This work was supported by a Wellcome Trust grant to L.P.

1. Equipment and Materials
<ol>
 <li>These experiments use a standard electrophysiology setup comprised of a stimulator, a stimulus isolation unit, two microelectrode amplifiers, a data acquisition system and a computer with collection software. Additional equipment includes a Faraday cage, a stereomicroscope on a boom stand, a vibration isolation table, a light source, and a recording platform. </li>
 <li>Five micromanipulators are used. Two micromanipulators require fine controls for positioning the recording electrodes, while the other three micromanipulators only require gross controls to position the two stimulation electrodes and the ground electrode. The micromanipulator for the DLM recording electrode is placed at the tail end of the preparation (left of experimenter) and the micromanipulator for the TTM recording electrode is placed between the experimenter and the side of the preparation (slightly to the left of the experimenter). The two micromanipulators that will hold the simulation electrodes are placed at the head of the preparation (right of experimenter). The micromanipulator for the ground electrode is placed at the far side of the preparation</li>
 <li>Pull glass recording microelectrodes with resistances of 40-60 M&Omega; and store flat in a dish supported by wax. For stimulation, two electrolytically (NaOH) sharpened tungsten electrodes are used. A tungsten wire, or a third electrolytically fabricated electrode is used as a ground. The stimulating and ground electrodes are prepared and attached to the micromanipulators before the start of the experimental session and need not be replaced for the duration of the session.</li>
</ol>
2. Preparing the D. melanogaster
<ol>
 <li>Once your equipment is set up, it's time to prepare the flies. Anesthetize the flies by cooling on them ice or by using CO2. If CO2 is used, then allow sufficient time (about 20mins) for the effects of the gas to wear off prior to beginning the experiment. </li>
 <li>Use forceps to transfer the flies gently by their legs to a dish containing a platform of soft wax sloped at an angle of approximately 45&deg;. The next four steps are done under a dissecting microscope away from (but close to) the recording equipment. </li>
 <li>The next step is to secure the fly in wax. Orient the fly ventral side down, with its anterior facing upwards on the slope. Using a pair of fine forceps, extend the legs outwards, in pairs, and push them into the wax.</li>
 <li>Familiarize yourself with the location of the muscles to be recorded from: the dorsal longitudinal muscle, or DLM, and the tergotrochanteral muscle, or TTM. The subcuticular attachment sites of the DLMs correspond with the region between the thoracic midline and the anterior dorsal bristles (or setae). The TTM attachment sites are located dorsally of the posterior and anterior supra-alar bristles. Making sure that the wings will not obstruct access to the DLM or TTM fibers, hold the wings outward and 'glue' them to the wax. </li>
 <li>Using a fine pair of forceps, pull the proboscis outward carefully, and secure it by immersing it into the wax. This is a critical step that requires some practice since the proboscis is soft and is easily detached from the rest of the head. If that happens, discard the fly and start over. Failure to secure the head in this way leads to problems when inserting the stimulating electrodes through the eyes.</li>
</ol>
3. Placing the Electrodes
<ol>
 <li>Once the fly is anchored to the wax, transfer the dish with the attached fly underneath the stereomicroscope that is located inside the Faraday cage. Orient the fly sideways with the head of the fly to the right of the experimenter.</li>
 <li>The next step is to insert the electrodes. Ground and stimulating electrodes can be inserted without looking through the microscope. Good recordings rely on precise impalement, so it is a good idea to practice handling the micromanipulators. Bring the electrodes close to the sites of insertion with the help of micromanipulators to facilitate their proper placing and subsequent recordings.</li>
 <li>Lower the ground electrode into the posterior end of the abdomen using the adjustment wheels on the micromanipulator. To place the sharpened tungsten stimulating electrodes in the brain, use the micromanipulator to position the tip of one of the electrodes so it just touches one of fly's eyes. Do the same with the other so both electrodes are just touching the outside of each eye. Then push the electrodes, in turn, through each eye so the tips of the electrodes reach the brain situated at the back of the head capsule (about 2-3mm). </li>
 <li>Correctly placed electrodes will activate the Giant Fiber System. To test that the stimulating electrodes are placed correctly, apply a short (0.03 ms) stimulus of 30-60 V across the stimulating electrodes, and look for movement of the wings and twitches of the flight/leg muscle'</li>
 <li>The next step is to back-fill the glass microelectrodes with 3M KCl using a Hamilton or heat-pulled plastic syringe, and place them into the fine-control micromanipulators. Properly inserted microelectrodes can be used for several rounds of experiments. </li>
 <li>The first recording electrode will be inserted into a DLM fiber. There are two bilaterally symmetric DLMs; each one is composed of six individual muscle fibers. The recordings can be done from any of the six fibers; however, the most commonly used are DLM fibers 45a and 45b due to their good accessibility through the dorsal side of the thoracic cuticle, and the fact that both fibers are innervated by the same motorneuron. </li>
 <li>Using the micromanipulator on the side farthest from you, insert a recording electrode into DLM fiber 45a or b. The slope of the platform allows the DLM electrode to enter the dorsal cuticle at a ~60-90&deg; angle, which aids penetration. Use the software in oscilloscope mode and look at the computer monitor while inserting recording electrodes into the thorax. When the electrode has entered a muscle the baseline will drop to near zero or a negative value. Test with a single stimulus to see if you can observe the muscle response. </li>
