This work has been supported by NINDS, RO1NS52699 and MH84874 to SA. 
We would like to thank Dr. Dave Featherstone (Department of Biological Sciences, University for Illinois at Chicago) for providing us with the suture glue used in the immunohistochemistry work. We thank Michael Alpert for his comments and proofreading of the manuscript.

1. Preparation of Poly-D-lysine Hydrobromide
<ol>
	<li>Prepare 1 mg/ml poly-D-lysine hydrobromide in 0.1 M borate buffer (pH 8.5).</li>
	<li>Aliquot and store at -20 &#176;C.</li>
</ol>
2. Poly-lysine Coating of Coverslips
Note: Carry out all cleaning and coating steps in a laminar flow chamber.
<ol>
	<li>Place coverslips in a Petri dish containing 1 N Hydrochloric acid (HCl) for 2 hr.</li>
	<li>Aspirate all HCl and rinse with 70% Ethanol (EtOH) 2-3x.</li>
	<li>Leave in 70% EtOH for 1 hr.</li>
	<li>Aspirate all 70% EtOH and rinse with 100% EtOH 2-3x.</li>
	<li>Leave in 100% EtOH for 2 hr. Aspirate all EtOH.</li>
	<li>Blot dry with filter paper and air dry for a few seconds.</li>
	<li>Place O/N in 1 mg/ml poly-lysine solution.</li>
	<li>Rinse coverslips next day with Millipore water 4-5x.</li>
	<li>Air dry poly-lysine coated coverslips on glass rollers in a clean Petri dish.</li>
	<li>For immunohistochemistry, use 35 x 10 mm Petri dish lids. Prepare sylgard (polydimethylsilioxane, PDMS) lined dishes by pouring PDMS (elastomer and curing agent 10:1 by weight) into the dish to a thickness of 0.3 cm and allow to set at 30 &#176;C O/N. Once this has set, cut a 2 cm by 1 cm inset into the PDMS. Clean and coat the inset with poly-lysine following the same steps as mentioned above for coverslips.</li>
	<li>Store poly-lysine coated coverslips and dishes covered inside the laminar flow chamber until use (up to 2 weeks, but achieve best adhesive results by preparing periodically every 3-4 days).</li>
	<li>Prepare a PDMS block, to pin the spinal cord in the vibrating tissue slicer. Mix Elastomer A and Elastomer B 1:1 by weight. Pour into a 100 x 15 mm Petri dish and allow to set O/N at 30 &#176;C. Cut a rectangular piece of the PMDS and glue it to the slicing base plate using epoxy glue. Allow the glue to set fixing the PDMS in place and slice multiple thin sections off the surface to obtain a flat surface to pin the tissue on.</li>
</ol>
3. Acute Dissociation of Lamprey Spinal Cord to Yield Isolated Reticulospinal Axons
<ol>
	<li>Anesthetize an ammoecoete or adult lamprey (Petromyzon marinus) with tricaine methanesulphonate (MS-222; 100 mg/L). Add anesthetic into the water, in a plastic cup covered by a lid, containing the lamprey to be sacrificed.</li>
	<li>Decapitate lamprey in cold (4 &#176;C) Ringer&#8217;s solution of the following composition (in mM): 130 NaCl, 2.1 KCl, 2.6 CaCl2, 1.8 MgCl2, 4 HEPES, 4 dextrose (pH 7.6, osmolarity 270 mOsm) and remove body wall muscles to expose dorsal surface of spinal cord.</li>
	<li>Remove meninx primitiva from the dorsal surface of the spinal cord using fine forceps. Do not remove ventral meninx primitiva at this stage.</li>
	<li>Cut spinal cord into 1 cm long pieces.</li>
	<li>Pin a spinal cord piece using fine insect pins, dorsal side facing up, on a PDMS lined slicing base plate (see 2.12) in a vibrating tissue slicer chamber containing ice-cold Ringer&#8217;s solution.</li>
	<li>Remove a central section of the dorsal column of the spinal cord by slicing along the rostro-caudal axis with the blade, leaving behind intact dorsal column sections on the rostral and caudal ends to serve as handles during the dissociation process Figure 2a&#160;and Figure 2b. Use slowest speed setting that permits slicing and a depth setting that removes only the dorsal column leaving the underlying reticulospinal axon column undamaged Figure 2b.</li>
	<li>Incubate sliced spinal cord pieces at RT for 45 min&#160;in a cocktail of 1 mg/ml protease (Type XIV from Streptomyces griseus) and 1 mg/ml collagenase prepared (Type IA from Clostridium histolyticum) in Ringer&#8217;s solution (adapted from El Manira &#38; Bussi&#232;res, 1997 17).</li>
	<li>Pin enzyme-treated spinal cord pieces using fine pins in a PDMS lined Petri dish containing cold (4 &#176;C) Ringer&#8217;s solution.</li>
	<li>Remove ventral meninx primitiva with fine forceps.</li>
