Testing Monosynaptic Connections Using Tetrodotoxin and Tetrodotoxin-Resistant Sodium Channels

研究方案

1. Prepare Flies and Identify GAL4 promoter Lines

1. Select a GAL4 promoter line and cross it with flies carrying both the UAS-NaChBac transgene and an optogenetic transgene for activation.
   NOTE: UAS-CsChrimson is selected due to its high conductance and the ease with which it activates tetrodotoxin (TTX)-resistant sodium channel (NaChBac)-mediated action potentials. Both UAS-NaChBac and UAS-CsChrimson are available from the Bloomington Stocks Repository.
2. Prepare all-trans retinal as a stock solution in ethanol (35 mM) and store the solution in freezer at -20 °C.
3. Mix 28 µL of this stock into approximately 5 mL of rehydrated potato flake food and add this mixture to the top of a vial of conventional Drosophila medium.
4. Raise adult flies expressing csChrimson on food containing 0.2 mM all-trans-retinal for 1–3 days.

2. Prepare TTX Stock

1. Create a stock solution of 1 mM TTX in distilled or deionized water. Store this solution in a lab refrigerator for several months. Check the institution's policies regarding the storage of TTX, as many institutions classify TTX as a hazardous or controlled substance.

3. Prepare Flies for Electrophysiology

1. Collect 1–3 day old adult flies that are raised on the food with all-trans-retinal.
2. Put the flies in a glass scintillation vial. Place the vial onto ice for 1 min to immobilize the flies.
3. Use coarse forceps to insert a fly into a custom-made recording chamber. The chamber consists of a 3 ½" circle laser cut from 1/16" acrylic. Cut a ¾" hole into the center, where a custom foil is attached with 2-part epoxy. The foil contains a triangular-shaped hope that the fly is inserted into.
4. Apply wax or ultraviolet (UV) curable epoxy around the animal to firmly secure it within the recording chamber.
5. Illuminate the preparation with a gooseneck optical fiber for visualization under a stereomicroscope. Set the gooseneck fibers for oblique illumination by adjusting them so that they illuminate the fly directly from the side.
6. Adjust the light level to lowest possible setting that still allows clear visualization of the fly. This light intensity will vary across individual experiments.
   NOTE: This protocol can be adapted by using an in situ brain explant method.
7. Retrieve external recording saline. The saline contains in mM: 103 sodium chloride, NaCl, 3 potassium chloride, KCl, 5 N-tris(hydroxymethyl)methyl-2- aminoethane-sulfonic acid, 8 trehalose, 10 glucose, 26 sodium bicarbonate, NaHCO₃, 1 sodium dihydrogen phosphate, NaH₂PO₄, 1.5 calcium chloride, CaCl₂, and 4 magnesium chloride, MgCl₂ (adjusted to 270–275 mΩ). The saline is bubbled with 95% O₂/5% CO₂ (carbogen) and has a pH adjusted to 7.3.
8. Place 4-6 drops of extracellular recording saline over the fly preparation. At this point, increase the light level at the dissection microscope for better visibility.
9. Sharpen a tungsten wire using electrolysis. To make the wire, prepare a saturated solution of potassium nitrate, KNO₃ and apply approximately 20 V of alternating current (AC) while dipping the tip of the wire into the solution 20 times for 100 ms each until the tip is sharpened. Use the sharpened tungsten wire to remove the cuticle on the head of the fly and thus exposing the brain.
10. Further expose the brain by removing trachea and fat bodies that surround it. If accessing the antennal lobes, use a bent tungsten wire or similar tool to gently tuck the antennae underneath the foil recording chamber to increase visibility. Use sharp forceps to gently tear away the glial sheathing covering the area of interest where recordings will be made.
11. Transfer the recording chamber to the electrophysiology rig. Immediately start perfusion with external recording saline bubbled with carbogen to preserve the brain. The saline solution reservoir is placed above the microscope stage and is gravity-fed to the preparation. Use a vacuum line and a 2 L Erlenmeyer flask to collect the waste saline.

