FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation

Summary

Synaptic vesicle (SV) cycling is the core mechanism of intercellular communication at neuronal synapses. FM dye uptake and release are the primary means of quantitatively assaying SV endo- and exocytosis. Here, we compare all the stimulation methods to drive FM1-43 cycling at the Drosophila neuromuscular junction (NMJ) model synapse.

Abstract

FM dyes are used to study the synaptic vesicle (SV) cycle. These amphipathic probes have a hydrophilic head and hydrophobic tail, making them water-soluble with the ability to reversibly enter and exit membrane lipid bilayers. These styryl dyes are relatively non-fluorescent in aqueous medium, but insertion into the outer leaflet of the plasma membrane causes a >40X increase in fluorescence. In neuronal synapses, FM dyes are internalized during SV endocytosis, trafficked both within and between SV pools, and released with SV exocytosis, providing a powerful tool to visualize presynaptic stages of neurotransmission. A primary genetic model of glutamatergic synapse development and function is the Drosophila neuromuscular junction (NMJ), where FM dye imaging has been used extensively to quantify SV dynamics in a wide range of mutant conditions. The NMJ synaptic terminal is easily accessible, with a beautiful array of large synaptic boutons ideal for imaging applications. Here, we compare and contrast the three ways to stimulate the Drosophila NMJ to drive activity-dependent FM1-43 dye uptake/release: 1) bath application of high [K⁺] to depolarize neuromuscular tissues, 2) suction electrode motor nerve stimulation to depolarize the presynaptic nerve terminal, and 3) targeted transgenic expression of channelrhodopsin variants for light-stimulated, spatial control of depolarization. Each of these methods has benefits and disadvantages for the study of genetic mutation effects on the SV cycle at the Drosophila NMJ. We will discuss these advantages and disadvantages to assist the selection of the stimulation approach, together with the methodologies specific to each strategy. In addition to fluorescent imaging, FM dyes can be photoconverted to electron-dense signals visualized using transmission electron microscopy (TEM) to study SV cycle mechanisms at an ultrastructural level. We provide the comparisons of confocal and electron microscopy imaging from the different methods of Drosophila NMJ stimulation, to help guide the selection of future experimental paradigms.

Introduction

The beautifully-characterized Drosophila larval neuromuscular junction (NMJ) glutamatergic synapse model has been used to study synapse formation and function with a vast spectrum of genetic perturbations¹. The motor neuron terminal consists of multiple axon branches, each with many enlarged synaptic boutons. These capacious varicosities (up to 5 µm in diameter) contain all of the neurotransmission machinery, including uniform glutamatergic synaptic vesicles (SVs; ~40 nm in diameter) in cytosolic reserve and readily-releasable pools². These vesicles dock at the presynaptic plasma membrane fusion site active zones (AZs), where exocytosis mediates the glutamate neurotransmitter release for trans-synaptic communication. Subsequently, the SVs are retrieved from the plasma membrane via kiss-and-run recycling or clathrin-mediated endocytosis (CME) for repeated exo/endocytosis cycles. The Drosophila NMJ is easily accessible and well-suited for both isolating and characterizing SV cycle mutants. Using forward genetic screens, novel mutations have led to the identification of new genes critical for the SV cycle³. Moreover, reverse genetic approaches starting with already known genes have led to the elucidation of new SV cycle mechanisms through the careful description of mutant cycling phenotypes⁴. The Drosophila NMJ is nearly ideal as an experimental synaptic preparation for dissecting SV endocytosis and exocytosis mechanisms via methods to optically track vesicle cycling during neurotransmission.

