Name: fm dye cycling at the synapse: comparing high potassium depolarization, electrical and channelrhodopsin stimulation
Uploaded: 2018-05-24T15:00:00.000+00:00
Duration: 8 min 31 s
Description: Watch this Scientific Journal Video about FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation at JoVE.com

We thank Broadie Lab members for contributions to this article. This work was supported by NIH R01s MH096832 and MH084989 to K.B., and NIH predoctoral fellowship F31 MH111144 to D.L.K.

1. Larval Glue Dissection
<ol>
	<li>Thoroughly mix 10 parts of silicone elastomer base with 1 part of silicone elastomer curing agent from the elastomer kit (Table of Materials).</li>
	<li>Coat 22 x 22 mm glass coverslips with the elastomer and cure on a hot plate at 75 &#730;C for several hours (until no longer sticky to the touch).</li>
	<li>Place a single elastomer-coated glass coverslip into the custom-made plexi glass dissection chamber (Figure 1, bottom) in preparation for the larval dissection.</li>
	<li>Prepare the glue pipettes from borosilicate glass capillary using a standard microelectrode puller to obtain the desired taper and tip size.</li>
	<li>Gently break off the micropipette tip, and to the other end, attach 2 ft of flexible plastic tube (1/32&#34; interior diameter, ID; 3/32&#34; outside diameter, OD; 1/32&#34; wall; Table of Materials) with mouth fitting (P2 pipette tip).</li>
	<li>Fill a small container (0.6 mL Eppendorf tube cap) with a small volume (~20 &#181;L) of glue (Table of Materials) in preparation for the larval dissection.</li>
	<li>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.</li>
	<li>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 dissection21,22.</li>
	<li>Using a fine paintbrush (size 2), remove a wandering third instar from the food vial and place onto the elastomer-coated cover glass.</li>
	<li>Fill the glass micropipette tip with a small volume of glue using negative air pressure generated by mouth with attachment (step 1.5).</li>
	<li>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.</li>
	<li>Repeat this procedure with the posterior end of the larva, making sure that the animal is stretched taut between the two glue attachments.</li>
	<li>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.</li>
	<li>Using fine forceps (#5, Table of Materials), gently remove dorsal trachea, gut, fat body and other internal organs covering the musculature.</li>
	<li>Repeat the gluing procedure for the four body wall flaps, making sure to gently stretch the body wall both horizontally and vertically.</li>
	<li>Lift the ventral nerve cord (VNC) using forceps, carefully cut the motor nerves with the scissors, and then completely remove the VNC.</li>
	<li>Replace the dissection saline with Ca2+-free saline (same as the above dissection saline without the CaCl2) to stop SV cycling.</li>
</ol>
2. Option 1: High [K+] FM Dye Loading
<ol>
	<li>From a FM1-43 stock solution (4 mM), add 1 &#181;L to 1 mL of 90 mM KCl solution (high [K+] in dissection saline) for a final concentration of 4&#160;&#181;M.</li>
	<li>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.</li>
	<li>Immediately start a digital timer for the pre-determined duration of the high [K+] depolarizing stimulation period (e.g., 5 min; Figure 2).</li>
	<li>To confirm a healthy larval preparation, note the strong contractions of the musculature for the duration of the high [K+] depolarization period.</li>
	<li>When the timer period ends, quickly remove the high [K+] FM dye solution and replace with Ca2+-free saline to stop SV cycling.</li>
	<li>Wash in quick succession with the Ca2+-free saline (5x for 1 min) to ensure the high [K+] FM dye solution is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging with the confocal microscope.</li>
</ol>
3. Imaging: Confocal Microscopy
<ol>
	<li>Use an upright confocal microscope with a 40X water immersion objective to image NMJ dye fluorescence (other microscopes can be used).</li>
	<li>Image muscle 4 NMJ of abdominal segments 2-4 (other NMJs can be imaged) and collect images using appropriate software (Table of Materials).</li>
	<li>Use a HeNe 633 nm laser to excite HRP:647 (with long-pass filter &#62; 635 nm) and an Argon 488 nm laser to excite FM1-43 (with bandpass filter 530-600&#160;nm).</li>
	<li>Operationally determine optimal gain and offset for both channels. 
	NOTE: These settings will remain constant throughout the rest of the experiment.</li>
	<li>Take a confocal Z-stack through the entire selected NMJ from the HRP-marked top to bottom of the synaptic terminal.</li>
	<li>Take careful note of the NMJ imaged (segment, side and muscle) to ensure excess to the exact same NMJ after FM dye unloading.</li>
</ol>
4. High [K+] Stimulation: FM Dye Unloading
<ol>
	<li>Replace Ca2+-free saline with the high [K+] saline (without FM1-43 dye) to drive depolarization, SV exocytosis and dye release.</li>
	<li>Immediately start a digital timer for the pre-determined duration of the high [K+] stimulation period (e.g., 2 min; Figure 2).</li>
	<li>When the timer period ends, immediately remove the high [K+] saline and replace with Ca2+-free saline to stop SV cycling.</li>
	<li>Wash in quick succession with Ca2+-free saline (5x for 1 min) to ensure the high [K+] saline is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging with the confocal microscope.</li>
	<li>Be certain to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.</li>
</ol>
