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

Podsumowanie

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.

Streszczenie

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca²⁺ 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⁺ currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca²⁺ and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca²⁺ entry has been characterized using imaging techniques and macroscopic current measurements – 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.

Wprowadzenie

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⁺ concentration by uncaging Ca^{2+ 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⁺ 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⁺ 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 ganglion^7-10 and rat calyces^11,12. 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⁺ imaging to study presynaptic Ca²⁺ fluxes.

Figure 1. Lamprey giant reticulospinal synapse. (a) Cross-section of lamprey spinal cord indicating dorso-ventral orientation. Reticulospinal axons are marked with green asterix. (b) 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 Figure 1b¹³. Macroscopic whole-cell Ca²⁺ currents have been recorded from reticulospinal axons in the intact spinal cord^13,15. However, previous blind attempts at direct measurement of Ca²⁺ 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 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.

Protokół

1. Preparation of Poly-D-lysine Hydrobromide

1. Prepare 1 mg/ml poly-D-lysine hydrobromide in 0.1 M borate buffer (pH 8.5).
2. Aliquot and store at -20 °C.

2. Poly-lysine Coating of Coverslips

Note: Carry out all cleaning and coating steps in a laminar flow chamber.

1. Place coverslips in a Petri dish containing 1 N Hydrochloric acid (HCl) for 2 hr.
2. Aspirate all HCl and rinse with 70% Ethanol (EtOH) 2-3x.
3. Leave in 70% EtOH for 1 hr.
4. Aspirate all 70% EtOH and rinse with 100% EtOH 2-3x.
5. Leave in 100% EtOH for 2 hr. Aspirate all EtOH.
6. Blot dry with filter paper and air dry for a few seconds.
7. Place O/N in 1 mg/ml poly-lysine solution.
8. Rinse coverslips next day with Millipore water 4-5x.
9. Air dry poly-lysine coated coverslips on glass rollers in a clean Petri dish.
10. 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 °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.
11. 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).
12. 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 °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.

3. Acute Dissociation of Lamprey Spinal Cord to Yield Isolated Reticulospinal Axons

1. 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.
2. Decapitate lamprey in cold (4 °C) Ringer’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.
3. Remove meninx primitiva from the dorsal surface of the spinal cord using fine forceps. Do not remove ventral meninx primitiva at this stage.
4. Cut spinal cord into 1 cm long pieces.
5. 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’s solution.
6. 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 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.
7. Incubate sliced spinal cord pieces at RT for 45 min 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’s solution (adapted from El Manira & Bussières, 1997 17).
8. Pin enzyme-treated spinal cord pieces using fine pins in a PDMS lined Petri dish containing cold (4 °C) Ringer’s solution.
9. Remove ventral meninx primitiva with fine forceps.
10. 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.
11. Place a drop of immersion oil on the lens.
12. 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.
13. Add Ringer’s solution into the recording chamber using a Pasteur pipette.
14. 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.
15. 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 and Figure 2f.
16. Bring the recording solution temperature to 10 °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.
17. Allow axons to recover for 1 hr post dissociation at 10 °C.

Figure 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 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.

Figure 3. Schematic of recording chamber for electrophysiology experiments. Dimensions provided are in inches. Red rectangle shown in basepiece diagram (a) 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) 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

1. Label presynaptic terminals, by incorporating FM 1-43 into vesicles during synaptic exo-endocytosis during high K+ depolarization. Perfuse preparation with 5 µM FM 1-43 (in 5 ml Ringer’s solution containing 30 mM KCl).
2. Perfuse preparation with 1 mg/ml Advasep-7 (in 5 ml Ringer’s solution) to remove excess FM dye)¹⁸.
3. Perfuse preparation with Ringer’s solution for 15 min to washout any remant dye and the Advasep.
4. 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.
5. Identify fluorescently labeled presynaptic terminals to target for recording Figure 4c and Figure 4d.

5. Immunohistochemistry of Isolated Reticulospinal Axons

1. Fill dish inset (Protocol section 2.10.) with divalent-ion (Ca²⁺ and Mg²⁺) free Ringer’s solution.
2. 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).
3. 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.
4. Exchange divalent free Ringer’s solution with regular Ringer’s solution by perfusion.
5. Allow axons to recover for 20 min post dissociation at 10 °C.
6. Fix dissociated axons in 4% paraformaldehyde (PFA) {prepared in Phosphate Buffer Saline (PBS, (mM) NaCl 137, 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 µM syringe filter.
7. Wash out the PFA by perfusing 0.1 M glycine (in PBS) for 10 min.
8. Incubate in 0.1% Triton-X (in PBS) for 10 min.
9. Wash by perfusing PBS for 20 min.
10. Block with 5% non-fat milk (in PBS) for 6 hr at 4 °C.
11. Add in primary antibody to VGCC of interest (1:200 dilution in PBS) and incubate for 20 hr at 4 °C.
12. Wash by perfusing PBS for 20 min.
13. Block with 5% non-fat milk (in PBS) for 10 min at 4 °C.
14. Add in secondary antibody (1:400 dilution in PBS) and incubate in dark for 2 hr at 4 °C.
15. Wash by perfusing with PBS for 20 min.
16. Block with 1% Bovine Serum Albumin (in PBS) for 20 min.
17. Add in Alexa Fluor 488 phalloidin (5 Units/µl working concentration; stock prepared in methanol 200 Units/ml).
18. Wash by perfusing with PBS for 20 min.
19. Image using 100X water immersion lens on confocal microscope.