 <li>Insert the other recording electrode into the TTM closest to you. This electrode is inserted laterally, in front of you, due to the location of the muscle attachment site. Again observe the monitor while doing this and test with a single stimulus once the trace indicates the electrode is in the muscle.</li>
</ol>
4. Stimulation and Recording
<ol>
 <li>You are now ready to begin stimulating the brain and recording responses from the leg and flight muscles. Apply a short (0.03 ms) stimulus across the stimulating electrodes starting at 30 V and increasing to 60 V until you observe a response (i.e. a muscle twitch, and a muscle cell depolarization as observed on the computer monitor). For the remainder of the experiment, set the voltage 5-10 V above the response threshold. </li>
 <li>To measure response latency, give at least 5 single stimuli with a 5 second rest period between each stimulus. </li>
 <li>Determine the "frequency of following" by providing trains of stimuli at different rates. Typically 10 trains of 10 stimuli are given at 100Hz (10ms between each stimulus), 200Hz (5ms between each stimulus) and 300Hz (3ms between each stimulus). Allow a rest period of 2 seconds between each train of stimuli.</li>
</ol>
5. Results: Response Latencies and Frequency of Following in the Giant Fiber Pathway
<ol>
 <li>The response latency is the time between stimulation of the brain and depolarization of the muscle. This figure compares the response latencies for DLM and TTM to a single stimulus. Latencies between 0.7 and 1.2 ms for the GF-TTM pathway and between 1.3 and 1.7 ms for the GF-DLM pathway indicate a healthy preparation and proper recording technique. The latencies can vary with genotype, genetic background, temperature and age. 
Figure 1 (A and B). Representative traces showing responses recorded from the TTMs and DLMs following a single stimulus applied to the brain.</li>
 <li>As shown here, recordings from the TTM show more variability in terms of amplitude and shape of the postsynaptic potential (PSP) compared to those from the large DLM fibers; this increased variability is due to the small size of the TTM muscle fibers. This variability, however, does not affect the response latency values for the Giant Fiber-to-TTM pathway. Figure 1 (C and D). Further 'response latency' traces from 4 individual flies for both the TTM and DLM. Note TTM traces exhibit variability in PSP shape but response latency is unaffected. For DLM there is less variability in PSP shape. <img src="/files/ftp_upload/2412/2412fig1.jpg" alt="Figure 1 A-D" /> </li>
 <li>Compare the "frequency of following" at 100 Hz, 200 Hz, and 300 Hz by calculating the proportion of successful responses (out of 10) for both DLM and TTM pathways at each stimulation frequency. At 100 Hz, both TTM and DLM follow the stimuli 1:1. At stimulation frequencies above 100 Hz, the DLM responses start to show failures because the intermediary chemical synapse between two interneurons does not have sufficient time to recover between stimuli. The TTM responses, however, remain 1:1 with stimuli even beyond 300Hz. Figure 2. Representative traces showing the "frequency of following" recordings. At 100Hz, both TTMs and DLMs respond to all 10 stimuli (left). At 200Hz, the DLM responses start to fail (asterisk). <img src="/files/ftp_upload/2412/2412fig2.jpg" alt="Figure 2" /> </li>
</ol>
6. Representative Results
Wild type short-latency responses (stimulated electrodes are placed in the eyes, bypassing sensory receptors and triggering the GF circuit directly) depend on genotype, genetic background, temperature and age, and range between 0.7 and 1.2 ms for the GF-TTM pathway and 1.3 and1.7 ms for the GF-DLM pathway (Tanouye and Wyman, 1980; Thomas and Wyman, 1984; Engel and Wu, 1992; Allen and Murphey, 2007; Phelan et al., 2008; Augustin et al., unpublished). This very short TTM latency is due to the robust GF-TTMn electrochemical synapse of the monosynaptic pathway and the longer DLM latency occurs because of the disynaptic nature of the pathway as well as the presence of a chemical synapse (PSI-DLMn). Intermediate- and long-latency responses (&gt;3 ms) result from the activation of the GF afferents and are achieved either by using a lower intensity stimulation or providing a visual ("light-off") signal. At 100Hz both TTM and DLM should follow the stimuli 1:1. Above 100Hz DLM responses will start to show failures as the chemical synapse between PSI and the DLMns does not have sufficient time to recover between stimuli less than 10ms apart. TTM responses, however, will remain 1:1 with stimuli even beyond 300Hz (Tanouye and Wyman, 1980; Engel and Wu, 1992; Allen et al., 2007; Martinez et al., 2007). Mutations in the shakB gene, encoding a Drosophila gap junction channel ("innexin"), significantly increase the response latency of the GF-TTM pathway (~1.5 ms) while the GF-DLM branch is unresponsive (Allen and Murphey, 2007; Phelan et al., 2008). The mutant response can be restored by stimulating thoracic ganglia directly, demonstrating that the delayed effect is not due to disrupted neuromuscular transmission. The ability to follow high frequency stimulation is also impaired in these mutants compared to wild type flies where the GF-DLM and GF-TTM pathways are usually able to follow 10 stimuli with 1:1 ratio up to 100 Hz and 300 Hz, respectively. It is important to note that these frequencies are considerably above normal stimulation frequencies received by the contracting muscles during the sustained flight (3-10 Hz) (Hummon and Costello, 1989).
Another parameter used to describe the stability of the GFS outputs is the "refractory period", or the minimal time between twin stimulus pulses that still produces two responses from the muscle. The refractory time varies between 1-4 ms for TTMs and 7-15 ms for DLMs. The comparatively long refractory period for DLMs is due to relatively labile chemical synapses at the PSI-DLMn junction (Tanouye and Wyman, 1980; Gorczyca and Hall, 1984; Engel and Wu, 1992; Banerjee et al., 2004; Allen and Godenschwege, 2010).

