We thank Hyewon Cooper for illustrations.

1. Dissection
Cockroach saline&#160;solution used throughout this protocol has the following composition:
Cockroach saline solution36: (grams for 100 ml) 
210 mM NaCl (1.227 g) 
2.9 mM KCl (0.0216 g) 
1.8 mM CaCl2 (0.0265 g) 
0.2 mM NaH2PO4&#8226;2H2O (0.0032 g) 
1.8 mM Na2HPO4&#8226;7H2O (0.0483 g) 
(pH 7.2. Adjust pH with 1 M NaOH or 1 M HCl).
<ol>
	<li>Select a male cockroach from the holding tank that has robust cerci (Figure 1). The last segments of the male are narrow compared to the female; and containing no ovaries and egg mass, males are easier to dissect. The males of P. americana have a pair of short styli between the cerci. These styli are not observed in the females.</li>
	<li>Cut off the wings, legs and head and pin the body, ventral side up, to a dish lined with silicone elastomer.</li>
	<li>With forceps pick up the ventral plates and cut them off with fine scissors, starting at the posterior end and working anteriorly. Always keep the internal organs moist with saline&#160;while trying to keep the cerci dry. One can use wax or pieces of rubber to position the abdomen upwards to prevent the saline from wetting the cerci. If they do get wet, dry them with a piece of tissue paper. Push to the side the internal organs and the white matter (fat body). The VNC is in the center of the field, runs the length of the abdomen and should be visible between the shiny tracheae. The nerve cord is translucent and may initially be difficult to see until the lighting is adjusted properly (Figure 2). DO NOT handle the VNC with forceps or insect pins; instead manipulate it using glass probes.</li>
	<li>Clear away the animal&#39;s tracheae system as best as possible from the nerve cord with forceps and with a pair of fine glass needles, very carefully split the VNC connectives longitudinally between A6 and A5 or A5 and A4 ganglia (Figure 3). Cradle the cerci and abdomen upwards out of the saline bath with shortened insect pins and wax or a wedge of the silicone elastomer that can be cut to fit the preparation (Figures 4A and B). Be extra careful in the last abdominal segment not to damage the cercal nerves that project into the ganglion (Figures 2D and 5).</li>
</ol>
2. Extracellular Recording
<ol>
	<li>The dissected preparation, microscope, and recording apparatus should be setup inside a Faraday cage to block external, particularly AC, electric fields that could override signals from neurons (Figure 6).</li>
	<li>Position the microscope so that it is overlooking the microscope stage. Once the preparation is placed on the stage, adjust the position of the high intensity illuminator beam for best visualization of it.</li>
	<li>Connect the AC/DC differential amplifier to the integrated data recording unit (details on the specific hardware and software settings have been previously described 25). The head stage holding a suction&#160;electrode should be connected to the amplifier. A silver ground wire that has been coated with Cl- inserted into the abdomen results in more stable recordings. The reason is if the solution in the body cavity is not in contact with the bathing fluid in the dish, the fluid associated with the recording electrode remains grounded.</li>
	<li>Set the recording frequency to 4 kHz. Set the voltage scale (y-axis) to 500 mV (this can be adjusted to optimize visualization of the trace). Run the recording software in continuous or oscilloscope mode to record neural activity in response to stimulations.</li>
	<li>Cut one of the VNC connectives close to A5 and place the cut end attached to A6 into a suction electrode. Be sure to pull saline&#160;into the suction electrode to cover the silver wire inside it before sucking in the nerve.</li>
	<li>With a dry pipette blow air on to the hairs located on each cercus. See if stimulating the hairs on the cercus ipsilateral to the recorded connective gives a different response than the contralateral one. Take note of the amplitude of the responses and the number of spikes in a given time interval during the stimulation.</li>
	<li>Move the suction electrode to a cercal nerve for recording. To get a better fit, switch to an electrode tip with a smaller opening if necessary.</li>
	<li>Cut the cercal nerve close to A6 and then suck up the nerve leading to the cercus. There should be spontaneous firing of action potentials. Now, blow air onto the cercus and note the responses.</li>
</ol>
3. Electrically Stimulating the Sensory Nerves to Determine Recruitment
<ol>
	<li>Change the recording software to sweep mode so that it records traces (100-500 msec.) each time a stimulus is triggered.</li>
	<li>Connect the stimulating electrode to the output of the stimulator.</li>
	<li>Connect the stimulator cable with the two mini-hook leads or clips.</li>
	<li>Connect the BNC trigger output from the stimulator to the trigger input on the recording unit.</li>
	<li>The following stimulation parameters should evoke a response: Duration: 0.3 sec; Delay: 10 msec; Frequency: 1 Hz; Voltage: adjust as needed to obtain a signal in the recordings (just over threshold and being able to obtain a maximal response). There is no reason to go to voltages much higher than maximal threshold for recruitment as a high voltage can be damaging to the nerve.</li>
	<li>Cut the cercal nerve as distal as possible so that a long nerve root can be pulled into the stimulating suction electrode (Figure 7, arrow head). The connective between A6 and A5 or another segment more anterior can be used.</li>
	<li>Set the recording suction electrode so you can pull up a cut connective into the electrode. Be sure to pull some saline&#160;into the suction electrodes to cover the silver wire inside it before sucking in the nerves. Make sure the stimulating electrode is also grounded in the bath saline (in the abdomen near A3 is ideal).</li>
	<li>Deliver a series of single stimuli of increasing voltage until an action potential appears on the screen. One should make a record of the minimal stimulating voltage and duration to recruit a response. Increase the intensity until a synaptic response in the connectives is observed. The large spike (extracellular APs) from the giant axons appears first, and then other smaller AP&#39;s may also be observed.</li>
</ol>