	<li>Cut lateral tracts of spinal cord with a scalpel blade at the midpoint of the sliced dorsal section leaving the central column of reticulospinal axons intact in the spinal cord Figure 2d.</li>
	<li>Place a drop of immersion oil on the lens.</li>
	<li>Place the poly-lysine coated coverslip (Figure 3a, top piece, red rectangle) in the slot of the recording chamber Figure 3 and apply vacuum grease to all edges (space between red and blue rectangles in Figure 3a, to facilitate a seal. Screw the top into place. Place the chamber in the inset on the recording rig.</li>
	<li>Add Ringer&#8217;s solution into the recording chamber using a Pasteur pipette.</li>
	<li>Connect the outflow tubing of the pressure bottle containing antifreeze solution to the thermoelectric cooling device input tubing. Connect the output tubing of the thermoelectric cooling device to the input end of the outer cooling jacket and the output end to the reservoir Figure 3b.</li>
	<li>Place one piece of spinal cord at a time in the recording chamber and gently separate the spinal cord maintaining it along the coverslip at all times using Teflon coated forceps until axons are isolated Figure 2e&#160;and Figure 2f.</li>
	<li>Bring the recording solution temperature to 10 &#176;C by passing the pressurized (nitrogen gas is pushed into the bottle) antifreeze solution (by displacement), via the thermoelectric cooling device, through the outer cooling jacket of the recording chamber Figure 3b.</li>
	<li>Allow axons to recover for 1 hr post dissociation at 10 &#176;C.</li>
</ol>
<img alt="Figure 2" fo:content-width="4in" src="/files/ftp_upload/51925/51925fig2highres.jpg" width="400" /> 
Figure&#160;2. Schematic description of dissociation protocol for isolation of reticulospinal axons. (a) Removal of dorsal column. The arrow indicates the direction of slicing of the tissue. The dorsal horns are marked by the alphabet D (red font color), while the ventral horns by the alphabet V (red font color). (b) Dorsal column removed in the central portion of the spinal cord exposing the reticulospinal axons (green lines). The ventral horns, which remain intact after the slicing process, are marked by the alphabet V (red font color). (c) 45 min&#160;treatment with protease and collagenase enzymes cocktail (1 mg/ml). (d) Cutting of lateral tracts of the spinal cord; indicating position, direction and extent of lateral cut. (e) Mechanical dissociation of spinal cord. Arrows indicate position of forceps and direction of separation force during dissociation. (f) Representative example of dissociated reticulospinal axon preparation. Green arrows mark regions of acutely dissociated reticulospinal axons without any postsynaptic processes.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51925/51925fig3highres.jpg" width="500" /> 
Figure&#160;3. Schematic of recording chamber for electrophysiology experiments. Dimensions provided are in inches. Red rectangle shown in basepiece diagram (a)&#160;indicates positioning of coverslip in the coverslip groove in the basepiece. The region between the red and blue rectangle, shown in the basepiece diagram (a)&#160;is where the high vacuum grease is applied. (b) shows the assembled recording chamber.
4. Labeling and Identification of Presynaptic Terminals with FM 1-43
<ol>
	<li>Label presynaptic terminals, by incorporating FM 1-43 into vesicles during synaptic exo-endocytosis during high K+ depolarization. Perfuse preparation with 5 &#181;M FM 1-43 (in 5 ml Ringer&#8217;s solution containing 30 mM KCl).</li>
	<li>Perfuse preparation with 1 mg/ml Advasep-7 (in 5 ml Ringer&#8217;s solution) to remove excess FM dye)18.</li>
	<li>Perfuse preparation with Ringer&#8217;s solution for 15 min&#160;to washout any remant dye and the Advasep.</li>
	<li>Image with 100 X oil immersion lens (NA 1.25) on an inverted fluorescence microscope, in conjunction with a digital CCD camera and image acquisition software Micromanager, using standard fluorescence imaging protocols.</li>
	<li>Identify fluorescently labeled presynaptic terminals to target for recording Figure 4c&#160;and Figure 4d.</li>
</ol>
5. Immunohistochemistry of Isolated Reticulospinal Axons
<ol>
	<li>Fill dish inset (Protocol section 2.10.) with divalent-ion (Ca2+ and Mg2+) free Ringer&#8217;s solution.</li>
