We thank Jos Burgert for helping with some of the figures. We are grateful to Ingo Selbach (and the group &#8220;Edelkrebsprojekt NRW&#8221;) for his efforts to supply the lab with experimental animals. We thank Anna C. Schneider for proofreading first versions of the manuscript. This research was supported by an Emmy Noether DFG grant SM 206/3-1 and a startup grant of the University of Cologne for female faculty.

This dissection procedure is in accordance with the European Communities Council Directive of 22nd September 2010 (2010/63/EU).
1. Preparation
<ol>
	<li>Obtain crayfish, Pacifastacus leniusculus (Dana), of both sexes &#8805;8 cm in size. Ensure that the animals are vital and the abdomen and abdominal limbs are intact.</li>
	<li>Take care to inspect the carapace and that this cuticle is hard and rigid. Pre- and postmolt animals have a soft carapace and are not suited for experiments because during the molting process many parameters change (e.g., decrease in locomotor activity).</li>
	<li>Assemble all tools and materials used during the dissection, pinning and desheathing of the nerve cord shown in Figure 2 and listed in the supplements provided.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/52109/52109fig2highres.jpg" /> 
Figure 2: Materials and tools used for the dissection, pinning and desheathing of the nerve cord.
(1) big bucket filled with ice; (2) crayfish saline; (3) saline dispenser; (4) dissection microscope; (5) dissection dish; (6) strong scissors; (7) forceps (8) spring scissors; (9) Petri dish lined with clear sylgard; (10) fixing pins; (11) cold lamp source.
2. Gross Dissection
<ol>
	<li>Anesthetize animal on ice for 15 - 20 min. Carry out the first part of the gross dissection at a lab bench near the sink since it includes an exsanguination step and the specimen needs to be rinsed regularly with crayfish saline during the procedure.</li>
	<li>Hold animal ventral side up and use strong scissors to cut both claws at their bases near the thorax (Figure 3-1). Remove left and right uropod (Figure 3-2).</li>
	<li>Place the animal ventral side up in the dissection dish lined with black sylgard. Elevate cephalothorax by inserting ice underneath and pin the abdomen at the telson (Figure 4A).</li>
	<li>Fill the saline dispenser with ~60 ml chilled crayfish saline. Perfuse crayfish with cold saline through the claw opening (Figure 4B). Excess saline will drain off through the cuts at the uropods. Cover crayfish with ice during exsanguination</li>
	<li>Decapitate the animal with a single transverse cut just posterior to the animal&#8217;s eyes using strong scissors (Figure 5A). Remove all walking legs near the base joints as indicated in Figure 5B.</li>
	<li>Isolate the abdomen with the last thoracic segments from the rest of the cephalothorax. Make a first cut at the level of the second walking legs (thoracic segment 3) by inserting the tip of the scissors into the opening of the second walking leg and cutting to the opposite side. (Figure 6A-1).</li>
	<li>Extend this first cut to both sides through the cephalothorax (Figure 6A-2).</li>
	<li>Flip the animal open to make some of the internal organs visible. Push the prominent digestive gland (Figure 6B-3) to the anterior part of the specimen and use forceps to remove the reproductive organs from the abdominal cavity.</li>
	<li>Remove the anterior part of the cephalothorax (Figure 6C). Use lateral cuts to remove the lateral parts of the carapace, which cover the gills, on both sides of the remaining thorax (Figure 6D-4). Remove the gills and rinse the specimen with cold saline.</li>
	<li>Continue the dissection with a cut through the whole length of the sternal plate as indicated in Figure 6E-5. Make this cut at maximal lateral positions between pleuron and swimmerets (Figure 6E red markings). Proceed on the other side with a same cut. Rinse the specimen with cold saline.</li>
	<li>Carry out the remaining dissection under a dissection microscope. Place the crayfish&#8217;s abdomen ventral side up in the dissection dish lined with black sylgard and filled with crayfish saline so that it covers the specimen. 
	NOTE: The following steps (2.12&#8211;4.8) contain directional instructions that apply to right handed experimenters. In the following steps (2.12-6.8) it is important to replace crayfish saline in regular intervals, every 20-30 min with cold saline, to keep the nervous system healthy.</li>
	<li>Fix the specimen with insect pins posteriorly at the telson and anteriorly at the remains of the carapace. Place the specimen so that the telson points to the left and is parallel to the table&#8217;s edge.</li>
	<li>Use coarse forceps in the left hand to grab through a walking leg opening (Figure 7A) and pull the specimen open (Figure 7B white arrow). Identify the large two dorsal flexor muscle strands (Figure 7B-1 and C-1) and cut their ventral basis as shown in Figure 7B.</li>