<ol>
	<li>Allen, M. J., Godenschwege, T., Zhang, B., Freeman, M. R., Wadde, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+Recordings+from+the+Drosophila+Giant+Fiber+System.">Electrophysiological Recordings from the Drosophila Giant Fiber System.</a> Drosophila Neurobiology: A Laboratory Manual. , (2010).</li><li>Allen, M. J., Godenschwege, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Making+an+escape:+development+and+function+of+the+Drosophila+giant+fibre+system.">Making an escape: development and function of the Drosophila giant fibre system.</a> Semin Cell Dev Biol. 17, 31-41 (2006).</li><li>Allen, M. J., Murphey, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chemical+component+of+the+mixed+GF-TTMn+synapse+in+Drosophila+melanogaster+uses+acetylcholine+as+its+neurotransmitter.">The chemical component of the mixed GF-TTMn synapse in Drosophila melanogaster uses acetylcholine as its neurotransmitter.</a> Eur J Neurosci. 26, 439-445 (2007).</li><li>Banerjee, S., Lee, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Loss+of+flight+and+associated+neuronal+rhythmicity+in+inositol+1,4,5-trisphosphate+receptor+mutants+of+Drosophila.">Loss of flight and associated neuronal rhythmicity in inositol 1,4,5-trisphosphate receptor mutants of Drosophila.</a> J Neurosci. 24, 7869-7878 (2004).</li><li>Engel, J. E., Wu, C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interactions+of+membrane+excitability+mutations+affecting+potassium+and+sodium+currents+in+the+flight+and+giant+fiber+escape+systems+of+Drosophila.">Interactions of membrane excitability mutations affecting potassium and sodium currents in the flight and giant fiber escape systems of Drosophila.</a> J Comp Physiol A. 171, 93-104 (1992).</li><li>Gorczyca, M., Hall, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+a+cholinergic+synapse+in+the+giant+fiber+pathway+of+Drosophila+using+conditional+mutations+of+acetylcholine+synthesis.">Identification of a cholinergic synapse in the giant fiber pathway of Drosophila using conditional mutations of acetylcholine synthesis.</a> J Neurogenet. 1, 289-313 (1984).</li><li>Hummon, M. R., Costello, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Giant+fiber+activation+of+flight+muscles+in+Drosophila:+asynchrony+in+latency+of+wing+depressor+fibers.">Giant fiber activation of flight muscles in Drosophila: asynchrony in latency of wing depressor fibers.</a> J Neurobiol. 20, 593-602 (1989).</li><li>Jacobs, K., Todman, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptogenesis+in+the+giant-fibre+system+of+Drosophila:+interaction+of+the+giant+fibre+and+its+major+motorneuronal+target.">Synaptogenesis in the giant-fibre system of Drosophila: interaction of the giant fibre and its major motorneuronal target.</a> Development. 127, 5203-5212 (2000).</li><li>King, D. G., Wyman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomy+of+the+giant+fibre+pathway+in+Drosophila.+I.+Three+thoracic+components+of+the+pathway.">Anatomy of the giant fibre pathway in Drosophila. I. Three thoracic components of the pathway.</a> J Neurocytol. 9, 753-770 (1980).</li><li>Martinez, V. G., Javadi, 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=Age-related+changes+in+climbing+behavior+and+neural+circuit+physiology+in+Drosophila.">Age-related changes in climbing behavior and neural circuit physiology in Drosophila.</a> Dev Neurobiol. 67, 778-791 (2007).</li><li>Miller, A., Demerec, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+internal+anatomy+and+histology+of+the+imago+of+Drosophila+melanogaster.">The internal anatomy and histology of the imago of Drosophila melanogaster.</a> Biology of Drosophila. , 502-503 (1965).</li><li>Phelan, P., Goulding, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+mechanism+of+rectification+at+identified+electrical+synapses+in+the+Drosophila+giant+fiber+system.">Molecular mechanism of rectification at identified electrical synapses in the Drosophila giant fiber system.</a> Curr Biol. 18, 1955-1960 (2008).</li><li>Power, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+thoracico-abdominal+nervous+system+of+an+adult+insect,+Drosophila+melanogaster.">The thoracico-abdominal nervous system of an adult insect, Drosophila melanogaster.</a> J Comp Neurol. 88, 347-409 (1948).</li><li>Tanouye, M. A., Wyman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+outputs+of+giant+nerve+fiber+in+Drosophila.">Motor outputs of giant nerve fiber in Drosophila.</a> J Neurophysiol. 44, 405-421 (1980).</li><li>Thomas, J. B., Wyman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+altering+synaptic+connectivity+between+identified+neurons+in+Drosophila.">Mutations altering synaptic connectivity between identified neurons in Drosophila.</a> J Neurosci. 4, 530-538 (1984).</li></ol>