<ol>
	<li>Full, R. J., Tu, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanics+of+a+rapid+running+insect:+two-,+four-+and+six-legged+locomotion.">Mechanics of a rapid running insect: two-, four- and six-legged locomotion.</a> J. Exp. Biol. 156, 215-231 (1991).</li><li>Ritzmann, R. E., Tobias, M. L., Fourtner, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flight+activity+initiated+via+giant+interneurons+of+the+cockroach:+Evidence+for+bifunctional+trigger+interneurons.">Flight activity initiated via giant interneurons of the cockroach: Evidence for bifunctional trigger interneurons.</a> Science. 210, 443-445 (1980).</li><li>Libersat, F., Camhi, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+cercal+position+during+flight+in+the+cockroach:+a+mechanism+for+regulating+sensory+feedback.">Control of cercal position during flight in the cockroach: a mechanism for regulating sensory feedback.</a> J. Exp. Biol. 136, 483-488 (1988).</li><li>Ganihar, D., Libersat, F., Wendler, G., Cambi, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wind-evoked+evasive+responses+in+flying+cockroaches.">Wind-evoked evasive responses in flying cockroaches.</a> Journal of Comparative Physiology. A, Sensory, Neural, and Behavioral Physiology. 175, 49-65 (1994).</li><li>Pipa, R. a. D., F, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The American Cockroach. , 175-216 (1981).</li><li>Westin, J., Langberg, J. J., Camhi, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Responses+of+giant+interneurons+of+the+cockroach;+Periplaneta+americana+to+wind+puffs+of+different+directions+and+velocities.">Responses of giant interneurons of the cockroach; Periplaneta americana to wind puffs of different directions and velocities.</a> Journal of Comparative Physiology A: Neuroethology, Sensory, Neural, and Behavioral Physiology. 121, 307-324 (1977).</li><li>Camhi, J. M., Tom, W., Volman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+escape+behavior+of+the+cockroach+Periplaneta+americana.">The escape behavior of the cockroach Periplaneta americana.</a> J. Comp. Physiol. A Neuroethol. Sens. Neural Physiol. 128, 203-212 (1978).</li><li>Nicklaus, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Die+Erregung+einzelner+Fadenhaare+von+Periplaneta+americana+in+Abh&#228;ngigkeit+von+der+Gr&#246;sse+und+Richtung+der+Auslenkung.">Die Erregung einzelner Fadenhaare von Periplaneta americana in Abh&#228;ngigkeit von der Gr&#246;sse und Richtung der Auslenkung.</a> Z. Vgl. Physiol. 50, 331-362 (1965).</li><li>Westin, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Responses+to+wind+recorded+from+the+cercal+nerve+of+the+cockroach+Periplaneta+americana.">Responses to wind recorded from the cercal nerve of the cockroach Periplaneta americana.</a> J. Comp. Physiol. A Neuroethol. Sens. Neural Physiol. 133, 97-102 (1979).</li><li>Ritzmann, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Neural Mechanisms of Startle Behavior. , 93-131 (1984).</li><li>Ritzmann, R. E., Pollack, A. J. <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+thoracic+interneurons+that+mediate+giant+interneuron-to-motor+pathways+in+the+cockroach.">Identification of thoracic interneurons that mediate giant interneuron-to-motor pathways in the cockroach.</a> J. Comp. Physiol. A Neuroethol. Sens. Neural Physiol. 159, 639-654 (1986).</li><li>Plummer, M. R., Camhi, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discrimination+of+sensory+signals+from+noise+in+the+escape+system+of+the+cockroach+-+the+role+of+wind+acceleration.">Discrimination of sensory signals from noise in the escape system of the cockroach - the role of wind acceleration.</a> J. Comp. Physiol. 142, 347-357 (1981).</li><li>Bullock, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Neural Mechanisms of Startle Behavior. , 1-14 (1984).</li><li>Pollack, A. J., Ritzmann, R. E., Watson, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+pathways+for+tactile+sensory+information+to+thoracic+interneurons+in+the+cockroach.">Dual pathways for tactile sensory information to thoracic interneurons in the cockroach.</a> J. Neurobiol. 26, 33-46 (1995).</li><li>Atwood, H. L., Wiersma, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Command+interneurons+in+the+crayfish+central+nervous+system.">Command interneurons in the crayfish central nervous system.</a> J. Exp. Biol. 46, 249-261 (1967).</li><li>Olson, G. C., Krasne, F. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+crayfish+lateral+giants+as+command+neurons+for+escape+behavior.">The crayfish lateral giants as command neurons for escape behavior.</a> Brain Res. 214, 89-100 (1981).</li><li>Comer, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analyzing+cockroach+escape+behavior+with+lesions+of+individual+giant+interneurons.">Analyzing cockroach escape behavior with lesions of individual giant interneurons.</a> Brain Res. 335, 342-346 (1985).</li><li>Comer, C. M., Dowd, J. P., Stubblefield, G. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Escape+responses+following+elimination+of+the+giant+interneuron+pathway+in+the+cockroach,+Periplaneta+americana.">Escape responses following elimination of the giant interneuron pathway in the cockroach, Periplaneta americana.</a> Brain Res. 445, 370-375 (1988).</li><li>Keegan, A. P., Comer, C. 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+wind-elicited+escape+response+of+cockroaches+(Periplaneta+americana)+is+influenced+by+lesions+rostral+to+the+escape+circuit.">The wind-elicited escape response of cockroaches (Periplaneta americana) is influenced by lesions rostral to the escape circuit.</a> Brain Res. 620, 310-316 (1993).