	<li>Fabricate patch pipettes in a P-87 micropipette puller. Place a small amount of suture glue at one end of the poly-lysine inset using a patch pipette (1.5 mm outer diameter glass) and suction (using a silicone tubing 0.89 mm inner diameter with a micropipette tip attached to the suction end).</li>
	<li>Carry out dissociations using the same procedure as mentioned in protocol section 3 Figure 2. Place one end of the spinal cord on the suture glue and press gently with forceps to adhere it strongly to the surface. Place another drop of suture glue at the other end of the inset and gently stretch the spinal cord until axons are dissociated and adhere the free spinal cord end by dragging it gently over the suture glue. Fix in place by gentle pressing down with the forceps.</li>
	<li>Exchange divalent free Ringer&#8217;s solution with regular Ringer&#8217;s solution by perfusion.</li>
	<li>Allow axons to recover for 20 min&#160;post dissociation at 10 &#176;C.</li>
	<li>Fix dissociated axons in 4% paraformaldehyde (PFA) {prepared in Phosphate Buffer Saline (PBS, (mM) NaCl 137,&#160;KCl 2.7, Na2HPO4 10, KH2PO4 1.8, pH 7.4)} for 20 min. Filter PFA solution prior to use by passing through a 0.2 &#181;M syringe filter.</li>
	<li>Wash out the PFA by perfusing 0.1 M glycine (in PBS) for 10 min.</li>
	<li>Incubate in 0.1% Triton-X (in PBS) for 10 min.</li>
	<li>Wash by perfusing PBS for 20 min.</li>
	<li>Block with 5% non-fat milk (in PBS) for 6 hr at 4 &#176;C.</li>
	<li>Add in primary antibody to VGCC of interest (1:200 dilution in PBS) and incubate for 20 hr at 4 &#176;C.</li>
	<li>Wash by perfusing PBS for 20 min.</li>
	<li>Block with 5% non-fat milk (in PBS) for 10 min&#160;at 4 &#176;C.</li>
	<li>Add in secondary antibody (1:400 dilution in PBS) and incubate in dark for 2 hr at 4 &#176;C.</li>
	<li>Wash by perfusing with PBS for 20 min.</li>
	<li>Block with 1% Bovine Serum Albumin (in PBS) for 20 min.</li>
	<li>Add in Alexa Fluor 488 phalloidin (5 Units/&#181;l working concentration; stock prepared in methanol 200 Units/ml).</li>
	<li>Wash by perfusing with PBS for 20 min.</li>
	<li>Image using 100X water immersion lens on confocal microscope.</li>
</ol>
6. Electrophysiological Recording
<ol>
	<li>Fabricate aluminosilicate glass patch pipettes (pipette resistance 2-5 M&#937;) in a P-87 micropipette puller. Design patch pipette such that pipette tip encompasses entire presynaptic terminal diameter.</li>
	<li>PDMS coat patch pipettes by dipping the patch pipette under 50-60 psi pressure into PDMS23 (attach a silicone tubing, 0.89 mm inner diameter, connected to a nitrogen gas cylinder, to the back end of the patch pipette) and dry using a heat gun. Alternately, manually coat the pipette with PDMS, applying coating as close to the tip as possible, under a compound microscope.</li>
	<li>Fire polish patch pipettes using a microforge (a custom built platinum filament fitted onto the stage of a compound microscope).</li>
	<li>Identify isolated axons demonstrating labeled presynaptic terminals by fluorescence microscopy.</li>
	<li>Fill recording solution (designed to isolate Ca2+ currents; 10 mM CaCl2 or 90 mM BaCl2 as charge carrier, HEPES buffered pH 7.6, osmolarity 270 mOsm) into the patch pipette using a syringe.</li>
	<li>Insert patch pipette into the pipette holder and position above bath using a motorized manipulator MP225. Gently lower the patch pipette into the bath and position against the face of a fluorescently identified presynaptic terminal.</li>
	<li>Advance the patch pipette slowly until contact is made with the membrane. At this point, achieve a gigaohm seal by gentle mouth suction through a tube attached to the pipette holder.</li>
	<li>To achieve the extremely low background noise levels required for recording single channel Ca2+ currents, use an Axopatch 200B with a cooled headstage for the recordings. Sample data at 20-50 kHz and filter using a 5 kHz Bessel filter. Carry out data acquisition using an Axograph X.</li>
	<li>Use a standard step protocol, in increments of 10 mV as stimulus. Incorporate a pre-pulse in the protocol, preceding the step protocol, to ensure maximal activation of Ca2+ channels. Incorporate a 10 mV leak step into the step protocol for post-analysis subtraction of leak currents.</li>
</ol>