A range of fluorescent markers allow visual tracking of vesicles during cycling dynamics, but the most versatile are FM dye analogs which is first synthesized by Mao, F., et al.⁵. Structurally, FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring, with a central region conferring spectral properties. These styryl dyes partition reversibly in membranes, do not 'flip-flop' between membrane leaflets and so are never free in the cytosol, and are far more fluorescent in membranes than water⁵. Reversible insertion into a lipid bilayer causes a 40-fold increase in fluorescence⁶. At neuronal synapses, classic FM dye labeling experiments consist of bathing the synaptic preparation with the dye during depolarizing stimulation to load dye via SV endocytosis. External dye is then washed away and the SV cycle is arrested in a calcium-free ringer solution to image loaded synapses⁷. A second round of stimulation in a dye-free bath triggers FM release through exocytosis, a process that can be followed by measuring the fluorescence intensity decrease. SV populations from a single vesicle to pools containing hundreds of vesicles can be quantitatively monitored^6,7. FM dyes have been used to dissect activity-dependent mobilization of functionally distinct SV pools, and to compare kiss-and-run vs. CME cycling^8,9. The method has been modified to separately assay evoked, spontaneous and miniature synaptic cycle activities (with highly sensitive equipment to detect very small fluorescence changes and reduce photobleaching)¹⁰. Assays can be extended to the ultrastructural level by photoconverting the fluorescent FM signal into an electron-dense label for transmission electron microscopy^11,12,13,14.

Historically, bathing synaptic preparations in a high concentration of potassium (hereafter referred to as "high [K⁺]") has been the method of choice for depolarizing stimulation to induce SV cycling; ranging from the frog cholinergic NMJ⁵, to cultured rodent brain hippocampal neurons¹⁵, to the Drosophila glutamatergic NMJ model^16,17. This high [K⁺] approach is simple, requires no specialized equipment, and is therefore accessible to most labs, but has limitations for both application and data interpretation. A much more physiologically appropriate method is to use suction electrode electrical stimulation of the nerve^4,5,12. This approach drives action potential propagation for direct stimulation of the presynaptic nerve terminal, and results can be directly compared to electrophysiological assays of neurotransmission function^13,14,15, but requires specialized equipment and is technically much more challenging. With the advent of optogenetics, the use of channelrhodopsin neuronal stimulation has additional advantages, including tight spatiotemporal control of channel expression using the binary Gal4/UAS system²⁰. This approach is technically much easier than suction electrode stimulation and requires nothing more than a very cheap LED light source. Here, we employ imaging of FM1-43 (N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl) pyridinium dibromide) to both compare and contrast these three different stimulation methods at the Drosophila NMJ: simple high [K⁺], challenging electrical and new channelrhodopsin approaches.

Protocol

1. Larval Glue Dissection

1. Thoroughly mix 10 parts of silicone elastomer base with 1 part of silicone elastomer curing agent from the elastomer kit (Table of Materials).
2. Coat 22 x 22 mm glass coverslips with the elastomer and cure on a hot plate at 75 ˚C for several hours (until no longer sticky to the touch).
3. Place a single elastomer-coated glass coverslip into the custom-made plexi glass dissection chamber (Figure 1, bottom) in preparation for the larval dissection.
4. Prepare the glue pipettes from borosilicate glass capillary using a standard microelectrode puller to obtain the desired taper and tip size.
5. Gently break off the micropipette tip, and to the other end, attach 2 ft of flexible plastic tube (1/32" interior diameter, ID; 3/32" outside diameter, OD; 1/32" wall; Table of Materials) with mouth fitting (P2 pipette tip).
6. Fill a small container (0.6 mL Eppendorf tube cap) with a small volume (~20 µL) of glue (Table of Materials) in preparation for the larval dissection.
7. Fill the chamber with saline (in mM): 128 NaCl, 2 KCl, 4 MgCl₂, 1 CaCl₂, 70 sucrose, and 5 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES) pH 7.2.
8. Add anti-horse radish peroxidase (HRP) antibody conjugated to Alexa Fluor 647 (anti-HRP:647; dilute 1:10 from a 1 mg/mL stock) for labeling the NMJ presynaptic terminal during dissection^21,22.
9. Using a fine paintbrush (size 2), remove a wandering third instar from the food vial and place onto the elastomer-coated cover glass.
10. Fill the glass micropipette tip with a small volume of glue using negative air pressure generated by mouth with attachment (step 1.5).
11. Position larva dorsal side up with forceps and glue the head to the elastomer-coated coverslip with a small drop of glue using positive air pressure by mouth.
12. Repeat this procedure with the posterior end of the larva, making sure that the animal is stretched taut between the two glue attachments.
13. Using scissors (blades 3 mm; Table of Materials), make a horizontal cut (~1 mm) at posterior and a vertical cut all along the dorsal midline.
14. Using fine forceps (#5, Table of Materials), gently remove dorsal trachea, gut, fat body and other internal organs covering the musculature.
15. Repeat the gluing procedure for the four body wall flaps, making sure to gently stretch the body wall both horizontally and vertically.
16. Lift the ventral nerve cord (VNC) using forceps, carefully cut the motor nerves with the scissors, and then completely remove the VNC.
17. Replace the dissection saline with Ca²⁺-free saline (same as the above dissection saline without the CaCl₂) to stop SV cycling.

