell+types+in+whole+mount+Drosophila+embryos+and+larvae+using+antibody+probes.">Imaging neuronal subsets and other cell types in whole mount Drosophila embryos and larvae using antibody probes.</a> Methods in cell biology, vol 44. Drosophila melanogaster: practical uses in cell biology. 44, 445-487 (1994).</li><li>Lee, H. K., Wright, A. 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(34), (2009).</li><li>Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+Embryonic+Development.">Stages of Embryonic Development.</a> Atlas of Drosophila. development. , 52 (1993).</li><li>Brady, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+technique+for+making+very+fine,+durable+dissecting+needles+by+sharpening+tungsten+wire+electrolytically.">A simple technique for making very fine, durable dissecting needles by sharpening tungsten wire electrolytically.</a> Bull. World Health Organ. 32 (1), 143-144 (1965).</li><li>Kolodkin, A. 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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plexin+A+is+a+neuronal+semaphorin+receptor+that+controls+axon+guidance.">Plexin A is a neuronal semaphorin receptor that controls axon guidance.</a> Cell. 95 (7), 903-916 (1998).</li><li>Yang, D. S., Roh, S., Jeong, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+axon+guidance+function+of+Rap1+small+GTPase+is+independent+of+PlexA+RasGAP+activity+in+Drosophila.">The axon guidance function of Rap1 small GTPase is independent of PlexA RasGAP activity in Drosophila.</a> Dev. Biol. 418 (2), 258-267 (2016).</li><li>Yu, H. H., Araj, H. H., Ralls, S. A., Kolodkin, A. 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Rev. Neurosci. 13 (9), 605-618 (2012).</li></ol>

The author declares that he has no competing financial interests.

The details of motor axon guidance defects are scored faster and with better accuracy by the filleted preparation of DAB-stained embryos than by laser scanning confocal microscopy of fluorescently labeled ones. Therefore, the filleted preparation of fixed and 1D4-stained embryos is best suited for the functional characterization of guidance cue molecules. Four major classes of guidance cues, including netrins, Slits, semaphorins (Semas), and ephrins, and their cognate receptors have evolutionarily conserved roles across worms, flies, and vertebrates20. Among roundabout (Robo) family molecules that act as receptors for Slits, Drosophila Robo was first identified to mediate repulsive guidance function in the CNS6,21. Semas are a large family of guidance molecules that bind to plexin-containing receptor complexes22. The axon guidance function of Semas was first discovered in the grasshopper CNS14. Subsequently, the class 1 transmembrane semaphorin Sema-1a was extensively characterized in Drosophila. Sema-1a, which functions not only as a ligand for plexin A, but also as a receptor for an unknown ligand, plays an important role in axon-axon repulsion at specific choice points during embryonic neural development15,16,17,18 (Figure 2C).
There are a few common troubleshooting steps that may occur with the current protocol. First, in steps 3.9-3.13, the embryos can be devitellinized with low efficiency if too many or too-small embryos are used in the methanol devitellinization procedure. In this case, it is recommended to use a bed volume of 40-80 &#181;L of fixed embryos for devitellinization at step 3.5. Second, in steps 3.9-3.13, weak X-Gal staining can happen in case of longer exposure to methanol or incomplete heptane removal, since methanol used as a fixative destroys &#946;-Gal activity. In this case, it is best that the devitellinization procedure using methanol in steps 3.9-3.13 is performed as quickly as possible. In addition, residual heptane that is solubilized in methanol may interfere with the staining; therefore, it is best to remove as much methanol as possible in steps 3.11-3.13.
The current filleted preparation protocol can be applied to loss-of-function and gain-of-function genetic screens to identify novel genes that are involved in motor axon pathfinding and target recognition. However, a disadvantage of this protocol is that it is quite time-consuming to dissect fixed embryos to generate filleted preparations. To overcome this problem, phenotypic characterization of the axonal projections can be monitored in a whole-mount preparation of 1D4-stained embryos, although this method provides low-resolution visualization of the axonal projections in the periphery. Nevertheless, the whole-mount preparation protocol has proven to be an efficient approach for screening axon guidance genes4. An alternative protocol using the live dissection of fluorescently labeled embryos was recently developed for systematic screening11. The live dissection protocol allows for relatively rapid preparation for phenotypic analysis11. However, this protocol is not adequate for embryos older than the beginning of stage 1711.
The filleted preparation protocol described here can be used to examine protein expression patterns with different antibodies. In addition, this protocol can also be applied to Drosophila embryos at various stages of development.