<ol>
	<li>Katz, B., Miledi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+timing+of+calcium+action+during+neuromuscular+transmission.">The timing of calcium action during neuromuscular transmission.</a> Journal of Physiology. 189 (3), 535-544 (1967).</li><li>Sabatini, B. L., Regehr, W. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Timing+of+neurotransmission+at+fast+synapses+in+the+mammalian+brain.">Timing of neurotransmission at fast synapses in the mammalian brain.</a> Nature. 384 (6605), 170-172 (1996).</li><li>Augustine, G. J., Adler, E. M., Charlton, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+calcium+signal+for+transmitter+secretion+from+presynaptic+nerve+terminals.">The calcium signal for transmitter secretion from presynaptic nerve terminals.</a> Annals of the New York Academy of Sciences. 635, 365-381 (1991).</li><li>Zucker, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+and+transmitter+release.">Calcium and transmitter release.</a> Journal of Physiology Paris. 87 (1), 25-36 (1993).</li><li>Delaney, K. R., Shahrezaei, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uncaging+Calcium+in+Neurons.">Uncaging Calcium in Neurons.</a> Cold Spring Harbor protocols. (12), (2013).</li><li>Paradiso, K., Wu, W., Wu, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+patch+clamp+capacitance+recordings+from+the+calyx.">Methods for patch clamp capacitance recordings from the calyx.</a> Journal of Visualized Experiments. (6), 244 (2007).</li><li>Stanley, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+calyx-type+synapse+of+the+chick+ciliary+ganglion+as+a+model+of+fast+cholinergic+transmission.">The calyx-type synapse of the chick ciliary ganglion as a model of fast cholinergic transmission.</a> Canadian Journal of Physiology and Pharmacology. 70, S73-S77 (1992).</li><li>Church, P. J., Stanley, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+L+type+calcium+channel+conductance+with+physiological+levels+of+calcium+in+chick+ciliary+ganglion+neurons.">Single L type calcium channel conductance with physiological levels of calcium in chick ciliary ganglion neurons.</a> The Journal of Physiology. 496 (Pt 1), 59-68 (1996).</li><li>Stanley, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+calcium+channels+and+acetylcholine+release+at+a+presynaptic+nerve+terminal.">Single calcium channels and acetylcholine release at a presynaptic nerve terminal.</a> Neuron. 11 (6), 1007-1011 (1993).</li><li>Weber, A. M., Wong, F. K., Tufford, A. R., Schlichter, L. C., Matveev, V., Stanley, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=N-type+Ca(2+)+channels+carry+the+largest+current+implications+for+nanodomains+and+transmitter+release.">N-type Ca(2+) channels carry the largest current implications for nanodomains and transmitter release.</a> Nature Neuroscience. 13 (11), 1348-1350 (2010).</li><li>He, L., Wu, X. S., Mohan, R., Wu, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+modes+of+fusion+pore+opening+revealed+by+cell+attached+recordings+at+a+synapse.">Two modes of fusion pore opening revealed by cell attached recordings at a synapse.</a> Nature. 444 (7115), 102-105 (2006).</li><li>Sheng, J., He, L., 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=Calcium+channel+number+critically+influences+synaptic+strength+and+plasticity+at+the+active+zone.">Calcium channel number critically influences synaptic strength and plasticity at the active zone.</a> Nature Neuroscience. 15 (7), 998-1006 (2012).</li><li>Photowala, H., Freed, R., Alford, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Location+and+function+of+vesicle+clusters,+active+zones+and+Ca2++channels+in+the+lamprey+presynaptic+terminal.">Location and function of vesicle clusters, active zones and Ca2+ channels in the lamprey presynaptic terminal.</a> The Journal of Physiology. 569. 569 (Pt 1), 119-135 (2005).</li><li>Rovainen, 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=Synaptic+interactions+of+reticulospinal+neurons+and+nerve+cells+in+the+spinal+cord+of+the+sea+lamprey.">Synaptic interactions of reticulospinal neurons and nerve cells in the spinal cord of the sea lamprey.</a> Journal of Comparative Neurology. 154 (2), 207-223 (1974).</li><li>Takahashi, M., Freed, R., Blackmer, T., Alford, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+influx+independent+depression+of+transmitter+release+by+5+HT+at+lamprey+spinal+cord+synapses.">Calcium influx independent depression of transmitter release by 5 HT at lamprey spinal cord synapses.</a> The Journal of Physiology. 532 (2), 323-336 (2001).</li><li>Stanley, E. F., Goping, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+a+calcium+current+in+a+vertebrate+cholinergic+presynaptic+nerve+terminal.">Characterization of a calcium current in a vertebrate cholinergic presynaptic nerve terminal.</a> The Journal of Neuroscience. 11 (4), 985-993 (1991).</li><li>El Manira, A., Bussi&#232;res, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+channel+subtypes+in+lamprey+sensory+and+motor+neurons.">Calcium channel subtypes in lamprey sensory and motor neurons.</a> Journal of Neurophysiology. 78 (3), 1334-1340 (1997).</li><li>Kay, A. R., Alfonso, A., 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=Imaging+synaptic+activity+in+intact+brain+and+slices+with+FM1+43+in+C+elegans+lamprey+and+rat.">Imaging synaptic activity in intact brain and slices with FM1 43 in C elegans lamprey and rat.</a> 24 (4), 809-817 (1999).</li><li>Betz, W. J., Bewick, G. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+analysis+of+synaptic+vesicle+recycling+at+the+frog+neuromuscular+junction.">Optical analysis of synaptic vesicle recycling at the frog neuromuscular junction.</a> Science. 255 (5041), 200-203 (1992).</li><li>Bleckert, A., Photowala, H., Alford, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+pools+of+actin+at+presynaptic+terminals.">Dual pools of actin at presynaptic terminals.</a> Journal of Neurophysiology. 107 (12), 3479-3492 (2012).</li><li>Gustafsson, J. S., Birinyi, A., Crum, J., Ellisman, M., Brodin, L., Shupliakov, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrastructural+organization+of+lamprey+reticulospinal+synapses+in+three+dimensions.">Ultrastructural organization of lamprey reticulospinal synapses in three dimensions.</a> The Journal of Comparative Neurology. 450 (2), 167-182 (2002).</li><li>Broadie, K., Featherstone, D. E., Chen, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+Recording+in+the+Drosophila+Embryo.">Electrophysiological Recording in the Drosophila Embryo.</a> Journal of Visualized Experiments. (27), (2009).</li><li>Rae, J. L., Levis, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+exceptionally+low+noise+single+channel+recordings.">A method for exceptionally low noise single channel recordings.</a> Pfl&#252;gers Archiv European Journal of Physiology. 420 (5-6), 618-620 (1992).</li></ol>