</li><li>Casagrand, J. L., Ritzmann, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biogenic+amines+modulate+synaptic+transmission+between+identified+giant+interneurons+and+thoracic+interneurons+in+the+escape+system+of+the+cockroach.">Biogenic amines modulate synaptic transmission between identified giant interneurons and thoracic interneurons in the escape system of the cockroach.</a> J. Neurobiol. 23, 644-655 (1992).</li><li>Roeder, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+of+the+ascending+giant+fiber+system+in+the+cockroach,+Periplaneta+americana.">Organization of the ascending giant fiber system in the cockroach, Periplaneta americana.</a> J. Exp. Zool. 108, 243-261 (1948).</li><li>Ramos, R. L., Moiseff, A., Brumberg, 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=Utility+and+versatility+of+extracellular+recordings+from+the+cockroach+for+neurophysiological+instruction+and+demonstration.">Utility and versatility of extracellular recordings from the cockroach for neurophysiological instruction and demonstration.</a> J. Undergrad. Neurosci. Educ. 5, (2007).</li><li>Oakley, B., Schafer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+neurobiology.">Experimental neurobiology.</a> , (1978).</li><li>Welsh, J. H., Smith, R. I., Kammer, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Laboratory exercises in invertebrate physiology. , (1968).</li><li>Leksrisawat, B., Cooper, A. S., Gilberts, A. B., Cooper, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscle+receptor+organs+in+the+crayfish+abdomen:+a+student+laboratory+exercise+in+proprioception.">Muscle receptor organs in the crayfish abdomen: a student laboratory exercise in proprioception.</a> J. Vis. Exp. (45), e2323 (2010).</li><li>Bacon, J. P., Blagburn, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ectopic+sensory+neurons+in+mutant+cockroaches+compete+with+normal+cells+for+central+targets.">Ectopic sensory neurons in mutant cockroaches compete with normal cells for central targets.</a> Development. 115, 773-784 (1992).</li><li>Blagburn, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Co-factors+and+co-repressors+of+Engrailed:+expression+in+the+central+nervous+system+and+cerci+of+the+cockroach,+Periplaneta+americana.">Co-factors and co-repressors of Engrailed: expression in the central nervous system and cerci of the cockroach, Periplaneta americana.</a> Cell Tiss. Res. 327, 177-187 (2007).</li><li>Blagburn, J. M., Gibbon, C. R., Bacon, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+engrailed+in+an+array+of+identified+sensory+neurons:+comparison+with+position,+axonal+arborization,+and+synaptic+connectivity.">Expression of engrailed in an array of identified sensory neurons: comparison with position, axonal arborization, and synaptic connectivity.</a> J. Neurobiol. 28, 493-505 (1995).</li><li>Booth, D., Marie, B., Domenici, P., Blagburn, J. M., Bacon, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptional+control+of+behavior:+engrailed+knock-out+changes+cockroach+escape+trajectories.">Transcriptional control of behavior: engrailed knock-out changes cockroach escape trajectories.</a> J. Neurosci. 29, 7181-7190 (2009).</li><li>Schrader, S., Horseman, G., Cokl, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directional+sensitivity+of+wind-sensitive+giant+interneurons+in+the+cave+cricket+Troglophilus+neglectus.">Directional sensitivity of wind-sensitive giant interneurons in the cave cricket Troglophilus neglectus.</a> J. Exp. Zool. 292, 73-81 (2002).</li><li>Libersat, F., Goldstein, R. S., Camhi, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonsynaptic+regulation+of+sensory+activity+during+movement+in+cockroaches.">Nonsynaptic regulation of sensory activity during movement in cockroaches.</a> Proc. Natl. Acad. Sci. U.S.A. 84, 8150-8154 (1987).</li><li>Stern, M., Ediger, V. L., Gibbon, C. R., Blagburn, J. M., Bacon, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regeneration+of+cercal+filiform+hair+sensory+neurons+in+the+first-instar+cockroach+restores+escape+behavior.">Regeneration of cercal filiform hair sensory neurons in the first-instar cockroach restores escape behavior.</a> J. Neurobiol. 33, 439-458 (1997).</li><li>Blagburn, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+specificity+in+the+first+instar+cockroach:+patterns+of+monosynaptic+input+from+filiform+hair+afferents+to+giant+interneurons.">Synaptic specificity in the first instar cockroach: patterns of monosynaptic input from filiform hair afferents to giant interneurons.</a> J. Comp. Physiol A. 166, 133-142 (1989).</li><li>Watanabe, H., Ai, H., Yokohari, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatio-temporal+activity+patterns+of+odor-induced+synchronized+potentials+revealed+by+voltage-sensitive+dye+imaging+and+intracellular+recording+in+the+antennal+lobe+of+the+cockroach.">Spatio-temporal activity patterns of odor-induced synchronized potentials revealed by voltage-sensitive dye imaging and intracellular recording in the antennal lobe of the cockroach.</a> Front. Sys. Neurosci. (6), 55 (2012).</li><li>Nishino, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visual+and+olfactory+input+segregation+in+the+mushroom+body+calyces+in+a+basal+neopteran,+the+American+cockroach.">Visual and olfactory input segregation in the mushroom body calyces in a basal neopteran, the American cockroach.</a> Arthropod Struct. Dev. 41, 3-16 (2012).</li><li>Elia, A. J., Gardner, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+effects+of+DDT+on+the+behavior+and+central+nervous+system+activity+in+Periplaneta+americana.">Long-term effects of DDT on the behavior and central nervous system activity in Periplaneta americana.</a> Pestic. Biochem. Physiol. 21, 326-335 (1984).</li></ol>