No conflicts of interest declared.

One of the most important things one has to pay attention to when trying to obtain high quality recordings is the proper orientation and health of the preparation. Ideally, the fly should still be alive at the end of the recording session and responsive to electrical stimuli. For the recording electrodes to most efficiently penetrate the thoracic exoskeleton, the fly should be glued to the surface in such a way as to form a right angle with the electrodes; if necessary, the insertion of electrodes can be facilitated by removing a portion of the dorsal thoracic cuticle with a tungsten scalpel thus exposing the DLM flight muscle (this step offers an additional advantage of making it harder for the tips of glass electrodes to break). Also, the care must be taken to avoid pushing the electrodes through the subcuticularly located DLMs and TTMs. The head of the fly should be well secured to allow for the stimulating electrodes to be properly inserted into the brain and to prevent them from being pulled out during the recording session.
Due to its size and well-described morphology, the GFS represents one of the most accessible neuronal pathways in Drosophila. The permeability of electrical synapses to small molecular weight tracer dyes allows for the visualisation of electrically coupled neurons, and several available GAL4 lines make it possible to manipulate gene expression levels in a subset of cells or cell groups (Jacobs et al., 2000; Allen et al., 2006) In addition to the above mentioned advantages, both afferent and thoracic components of the circuit display properties such as habituation, spontaneous recovery and dishabituation, making the Drosophila GFS a convenient model system for studying neuronal plasticity (Engel and Wu, 1996).