	<li>Identify the sternal artery (Figure 7C-2), that descends from dorsal (the heart) to ventral, at the 4th thoracic segment. This artery lies right above the nerve cord (Figure 7C-3), before it projects under the nerve cord, forming the ventral artery.</li>
	<li>Transect the sternal artery. As demonstrated in Figure 7C, lift the artery first, using one blade of the scissors, and only cut when the ventrally located nerve cord is visible.</li>
	<li>Fix the dorsal flexor muscles anteriorly (to the right side). This should be done in maximum stretched position with pins so they will not block vision and the specimen remains stretched. When the dorsal muscle strands are fixed (Figure 8-1), the first abdominal ganglia, A1 and A2, with associated nerves N1, N2, and N3 are visible (Figure 8).</li>
	<li>Use forceps in the left hand to grab the specimen at one opening of the walking legs. Throughout the following dissection steps pull gently to keep the specimen open.</li>
	<li>Transect the nerves N3 at the most distant position from the nerve cord. (Figure 8-3).</li>
	<li>Cut the flexor muscles close to the ventral apodeme as shown in Figure 8-4. Take care not to damage the nerve cord or the nerves N2.</li>
	<li>Repeat the steps 2.18 and 2.19 for the nerves N3 and the flexor muscles of the remaining abdominal ganglia A2 to A5.</li>
	<li>At the last abdominal ganglion, A6, cut the dorsal flexor muscles from the ventral apodeme and the specimen should look as shown in Figure 9.</li>
	<li>Cut the sternal plate posterior to the nerves of A6 (Figure 9-1) and keep the ventral part (Figure 9-2). Discard the dorsal part with flexor muscles (Figure 9-3). Fix the sternal plate anteriorly with pins through the openings of the walking legs, and posteriorly to A6.</li>
</ol>
3. Fine Dissection
<ol>
	<li>Place the specimen under the microscope with the anterior part directed away and the posterior part towards the tables&#8217; edge.</li>
	<li>Use forceps as shown in Figure 10A to remove the most anterior parts of the cephalothoracic sterna. 
	NOTE: The thoracic ganglia and associated nerves are partly covered by the leg musculature and cephalothoracic sterna. The cephalothoracic sterna form a skeleton that separates the laterally located cavities of the walking legs from each other and from the medial cavity in which the ventral nerve cord resides.</li>
	<li>Cut the muscles between the remaining exoskeletal structures as indicated in Figure 10B-1 and B2. Use forceps to grab and lift the anterior end of the ventral nerve cord (Figure 10C). 
	NOTE: The nerve cord will be damaged in the process so avoid picking up the nerve cord multiple times.</li>
	<li>Cut the thoracic nerves laterally while lifting the nerve cord (Figure 10C-3). Keep these nerves at suitable length for pinning. Remove the squeezed part of the chain of ganglia, which was picked up with forceps, by cutting away all the tissue anterior to T4 (Figure 10C-4).</li>
	<li>Place the specimen with the anterior part to the left and focus on A1. Cut the nerves N1 and N2 of A1 at a suitable length (max. 1 cm) for pinning them out.</li>
	<li>Focus on A2 and identify the nerves N1, N2, and N3 of this segment (Figure 11). The nerves N1 of abdominal ganglia A2-A5 reside between two sternal cuticular infoldings in each segment (Figure 11A-1) and are covered by musculature. Make one cut along the posterior sternal cuticular infolding. Start at the lateral rim of the abdomen and proceed towards the midline as shown in Figure 11A.</li>
	<li>If the target N1 is still covered with tissue as shown in Figure 11B (red arrow), cut across the muscle bundle, but this time anterior to both sternal cuticular infoldings and the nerve N1 (Figure 11B-2).</li>
	<li>Cut nerve N1 as distally as possible (Figure 11C-3). Nerve N1 is fully visible and the anterior and posterior branch can be identified (Figure 11C).</li>
	<li>Proceed to the contralateral nerve N1 and first cut the muscles along the posterior sternal cuticular infolding, starting medially, near the ganglion (Figure 11D). If the nerve is still covered by tissue, cut across the muscle bundle, but this time anterior to both sternal cuticular infoldings and the nerve N1, similar to Figure 11B-2. Cut the nerve N1 as distally as possible.</li>
	<li>Cut the nerves N2 of this ganglion to a suitable length (approx. 0.5 cm) for pinning.</li>
	<li>Repeat steps 3.7-3.11 for the nerves of A3-A5.</li>
	<li>Cut the nerves of ganglion A6 as distally as possible (Figure 12A). Use forceps to grab multiple nerves of A6 to lift this ganglion and start isolating the chain of ganglia from the sternal plate.</li>
	<li>While lifting the nerve cord, pull it gently in the anterior direction as demonstrated in Figure 12B (white arrow). As the individual ganglia are lifted, remove the ventral artery that may be attached to the ventral side of the nerve cord (Figure 12C). Continue this (gently) pull-cut sequence until the nerve cord is completely isolated.</li>