The authors declare that there are no conflicts of interest.

One of the reasons for exhibiting techniques for this classical preparation is that the cerci system has been and still is an active area of research in addressing questions of the development of neural circuitry as well as questions regarding synaptic repair and regeneration 26-31. Either method of evoking activity in the cockroach ventral nerve cord can be used to examine the effects of pharmacological agents or insecticides on nervous system function. These experiments are done by simply dissolving neuroactive chemicals into saline. After exchanging this solution with the normal bathing medium, changes in evoked or spontaneous activity may be observed while recording from connectives or a motor nerve to give a consistent readout of the chemical&#39;s effect on CNS function.
As in all neurophysiological experiments a common problem is electrical noise. Probably the biggest factor in signal quality for these preparations is the suction electrode seal on the nerve tissue. A tight seal that does not completely draw in the cercal nerve or connective is ideal. Recordings can also be made with dual hook electrodes placed under the nerve cord and insulating the VNC with a mixture of mineral oil and petrolatum. The mixture can be loaded into a syringe and expelled around the nerve cord32. Also a careful dissection is as critical here as in any CNS preparation. Some may find it easier to access the CNS by dissecting the dorsal cuticle. While this reduces the possibility of damaging the ventral nerve cord it can be more difficult to remove all of the viscera using this approach.
It is not described here but this preparation is amenable to intracellular recording in the giant interneurons 32,33. The entire nerve cord can also be removed to accommodate several recording and stimulating electrodes simultaneously. In fact exploration of the antennal lobe, mushroom body, and other anterior CNS structures is still in progress34-35. While the cockroach CNS continues to shed light on modern neurobiological research this particular preparation is simple enough to be used in undergraduate academic laboratories.

