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>

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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

8.8K 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

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

Biology

The Identification of Sea Lamprey Pheromones Using Bioassay-Guided Fractionation

Bioassay-guided fractionation is an iterative approach that uses the results of physiological and behavioral bioassays to guide the isolation and identification of an active pheromone compound. This method has resulted in the successful characterization of the chemical signals that function as pheromones in a wide range of animal species. Sea lampreys rely on olfaction to detect pheromones that mediate behavioral or physiological responses. We use this knowledge of fish biology to posit functions of putative pheromones and to guide the isolation and identification of active pheromone components. Chromatography is used to extract, concentrate, and separate compounds from the conditioned water. Electro-olfactogram (EOG) recordings are conducted to determine which fractions elicit olfactory responses. Two-choice maze behavioral assays are then used to determine if any of the odorous fractions are also behaviorally active and induce a preference. Spectrometric and spectroscopic methods provide the molecular weight and structural information to assist with the structure elucidation. The bioactivity of the pure compounds is confirmed with EOG and behavioral assays. The behavioral responses observed in the maze should ultimately be validated in a field setting to confirm their function in a natural stream setting. These bioassays play a dual role to 1) guide the fractionation process and 2) confirm and further define the bioactivity of isolated components. Here, we report the representative results of a sea lamprey pheromone identification that exemplify the utility of the bioassay-guided fractionation approach. The identification of sea lamprey pheromones is particularly important because a modulation of its pheromone communication system is among the options considered to control the invasive sea lamprey in the Laurentian Great Lakes. This method can be readily adapted to characterize the chemical communication in a broad array of taxa and shed light on waterborne chemical ecology.

Here, we present a protocol to isolate and characterize the structure, olfactory potency, and behavioral response of putative pheromone compounds of sea lampreys.

Bioassay-guided fractionation is an iterative approach that uses the results of physiological and behavioral bioassays to guide the isolation and ...

Biochemistry

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 ...

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of neurophysiology, neuroethology, and developmental biology. The goal of this report is to consolidate experimental techniques from the leech system into a single article that will be of use to physiologists with expertise in other nervous system preparations, or to biology students with little or no electrophysiology experience. We demonstrate how to dissect the leech for recording intracellularly from identified neural circuits in the ganglion. Next we show how individual cells of known function can be removed from the ganglion to be cultured in a Petri dish, and how to record from those neurons in culture. Then we demonstrate how to prepare a patch of innervated skin to be used for mapping sensory or motor fields. These leech preparations are still widely used to address basic electrical properties of neural networks, behavior, synaptogenesis, and development. They are also an appropriate training module for neuroscience or physiology teaching laboratories.

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

This article describes three nervous system preparations using leeches: intracellular recording from neurons in ventral ganglia, culturing neurons from ventral ganglia, and recording from a patch of innervated skin to map sensory fields.

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of ...

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several weeks, and this is accompanied by short distance regeneration (a few mm) of propriospinal axons and spinal-projecting axons from the brainstem. Among the 36 large identifiable spinal-projecting neurons, some are good regenerators and others are bad regenerators. These neurons can most easily be identified in wholemount CNS preparations. In order to understand the neuron-intrinsic mechanisms that favor or inhibit axon regeneration after injury in the vertebrates CNS, we determine differences in gene expression between the good and bad regenerators, and how expression is influenced by spinal cord transection. This paper illustrates the techniques for housing larval and recently transformed adult sea lampreys in fresh water tanks, producing complete spinal cord transections under microscopic vision, and preparing brain and spinal cord wholemounts for in situ hybridization. Briefly, animals are kept at 16&#160;&#176;C and anesthetized in 1% Benzocaine in lamprey Ringer. The spinal cord is transected with iridectomy scissors via a dorsal approach and the animal is allowed to recover in fresh water tanks at 23 &#176;C. For in situ hybridization, animals are reanesthetized and the brain and cord removed via a dorsal approach.

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

Lampreys recover locomotion after a complete spinal cord injury. However, some spinal-projecting neurons are good regenerators and others are not. This paper illustrates the techniques for housing sea lamprey larvae (and recently transformed adults), producing complete spinal cord transections and preparing wholemount brains and spinal cords for in situ hybridization.

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several ...

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

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 ...

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ganglion cells and project it to the visual cortex. In addition, relay neurons receive top-down excitation from the cortex. The two main excitatory inputs to the relay neurons differ in several aspects. Each relay neuron receives input from only a few retinogeniculate synapses, which are large terminals with many release sites. This is reflected by the comparably strong excitation, the relay neurons receive, from retinal ganglion cells. Corticogeniculate synapses, in contrast, are simpler with few release sites and weaker synaptic strength. The two synapses also differ in their synaptic short-term plasticity. Retinogeniculate synapses have a high release probability and consequently display a short-term depression. In contrast, corticogeniculate synapses have a low release probability. Corticogeniculate fibers traverse the reticular thalamic nuclei before entering the lateral geniculate nucleus. The different locations of the reticular thalamic nucleus (rostrally from the lateral geniculate nucleus) and optic tract (ventro-laterally from the lateral geniculate nucleus) allow stimulating corticogeniculate or retinogeniculate synapses separately with extracellular stimulation electrodes. This makes the lateral geniculate nucleus an ideal brain area where two excitatory synapses with very different properties impinging onto the same cell type, can be studied simultaneously. Here, we describe a method to investigate the recording from relay neurons and to perform detailed analysis of the retinogeniculate and corticogeniculate synapse function in acute brain slices. The article contains a step-by-step protocol for the generation of acute brain slices of the lateral geniculate nucleus and steps for recording activity from relay neurons by stimulating the optic tract and corticogeniculate fibers separately.

Here, we present protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and the electrophysiological investigation of retinogeniculate and corticogeniculate synapses function. This protocol provides an efficient way to study synapses with the high-release and low-release probability in the same acute brain slices.

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ...

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates reafferent cues that animals must disentangle from relevant sensory cues of the surrounding environment. For example, when a fish swims, flow generated from body undulations is detected by the mechanoreceptive neuromasts, comprising hair cells, that compose the lateral line system. The hair cells then transmit fluid motion information from the sensor to the brain via the sensory afferent neurons. Concurrently, corollary discharge of the motor command is relayed to hair cells to prevent sensory overload. Accounting for the inhibitory effect of predictive motor signals during locomotion is, therefore, critical when evaluating the sensitivity of the lateral line system. We have developed an in vivo electrophysiological approach to simultaneously monitor posterior lateral line afferent neuron and ventral motor root activity in zebrafish larvae (4-7 days post fertilization) that can last for several hours. Extracellular recordings of afferent neurons are achieved using the loose patch clamp technique, which can detect activity from single or multiple neurons. Ventral root recordings are performed through the skin with glass electrodes to detect motor neuron activity. Our experimental protocol provides the potential to monitor endogenous or evoked changes in sensory input across motor behaviors in an intact, behaving vertebrate.

We describe a protocol to monitor changes in the afferent neuron activity during motor commands in a model vertebrate hair cell system.

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates ...

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

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