5. Option 2: Electrical Stimulation FM Dye Loading
<ol>
	<li>Prepare a suction pipette using the microelectrode puller (Table of Materials) to obtain the required taper and tip size.</li>
	<li>Fire-polish the microelectrode tip with a micro-forge until a single motor nerve can be sucked up with a tight fit.</li>
	<li>Slide suction pipette onto the electrode holder on a micromanipulator and attach to the long flexible plastic tube and a syringe.</li>
	<li>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)).</li>
	<li>Replace the Ca2+-free saline on the larval preparation with above FM1-43 saline (4 &#181;M; 1 mM CaCl2) on the electrophysiology rig.</li>
	<li>Put the preparation on the microscope stage and raise the stage until the larva and suction pipette are in focus (40X water-immersion objective).</li>
	<li>Suck up a loop of cut motor nerve innervating the selected hemisegment with negative air pressure generated by the syringe into the suction electrode.</li>
	<li>Test the suction electrode function with a short burst of stimulation while visually monitoring for the muscle contraction in the selected hemisegment.</li>
	<li>Stimulate the motor nerve using selected parameters (step 5.4) to drive SV endocytosis and FM1-43 dye uptake (Figure 2).</li>
	<li>Wash in quick succession with Ca2+-free saline (5x for 1 min) to ensure the FM1-43 dye solution is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging using the confocal imaging protocol from above.</li>
	<li>Take careful note of the NMJ imaged (segment, side and muscle) to ensure access to the exact same NMJ after FM dye unloading.</li>
</ol>
6. Electrical Stimulation: FM Dye Unloading
<ol>
	<li>Replace the Ca2+-free saline with regular saline (without FM1-43 dye)and place the preparation back on the electrophysiology rig stage.</li>
	<li>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)).</li>
	<li>Suck the same motor nerve into the same electrode as above, and then stimulate to activate SV exocytosis and FM1-43 dye release.</li>
	<li>Wash in quick succession with Ca2+-free saline (5x for 1 min) to ensure the external dye is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging with the confocal microscope.</li>
	<li>Ensure to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.</li>
</ol>
7. Option 3: Channelrhodopsin Stimulation FM Dye Loading
<ol>
	<li>Raise ChR2-expressing larvae on food containing the ChR2 co-factor all-trans retinal (dissolved in ethanol; 100 &#181;M final concentration).</li>
	<li>Place the larval preparation in the plexiglass chamber on a dissection microscope stage equipped with a camera port.</li>
	<li>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.</li>
	<li>Focus the blue LED light beam onto the dissected larval function using the microscope zoom function.</li>
	<li>Replace the Ca2+-free saline on the larval preparation with above FM1-43 saline (4 &#181;M; 1 mM CaCl2) on the optogenetic stage.</li>
	<li>Set the LED parameters using the stimulator (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 5 min (Figure 2)).</li>
	<li>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).</li>
	<li>When the timer stops, quickly remove the FM dye solution and replace with Ca2+-free saline to stop the SV cycling.</li>
	<li>Wash in quick succession with the Ca2+-free saline (5x for 1 min) to ensure the FM dye solution is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging with the confocal microscope using imaging protocol from above.</li>
	<li>Take careful note of the NMJ imaged (segment, side and muscle) to ensure access to the exact same NMJ after FM dye unloading.</li>
</ol>
8. Channelrhodopsin Stimulation: FM Dye Unloading
<ol>
	<li>Replace Ca2+-free saline with regular saline (without FM1-43 dye)on the dissection microscope stage with camera port LED focused on the larva.</li>
	<li>Set the stimulator parameters for unloading (e.g., 15 V, 20 Hz frequency, 20 ms duration and time of 2 min (Figure 2)).</li>
	<li>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).</li>
	<li>When the timer period ends, quickly remove the FM dye solution and replace with Ca2+-free saline to stop the SV cycling.</li>
	<li>Wash in quick succession with Ca2+-free saline (5x for 1 min) to ensure the external dye is completely removed.</li>
	<li>Maintain the larval preparation in fresh Ca2+-free saline for immediate imaging with the confocal microscope.</li>
	<li>Ensure to image the FM1-43 dye fluorescence at the same NMJ noted above using the same confocal settings.</li>
</ol>
9. Fluorescence Quantification
<ol>
	<li>Load the image in Image J (NIH open source) and create a maximum intensity projection by clicking Image | Stacks | Z Project.</li>
	<li>Using the anti-HRP:647 channel, go to Image | Adjust | Threshold and slide the top tool bar until just the NMJ is highlighted.</li>
	<li>Using the wand tool, click on the NMJ. If the NMJ is discontinuous, hold the Shift button and select all parts.</li>
	<li>Change the image to the FM1-43 dye channel and go to Analyze | Measure to obtain the fluorescence measurement.</li>
	<li>Repeat steps 9.1-9.4 for the &#34;unloaded&#34; image from the same NMJ (identified segment, side and muscle).</li>
	<li>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 &#34;oval&#34; or &#34;freehand&#34; selection tools. Background fluorescence can be subtracted by sampling the muscle fluorescence. Agents can also be added to reduce this background.</li>
</ol>