There are more than 4,000 cockroach species but only about 30 are household pests. Perhaps the most recognized is the misnamed American cockroach Periplaneta americana which originated in Africa, and is now found nearly everywhere on the planet. In addition to its rapid running speed1 and evasive behavior, in the tropics P. americana is capable of flight2,3.
The predominant characteristics of the cockroach central nervous system (CNS) are its segmented nature and decentralization of control processes4,5. The brain, thoracic, and abdominal ganglia are joined together by paired interganglionic connectives to form the ventral nerve cord (VNC).
The ganglia at each segment are integrating centers. They are composed of an outer, cortical region containing cells responsible for the blood-brain permeability barrier just beneath them, and the somata of neurons originating in that ganglion. These somata may belong to interneurons, modulatory neurons, or motor neurons. They supply axons that remain within the ganglion of origin (local interneuron), or axons that project between the ganglia of the CNS (interganglionic interneurons) or that terminate on peripheral muscle cells (motor neurons). Most somata are positioned ventrally or ventrolaterally in the ganglionic cortex5. The paired, interganglionic connectives contain only axons and no neuronal cell bodies.
The neuropil of a ganglion contains glial cells (neuroglia), axon tracts, bundles of axons, and dendrites (neurites) of neurons. The neuropil is devoid of neuronal cell bodies. This is the region within the ganglion where direct synaptic communication among nerve cells and integration of inputs occur.
The ability of the American cockroach P. americana to detect and suddenly respond to an approaching predator (foot, hand, etc.) has been attributed to a reflex circuit that consists of the cerci and giant fiber system6,7. The cerci are a pair of horn-like, wind-sensitive structures located on the end of the abdomen (Figure 1). In P. americana the ventral surface of each cercus contains about 200 filiform (thread) hairs that are organized into 14 columns. Nine of these columns can be consistently identified in different animals according to the response properties of the associated receptor cell and axon. Each hair is in a socket that allows it to bend most readily in one plane that is column specific. Movement of the hair in one direction along its plane induces a depolarization in the receptor cell and a burst of action potentials (APs) in the sensory neuron. Movement in the opposite direction inhibits any ongoing spontaneous APs8. The preferred plane of deflection and directionality of the response is different in each column. Thus, the filiform hair-receptor complexes are responsible not only for detecting the movement of air but also for &#39;coding&#39;, in the form of APs, the direction from which the air current originated. Processing of this information by the CNS results in an &#39;appropriate&#39; escape response6,7. This functional, columnar specificity of the sensory hairs is preserved from animal to animal.
The receptor cell of each filiform hair is responsible for transducing the mechanical deflection of the hair into a neural event (resulting in a burst or inhibition of APs in the receptor cell&#39;s axon9. The APs travel to the terminal abdominal ganglion (A6) via cercal nerve XI, where they synapse with giant axons of the ventral nerve cord (VNC). The giant axons are believed to be responsible for the transmission and subsequent excitation of motor neurons that results in an escape behavior6,10,11.
The behavioral latency of the escape response of P. americana is one of the shortest of any animal7. Behavioral latency is the time between the arrival of a stimulus at a mechanoreceptor and the initiation of an escape response. In experiments using high speed cinematography to record the attempted escape from an attacking toad, the cockroach was observed to begin its turn away from the toad in about 40 msec (time from beginning of tongue extension to cockroach movement7,12. Using controlled wind puffs, the behavioral latency could be reduced to 11 msec. Other experiments revealed that a minimum wind puff velocity of 12 mm/msec&#160;(with an acceleration of 600 mm/msec2) can evoke an escape response, while even lower velocities (3 mm/sec) caused slowly walking cockroaches to stop moving12.
The strong correlation that typically exists between giant fiber systems and escape behavior has been well documented13,14. In instances where a particular cell is necessary and sufficient to evoke a particular behavior the cell is referred to as a command neuron15,16. Giant interneurons (GIs) in the wind escape circuit of P. americana are not necessary for the reflex. Animals that have experimentally ablated GIs still exhibit the escape behavior therefore these GIs are not considered command neurons17,18. Severing cervical connectives that are rostral to the sensorimotor circuit also influences the behavior, indicating that descending input from the brain has an effect on the direction of escape19. These aspects of fine control and redundancy are paramount to the organism&#39;s survival and are complemented by neurochemical modulation via biogenic amines20.
The P. americana&#160;nerve cord preparation has been an elegant model system for neuroethologists over the past many decades starting with the pioneering work of Roeder21. It permits students to record, display and analyze primary sensory activity and the resultant responses by giant interneurons to their input22,23,24. In addition to conveying the idea that identifiable neural circuits underlie behavioral responses to the environment, these exercises should instill an appreciation for the biological contributions made by this common household pest.