<table border="1">
 <tbody>
 <tr>
 <td>NAME</td>
 <td>COMPANY</td>
 <td>CAT. #</td>
 <td>COMMENTS</td>
 </tr>
 <tr>
 <td>S48 Square Pulse Stimulator</td>
 <td>Grass Instruments</td>
 <td>&nbsp;</td>
 <td> http://www.grasstechnologies.com/</td>
 </tr>
 <tr>
 <td>Stimulation unit</td>
 <td>Grass Instruments</td>
 <td>&nbsp;</td>
 <td>http://www.grasstechnologies.com/</td>
 </tr>
 <tr>
 <td>SIU5 RF Transformer Isolation Unit</td>
 <td>Grass Instruments</td>
 <td>&nbsp;</td>
 <td>http://www.grasstechnologies.com/</td>
 </tr>
 <tr>
 <td>5A two-channel intracellular Microelectrode Amplifier</td>
 <td>Getting Instruments</td>
 <td>&nbsp;</td>
 <td>http://www.gettinginstruments.com/</td>
 </tr>
 <tr>
 <td>Digidata 1440A data acquisition system</td>
 <td>Molecular Devices</td>
 <td>&nbsp;</td>
 <td>http://www.moleculardevices.com/</td>
 </tr>
 <tr>
 <td>Analogue-digital Digidata 1320 and Axoscope 9.0 software</td>
 <td>Molecular Devices</td>
 <td>&nbsp;</td>
 <td>http://www.moleculardevices.com/</td>
 </tr>
 <tr>
 <td>Recording platform with manual micromanipulators</td>
 <td>Narishige, Sutter Ins., World Precision Ins.</td>
 <td>&nbsp;</td>
 <td>http://narishige-group.com/ 
 http://www.sutter.com/index.html 
 http://www.wpi-europe.com/en/</td>
 </tr>
 <tr>
 <td>Light source</td>
 <td>Fostec</td>
 <td>&nbsp;</td>
 <td>http://www.nuhsbaum.com/FOSTEC.htm</td>
 </tr>
 <tr>
 <td>Wild M5 stereomicroscope</td>
 <td>Wild Heerbrugg</td>
 <td>&nbsp;</td>
 <td>http://www.wild-heerbrugg.com/</td>
 </tr>
 <tr>
 <td>Vibration isolation table</td>
 <td>TMC</td>
 <td>&nbsp;</td>
 <td>http://www.techmfg.com/</td>
 </tr>
 <tr>
 <td>Borosilicate tubing for microelectrodes</td>
 <td>Sutter Instrument</td>
 <td>&nbsp;</td>
 <td>http://www.sutter.com/index.html</td>
 </tr>
 <tr>
 <td>P-95 Micropipette puller</td>
 <td>Sutter Instrument</td>
 <td>&nbsp;</td>
 <td>http://www.sutter.com/index.html</td>
 </tr>
 <tr>
 <td>Microfil 34 gauge, 67 mm (electrode filler)</td>
 <td>World Precision Instruments</td>
 <td>MF34G-5</td>
 <td>http://www.wpi-europe.com/en/</td>
 </tr>
 <tr>
 <td>Microdissection tools (forceps,&hellip;)</td>
 <td>Fine Science Tools </td>
 <td>&nbsp;</td>
 <td>www.finescience.com</td>
 </tr>
 <tr>
 <td>Dissecting (stereo) microscope</td>
 <td>Leica</td>
 <td>&nbsp;</td>
 <td>http://www.leica-microsystems.com/</td>
 </tr>
 <tr>
 <td>Faraday cage</td>
 <td>Unknown manufacturer</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>
Other: plastic syringes, tungsten earth wire and NaOH-sharpened tungsten electrodes, KCl, wax platform, a PC with monitor...