The authors do not have any competing financial interests or other conflicts of interest to disclose.

Our dissociation protocol is significant by yielding isolated reticulospinal axons devoid of postsynaptic projections Figure 2f, but which nevertheless retain functional presynaptic terminals Figure 4c&#160;and Figure 4d. The absence of postsynaptic processes opposing the presynaptic terminal permits direct recording access to the presynaptic release face membrane at single presynaptic terminals, previously not possible in central synapses and successfully achieved in on...

Synaptic transmission is an extremely rapid and precise process. Action potential invasion of the presynaptic terminal leads to opening of VGCCs located in the release face membrane, the resulting increase in presynaptic Ca2+ acting as the trigger for vesicle fusion and neurotransmitter release1. All of these steps occur within hundreds of microseconds2, and hence require tight spatial coupling of VGCCs to the vesicle fusion machinery3. Presynaptic Ca2+ fluxes have been primarily characterized through imaging approaches using Ca2+ sensitive dyes4. Incorporating Ca2+ buffers that modulate Ca2+ in presynaptic neurons has been used to indirectly characterize the relationship between presynaptic calcium and neurotransmission3. In addition, modulating the presynaptic free Ca2+&#160;concentration by uncaging Ca2+ 5 or recording macroscopic Ca2+ currents have been used in conjunction with measures of vesicle fusion and/or release; such as capacitance measurements6 or postsynaptic responses2 to address the same question. However, characterizing Ca2+ currents directly at the release face, the specialized section of the presynaptic membrane where membrane depolarization is translated into Ca2+&#160;currents triggering synaptic vesicle fusion and neurotransmitter release, is integral to obtaining a precise measure of the Ca2+ requirement for synaptic vesicle fusion. In addition, the ability to directly characterize Ca2+ currents at individual presynaptic terminals, coupled with accurate simultaneous measurements of vesicle fusion and release allows a precise elucidation of the timing relationship between the time course of the action potential, presynaptic Ca2+&#160;current, vesicle fusion and release. Access to the release face membrane is not available in the majority of presynaptic terminals due to close apposition by the postsynaptic dendrites. This inaccessibility has been a major obstacle in the characterization of VGCCs since it prevents direct measurements of current at individual presynaptic terminals. Direct characterization of presynaptic Ca2+ currents at individual presynaptic terminals has thus far not been possible in central synapses and has only been achieved in two calyceal type presynaptic terminals; calyx-type synapse of the chick ciliary ganglion7-10 and rat calyces11,12. In all other presynaptic terminals including the giant reticulospinal synapse in the lamprey spinal cord13, the lack of access to the presynaptic release face membrane has necessitated the use of indirect approaches such as Ca2+&#160;imaging to study presynaptic Ca2+&#160;fluxes.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51925/51925fig1highres.jpg" width="500" /> 
Figure&#160;1. Lamprey giant reticulospinal synapse. (a) Cross-section of lamprey spinal cord indicating dorso-ventral orientation. Reticulospinal axons are marked with green asterix. (b)&#160;3-D reconstruction of the reticulospinal synapse in the lamprey spinal cord showing presynaptic reticulospinal axon making numerous en passant contacts (marked by green arrows) onto the postsynaptic neuron13. Presynaptic terminals have been labeled with Alexa Fluor 488 hydrazide conjugated phalloidin (green), while the postsynaptic neuron has been filled with Alexa Fluor 568 hydrazide (red).
Lamprey giant reticulospinal axons, located in the ventral region of the spinal cord parallel to the rostral-caudal axis Figure 1a, form multiple enpassant synaptic contacts onto neurons of the spinal ventral horn14&#160;Figure 1b13. Macroscopic whole-cell Ca2+ currents have been recorded from reticulospinal axons in the intact spinal cord13,15. However, previous blind attempts at direct measurement of Ca2+&#160;currents in reticulospinal axons in the intact lamprey spinal cord using cell-attached patch clamp technique have proven unsuccessful13 due to lack of access to the presynaptic release face membrane owing to the opposing postsynaptic processes Figure 1b. The release face membrane has been previously made accessible by removal of the postsynaptic neuron11, mechanical perturbation of the synapse prior to recording12 or enzymatic treatment coupled with mechanical dissociation16. Given the complex organization of the spinal cord, it would prove extremely difficult to identify the postsynaptic neuron and retract it mechanically or perturb the synapse. Hence, we decided to use enzymatic treatment17 followed by mechanical dissociation.
Using this approach, we have developed an acutely dissociated preparation of the lamprey spinal cord that yields viable isolated reticulospinal axons with functional presynaptic terminals devoid of any postsynaptic processes, thereby providing unrestricted access to individual presynaptic terminals. In conjunction with a standard inverted microscope and fluorescence imaging, it enables us to identify and target individual fluorescently-identified presynaptic terminals, with a patch pipette containing a recording solution that isolates Ca2+ currents FIgure 4c&#160;and Figure 4d, for recording using cell-attached voltage-clamp technique. Ca2+ currents have been recorded directly at the presynaptic release face membrane of individual presynaptic terminals Figure 4f. This is a significant breakthrough in the field of synaptic transmission since it is the first such recording to be carried out at central synapses.