The authors declare no competing interests.

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 ...

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 &#181;m 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 &#39;flip-flop&#39; 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 monitored6,7. FM dyes have been used to dissect activity-dependent mobilization of functionally distinct SV pools, and to compare kiss-and-run vs. CME cycling8,9. 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 microscopy11,12,13,14.
Historically, bathing synaptic preparations in a high concentration of potassium (hereafter referred to as &#34;high [K+]&#34;) 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 model16,17. 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 nerve4,5,12. 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 function13,14,15, 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.

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

fm dye cycling at the synapse: comparing high potassium depolarization, electrical and channelrhodopsin stimulation

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 &#62;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.

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 &#181;M solution of FM1-43 (<st...

Watch this Scientific Journal Video about FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation at JoVE.com

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

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.

FM dyes are used to study the synaptic vesicle (SV) cycle. These amphipathic probes have a hydrophilic head and hydrophobic tail, making them ...

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Channels and the Electrical Properties of Membranes

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8.4K Views.  Vanderbilt University. Synaptic Vesikel (SV) fietsen is het mechanisme van de kern van de intercellulaire communicatie in neuronale synapsen. FM kleurstof opname- en release zijn het belangrijkste middel van kwantitatief onderzoek SV endo- en exocytose. Hier, vergelijken wij alle stimulatie methoden om te rijden FM1-43 fietsen in de Drosophila motorische eindplaat (NMJ) model synaps.