electrophysiological recordings from the giant fiber pathway of d. melanogaster

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" (or "startle") response during the adult stage is mediated by the multi-component neuronal circuit called the Giant Fiber System (GFS). The comparative large size of the neurons, their distinctive morphology and simple connectivity make the GFS an attractive model system for studying neuronal circuitry. The GFS pathway is composed of two bilaterally symmetrical Giant Fiber (GF) interneurons whose axons descend from the brain along the midline into the thoracic ganglion via the cervical connective. In the mesothoracic neuromere (T2) of the ventral ganglia the GFs form electro-chemical synapses with 1) the large medial dendrite of the ipsilateral motorneuron (TTMn) which drives the tergotrochanteral muscle (TTM), the main extensor for the mesothoracic femur/leg, and 2) the contralateral peripherally synapsing interneuron (PSI) which in turn forms chemical (cholinergic) synapses with the motorneurons (DLMns) of the dorsal longitudinal muscles (DLMs), the wing depressors. The neuronal pathway(s) to the dorsovental muscles (DVMs), the wing elevators, has not yet been worked out (the DLMs and DVMs are known jointly as indirect flight muscles - they are not attached directly to the wings, but rather move the wings indirectly by distorting the nearby thoracic cuticle) (King and Wyman, 1980; Allen et al., 2006). The di-synaptic activation of the DLMs (via PSI) causes a small but important delay in the timing of the contraction of these muscles relative to the monosynaptic activation of TTM (~0.5 ms) allowing the TTMs to first extend the femur and propel the fly off the ground. The TTMs simultaneously stretch-activate the DLMs which in turn mutually stretch-activate the DVMs for the duration of the flight. The GF pathway can be activated either indirectly by applying a sensory (e.g."air-puff" or "lights-off") stimulus, or directly by a supra-threshold electrical stimulus to the brain (described here). In both cases, an action potential reaches the TTMs and DLMs solely via the GFs, PSIs, and TTM/DLM motoneurons, although the TTMns and DLMns do have other, as yet unidentified, sensory inputs. Measuring "latency response" (the time between the stimulation and muscle depolarization) and the "following to high frequency stimulation" (the number of successful responses to a certain number of high frequency stimuli) provides a way to reproducibly and quantitatively assess the functional status of the GFS components, including both central synapses (GF-TTMn, GF-PSI, PSI-DLMn) and the chemical (glutamatergic) neuromuscular junctions (TTMn-TTM and DLMn-DLM). It has been used to identify genes involved in central synapse formation and to assess CNS function.

Watch this Scientific Journal Video about Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster at JoVE.com

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

The Giant Fiber System is a simple neuronal circuit of adult Drosophila melanogaster containing the largest neurons in the fly. We describe the protocol for monitoring synaptic transmission through this pathway by recording post synaptic potentials in dorsal longitudinal (DLM) and tergotrochanteral (TTM) muscles following direct stimulation of the Giant Fiber interneurons.

Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

When startled adult D. melanogaster react by jumping into the air and flying away. In many invertebrate species, including D. melanogaster, the "escape" ...

electrophysiological-recordings-from-giant-fiber-pathway-d

Research

JoVE Journal

Neuroscience

14.4K Views.  University College London - UCL. The Giant Fiber System is a simple neuronal circuit of adult Drosophila melanogaster containing the largest neurons in the fly. We describe the protocol for monitoring synaptic transmission through this pathway by recording post synaptic potentials in dorsal longitudinal (DLM) and tergotrochanteral (TTM) muscles following direct stimulation of the Giant Fiber interneurons.

Video: Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

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 ...

Paired Nanoinjection and Electrophysiology Assay to Screen for Bioactivity of Compounds using the Drosophila melanogaster Giant Fiber System

Screening compounds for in vivo activity can be used as a first step to identify candidates that may be developed into pharmacological agents1,2. We developed a novel nanoinjection/electrophysiology assay that allows the detection of bioactive modulatory effects of compounds on the function of a neuronal circuit that mediates the escape response in Drosophila melanogaster3,4. Our in vivo assay, which uses the Drosophila Giant Fiber System (GFS, Figure 1) allows screening of different types of compounds, such as small molecules or peptides, and requires only minimal quantities to elicit an effect. In addition, the Drosophila GFS offers a large variety of potential molecular targets on neurons or muscles. The Giant Fibers (GFs) synapse electrically (Gap Junctions) as well as chemically (cholinergic) onto a Peripheral Synapsing Interneuron (PSI) and the Tergo Trochanteral Muscle neuron (TTMn)5. The PSI to DLMn (Dorsal Longitudinal Muscle neuron) connection is dependent on D&alpha;7 nicotinic acetylcholine receptors (nAChRs)6. Finally, the neuromuscular junctions (NMJ) of the TTMn and the DLMn with the jump (TTM) and flight muscles (DLM) are glutamatergic7-12. Here, we demonstrate how to inject nanoliter quantities of a compound, while obtaining electrophysiological intracellular recordings from the Giant Fiber System13 and how to monitor the effects of the compound on the function of this circuit. We show specificity of the assay with methyllycaconitine citrate (MLA), a nAChR antagonist, which disrupts the PSI to DLMn connection but not the GF to TTMn connection or the function of the NMJ at the jump or flight muscles.
Before beginning this video it is critical that you carefully watch and become familiar with the JoVE video titled "Electrophysiological Recordings from the Giant Fiber Pathway of D. melanogaster " from Augustin et al7, as the video presented here is intended as an expansion to this existing technique. Here we use the electrophysiological recordings method and focus in detail only on the addition of the paired nanoinjections and monitoring technique.