4. Obtain Whole-cell Recording

1. Pull patch pipettes on a commercial pipette puller. Consult the manual for settings to achieve an appropriate pipette with a tip resistance near 7 mΩ.
2. Add 4 µL of internal recording solution into a patch pipette. The internal saline consists of 140 potassium aspartate, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid  (HEPES), 4 magnesium adenosine triphosphate (MgATP), 0.5 Guanosine 5'-triphosphate trisodium salt hydrate (Na₃GTP), 1 ethylene glycol tetraacetic acid (EGTA), and 1 potassium chloride (KCl) in mM.
3. Set up patch clamp amplifier for whole-cell recording. Zero the amplifier's offset and set the amplifier to deliver test pulses. Here, an AM systems Patch clamp amplifier 2,400 is used.
4. Apply positive pressure through the pipette and approach the cell. Use a micromanipulator to direct the pipette and contact the cell. Release positive pressure. Set the holding potential of the membrane to -60 mV. The pipette should form a gigaohm seal with the cell's membrane as evidenced by a low sub-picoamp holding current.
   NOTE: If using green fluorescent protein (GFP) guided cell targeting, try to minimize use of epifluorescent light to prevent ChR2 activation and potential synaptic depression. Also wait for a few minutes before applying step 5.
5. Set the amplifier to deliver test pulses from -50 mV to -60 mV. This setting is standard on patch-clamp amplifiers. Test pulses will reveal a large capacitative transient representing the current necessary to charge the patch pipette. Remove capacitative transients by adjusting the capacity compensation knobs on the AM 2400 or similar amplifier.
6. Apply brief pulses of negative pressure to rupture the cell membrane and obtain whole-cell configuration.  
   NOTE: A whole cell configuration is observed when there is a large increase in the capacitative transients showing the capacitance of the neuron. A small increase in the holding current (baseline) will likely also be observed. Set the amplifier to the current clamp mode and adjust the cell's potential to -50 to -60 mV.

5. Test Synaptic Connectivity

1. Configure the data acquisition (DAQ) system to trigger a light-emitting diode (LED) driver to generate light pulses. Here, a National Instruments DAQ system is used with Matlab to deliver an analog signal to a LED driver. The LED intensity is adjusted via the analog signal to the driver. The LED is a 620-630 nm wavelength LED.
2. Position the high-powered red LED directly underneath the preparation. Adjust the light intensity so that 0.238 mW/mm² reaches the fly.
3. Activate presynaptic neurons while recording from the postsynaptic cell of interest. Achieve cell activation optogenetically, pharmacogentically, or via nerve stimulation. Here, csChrimson and brief pulses of red light are applied.
   NOTE: Synaptic connections can be monitored either in the current clamp as excitatory postsynaptic potentials (EPSPs) or voltage clamp as excitatory postsynaptic currents (EPSCs). A synaptic connection should be visible before applying TTX in response to a 40 ms light pulse. If no connection is observed, it is unlikely that the neurons are synaptically connected either mono- or polysynaptically. The holding potential may be adjusted to emphasize excitatory or inhibitory connections accordingly.
4. If a connection is observed in normal saline, add TTX to the perfusion system to achieve a final concentration of 1 µM. Use a recirculating pump to capture and reuse the saline to conserve TTX. The peristaltic pump will draw the saline solution from the bath and return it to the saline reservoir.
5. Adjust the pump's speed to provide a strong constant vacuum that does not allow the saline level in the bath to rise and fall.
   NOTE: The application of TTX should result in the cessation of spiking activity in the neuron that is being monitored in whole-cell. This confirms that enough TTX has been added to the saline.
6. Apply brief pulses of red light (40 ms for each pulse) again. At this point, any synaptic connection that remains is likely monosynaptic. 