6. Electrophysiological Recording

1. Fabricate aluminosilicate glass patch pipettes (pipette resistance 2-5 MΩ) in a P-87 micropipette puller. Design patch pipette such that pipette tip encompasses entire presynaptic terminal diameter.
2. 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.
3. Fire polish patch pipettes using a microforge (a custom built platinum filament fitted onto the stage of a compound microscope).
4. Identify isolated axons demonstrating labeled presynaptic terminals by fluorescence microscopy.
5. 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.
6. 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.
7. 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.
8. 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.
9. 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.

Wyniki

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 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...

Dyskusje

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 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...

Materiały

Name	Company	Catalog Number	Comments
0.2 µm Syringe filter	EMD Millipore	SLGV004SL
22 x 60 mm Coverslips	Fisherbrand	12545J
Advasep-7	Cydex Pharmaceuticals	ADV7
Alexa Fluor 488 Phalloidin	Invitrogen/Life Technologies	A12379
Alexa-633 conjugated goat anti-rabbit secondary antibody	Invitrogen/Life Technologies	A21070
Antifreeze	Prestone
Boric acid	Sigma-Aldrich	B7660
Bovine serum albumin	Sigma-Aldrich	A7906
Bright field light source	Dolan-Jenner	Fiberlite 180
Calcium chloride	Sigma Aldrich	C4901
Collagenase Type IA from Cloristridium Histolyticum	Sigma-Aldrich	C9891
Cover slip	Fisher-Scientific	12-545-J
Dextrose	Sigma-Aldrich	D9559
Digital CCD Camera	Hamamatsu	C8484-03G01
Dissection fine forceps	Fine Science Tools	91150-20
Dissection forceps	Fine Science Tools	11251-20
Dissection microscope	Leica Biosystems	Leica MZ 12
Dissection scissors	Fine Science Tools	15025-10
Dissection scissors fine	Fine Science Tools	91500-09
Dissection scissors ultra fine	Fine Science Tools	15000-08
FM 1-43	Invitrogen/Life Technologies	T3163
Glycine	Sigma-Aldrich	G7126
HEPES	Sigma-Aldrich	H7523
High vacuum grease	Dow-Corning
Hydrochloric acid	Fisherbrand	SA-56-500
Immersion oil	Fisher-Scientific	M2000
Industrial grade nitrogen gas tank	Praxair	UN1066
Insect pins	Fine Science Tools	26002-10
Liquid suture glue	Braun Veterinary Cair Division	8V0305	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)
Magnesium chloride	Sigma-Aldrich	M2670
Methanol	Sigma-Aldrich	154903
Non-fat dry milk	Cell Signaling Technology	9999S
P-87 Micropipette puller	Sutter Instruments
Paraformaldehyde	Sigma-Aldrich	P6148
Perfusion pump	Cole-Palmer	Masterflex C/L
Petri dish 100 x 15 mm	Fisher-Scientific	875712
Petri dish 35 x 10 mm	Fisher-scientific	875712
Poly-D-lysine hydrobromide	Sigma-Aldrich	P1024	MW > 300,000
Potassium chloride	Sigma-Aldrich	P9333
Potassium phosphate monobasic	Sigma-Aldrich	P5379
Primary antibodies R-type calcium channel	Alomone Labs	ACC-006
Protease Type XIV from Streptomyces Griseus	Sigma-Aldrich	P5147
Scalpel blades	World Precision Instruments	500240
Schot Duran Pressure Bottle	Fisher-Scientific	09-841-006
Silicone tubing for glue application	Cole-Palmer	07625-26
Slicing base plate	Leica Biosystems	14046327404
Slicing chamber	Leica Biosystems	14046230132
Sodium chloride	Sigma-Aldrich	S7653
Sodium hydroxide		S8045
Sodium phosphate dibasic	Sigma-Aldrich	S9763
Sodium tetraborate	Sigma-Aldrich	B3545
Sylgard 160 Silicone Elastomer Kit	Dow Corning	SYLGARD® 160	To prepare, mix elastomer A and elastomer B 10:1 by weight
Sylgard 184 Silicone Elastomer Kit	Dow Corning	SYLGARD® 184	To prepare, mix elastomer and curing agent 10:1 by weight
Teflon coated forceps	Fine Science Tools	11626-11
Tricaine methanesulphonate	Sigma-Aldrich	A5040
Vibratome blades	World Precision Instruments	BLADES
Xenon lamp	Nikon