Analyzing Synaptic Vesicle Recycling and Fluorescent Dye Uptake in Drosophila Larvae

Protokół

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 plexiglass 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 neuromuscular junction (NMJ) presynaptic terminal during dissection.
9. Using a fine paintbrush (size 2), remove a wandering third instar larva from the food vial and place it 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 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 synaptic vesicle (SV) cycling.

2. 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.

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.
11. Take careful note of the NMJ imaged (segment, side and muscle) to ensure access to the exact same NMJ after FM dye unloading.

4. 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.

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Wyniki

Figure 1: Flowchart of FM1-43 dye loading protocol at the Drosophila NMJ. The larval glue dissection produces a flattened neuromusculature preparation, with the ventral nerve cord (VNC) projecting segmental nerves from the ventral midline (Vm) to the hemisegmentally repeated body wall muscle arrays (step a). The VNC is cut free, and the entire larval dissection is then incubated in the pink FM1-43 solution (4 µM) in preparation for stimulation (step b). FM1-43 is then loaded with a selected stimulation paradigm (step 3); with the options of high [K+] depolarization of the entire larva (cI), suction electrode stimulation of a single motor nerve (cII), or light-driven activation of highly targeted channelrhodopsin (cIII). FM1-43 incorporation is arrested using Ca2+-free saline and the dye-loaded NMJ imaged (step d). A second stimulation is then done without FM1-43 in the bath to drive dye synaptic vesicle exocytosis (step e). The same NMJ is then re-imaged to assay the unloaded synaptic terminal (step f). Fluorescent intensity is measured from both loaded and unloaded NMJs to quantify SV endocytosis and SV exocytosis levels. The bottom panel shows the construction parameters and dimensions for the transparent acrylic chamber used for these studies.

Figure 2: FM dye loading and unloading comparison of all stimulation methods. Comparison of FM1-43 dye loading and unloading in the wandering third instar NMJ with 1) high [K+] depolarization of the entire larval preparation (top), 2) suction electrode electrical stimulation of the motor nerve (middle) and 3) light-driven activation of the targeted channelrhodopsin (ChR2) only in motor neurons (bottom). (A) The larval NMJ labeled with the anti-HRP:647 presynaptic membrane marker (blue, left), loaded with FM1-43 via high [K+] depolarization for 5 min (middle) and then unloaded via high [K+] depolarization for 2 min. (B) Comparison with suction electrode electrical nerve stimulation with the same stimuli periods for both FM1-43 dye loading and unloading. (C) Targeted vglut-Gal4>UAS-ChR2-H134R expression in motor neurons activated with blue (470 nm) light for the same stimuli periods of FM1-43 dye loading and unloading. Asterisks refer to insets displaying higher magnification boutons. The scale bar is 10 µm, with inset synaptic boutons enlarged 3.5X from main panels.

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Materiały

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
HRP:Alexa Fluor 647	Jackson ImmunoResearch	123-605-021	To label neuronal membranes
HEPES	Millipore Sigma	H3375	For saline
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
Stimulator	Grass	S48	To control the LED and electrical stimulation
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
40X Achroplan Water Immersion Objective	Zeiss	Used during electrical stimulation and confocal imaging