材料

Name	Company	Catalog Number	Comments
UAS-csChrimson	Bloomington Drosophila Stock Center	55135	Used as a neural activator
UAS-NaChBac	Bloomington Drosophila Stock Center	9466	Resotores excitibility in cells in TTX
Tetrodotoxin	Tochris	1078	Special permission may be needed to purchase TTX as it is a controlled substance
all trans-Retinal	Sigma-Aldrichall trans-Retinal	R2500	Require co-factor for channelrhodopsin
Weldable 321 Stainless Steel Sheet, 0.002" Thick, 10" Wide	McMaster Carr	3254K7	Used to make custom fly holder. Custom foil can be laser cut at pololu.com from the provided PDF file
Dissection Microscope	Zeiss	Stemi 2000-C	Used for dissection of preparation
Waxer	Almore	Eectra Waxer 66000	Used during dissection to secure fly in foil
Paraffin Wax	Joann	4917217	Used with waxer
Number 5 forceps	Fine Science Tools	#5CO	Used for dissection and desheathing
Dissection Scissors	Fine Science Tools	15001-08	Used to remove parts of the cuticle during dissection
Tungsten wire	A-M Systems	797500	Use with electrolysis to make sharpened needles for dissection
Reciculating Peristaltic Pump	Simply Pumps (Amazon)	PM200S	for recirculating TTX
Speed Controller for peristaltic pump	Zitrades (Amazon)	N/A	PWM Dimming Controller For LED Lights or Ribbon, 12 Volt 8 Amp,Adjustable Brightness Light Switch Dimmer Controller DC12V 8A 96W for Led Strip Light B
Versa-Mount Precision Compressed Air Regulator	McMaster Carr	1804T1	For applying positive pressure during patching
Glass capilaries	World Precision Instruments	TW150F-3	For patch pipettes
Multipurpose Gauge	McMaster Carr	3846K431	Gauge for pressure regulator
Electrophysiology Camera	Dage MTI	IR-1000	Any camera that works in the IR range (850 nm) will work. You do not want to use red illumination as this can activate csChrimson
IR LED	Thorlabs	M850F2	For oblique illumination
Fiber optic for IR LED	Thorlabs	M89L01	Couples to IR LED
Objective lens	Olympus 40X	LUMPlanFLN	This can be used on most microscipes and works well for visualizing fly neurons.
Amber LED	Thorlabs	M590L3d	For visualizing RFP and mCherry
Blue LED	Thorlabs	M470L3d	For visualizing GFP
GFP filter set	Chroma	49011	For visualizing GFP or stimulating channel2rhodopsin
Custom mCherry Filter Set	Chroma	et580/25x and t600lpxr (from the 49306 set) but with an et610lp barrier/emission optic	Use only if you wish to patch identified neurons with channel2rhodopsin
Dichroic to combine Amber and blue LED	Thorlabs	DMLP550R	Use only patch under mCherry and excite channel2rhodopsin with blue light.
Red LED	LEDSupply	Cree XPE 620 - 630 nm	Used to drive csChrimson
LED Driver	LEDSupply	3021-D-E-1000	Used to drive LEDs for optogenetic stimulation
Manipulator	Sutter Instruments	MP-225	Used to position pipette during recordings
Patchclamp Amplifier	A-M Systems	Model 2400	An equivalent amplifier is suitable
Bessel Filter	Warner Instruments	LPF 202A	Auxillary filter used to filter current trace to oscilloscope during patching.
Data Acquisition System	National Instruments	NI PCIe-6321       781044-01	Used to record data from amplifier to computer
Connector Block - BNC Terminal BNC-2090A	National Instruments	779556-01	Used to connect amplifier to DAQ card.
Steel Foil	McMaster Carr	3254K7	Steel foil for custom recording chamber
Magnets	K&J Magnetics	D42	To secure recording chamber to ring stand
1/16" Cell Cast Acrylic Clear	Pololu		Used to make custom recording chamber. Acrylic can be laser cut at pololu.com from the provided PDF file