	<li>Transfer the isolated chain of ganglia to a Petri dish lined with clear sylgard and filled with crayfish saline (Figure 12D).</li>
</ol>
4. Pinning the Nerve Cord into the Petri Dish
NOTE: Use small pins cut from stainless steel wire (see supplements) to pin the nerve cord. Touch only the nerve ending with the forceps and do not squeeze the connectives or ganglia.
<ol>
	<li>Pin the chain of ganglia in a straight line, while applying gentle stretch.</li>
	<li>Arrange the nerve cord in the Petri dish with the dorsal side facing upwards (Figure 13, black line). The ventral side of the ganglia can be identified by its convexity; the dorsal side is flat. Pin the thoracic nerves to the sides. Continue with the nerves of A6, stretching the nerve cord along its longitudinal axis.</li>
	<li>Pin out the nerves of A1 at a 90&#176; angle relative to the nerve cord.</li>
	<li>Proceed to A2 and pin the nerves N2 at an angle of 35&#8211;45&#176; relative to the nerve cord (Figure 1A).</li>
	<li>Separate the nerves N1 in their anterior and posterior branches before pinning as demonstrated in Figure 14. Use two pairs of fine forceps to pick up with one pair of forceps the anterior and with the other the posterior branch of the nerve N1. Take care to pick only the most distal ends of the nerve branches. Now pull them carefully apart.</li>
	<li>Pin the anterior branch of the nerve N1 at a 90&#176; angle relative to the nerve cord (Figure 1A). Pin the posterior branch of nerve N1 between the anterior N1 branch and the nerve N2.</li>
	<li>Repeat the steps 4.4&#8211;4.6 for the nerves of ganglia A3&#8211;A5. While fixating the nerve cord stretch it in longitudinal as well as transversal directions.</li>
</ol>
5. Desheathing the Ganglia
<ol>
	<li>Place the preparation in such a manner that the experimenter&#8217;s hands are always resting on a stable plane to avoid shaking. In order to desheath the ganglia illuminate the nerve cord from below.</li>
	<li>Focus on any abdominal ganglion A1 to A5. Use fine spring scissors to make a small lateral cut through the ganglion sheath, posterior to the ganglion and between the nerves N2 and N3 (Figure 15A red arrow).</li>
	<li>Pick up the ganglion sheath using very fine forceps and cut transversely across the sheath above the connectives, as indicated in Figure 15A-1. Take care not to squeeze or cut the nerve cord with the scissors.</li>
	<li>Still holding and lifting the ganglion sheath with forceps continue to cut it along the lateral borders of the ganglion (Figure 15B-2 and -3). Remove the sheath. Alternatively pin it to both sides of the connectives in such a way that it is fixed but the nerve cord is not squeezed.</li>
	<li>Repeat steps 5.2-5.4 for all abdominal ganglia A1 to A5.</li>
	<li>Desheathe the thoracic ganglia and the ganglion A6 in a similar fashion. In order to desheathe A6 start anterior to the ganglion and proceed in posterior direction. Pin the ganglion sheath of A6 to the posterior end of the chain of ganglia.</li>
</ol>
6. Extracellular Recordings from Motor Neurons
<ol>
	<li>Assemble all tools and materials used for extracellular recordings shown in Figure 16B and listed in the supplements. An overview of the recording setup is shown in Figure 16A. Start all electronic equipment used in this experiment (Figure 16C), so that the amplifiers can warm up for at least 30 min prior to recording. Turn on the computer and start the recording software.</li>
	<li>Place the chain of ganglia on the microscope table and illuminate from below. Insert the recording electrode into the sylgard close to the target nerve and the reference electrode at a position nearby, but lateral to the ganglia (Figure 17A and B). Bend the target nerve around the recording electrode (Figure 17C).</li>
	<li>Stretch the nerve slightly, to ensure contact between the electrode and the nerve and pin it to the side (Figure 17D). Fix the electrode cables to the microscope table using modeling clay, so they stay in the desired position.</li>
	<li>Use a syringe filled with petroleum jelly and a 20 gauge needle (with rounded tip) (Figure 17E-3) to isolate the target nerve from the bathing solution. First dab some petroleum jelly on the sylgard around the recording electrode. The result is a layer of petroleum jelly covering the sylgard in the proximity of the recording electrode (Figure 17E-4). Take care not to directly dab onto the nerve and avoid air bubbles in this layer.</li>
	<li>Seal the recording electrode with petroleum jelly from all sides up to the surface level of the saline (Figure 17F).</li>
	<li>Repeat this procedure for all target nerves whose activity should be monitored.</li>
	<li>Start recording. Use a continuous or gap free acquisition mode and a sampling rate of 5 kHz. Set the extracellular amplifier to the following parameters; gain to 1,000 (amplifies the signal 1,000 times, take care to include this amplification parameter in the acquisition software settings) and a bandpass filter range of 300 Hz (low cut) to 2,000 Hz (high cut).</li>
</ol>