<table><tbody><tr><td>0.2 &amp;micro;m Syringe filter</td><td>EMD Millipore</td><td>SLGV004SL</td><td></td></tr><tr><td>22 x 60 mm Coverslips</td><td>Fisherbrand</td><td>12545J</td><td></td></tr><tr><td>Advasep-7</td><td>Cydex Pharmaceuticals</td><td>ADV7</td><td></td></tr><tr><td>Alexa Fluor 488 Phalloidin</td><td>Invitrogen/Life Technologies</td><td>A12379</td><td></td></tr><tr><td>Alexa-633 conjugated goat anti-rabbit secondary antibody</td><td>Invitrogen/Life Technologies</td><td>A21070</td><td></td></tr><tr><td>Antifreeze</td><td>Prestone</td><td></td><td></td></tr><tr><td>Boric acid</td><td>Sigma-Aldrich</td><td>B7660</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>A7906</td><td></td></tr><tr><td>Bright field light source</td><td>Dolan-Jenner</td><td>Fiberlite 180</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma Aldrich</td><td>C4901</td><td></td></tr><tr><td>Collagenase Type IA from Cloristridium Histolyticum</td><td>Sigma-Aldrich</td><td>C9891</td><td></td></tr><tr><td>Cover slip</td><td>Fisher-Scientific</td><td>12-545-J</td><td></td></tr><tr><td>Dextrose</td><td>Sigma-Aldrich</td><td>D9559</td><td></td></tr><tr><td>Digital CCD Camera</td><td>Hamamatsu</td><td>C8484-03G01</td><td></td></tr><tr><td>Dissection fine forceps</td><td>Fine Science Tools</td><td>91150-20</td><td></td></tr><tr><td>Dissection forceps</td><td>Fine Science Tools</td><td>11251-20</td><td></td></tr><tr><td>Dissection microscope</td><td>Leica Biosystems</td><td>Leica MZ 12</td><td></td></tr><tr><td>Dissection scissors</td><td>Fine Science Tools</td><td>15025-10</td><td></td></tr><tr><td>Dissection scissors fine</td><td>Fine Science Tools</td><td>91500-09</td><td></td></tr><tr><td>Dissection scissors ultra fine</td><td>Fine Science Tools</td><td>15000-08</td><td></td></tr><tr><td>FM 1-43</td><td>Invitrogen/Life Technologies</td><td>T3163</td><td></td></tr><tr><td>Glycine</td><td>Sigma-Aldrich</td><td>G7126</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H7523</td><td></td></tr><tr><td>High vacuum grease</td><td>Dow-Corning</td><td></td><td></td></tr><tr><td>Hydrochloric acid</td><td>Fisherbrand</td><td>SA-56-500</td><td></td></tr><tr><td>Immersion oil</td><td>Fisher-Scientific</td><td>M2000</td><td></td></tr><tr><td>Industrial grade nitrogen gas tank</td><td>Praxair</td><td>UN1066</td><td></td></tr><tr><td>Insect pins</td><td>Fine Science Tools</td><td>26002-10</td><td></td></tr><tr><td>Liquid suture glue</td><td>Braun Veterinary Cair Division</td><td>8V0305</td><td>The suture glue we used in our experiments was provided to us by another lab. It is no longer manufactured. We have sourced a Histoacryl Suture Glue for future use from Aesculap (Ts1050071FP)</td></tr><tr><td>Magnesium chloride</td><td>Sigma-Aldrich</td><td>M2670</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>154903</td><td></td></tr><tr><td>Non-fat dry milk</td><td>Cell Signaling Technology</td><td>9999S</td><td></td></tr><tr><td>P-87 Micropipette puller</td><td>Sutter Instruments</td><td></td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>P6148</td><td></td></tr><tr><td>Perfusion pump</td><td>Cole-Palmer</td><td>Masterflex C/L</td><td></td></tr><tr><td>Petri dish 100 x 15 mm</td><td>Fisher-Scientific</td><td>875712</td><td></td></tr><tr><td>Petri dish 35 x 10 mm</td><td>Fisher-scientific</td><td>875712</td><td></td></tr><tr><td>Poly-D-lysine hydrobromide</td><td>Sigma-Aldrich</td><td>P1024</td><td>MW &gt; 300,000</td></tr><tr><td>Potassium chloride</td><td>Sigma-Aldrich</td><td>P9333</td><td></td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma-Aldrich</td><td>P5379</td><td></td></tr><tr><td>Primary antibodies R-type calcium channel</td><td>Alomone Labs</td><td>ACC-006</td><td></td></tr><tr><td>Protease Type XIV from Streptomyces Griseus</td><td>Sigma-Aldrich</td><td>P5147</td><td></td></tr><tr><td>Scalpel blades</td><td>World Precision Instruments&amp;nbsp;</td><td>500240</td><td></td></tr><tr><td>Schot Duran Pressure Bottle</td><td>Fisher-Scientific</td><td>09-841-006</td><td></td></tr><tr><td>Silicone tubing for glue application</td><td>Cole-Palmer</td><td>07625-26</td><td></td></tr><tr><td>Slicing base plate</td><td>Leica Biosystems</td><td>14046327404</td><td></td></tr><tr><td>Slicing chamber</td><td>Leica Biosystems</td><td>14046230132</td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>S7653</td><td></td></tr><tr><td>Sodium hydroxide</td><td></td><td>S8045</td><td></td></tr><tr><td>Sodium phosphate dibasic</td><td>Sigma-Aldrich</td><td>S9763</td><td></td></tr><tr><td>Sodium tetraborate</td><td>Sigma-Aldrich</td><td>B3545</td><td></td></tr><tr><td>Sylgard 160 Silicone Elastomer Kit</td><td>Dow Corning&amp;nbsp;</td><td>SYLGARD&amp;reg; 160</td><td>To prepare, mix elastomer A and elastomer B 10:1 by weight</td></tr><tr><td>Sylgard 184 Silicone Elastomer Kit</td><td>Dow Corning</td><td>SYLGARD&amp;reg; 184&amp;nbsp;</td><td>To prepare, mix elastomer and curing agent 10:1 by weight</td></tr><tr><td>Teflon coated forceps</td><td>Fine Science Tools</td><td>11626-11</td><td></td></tr><tr><td>Tricaine methanesulphonate</td><td>Sigma-Aldrich</td><td>A5040</td><td></td></tr><tr><td>Vibratome blades</td><td>World Precision Instruments&amp;nbsp;</td><td>BLADES</td><td></td></tr><tr><td>Xenon lamp</td><td>Nikon</td><td></td><td></td></tr></tbody></table>

acute dissociation of lamprey reticulospinal axons to enable recording from the release face membrane of individual functional presynaptic terminals

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+&#160;currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+&#160;and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+&#160;entry has been characterized using imaging techniques and macroscopic current measurements &#8211; techniques that do not have sufficient temporal resolution to visualize Ca2+ entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+ currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.