Precise and selective connections between motor axons and target muscles during embryonic development are essential for normal locomotion in Drosophila larvae. The embryonic patterning of 30 muscle fibers in each of the abdominal hemisegments A2-A7 is established by stage 161. The 36 motor neurons that are generated in the ventral nerve cord extend their axons into the periphery to innervate specific target muscles2. Motor axon pathfinding and target recognition can be visualized by immunohistochemistry with an antibody (mouse monoclonal antibody 1D4)3,4. Multiple images of the motor axon projection patterns in wildtype embryos are available on the web5. The 1D4 antibody labels all motor axons and three longitudinal axon fascicles on each side of the midline of the embryonic central nervous system (CNS)4,6 (Figure 1C and Figure 2A). Therefore, immunohistochemistry with FasII antibody provides a powerful tool for identifying genes required for neuromuscular connectivity for demonstrating the molecular mechanisms underlying motor axon guidance and target recognition.
In each of the abdominal hemisegments A2-A7, motor axons project and selectively fasciculate into two principal nerve branches, the segmental nerve (SN) and the intersegmental nerve (ISN)2,4, and a minor nerve branch, the transverse nerve (TN)7. The SN selectively defasciculates to give rise to two nerve branches called the SNa and SNc, whereas the ISN splits into three nerve branches called the ISN, ISNb, and ISNd2,4. Among them, ISN, ISNb, and SNa motor axon projection patterns are most precisely visualized when late stage-16 embryos are stained with FasII antibody and are filleted (Figure 1C and Figure 2A). The ISN motor neurons extend their axons to innervate dorsal muscles 1, 2, 3, 4, 9, 10, 11, 18, 19, and 202,4 (Figure 2A). The ISNb motor neurons innervate ventrolateral muscles 6, 7, 12, 13, 14, 28, and 302,4 (Figure 2A and 2B). The SNa nerve branch projects to innervate lateral muscles 5, 8, 21, 22, 23, and 242,4 (Figure 2A). The TN, which consists of two motor axons, projects ipsilaterally along the segmental border to innervate muscle 25 and makes synapses with the lateral bipolar dendritic neuron (LBD) in the periphery7 (Figure 2A). These target muscle innervations require not only selective defasciculation of motor axons at specific choice points, but also target muscle recognition. In addition, some putative mesodermal guidepost cells that act as intermediate targets were found in both the ISN and SNa pathways, but not along the ISNb pathway4. This might suggest that ISNb motor axon pathfinding can be regulated in a distinct manner compared to ISN and SNa motor axon guidance, and it also indicates that peripheral motor axon guidance provides an attractive experimental model to study the differential or conserved roles of a single guidance cue molecule8.
This work presents a standard method to visualize the axonal projection patterns of embryonic motor neurons in Drosophila. The described protocols include how to dissect fixed embryos stained with 1D4 antibody and processed in 3,3&#8242;-diaminobenzidine (DAB) for filleted preparations. One critical advantage of the flat preparations of fixed embryos is the better visualization of the axonal projections and muscle patterns in the periphery. Furthermore, this work also shows how to genotype fixed embryos to sort the desired mutant embryos using the LacZ staining method.