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The authors declare that they have no competing financial interests.

The anatomy of crayfish and their abdominal ganglia has been described previously 5, 18, 19, 20 and it is recommended to become familiar with them prior to the dissection in order to avoid cutting of important nerves.
It is critical to keep the preparation at temperatures below 23 &#176;C to prevent degradation of the isolated nerve cord. This can be achieved easily by replacement of the bathing solution every 20&#8211;30 min with cold crayfish saline. Under these circumstances the ...

The swimmerets of crayfish serve a function in posture control and beat rhythmically when the animals swim forward, ventilate their burrows or females aerate their eggs 5, 6. The swimmerets of the signal crayfish, Pacifastacus leniusculus, occur in pairs from the second to the fifth abdominal segment, with one limb on each side of the abdomen 7. The central nervous system produces on its own the rhythmic motor patter which drives the swimmeret movement in the intact animal as well as in the isolated nerve cord preparation. When there is no sensory feedback or descending input present the rhythmic motor pattern produced is called fictive locomotion 1, 2. In the swimmeret system this motor pattern does not differ in any parameter from the activity of the swimmerets measured in the intact animal.
The movement of each swimmeret is driven by a microcircuit that is located in and restricted to one corresponding hemiganglion 1 - 3. In each microcircuit there is a pattern generating kernel that comprises five identified non spiking interneurons. They can be functionally characterized as being either Inhibitor of Power Stroke (IPS) or Inhibitor of Return Stroke (IRS) 8. These IPS and IRS interneurons are not endogenous oscillators, rather their alternating activity is driven by reciprocal inhibition 9. Because these interneurons inhibit the swimmeret motor neurons directly, the alternating PS-RS movement is generated 10. Locomotion however, does not only require the generation of activity, but also coordination of the different independent microcircuits. In the swimmeret system such coordination is established by the coordinating microcircuit which ensures that limbs are active at correct times. This microcircuit is built by three identified neurons in each segment 11-15.
This protocol provides for the first time a step-by-step dissection guide to isolate the chain of ganglia (T4 to A6, Figure 1). We show how to pin the isolated abdominal nerve cord and desheathe each ganglion. In this isolated nervous system preparation, the neurons responsible for swimmeret movement are ready for use in electrophysiological and morphological experiments. The second part of this protocol demonstrates the main features of the swimmeret motor pattern. This includes a step-by-step guide to extracellularly record the activity of swimmeret motor neurons. Axons of RS motor neurons project through the anterior branch of nerve N1, while axons of PS motor neurons project through the posterior branch of the same nerve (Figure 1) 4. Therefore their activity can be recorded from these branches with differential pin electrodes.
<img alt="Figure 1" fo:content-width="2in" src="/files/ftp_upload/52109/52109fig1highres.jpg" width="200" /> 
Figure 1: Isolated nervous system from thoracic ganglion 4 (T4) to abdominal ganglion 6 (A6) and a schematic diagram of it. T4: thoracic ganglion 4; T5: thoracic ganglion 5; A1, A2 &#8230; A6 abdominal ganglion 1, abdominal ganglion 2 &#8230; abdominal ganglion 6; N1: nerve N1; N2: nerve N2; N3: nerve N3; PS: power-stroke; RS: return-stroke. Directional abbreviations: A = anterior; P = posterior.
This dissection procedure and the electrophysiological technique demonstrated are convenient for undergraduate students and may complement student practical courses in physiology. The isolated chain of ganglia has been used in a number of experiments to study nervous system function, coordination, or modulation of swimmeret microcircuits 6 as well as neuronal control of adaptive behavior in locomotion 16, 17. The crayfish swimmeret system thus provides an enormous amount of interesting teaching or training opportunities which all begin with the dissection of the ventral nerve cord of crayfish and extracellular recording of the fictive motor pattern.