<table border="1">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Sylgard</td>
			<td>Dow Corning</td>
			<td>182 silicone kit</td>
			<td>182 silicone elastomer kit</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma-Aldrich</td>
			<td>P9333</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>C5670</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4&#8226;2H2O</td>
			<td>Sigma-Aldrich</td>
			<td>71505</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4&#8226;7H2O</td>
			<td>Sigma-Aldrich</td>
			<td>S9390</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma-Aldrich</td>
			<td>221465</td>
			<td>To adjust pH</td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>Sigma-Aldrich</td>
			<td>H1758</td>
			<td>To adjust pH</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Material Name</td>
		</tr>
		<tr>
			<td>Dissecting tools</td>
			<td>World Precision Instruments</td>
			<td>assortment</td>
			<td></td>
		</tr>
		<tr>
			<td>Insect Pins</td>
			<td>Fine Science Tools, Inc</td>
			<td>26001-60</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>World Precision Instruments</td>
			<td>PZMIII-BS</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass electrodes</td>
			<td>Sigma-Aldrich</td>
			<td>CLS7095B5X</td>
			<td>Box of 200, suction electrodes</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>MD4-M3-R</td>
			<td>Can fix for base or on a metal rod</td>
		</tr>
		<tr>
			<td>Silver wire (10/1,000 inch)</td>
			<td>A-M Systems</td>
			<td>782500</td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>any company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AC/DC differential amplifier</td>
			<td>A-M Systems</td>
			<td>Model 3000</td>
			<td></td>
		</tr>
		<tr>
			<td>PowerLab 26T</td>
			<td>AD Instruments</td>
			<td>27T</td>
			<td></td>
		</tr>
		<tr>
			<td>Head stage</td>
			<td>AD Instruments</td>
			<td></td>
			<td>Comes with AC/DC amplifier</td>
		</tr>
		<tr>
			<td>LabChart7</td>
			<td>AD Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical leads</td>
			<td>any company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass tools</td>
			<td>make yourself</td>
			<td></td>
			<td>For manipulating nerves</td>
		</tr>
		<tr>
			<td>Cable and connectors</td>
			<td>any company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes with bulbs</td>
			<td>Fisher Scientific</td>
			<td>13-711-7</td>
			<td>Box of 500</td>
		</tr>
		<tr>
			<td>Beakers</td>
			<td>any company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Wax or modeling clay</td>
			<td>any company or local stores</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stimulator</td>
			<td>Grass Instruments</td>
			<td>SD9 or S88</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic tip for suction electrode</td>
			<td>local hardware store (Watt&#39;s brand)</td>
			<td>&#188; inch OD x 0.170 inch ID</td>
			<td>Cut in small pieces. Pull out over a flame and cut back the tip to the correct size.</td>
		</tr>
	</tbody>
</table>

neural circuit recording from an intact cockroach nervous system

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological effects of insecticides. This article describes the scope of cockroach sensory modalities that can be used to assay how an insect nervous system responds to environmental perturbations. Emphasis here is on the escape behavior mediated by cerci to giant fiber transmission in Periplaneta americana. This in situ preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of neuronal activity. Peptides or other chemical reagents can then be applied directly to the nervous system in solution with the physiological saline. Insecticides could also be administered prior to dissection and the escape circuit can serve as a proxy for the excitable state of the central nervous system. In this context the assays described herein would also be useful to researchers interested in limb regeneration and the evolution of nervous system development for which P. americana is an established model organism.

Stimulation of hairs on the cerci by a puff of air causes discharges of primary sensory neurons that can be recorded using extracellular suction electrodes attached either to connectives between abdominal ganglia or the cercal nerve itself (Figure 8). Spike amplitudes recorded from the two regions are on the order of several micro-volts to millivolts.&#160;Because of sensory integration in the ganglion the number of spikes observed in the compound action potential or as individual spikes recorded from the cercal nerve is remarkably greater than observed in recordings from the connectives. However also note that there is substantially less noise in the recording at the connective due to the tighter seal between the electrode and the nervous tissue.
By puffing air at the cerci large spikes can be observed in the connectives (Figure 8A). Using this stimulating method, recordings between A3 and A4 typically show a large spike characteristic of the giant interneuron(s). Recording from a cercal nerve while physically rubbing the cerci with forceps produced a strong bust of activity (Figure 8B1). In another recording, 2 puffs of air each produced a rapid busting response in the cercal nerve (Figure 8B2). When electrically stimulating the cercal nerve with a suction electrode and recording in the connective between A3 and A4, one can observe a threshold in the stimulation to evoked responses (Figure 8C1). The electrical stimulation of the cercal nerve clearly elicits a response in connectives which can be quantified for manipulative studies with pharmacological agents or the local environmental surrounds, such as temperature (Figure 8C2).
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50584/50584fig1.jpg" /> 
Figure 1. Periplaneta americana with intact cerci.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50584/50584fig2.jpg" /> 
Figure 2. Ventral view of cockroach nerve cord as seen with the ventral cuticle removed (A). An enlarged view of the segment outlined by arrows is seen in (B). In (C) the connectives were spilt between A4 and A3 with a glass probe. The 6th abdominal ganglion is shown in (D) with the two cercal nerves leaving at the caudal end.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/50584/50584fig3.jpg" /> 
Figure 3. Schematic ventral view of cockroach nerve cord.
<img alt="Figure 4" src="/files/ftp_upload/50584/50584fig4.jpg" /> 
Figure 4. The cerci are positioned upwards out of the saline bath. The opened abdomen can be flooded with saline (A) with the caudal end of the roach being elevated with a small wedged shaped piece of silicone elastomer in order to keep the cerci out of the bath (B).
<img alt="Figure 5" src="/files/ftp_upload/50584/50584fig5.jpg" /> 
Figure 5. The 6th abdominal ganglion with the cercal nerve (outlined by arrows).
<img alt="Figure 6" fo:content-width="6in" src="/files/ftp_upload/50584/50584fig6.jpg" /> 
Figure 6. The equipment set up. <a href="https://www.jove.com/files/ftp_upload/50584/50584fig6large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 7" src="/files/ftp_upload/50584/50584fig7.jpg" /> 
Figure 7. Stimulating and recording electrode set up.
<img alt="Figure 8" fo:content-width="5.75in" src="/files/ftp_upload/50584/50584fig8.jpg" /> 
Figure 8. Neural recordings of the connectives and cercal nerve with various stimulation procedures. Recording with a suction electrode from the connectives between A3 and A4 while puffing air at the cerci (A). Recording from the primary cercal neurons with a suction electrode while either physically rubbing (B1) or providing air puffs (B2) results in rapid bursts of activity in the cercal nerve. Electrically stimulating the cercal nerve produces responses in connectives (C1). Note the gradual increase in the stimulating intensity (arrows indicate the amplitude of the stimulating artifact) and the intensity of the following evoked responses. The electrical stimulation of the cercal nerve provides a relatively more controlled means of stimulating the cercal nerve for consistency in stimulation for quantifying the responses (C2).