2. Option 1: High [K⁺] FM Dye Loading

1. From a FM1-43 stock solution (4 mM), add 1 µL to 1 mL of 90 mM KCl solution (high [K⁺] in dissection saline) for a final concentration of 4 µM.
2. Using a pipette, replace the Ca²⁺-free saline in the imaging chamber with the high [K⁺] FM dye solution to stimulate SV endocytosis dye uptake.
3. Immediately start a digital timer for the pre-determined duration of the high [K⁺] depolarizing stimulation period (e.g., 5 min; Figure 2).
4. To confirm a healthy larval preparation, note the strong contractions of the musculature for the duration of the high [K⁺] depolarization period.
5. When the timer period ends, quickly remove the high [K⁺] FM dye solution and replace with Ca²⁺-free saline to stop SV cycling.
6. Wash in quick succession with the Ca²⁺-free saline (5x for 1 min) to ensure the high [K⁺] FM dye solution is completely removed.
7. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging with the confocal microscope.

3. Imaging: Confocal Microscopy

1. Use an upright confocal microscope with a 40X water immersion objective to image NMJ dye fluorescence (other microscopes can be used).
2. Image muscle 4 NMJ of abdominal segments 2-4 (other NMJs can be imaged) and collect images using appropriate software (Table of Materials).
3. Use a HeNe 633 nm laser to excite HRP:647 (with long-pass filter > 635 nm) and an Argon 488 nm laser to excite FM1-43 (with bandpass filter 530-600 nm).
4. Operationally determine optimal gain and offset for both channels.
   NOTE: These settings will remain constant throughout the rest of the experiment.
5. Take a confocal Z-stack through the entire selected NMJ from the HRP-marked top to bottom of the synaptic terminal.
6. Take careful note of the NMJ imaged (segment, side and muscle) to ensure excess to the exact same NMJ after FM dye unloading.

4. High [K⁺] Stimulation: FM Dye Unloading

1. Replace Ca²⁺-free saline with the high [K⁺] saline (without FM1-43 dye) to drive depolarization, SV exocytosis and dye release.
2. Immediately start a digital timer for the pre-determined duration of the high [K⁺] stimulation period (e.g., 2 min; Figure 2).
3. When the timer period ends, immediately remove the high [K⁺] saline and replace with Ca²⁺-free saline to stop SV cycling.
4. Wash in quick succession with Ca²⁺-free saline (5x for 1 min) to ensure the high [K⁺] saline is completely removed.
5. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging with the confocal microscope.
6. Be certain to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.

5. Option 2: Electrical Stimulation FM Dye Loading

1. Prepare a suction pipette using the microelectrode puller (Table of Materials) to obtain the required taper and tip size.
2. Fire-polish the microelectrode tip with a micro-forge until a single motor nerve can be sucked up with a tight fit.
3. Slide suction pipette onto the electrode holder on a micromanipulator and attach to the long flexible plastic tube and a syringe.
4. Set stimulator parameters (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 5 min (Figure 2) or 1 min (Figure 3)).
5. Replace the Ca²⁺-free saline on the larval preparation with above FM1-43 saline (4 µM; 1 mM CaCl₂) on the electrophysiology rig.
6. Put the preparation on the microscope stage and raise the stage until the larva and suction pipette are in focus (40X water-immersion objective).
7. Suck up a loop of cut motor nerve innervating the selected hemisegment with negative air pressure generated by the syringe into the suction electrode.
8. Test the suction electrode function with a short burst of stimulation while visually monitoring for the muscle contraction in the selected hemisegment.
9. Stimulate the motor nerve using selected parameters (step 5.4) to drive SV endocytosis and FM1-43 dye uptake (Figure 2).
10. Wash in quick succession with Ca²⁺-free saline (5x for 1 min) to ensure the FM1-43 dye solution is completely removed.
11. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging using the confocal imaging protocol from above.
12. Take careful note of the NMJ imaged (segment, side and muscle) to ensure access to the exact same NMJ after FM dye unloading.