<table border="0" cellpadding="0" cellspacing="0" width="786">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:284px;">Bovine Serum Albumin</td>
			<td style="width:144px;">Sigma-Aldrich</td>
			<td style="width:131px;">A7906</td>
			<td style="width:228px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:284px;">Triton X-100</td>
			<td style="width:144px;">Sigma-Aldrich</td>
			<td style="width:131px;">X100</td>
			<td style="width:228px;">t-Octylphenoxypolyethoxyethanol</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:284px;">16% Paraformaldehyde Solution</td>
			<td style="width:144px;">Ted Pella</td>
			<td style="width:131px;">18505</td>
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			<td height="63" style="height:63px;width:284px;">Anti-FasciculinII Antibody</td>
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			<td style="width:131px;">1D4 anti-Fasciclin II</td>
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			<td style="width:144px;">Ted Pella</td>
			<td style="width:131px;">#27-11</td>
			<td style="width:228px;">Tungsten Wire, &#248;0.13mm/6.1m (&#248;.005&#34;/20 ft.)</td>
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			<td height="42" style="height:42px;width:284px;">Nutator (Mini twister)</td>
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			<td style="width:131px;">KO.VS-96TWS</td>
			<td style="width:228px;">Alternatively, BD Clay Adams Brand Nutator (BD 421125)</td>
		</tr>
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</table>

visualization of the axonal projection pattern of embryonic motor neurons in drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Precise connections between motor axons and target muscles during neural development depend on selective axon-axon repulsion and target recognition at specific choice points4. In Drosophila, selective repulsion between motor axons is in part regulated by the combined action of class 1 and 2 semaphorins (Semas), including Sema-1a, Sema-2a, and Sema-2b8,14,15,16,17,18. Therefore, loss of Sema-1a function frequently results in the failure of ISNb axons to defasciculate at specific choice points, causing them to exhibit an abnormally thick morphology (arrowhead in Figure 2C). In wildtype embryos, at least 7 motor neurons extend and fasciculate their axons to form an ISNb nerve branch (Figure 2A). When the ISNb nerve branch reaches a choice point, which is located between muscles 6 and 7, two axons selectively defasciculate from the main ISNb bundle and subsequently innervate muscles 6 and 7 (Figure 2B). At the next choice point, which is located between muscles 6, 13, and 30, another three motor axons selectively defasciculate to innervate muscles 13, 14, and 30 (Figure 2B). The remaining two axons extend dorsally to reach the last choice point near the edge of muscle 12, thereby innervating muscle 12 in the opposite direction (Figure 2B). These results clearly show that the filleted preparation protocol provides a useful tool to study the genes required for motor axon guidance.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55830/55830fig1.jpg" /> 
Figure 1: Staging of embryos based on the projection patterns of ISN and ISNb motor axons. (A) In late stage-16 wildtype embryos, A7, A8, and A9 ISN nerves (arrows) converge to touch each other (arrowhead). Anterior is left and ventral is down. The Scale bar = 15 &#181;m. (B) In mid-stage-16 wildtype embryos, the A7 ISN nerve extends in parallel with A8/A9 nerves (square bracket). Arrows indicate A7, A8, and A9 ISN nerves. Anterior is left and ventral is down. The Scale bar = 15 &#181;m. (C) Filleted preparation of late stage-16 wildtype embryos stained with FasII antibody. The projection axes (arrows) of the left and right A6 ISNb nerves are roughly aligned with the posterior end of the CNS longitudinal axon fascicles (black line). Abdominal segments A1-A7 are labelled at the top. Anterior is left. The Scale bar = 15 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55830/55830fig1large.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/55830/55830fig2.jpg" /> 
Figure 2: Motor axon projection patterns in wild-type and Sema-1a mutant embryos. (A-C) Filleted preparation of late stage-16 embryos stained with FasII antibody. Anterior is left and ventral is down. Schematic diagrams showing the axonal projection patterns represented in each panel. The Scale bars = 15 &#181;m. (A) A bright field photograph of two sequential abdominal hemisegments, A3 and A4, in wildtype embryos. Three bilaterally symmetric longitudinal axon bundles are present in the CNS. The normal innervation patterns of ISNb, SNa, ISN, and TN motor axons are indicated by the boxes and an arrow in the periphery. The lateral bipolar dendrite neuron (LBD), which innervates the alary muscles, is indicated by an arrowhead. In the schematic diagram, the nerve branches and innervation patterns of the ISNb (in brown), SNa (in blue), ISN (in green), and TN (in yellow) motor neurons and their target muscles are presented. The cell body positions of these motor neurons in the CNS are also shown in their own colors. (B) In wildtype embryos, the ISNb axons project to the ventrolateral muscles, which are numbered, and innervate them at specific choice points (arrows in the DIC image). (C) In Sema-1a loss-of-function mutants, ISNb axons frequently fail to defasciculate from one another between muscles 6 and 13 (arrowhead in the DIC image). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55830/55830fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila at JoVE.com