<table border="0" cellpadding="0" cellspacing="0" width="606">
	<tbody>
		<tr height="38">
			<td height="38" style="height:38px;width:131px;">Name of Material/ Equipment</td>
			<td style="width:131px;">Type</td>
			<td style="width:139px;">Company</td>
			<td style="width:103px;">Catalog Number</td>
			<td style="width:103px;">Comments/&#160;&#160; Description</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">4-channel extracellular amplifier: MA 102&#160;</td>
			<td style="width:131px;">Amplifier</td>
			<td style="width:139px;">Elektroniklabor, Zoologie, Universit&#228;t zu K&#246;ln, Germany</td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="160">
			<td height="160" style="height:160px;width:131px;">air-table</td>
			<td style="width:131px;"></td>
			<td style="width:139px;">Technical Manufacturing Corporation 
			(TMC) a unit of AMETEK Ultra Precision Technologies, Peabody, MA, USA</td>
			<td style="width:103px;">63-534</td>
			<td style="width:103px;">for intracellular recording</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:131px;">Axon Digidata 1440A</td>
			<td style="width:131px;">Digitizer</td>
			<td style="width:139px;">Axon Instruments, Molecular Devices Design, Union City, CA</td>
			<td style="width:103px;">DD1440A</td>
			<td style="width:103px;">digitizes recorded signals&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:131px;">big bucket&#160;</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">filled with ice</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">Clampex &#38; Clampfit</td>
			<td style="width:131px;">pClamp 10, recording and analysis software</td>
			<td style="width:139px;">Molecular Devices Design, Union City, CA</td>
			<td style="width:103px;">pClamps 10 Standard</td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">cold lamp source</td>
			<td style="width:131px;">with flexible light guide (fiber optic bundle)&#160;</td>
			<td style="width:139px;">Euromex microscopes holland, Arnhem, BD</td>
			<td style="width:103px;">LE.5211 &#38; LE.5235</td>
			<td style="width:103px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">computer and monitor</td>
			<td style="width:131px;">equipped with recording software</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">container and pipette for liquid waste&#160;</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;"></td>
		</tr>
		<tr height="160">
			<td height="160" style="height:160px;width:131px;">crayfish saline&#160;</td>
			<td style="width:131px;">contains (in mM): 5.4 KCl, 2.6 MgCl2, 13.5 CaCl2, and 195 NaCl, buffered with 10mM Tris base and 4.7mM maleic acid; aerated for 3 hours. Adjust at pH of 7.4.&#160;</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">always keep at temperatures ~ 4&#176; C</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">dextran, Texas Red (3000MW, lysine fixable)</td>
			<td style="width:131px;">fluorescent dye, lysine fixable</td>
			<td style="width:139px;">Life Technologies GmbH, Darmstadt, Germany</td>
			<td style="width:103px;">D3328</td>
			<td style="width:103px;">for intracellular dyefill of neurons</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">differential pin electrodes</td>
			<td style="width:131px;">made from stainless steel &#632; 0.2 mm</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">dissection dish&#160;</td>
			<td style="width:131px;">(l x w x h) 15x7x5 cm; linned with black silicone</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">used in the gross disection</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">faraday cage</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">fixing pins</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for pinning the specimen</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">forceps (biology, Dumont #5)</td>
			<td style="width:131px;">Forceps: Biology, tip 0.05 x 0.02 mm, length 11cm, INOX</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">11252-20</td>
			<td style="width:103px;">fine forceps: used to pick nerves</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">forceps (biology, Dumont #55)</td>
			<td style="width:131px;">Forceps: Biology, tip 0.05 x 0.02 mm, length 11cm, INOX</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">11255-20</td>
			<td style="width:103px;">extra fine forceps: used for desheathing</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:131px;">forceps (electronic, Dumont #5)</td>
			<td style="width:131px;">Forceps: Standard, tip 0.1 x 0.06 mm, length 11cm, INOX</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">11251-20</td>