Paired Nanoinjection and Electrophysiology Assay to Screen for Bioactivity of Compounds using the Drosophila melanogaster Giant Fiber System

A rapid in vivo assay to test for neuromodulatory compounds using the Giant Fiber System (GFS) of Drosophila melanogaster is described. Nanoinjections in the head of the animal along with electrophysiological recordings of the GFS can reveal bioactivity of compounds on neurons or muscles.

Screening compounds for in vivo activity can be used as a first step to identify candidates that may be developed into pharmacological agents1,2. We ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Design and Fabrication of Ultralight Weight, Adjustable Multi-electrode Probes for Electrophysiological Recordings in Mice

The number of physiological investigations in the mouse, mus musculus, has experienced a recent surge, paralleling the growth in methods of genetic targeting for microcircuit dissection and disease modeling. The introduction of optogenetics, for example, has allowed for bidirectional manipulation of genetically-identified neurons, at an unprecedented temporal resolution. To capitalize on these tools and gain insight into dynamic interactions among brain microcircuits, it is essential that one has the ability to record from ensembles of neurons deep within the brain of this small rodent, in both head-fixed and freely behaving preparations. To record from deep structures and distinct cell layers requires a preparation that allows precise advancement of electrodes towards desired brain regions. To record neural ensembles, it is necessary that each electrode be independently movable, allowing the experimenter to resolve individual cells while leaving neighboring electrodes undisturbed. To do both in a freely behaving mouse requires an electrode drive that is lightweight, resilient, and highly customizable for targeting specific brain structures.
A technique for designing and fabricating miniature, ultralight weight, microdrive electrode arrays that are individually customizable and easily assembled from commercially available parts is presented. These devices are easily scalable and can be customized to the structure being targeted; it has been used successfully to record from thalamic and cortical regions in a freely behaving animal during natural behavior.

Understanding the neural substrates of behavior requires brain circuit ensemble recording. Because of its genetic tractability, the mouse offers a model for circuit dissection and disease mimicry. Here, a method of designing and fabricating miniaturized probes is described that is suitable for targeting deep brain structure in the mouse.

The number of physiological investigations in the mouse, mus musculus, has experienced a recent surge, paralleling the growth in methods of genetic ...

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...

A Visual Guide to Sorting Electrophysiological Recordings Using 'SpikeSorter'

Few stand-alone software applications are available for sorting spikes from recordings made with multi-electrode arrays. Ideally, an application should be user friendly with a graphical user interface, able to read data files in a variety of formats, and provide users with a flexible set of tools giving them the ability to detect and sort extracellular voltage waveforms from different units with some degree of reliability. Previously published spike sorting methods are now available in a software program, SpikeSorter, intended to provide electrophysiologists with a complete set of tools for sorting, starting from raw recorded data file and ending with the export of sorted spikes times. Procedures are automated to the extent this is currently possible. The article explains and illustrates the use of the program. A representative data file is opened, extracellular traces are filtered, events are detected and then clustered. A number of problems that commonly occur during sorting are illustrated, including the artefactual over-splitting of units due to the tendency of some units to fire spikes in pairs where the second spike is significantly smaller than the first, and over-splitting caused by slow variation in spike height over time encountered in some units. The accuracy of SpikeSorter's performance has been tested with surrogate ground truth data and found to be comparable to that of other algorithms in current development.

A Visual Guide to Sorting Electrophysiological Recordings Using &#039;SpikeSorter&#039;

The article shows how to use the program SpikeSorter to detect and sort spikes in extracellular recordings made with multi-electrode arrays.

Few stand-alone software applications are available for sorting spikes from recordings made with multi-electrode arrays. Ideally, an application should be ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...