Watch this Scientific Journal Video about Neural Circuit Recording from an Intact Cockroach Nervous System at JoVE.com

Neural Circuit Recording from an Intact Cockroach Nervous System

This article describes the cockroach ventral nerve cord dissection and extracellular recordings from the cercal nerve and connectives. Evoked responses are generated by electrical stimulation of the cercal nerve or direct mechanical stimulation of the cerci.

The cockroach ventral nerve cord preparation is a tractable system for neuroethology experiments, neural network modeling, and testing the physiological ...

neural-circuit-recording-from-an-intact-cockroach-nervous-system

Research

JoVE Journal

Neuroscience

32.2K visualizações.  University of Kentucky.  Este artigo descreve a barata ventral do nervo espinhal dissecção e gravações extracelular do nervo Cercal e conectivos. Respostas evocadas são gerados por estimulação elétrica do nervo Cercal ou estimulação mecânica direta da Cerci. 

Vídeo: Neural Circuit Recording from an Intact Cockroach Nervous System

Revealing Neural Circuit Topography in Multi-Color

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably label the global patterning of multiple topographic projections. The cerebellum is an ideal model to study the orderly arrangement of neural circuits. For example, the compartmental organization of spinocerebellar mossy fibers has proven to be an indispensable system for studying mossy fiber patterning. We recently showed that wheat germ agglutinin (WGA) conjugated to Alexa 555 and 488 can be used for tracing spinocerebellar mossy fiber projections in developing and adult mice (Reeber et al. 2011). We found three major properties that make the WGA-Alexa tracers desirable tools for labeling neural projections. First, Alexa fluorophores are intense and their brightness allows for wholemount imaging directly after tracing. Second, WGA-Alexa tracers label the entire trajectory of developing and adult neural projections. Third, WGA-Alexa tracers are rapidly transported in both retrograde and anterograde directions. Here, we describe in detail how to prepare the tracers and other required tools, how to perform the surgery for spinocerebellar tracing and how best to image traced projections in three dimensions. In summary, we provide a step-by-step tracing protocol that will be useful for deciphering the organization and connectivity of functional maps not only in the cerebellum but also in the cortex, brainstem, and spinal cord.

We provide a practical guide for delivering tracers in vivo and use the spinocerebellar pathway as a model system to demonstrate essential steps for successful neuronal circuit analysis in mice. We describe in detail our versatile tracing protocol that exploits wheat germ agglutinin (WGA) conjugated to Alexa fluorophores.

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably ...

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Anatomically Inspired Three-dimensional Micro-tissue Engineered Neural Networks for Nervous System Reconstruction, Modulation, and Modeling