			<td style="width:103px;">coarse forceps:&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; used to grab specimen and pins</td>
		</tr>
		<tr height="120">
			<td height="120" style="height:120px;width:131px;">intracellular electrode</td>
			<td style="width:131px;">Borosilicate glass capillaries (outer/inner diameter: 1mm/0.5mm), with filament</td>
			<td style="width:139px;">Sutter Instruments, Novato, CA</td>
			<td style="width:103px;">BF100-50-10</td>
			<td style="width:103px;">for intracellular recording and dyefill of neurons</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">Leica S8 Apo StereoZoom</td>
			<td style="width:131px;">Dissection Microscope&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; Zoom 1x - 8x</td>
			<td style="width:139px;">Leica, Germany</td>
			<td style="width:103px;">10446298</td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">microscope table</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">mirror</td>
			<td style="width:131px;">to illuminate preparation from below</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:131px;">modeling clay</td>
			<td style="width:131px;"></td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">Olympus SZ61</td>
			<td style="width:131px;">Dissection Microscope&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; Zoom 0.67x - 4.5x</td>
			<td style="width:139px;">Olympus, Germany</td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for the dissection</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">petri dish&#160;</td>
			<td style="width:131px;">94 x 16 mm; lined with clear silicone</td>
			<td style="width:139px;">Greiner bio-one, Germany</td>
			<td style="width:103px;">633180</td>
			<td style="width:103px;">used to pin the isolated chain of ganglia</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:131px;">ring scissors</td>
			<td style="width:131px;">ThoughCut, cutting edge: sharp/blunt, straight: 13cm</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">14054-13</td>
			<td style="width:103px;">for gross dissection&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; (steps 2.1 - 2.11)</td>
		</tr>
		<tr height="100">
			<td height="100" style="height:100px;width:131px;">saline dispenser&#160; with a 16 gauge needle (outer &#632; 1.6mm) attached via a flexible tube.</td>
			<td style="width:131px;">Volume ~ 60ml,</td>
			<td style="width:139px;"></td>
			<td style="width:103px;"></td>
			<td style="width:103px;">used for exsanguination</td>
		</tr>
		<tr height="120">
			<td height="120" style="height:120px;width:131px;">spring scissors or alternative: Vannas spring scissors</td>
			<td style="width:131px;">cutting edge: 8 mm, tip diameter: 0.2mm, straight: 10cm or cutting edge 2.5 mm, tip diameter 0.075 mm, straight: 8cm</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">15024-10 or&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 15000-08</td>
			<td style="width:103px;">for desheathing&#160;</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:131px;">stainless steel wire &#632; 0.125 mm&#160;</td>
			<td style="width:131px;">to cut pins of 4-7 mm length</td>
			<td style="width:139px;">Goodfellow GmbH, Bad Nauheim, Germany</td>
			<td style="width:103px;"></td>
			<td style="width:103px;">for pinning of the nerve cord</td>
		</tr>
		<tr height="140">
			<td height="140" style="height:140px;width:131px;">student Vannas&#160; spring scissors or alternative:&#160; Moria Spring Scissors</td>
			<td style="width:131px;">cutting edge: 5mm, tip diameter: 0.35mm, straight: 9cm or cutting edge: 5mm, tip diameter 0,1 mm, straight: 8 cm</td>
			<td style="width:139px;">Fine Science Tools (FST), Germany</td>
			<td style="width:103px;">91500-09 or&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 15396-00</td>
			<td style="width:103px;">for gross and fine disection&#160;&#160;&#160;&#160;&#160;&#160;&#160; (steps 2.11 - 3.14)</td>
		</tr>
		<tr height="100">
			<td height="100" style="height:100px;width:131px;">sylgard</td>
			<td style="width:131px;">184 Silicone Elastomer Base and Curing Agent; for black sylgard add activated carbon</td>
			<td style="width:139px;">Dow Corning, Midland, MI, USA</td>
			<td style="width:103px;"></td>
			<td style="width:103px;"></td>
		</tr>
		<tr height="100">
			<td height="100" style="height:100px;width:131px;">syringe filled with petroleum jelly and equipped with a 20 gauche needle with rounded tip</td>
			<td></td>
			<td></td>
			<td></td>
			<td style="width:103px;">for extracellular recording</td>
		</tr>
	</tbody>
</table>