This dissociation protocol yields healthy and functional isolated reticulospinal axons devoid of postsynaptic projections Figure 2f, but which nevertheless retain functional presynaptic terminals capable of evoked synaptic vesicle exo-and endocytosis Figure 4c&#160;and Figure 4d. Sections of the isolated regions of the reticulospinal axons can be clearly identified under light microscopy to be clear of any other neuronal processes allowing unrestricted access to the reti...

Watch this Scientific Journal Video about Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals at JoVE.c

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals.

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated ...

acute-dissociation-lamprey-reticulospinal-axons-to-enable-recording

Research

JoVE Journal

Neuroscience

8.7K Views.  University of Illinois at Chicago. Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals. 

Video: Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of recordings are conducted on dissociated DRG neurons, which is a standard preparation for most laboratories. Neuronal properties, however, can be altered by axonal injury resulting from enzyme digestion used in acquiring dissociated neurons. Further, dissociated neuron preparations cannot fully represent the microenvironment of the DRG since loss of contact with satellite glial cells that surround the primary sensory neurons is an unavoidable consequence of this method. To overcome the limitations in using conventional dissociated DRG neurons for patch clamp recordings, in this report we describe a method to prepare intact DRGs and conduct patch clamp recordings on individual primary sensory neurons ex vivo. This approach permits the fast and straightforward preparation of intact DRGs, mimicking in vivo conditions by keeping DRG neurons associated with their surrounding satellite glial cells and basement membrane. Furthermore, the method avoids axonal injury from manipulation and enzyme digestion such as when dissociating DRGs. This ex vivo preparation can additionally be used to study the interaction between primary sensory neurons and satellite glial cells.

This manuscript describes how to prepare intact dorsal root ganglia for patch clamp recordings. This preparation maintains the microenvironment for neurons and satellite glial cells, thus avoiding the phenotypic and functional changes seen using dissociated DRG neurons.

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of ...

Biochemistry

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

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

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

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

Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle. This is a direct consequence of the accessibility to both cell types and ability to visually distinguish the single segmental CaP motor-neuron on the basis of morphology and location. This video demonstrates the microscopic methods used to identify a CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type. Identification of the CaP motor-neuron type is confirmed by either dye filling or by the biophysical features such as action potential waveform and cell input resistance. Motor-neuron recordings routinely last for one hour permitting long-term recordings from multiple different target muscle cells. Control over the motor-neuron firing pattern enables measurements of the frequency-dependence of synaptic transmission at the neuromuscular junction. Owing to a large quantal size and the low noise provided by whole cell voltage clamp, all of the unitary events can be resolved in muscle. This feature permits study of basic synaptic properties such as release properties, vesicle recycling, as well as synaptic depression and facilitation. The advantages offered by this in vivo preparation eclipse previous neuromuscular model systems studied wherein the motor-neurons are usually stimulated by extracellular electrodes and the muscles are too large for whole cell patch clamp. The zebrafish preparation is amenable to combining electrophysiological analysis with a wide range of approaches including transgenic lines, morpholino knockdown, pharmacological intervention and in vivo imaging. These approaches, coupled with the growing number of neuromuscular disease models provided by mutant lines of zebrafish, open the door for new understanding of human neuromuscular disorders.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target skeletal muscle. This video demonstrates the microscopic methods used to identify a segmental CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle.  ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale. This complex process requires the coordinated activity of many pre- and post-synaptic proteins to ensure appropriate synaptic connectivity, conduction of electrical signals, targeting and priming of secretory vesicles, calcium sensing, vesicle fusion, localization and function of postsynaptic receptors and finally, recycling mechanisms. As neuroscientists it is our goal to elucidate which proteins function in each of these steps and understand their mechanisms of action. Electrophysiological recordings from synapses provide a quantifiable read out of the underlying electrical events that occur during synaptic transmission. By combining this technique with the powerful array of molecular and genetic tools available to manipulate synaptic proteins in C. elegans, we can analyze the resulting functional changes in synaptic transmission. 
The C. elegans NMJs formed between motor neurons and body wall muscles control locomotion, therefore, mutants with uncoordinated locomotory phenotypes (known as unc s) often perturb synaptic transmission at these synapses 1. Since unc mutants are maintained on a rich supply of a bacterial food source, they remain viable as long as they retain some pharyngeal pumping ability to ingest food. This, together with the fact that C. elegans exist as hermaphrodites, allows them to pass on mutant progeny without the need for elaborate mating behaviors. These attributes, coupled with our recent ability to record from the worms NMJs 2,3,7 make this an excellent model organism in which to address precisely how unc mutants impact neurotransmission. 
The dissection method involves immobilizing adult worms using a cyanoacrylic glue in order to make an incision in the worm cuticle exposing the NMJs. Since C. elegans adults are only 1 mm in length the dissection is performed with the use of a dissecting microscope and requires excellent hand-eye coordination. NMJ recordings are made by whole-cell voltage clamping individual body wall muscle cells and neurotransmitter release can be evoked using a variety of stimulation protocols including electrical stimulation, light-activated channel-rhodopsin-mediated depolarization 4 and hyperosmotic saline, all of which will be briefly described.