Functional recovery rarely occurs&#160;following injury or disease-induced degeneration within the central nervous system (CNS)&#160;due to the inhibitory environment and the limited capacity for neurogenesis. We are developing&#160;a strategy to simultaneously address&#160;neuronal and axonal pathway loss within the damaged CNS. This manuscript presents the fabrication protocol for micro-tissue engineered neural networks (micro-TENNs), implantable constructs consisting of neurons and aligned axonal tracts spanning the extracellular matrix (ECM) lumen of a preformed hydrogel cylinder hundreds of microns in diameter that may extend centimeters in length. Neuronal aggregates are delimited to the extremes of the three-dimensional encasement and are spanned by axonal projections. Micro-TENNs are uniquely poised as a strategy for CNS reconstruction, emulating aspects of brain connectome cytoarchitecture and potentially providing means for network replacement. The neuronal aggregates may synapse with host tissue to form new functional relays to restore and/or modulate missing or damaged circuitry. These constructs may also act as pro-regenerative&#160;&#34;living scaffolds&#34; capable of exploiting developmental mechanisms for cell migration and axonal pathfinding, providing synergistic structural and soluble cues based on the state of regeneration. Micro-TENNs are fabricated by pouring liquid hydrogel into a cylindrical mold containing a longitudinally centered needle. Once the hydrogel has gelled, the needle is removed, leaving a hollow micro-column. An ECM solution is added to the lumen to provide an environment suitable for neuronal adhesion and axonal outgrowth. Dissociated neurons are mechanically aggregated for precise seeding within one or both ends of the micro-column. This methodology reliably produces self-contained miniature constructs with long-projecting axonal tracts that may recapitulate features of brain neuroanatomy. Synaptic immunolabeling and genetically encoded calcium indicators suggest that micro-TENNs possess extensive synaptic distribution and intrinsic electrical activity. Consequently, micro-TENNs represent a promising strategy for targeted neurosurgical reconstruction of brain pathways and may also be applied as biofidelic models to study neurobiological phenomena in vitro.

This manuscript details the fabrication of micro-tissue engineered neural networks: three-dimensional micron-sized constructs comprised of long aligned axonal tracts spanning aggregated neuronal population(s) encased in a tubular hydrogel. These living scaffolds can serve as functional relays to reconstruct or modulate neural circuitry or as biofidelic test-beds mimicking gray-white matter neuroanatomy.

Functional recovery rarely occurs&#160;following injury or disease-induced degeneration within the central nervous system (CNS)&#160;due to the inhibitory ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

Transplantation of Chemogenetically Engineered Cortical Interneuron Progenitors into Early Postnatal Mouse Brains

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is a complicated task, which is particularly difficult in regards to the development of &#947;-aminobutyric acid (GABA)ergic cortical interneurons (CIs). CIs are the main inhibitory neurons in the cerebral cortex, and they play key roles in neuronal networks, by regulating both the activity of individual pyramidal neurons, as well as the oscillatory behavior of neuronal ensembles. They are generated in transient embryonic structures (medial and caudal ganglionic eminences -&#160;MGE and CGE) that are very difficult to efficiently target using in utero electroporation approaches. Interneuron progenitors migrate long distances during normal embryonic development, before they integrate in the cortical circuit. This remarkable ability to disperse and integrate into a developing network can be hijacked by transplanting embryonic interneuron precursors into early post-natal host cortices. Here, we present a protocol that allows genetic modification of embryonic interneuron progenitors using focal ex vivo electroporation. These engineered interneuron precursors are then transplanted into early post-natal host cortices, where they will mature into easily identifiable CIs. This protocol allows the use of multiple genetically encoded tools, or the ability to regulate the expression of specific genes in interneuron progenitors, in order to investigate the impact of either genetic or environmental variables on the maturation and integration of CIs.

Here we present a protocol, designed to use chemogenetic tools to manipulate the activity of cortical interneuron progenitors transplanted into the cortex of early postnatal mice.

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Dissection and Isolation of Murine Glia from Multiple Central Nervous System Regions

The methods presented here demonstrate laboratory procedures for the dissection of four different regions of the central nervous system (CNS) from murine neonates for the isolation of glial subpopulations. The purpose of the procedure is to dissociate microglia, oligodendrocyte progenitor cells (OPCs), and astrocytes from cortical, cerebellar, brainstem, and spinal cord tissue to facilitate further in vitro analysis. The CNS region isolation procedures allow for the determination of regional heterogeneity among glia in multiple cell culture systems. Rapid CNS region isolation is performed, followed by the mechanical removal of meninges to prevent meningeal cell contamination of glia. This protocol combines gentle tissue dissociation and plating on a specified matrix designed to preserve cell integrity and adherence. Isolating mixed glia from multiple CNS regions provides a comprehensive analysis of potentially heterogenous glia while maximizing the use of individual experimental animals. Additionally, following dissociation of regional tissue, mixed glia are further divided into multiple cell types including microglia, OPCs, and astrocytes for use in either single cell type, cell culture plate inserts, or co-culture systems. Overall, the demonstrated techniques provide a comprehensive protocol of broad applicability for careful dissection of four individual CNS regions from murine neonates and includes methods for the isolation of three individual glia cell types to examine regional heterogeneity in any number of in vitro cell culture systems or assays.

Here we present a protocol for in vitro isolation of multiple glial cell populations from a mouse CNS. This method allows for the segregation of regional microglia, oligodendrocyte precursor cells, and astrocytes to study the phenotypes of each in a variety of culture systems.

The methods presented here demonstrate laboratory procedures for the dissection of four different regions of the central nervous system (CNS) from murine ...