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.

With the simultaneous extracellular recordings from RS and PS, motor neurons of one ganglion, the alternating activity of these motor neuron pools, can be monitored (Figure 18), representing the fictive locomotion pattern.
<img alt="Figure 18" src="/files/ftp_upload/52109/52109fig18highres.jpg" /> 
Figure 18: Schematic of one ganglion and placement of differential pin electrode. Extracellular recording of RS motor neurons (upper trace) an...

Watch this Scientific Journal Video about The Swimmeret System of Crayfish: A Practical Guide for the Dissection of the Nerve Cord and Extracellular Recordings of the Motor Pattern at JoVE.com

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

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 ...

the-swimmeret-system-crayfish-practical-guide-for-dissection-nerve

Research

JoVE Journal

Neuroscience

12.0K Views.  University of Cologne. 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. 

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

Physiological Recordings of High and Low Output NMJs on the Crayfish Leg Extensor Muscle

We explain in detail how to expose and conduct electrophysiological recordings of synaptic responses for high (phasic) and low (tonic) output motor neurons innervating the extensor muscle in the walking leg of a crayfish. Distinct differences are present in the physiology and morphology of the phasic and tonic nerve terminals. The tonic axon contains many more mitochondria, enabling it to take a vital stain more intensely than the phasic axon. The tonic terminals have varicosities, and the phasic terminal is filiform. The tonic terminals are low in synaptic efficacy but show dramatic facilitated responses. In contrast, the phasic terminals are high in quantal efficacy but show synaptic depression with high frequency stimulation. The quantal output is measured with a focal macropatch electrode placed directly over the visualized nerve terminals. Both phasic and tonic terminals innervate the same muscle fibers, which suggests that inherent differences in the neurons, rather than differential retrograde feedback from the muscle, account for the morphological and physiological differentiation.

This article demonstrates how to conduct electrophysiological recordings of synaptic responses on the extensor muscle in the walking leg of a crayfish and how the nerve terminals are visualized to show the gross morphological differences of high- and low-output nerve terminals.

We explain in detail how to expose and conduct electrophysiological recordings of synaptic responses for high (phasic) and low (tonic) output motor ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled samples with little to no tissue damage enables us to analyze cell morphology and to compare individual samples to each other, hence allowing the identification of mutant anomalies.
In the demonstrated dissection method the nervous system remains mostly inside the adult fly. Through a dorsal incision, the abdomen and thorax are opened and most of the internal organs are removed. Only the dorsal side of the ventral nerve cord (VNC) and the cervical connective 
(CvC) containing the big axons of the giant fibers (GFs)1 are exposed, while the brain containing the GF cell body and dendrites remains2 in the 
intact head. In this preparation most nerves of the VNC should remain attached to their muscles. 
Following the dissection, the intracellular filling of the giant fiber (GF) with a fluorescent dye is demonstrated. In the CvC the GF axons are located at the dorsal surface and thus can be easily visualized under a microscope with differential interference contrast (DIC) optics. This allows the injection of the GF axons with dye at this site to label the entire GF including the axons and their terminals in the VNC. This method results in reliable and strong staining of the GFs allowing the neurons to be imaged immediately after filling with an epifluorescent microscope. Alternatively, the fluorescent signal can be enhanced using standard immunohistochemistry procedures3 suitable for high resolution confocal microscopy.