6. Electrical Stimulation: FM Dye Unloading

1. Replace the Ca²⁺-free saline with regular saline (without FM1-43 dye)and place the preparation back on the electrophysiology rig stage.
2. Set the stimulator parameters for unloading (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 2 min (Figure 2) or 20 s (Figure 3)).
3. Suck the same motor nerve into the same electrode as above, and then stimulate to activate SV exocytosis and FM1-43 dye release.
4. Wash in quick succession with Ca²⁺-free saline (5x for 1 min) to ensure the external dye is completely removed.
5. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging with the confocal microscope.
6. Ensure to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.

7. Option 3: Channelrhodopsin Stimulation FM Dye Loading

1. Raise ChR2-expressing larvae on food containing the ChR2 co-factor all-trans retinal (dissolved in ethanol; 100 µM final concentration).
2. Place the larval preparation in the plexiglass chamber on a dissection microscope stage equipped with a camera port.
3. Attach a blue LED (470 nm; Table of Materials) to a programmable stimulator using a coaxial cable and place the LED into the camera port.
4. Focus the blue LED light beam onto the dissected larval function using the microscope zoom function.
5. Replace the Ca²⁺-free saline on the larval preparation with above FM1-43 saline (4 µM; 1 mM CaCl₂) on the optogenetic stage.
6. Set the LED parameters using the stimulator (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 5 min (Figure 2)).
7. Start the light stimulation and track with a timer for the pre-determined duration of the optogenetic stimulation period (e.g., 5 min; Figure 2).
8. When the timer stops, quickly remove the FM dye solution and replace with Ca²⁺-free saline to stop the SV cycling.
9. Wash in quick succession with the Ca²⁺-free saline (5x for 1 min) to ensure the FM dye solution is completely removed.
10. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging with the confocal microscope using imaging protocol from above.
11. Take careful note of the NMJ imaged (segment, side and muscle) to ensure access to the exact same NMJ after FM dye unloading.

8. Channelrhodopsin Stimulation: FM Dye Unloading

1. Replace Ca²⁺-free saline with regular saline (without FM1-43 dye)on the dissection microscope stage with camera port LED focused on the larva.
2. Set the stimulator parameters for unloading (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 2 min (Figure 2)).
3. Start the light stimulation and track with a timer for the pre-determined duration of the optogenetic stimulation period (e.g., 2 min; Figure 2).
4. When the timer period ends, quickly remove the FM dye solution and replace with Ca²⁺-free saline to stop the SV cycling.
5. Wash in quick succession with Ca²⁺-free saline (5x for 1 min) to ensure the external dye is completely removed.
6. Maintain the larval preparation in fresh Ca²⁺-free saline for immediate imaging with the confocal microscope.
7. Ensure to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.

9. Fluorescence Quantification

1. Load the image in Image J (NIH open source) and create a maximum intensity projection by clicking Image | Stacks | Z Project.
2. Using the anti-HRP:647 channel, go to Image | Adjust | Threshold and slide the top tool bar until just the NMJ is highlighted.
3. Using the wand tool, click on the NMJ. If the NMJ is discontinuous, hold the Shift button and select all parts.
4. Change the image to the FM1-43 dye channel and go to Analyze | Measure to obtain the fluorescence measurement.
5. Repeat steps 9.1-9.4 for the "unloaded" image from the same NMJ (identified segment, side and muscle).
6. To obtain the percentage of dye that was unloaded, take the ratio of the unloaded/loaded fluorescence intensities.
   NOTE: This procedure can be modified to analyze fluorescence on a bouton-per-bouton basis using either the "oval" or "freehand" selection tools. Background fluorescence can be subtracted by sampling the muscle fluorescence. Agents can also be added to reduce this background.