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

visualization-axonal-projection-pattern-embryonic-motor-neurons

Nanyang Technological University

Research

JoVE Journal

Neuroscience

7.7K Views.  Chonbuk National University. This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

Video: Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

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

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

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

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

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

Mechanical Manipulation of Neurons to Control Axonal Development

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the 10-1000 &mu;dyne range1,2. Force measurements are obtained through observation of the Hookean bending of the glass needles, which are calibrated by a direct and empirical method3. Equipment requirements and procedures for fabricating, calibrating, treating, and using the needles on cells are fully described. The force regimes previously used and different cell types to which these techniques have been applied demonstrate the flexibility of the methodology and are given as examples for future investigation4-6. The technical advantages are the continuous 'visualization' of the forces produced by the manipulations and the ability to directly intervene in a variety of cellular events. These include direct stimulation and regulation of axonal growth and retraction7; as well as detachment and mechanical measurements on any type of cultured cell8.

Application and direct measurements of forces on neurons in the 2-1000 microdyne range are achieved with high precision using calibrated glass needles. This methodology can be used to control and measure several aspects of axonal development, including axonal initiation, axonal tension, velocity of axonal elongation, and force vectors.

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the ...

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

The Swimmeret System of Crayfish: A Practical Guide for the Dissection of the Nerve Cord and Extracellular Recordings of the Motor Pattern

Here we demonstrate the dissection of the crayfish abdominal nerve cord. The preparation comprises the last two thoracic ganglia (T4, T5) and the chain of abdominal ganglia (A1 to A6). This chain of ganglia includes the part of the central nervous system (CNS) that drives coordinated locomotion of the pleopods (swimmerets): the swimmeret system. It is known for over five decades that in crayfish each swimmeret is driven by its own independent pattern generating kernel that generates rhythmic alternating activity 1-3. The motor neurons innervating the musculature of each swimmeret comprise two anatomically and functionally distinct populations 4. One is responsible for the retraction (power stroke, PS) of the swimmeret. The other drives the protraction (return stroke, RS) of the swimmeret. Motor neurons of the swimmeret system are able to produce spontaneously a fictive motor pattern, which is identical to the pattern recorded in vivo 1.
The aim of this report is to introduce an interesting and convenient model system for studying rhythm generating networks and coordination of independent microcircuits for students&#8217; practical laboratory courses. The protocol provided includes step-by-step instructions for the dissection of the crayfish&#8217;s abdominal nerve cord, pinning of the isolated chain of ganglia, desheathing the ganglia and recording the swimmerets fictive motor pattern extracellularly from the isolated nervous system.
Additionally, we can monitor the activity of swimmeret neurons recorded intracellularly from dendrites. Here we also describe briefly these techniques and provide some examples. Furthermore, the morphology of swimmeret neurons can be assessed using various staining techniques. Here we provide examples of intracellular (by iontophoresis) dye filled neurons and backfills of pools of swimmeret motor neurons. In our lab we use this preparation to study basic functions of fictive locomotion, the effect of sensory feedback on the activity of the CNS, and coordination between microcircuits on a cellular level.

Here we describe the dissection of the crayfish abdominal nerve cord. We also demonstrate an electrophysiological technique to record fictive locomotion from swimmeret motor neurons.

Here we demonstrate the dissection of the crayfish abdominal nerve cord. The preparation comprises the last two thoracic ganglia (T4, T5) and the chain of ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...