Application of electrophysiology to accessible synapses provides a quantifiable measure of synaptic activity, useful in analyzing synaptic mutants. This article describes a dissection method used to expose the neuromuscular junctions (NMJ) of Caenorhabditis elegans (C. elegans) and briefly discusses some of the uses to which this preparation can be applied.

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale.   This ...

Biology

Combined Recording of Mechanically Stimulated Afferent Output and Nerve Terminal Labelling in Mouse Hair Follicle Lanceolate Endings

A novel dissection and recording technique is described for monitoring afferent firing evoked by mechanical displacement of hairs in the mouse pinna. The technique is very cost-effective and easily undertaken with materials commonly found in most electrophysiology laboratories, or easily purchased. The dissection is simple and fast, with the mechanical displacement provided by a generic electroceramic wafer controlled by proprietary software. The same software also records and analyses the electroneurogram output. The recording of the evoked nerve activity is through a commercial differential amplifier connected to fire-polished standard glass microelectrodes. Helpful tips are given for improving the quality of the preparation, the stimulation and the recording conditions to optimize recording quality. The system is suitable for assaying the electrophysiological and optical properties of lanceolate terminals of palisade endings of hair follicles, as well as the outcomes from their pharmacological and/or genetic manipulation. An example of combining electrical recording with mechanical stimulation and labeling with a styryl pyridinium vital dye is given.

A simple and novel technique for recording afferent discharge due to mechanical stimulation of lanceolate terminals of palisade endings innervating mouse ear skin hair follicles is presented.

A novel dissection and recording technique is described for monitoring afferent firing evoked by mechanical displacement of hairs in the mouse pinna. The ...

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal macropatch recordings of synaptic currents from visualized boutons at the Drosophila larval NMJ. This technique requires customized fabrication of recording micropipettes, as well as a compound microscope equipped with a high magnification, long-distance water immersion objective, differential interference contrast (DIC) optics, and a fluorescent attachment. The recording electrode is positioned on the top of a selected synaptic bouton visualized with DIC optics, epi-fluorescence, or both. The advantage of this technique is that it allows monitoring the synaptic activity of a limited number of sites of release. The recording electrode has&#160;a diameter of several microns, and the release sites positioned outside of the electrode rim do&#160;not significantly affect the recorded currents. The recorded synaptic currents have fast kinetics and can be readily resolved. These advantages are especially important for the studies of mutant fly lines with enhanced spontaneous or asynchronous synaptic activity.

Focal Macropatch Recordings of Synaptic Currents from the Drosophila Larval Neuromuscular Junction

Synaptic currents can be recorded focally from visualized synaptic boutons at the Drosophila third instar larvae neuromuscular junction. This technique enables monitoring the activity of a single synaptic bouton.

Drosophila neuromuscular junction (NMJ) is an excellent model system to study glutamatergic synaptic transmission. We describe the technique of focal ...

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

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

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

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

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

Live Imaging of Synaptic Vesicle Recycling in the Neuromuscular Junction of Dissected Larval Zebrafish

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) in response to an action potential (AP). Since SVs are recycled locally at the presynaptic terminal, coordination of SV exocytosis and endocytosis is important for sustained synaptic transmission. A pH-sensitive green fluorescent protein, called pHluorin, provides a powerful tool to monitor SV exo/endocytosis by targeting it to the SV lumen. However, tracking AP-driven SV recycling with the pHluorin-based probes is still largely limited to in vitro culture preparations because the introduction of genetically encoded probes and subsequent optical imaging is technically challenging in general for in vivo animal models or tissue preparations. Zebrafish is a model system offering valuable features, including ease of genetic manipulation, optical clarity, and rapid external development. We recently generated a transgenic zebrafish that highly expresses a pHluorin-labeled probe at motor neuron terminals and developed a protocol to monitor AP-driven SV exo/endocytosis at the neuromuscular junction (NMJ), a well-established synapse model that forms in vivo. In this article, we show how to prepare larval zebrafish NMJ preparation suitable for pHluorin imaging. We also show that the preparation allows time-lapse imaging under conventional upright epifluorescence microscope, providing a cost-effective platform for analyzing NMJ function.

The zebrafish larval neuromuscular junction is an attractive model for studying synaptic physiology. It is amenable to many experimental techniques, including electrophysiology and optical imaging. Here, we describe a protocol for imaging synaptic transmission using a pHluorin-based probe under an upright epifluorescence microscope.

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) ...

Optogenetic Stimulation and Electrophysiological Recording of Synaptic Events in Rat Brain Slices

Source: Kinnavane, L., et al. <a href="https://app.jove.com/v/63077">Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex.</a> J. Vis. Exp. (2022)
This video demonstrates the procedure for studying synaptic efficiency in the lateral entorhinal cortex (LEC) by optogenetically stimulating medial prefrontal cortex (mPFC) axons and recording the responses of pyramidal neurons in the LEC. This method combines patch-clamp recordings with synaptic activation by light stimulation to study synaptic plasticity.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Electrophysiology and ...

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Summary

Explore More Videos

PLAYLIST