Results

Figure 1 shows the work-flow for the activity-dependent FM dye imaging protocol. The experiment always begins with the same larval glue dissection, regardless of the stimulation method subsequently used. Figure 1a is a schematic of a dissected larva, showing the ventral nerve cord (VNC), radiating nerves and repeated hemisegmental muscle pattern. The VNC is removed and the preparation bathed in a 4 µM solution of FM1-43 (

Discussion

High [K⁺] saline depolarizing stimulation is by far the easiest of the three options for activity-dependent FM dye cycling, but likely the least physiological²⁹. This simple method depolarizes every accessible cell in the entire animal, and so does not allow directed studies. It may be possible to locally apply high [K⁺] saline with a micropipette, but this will still depolarize pre/postsynaptic cells and likely synapse-associated glia¹. Another major ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
SylGard 184 Silicone Elastomer Kit	Fisher Scientific	NC9644388	To put on cover glass for dissections
Microscope Cover Glass 22x22-1	Fisherbrand	12-542-B	To put SylGard on for dissections
Aluminum Top Hot Plate Type 2200	Thermolyne	HPA2235M	To cure the SylGard
Plexi glass dissection chamber	N/A	N/A	Handmade
Borosilicate Glass Capillaries	WPI	1B100F-4	To make suction and glue micropipettes
Laser-Based Micropipette Puller	Sutter Instrument	P-2000	To make suction and glue micropipettes
Tygon E-3603 Laboratory Tubing	Component Supply Co.	TET-031A	For mouth and suction pipette
P2 pipette tip	USA Scientific	1111-3700	For mouth pipette
0.6-mL Eppendorf tube cap	Fisher Scientific	05-408-120	Used to put glue in for dissection
Vetbond 3M	WPI	vetbond	Glue used for dissections
Potassium Chloride	Fisher Scientific	P-217	Forsaline
Sodium Chloride	Millipore Sigma	S5886	For saline
Magnesium Chloride	Fisher Scientific	M35-500	For saline
Calcium Chloride Dihydrate	Millipore Sigma	C7902	For saline
Sucrose	Fisher Scientific	S5-3	For saline
HEPES	Millipore Sigma	H3375	For saline
HRP:Alexa Fluor 647	Jackson ImmunoResearch	123-605-021	To label neuronal membranes
Paintbrush	Winsor & Newton	94376864793	To manipulate the larvae
Dumont Dumostar Tweezers #5	WPI	500233	Used during dissection
7 cm McPherson-Vannas Microscissors (blades 3 mm)	WPI	14177	Used during dissection
FM1-43	Fisher Scientific	T35356	Fluorescent styryl dye
Digital Timer	VWR	62344-641	For timing FM dye load/unload
LSM 510 META laser-scanning confocal microscope	Zeiss		For imaging the fluorescent markers
Zen 2009 SP2 version 6.0	Zeiss		Software for imaging on confocal
HeNe 633nm laser	Lasos		To excite HRP:647 during imaging
Argon 488nm laser	Lasos		To excite the FM dye during imaging
Micro-Forge	WPI	MF200	To fire polish glass micropipettes
20mL Syringe Slip Tip	BD	301625	To suck up the axon for electrical stimulation.
Micro Manipulator (magnetic base)	Narishige	MMN-9	To control the suction electrode for electrical stimulation
Stimulator	Grass	S48	To control the LED and electrical stimulation
Zeiss Axioskop Microscope	Zeiss		Used during electrical stimulation.
40X Achroplan Water Immersion Objective	Zeiss		Used during electrical stimulation and confocal imaging
All-trans Retinal	Millipore Sigma	R2500	Essential co-factor for ChR2
Zeiss Stemi Microscope with camera port	Zeiss	2000-C	Used during channelrhodopsin stimulation
LED 470nm	ThorLabs	M470L2	Used for ChR activation
T-Cube LED Driver	ThorLabs	LEDD1B	To control the LED
LED Power Supply	Cincon Electronics Co.	TR15RA150	To power the LED
Optical Power and Energy Meter	ThorLabs	PM100D	To measure LED intensity