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

An in vivo dissection of the adult Drosophila ventral nerve cord (VNC) is demonstrated. This particular dissection method causes little damage to the VNC allowing the subsequent labeling of the giant fiber neurons with fluorescent dye for high resolution imaging.

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

A Visual Description of the Dissection of the Cerebral Surface Vasculature and Associated Meninges and the Choroid Plexus from Rat Brain

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain proper, that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis. The tissue harvested is suitable for biochemical and physiological analysis, and the MAV has been shown to be sensitive to damage produced by amphetamine and hyperthermia 1,2. As well, the major and minor cerebral vasculatures harvested in MAV are of potentially high interest when investigating concussive types of head trauma. The MAV dissected in this presentation consists of the pial and some of the arachnoid membrane (less dura) of the meninges and the major and minor cerebral surface vasculature. The choroid plexus dissected is the structure that resides in the lateral ventricles as described by Oldfield and McKinley3,4,5,6. The methods used for harvesting these two tissues also facilitate the harvesting of regional cortical tissue devoid of meninges and larger cerebral surface vasculature, and is compatible with harvesting other brain tissues such as striatum, hypothalamus, hippocampus, etc. The dissection of the two tissues takes from 5 to 10 min total. The gene expression levels for the dissected MAV and choroid plexus, as shown and described in this presentation can be found at GSE23093 (MAV) and GSE29733 (choroid plexus) at the NCBI GEO repository. This data has been, and is being, used to help further understand the functioning of the MAV and choroid plexus and how neurotoxic events such as severe hyperthermia and AMPH adversely affect their function.

This video presentation shows a method of harvesting the two most important highly vascular structures that support forebrain function. They are the cerebral surface (superficial) vasculature along with associated meninges (MAV) and the choroid plexus which are necessary for cerebral blood flow and cerebrospinal fluid (CSF) homeostasis.

This video presentation was created to show a method of harvesting the two most important highly vascular structures, not residing within the brain ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Assay to Measure Nucleocytoplasmic Transport in Real Time within Motor Neuron-like NSC-34 Cells

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a connection between amyotrophic lateral sclerosis (ALS) and impairments in the nucleocytoplasmic pathway. ALS is a neurodegenerative disease affecting the motor neurons and resulting in paralysis and ultimately in death, within 2-5 years on average. Most cases of ALS are sporadic, lacking any apparent genetic linkage, but 10% are inherited in a dominant manner. Recently, hexanucleotide repeat expansions (HREs) in the chromosome 9 open reading frame 72 (C9orf72) gene were identified as a genetic cause of ALS and frontotemporal dementia (FTD). Importantly, different groups have recently proposed that these mutants affect nucleocytoplasmic transport. These studies have mostly shown the final outcome and manifestations caused by HREs on nucleocytoplasmic transport, but they do not demonstrate nuclear transport dysfunction in real time. As a result, only severe nucleocytoplasmic transport deficiency can be determined, mostly due to high overexpression or exogenous protein insertion. 
This protocol describes a new and very sensitive assay to evaluate and quantify nucleocytoplasmic transport dysfunction in real time. The rate of import of a NLS-NES-GFP protein (shuttle-GFP) can be quantified in real time using fluorescent microscopy. This is performed by using an exportin inhibitor, thus allowing the shuttle GFP only to enter the nucleus. To validate the assay, the C9orf72 HRE translated dipeptide repeats, poly(GR) and poly(PR), which have been previously shown to disrupt nucleocytoplasmic transport, were used. Using the described assay, a 50% decrease in the nuclear import rate was observed compared to the control. Using this system, minute changes in nucleocytoplasmic transport can be examined and the ability of different factors to rescue (even partially) a nucleocytoplasmic transport defect can be determined.

This protocol describes a method to sensitively measure the nucleocytoplasmic transport rate within motor neuron-like NSC-34 cells by quantifying the real-time change in the nuclear import of a NLS-NES-GFP protein.

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a ...

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.

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.

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

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.