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

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available to evoke excitatory junctional potentials (EJPs) in this preparation involves the use of suction electrodes. In both research and teaching labs, students often have difficulty maneuvering and manipulating this type of stimulating electrode. In the present work, we show how to remotely stimulate synaptic potentials at the larval NMJ without the use of suction electrodes. By expressing channelrhodopsin2 (ChR2) 4,5,6 in Drosophila motor neurons using the GAL4-UAS system 7, and making minor changes to a basic electrophysiology rig, we were able to reliably evoke EJPs with pulses of blue light. This technique could be of particular use in neurophysiology teaching labs where student rig practice time and resources are limited.

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

This procedure uses a blue light-activated algal channel and cell-specific genetic expression tools to evoke synaptic potentials with light pulses at the neuromuscular junction (NMJ) in Drosophila larvae. This technique is an inexpensive and easy-to-use alternative to suction electrode stimulation for synaptic physiology studies in research and teaching laboratories.

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available ...

Biology

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...

TIRFM and pH-sensitive GFP-probes to Evaluate Neurotransmitter Vesicle Dynamics in SH-SY5Y Neuroblastoma Cells: Cell Imaging and Data Analysis

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real time and dissecting the different steps of exo-endocytosis at the single-vesicle level are crucial for understanding synaptic functions in health and disease.
Genetically-encoded pH-sensitive probes directly targeted to synaptic vesicles and Total Internal Reflection Fluorescence Microscopy (TIRFM) provide the spatio-temporal resolution necessary to follow vesicle dynamics. The evanescent field generated by total internal reflection can only excite fluorophores placed in a thin layer (&#60;150 nm) above the glass cover on which cells adhere, exactly where the processes of exo-endocytosis take place. The resulting high-contrast images are ideally suited for vesicles tracking and quantitative analysis of fusion events.
In this protocol, SH-SY5Y human neuroblastoma cells are proposed as a valuable model for studying neurotransmitter release at the single-vesicle level by TIRFM, because of their flat surface and the presence of dispersed vesicles. The methods for growing SH-SY5Y as adherent cells and for transfecting them with synapto-pHluorin are provided, as well as the technique&#160;to perform TIRFM and imaging. Finally, a strategy aiming to select, count, and analyze fusion events at whole-cell and single-vesicle levels is presented.
To validate the imaging procedure and data analysis approach, the dynamics of pHluorin-tagged vesicles are&#160;analyzed under resting and stimulated (depolarizing potassium concentrations) conditions. Membrane depolarization increases the frequency of fusion events and causes a parallel raise of the net fluorescence signal recorded in whole cell. Single-vesicle analysis reveals modifications of fusion-event behavior (increased peak height and width). These data suggest that potassium depolarization not only induces a massive neurotransmitter release but also modifies the mechanism of vesicle fusion and recycling.
With the appropriate fluorescent probe, this technique can be employed in different cellular systems to dissect the mechanisms of constitutive and stimulated secretion.

This paper provides a method for investigating neurotransmitter vesicle dynamics in neuroblastoma cells, using a synaptobrevin2-pHluorin construct and Total Internal Reflection Fluorescence Microscopy. The strategy developed for image processing and data analysis is also reported.

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real ...

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to blindness in many retinal diseases. Calcium (Ca2+), a key second messenger in photoreceptor signaling and metabolism, has been proposed to be indirectly linked with photoreceptor degeneration in various animal models. Systematically studying these aspects of cone physiology and pathophysiology has been hampered by the difficulties of electrically recording from these small cells, in particular in the mouse where the retina is dominated by rod photoreceptors. To circumvent this issue, we established a two-photon Ca2+ imaging protocol using a transgenic mouse line that expresses the genetically encoded Ca2+ biosensor TN-XL exclusively in cones and can be crossbred with mouse models for photoreceptor degeneration. The protocol described here involves preparing vertical sections (&#8220;slices&#8221;) of retinas from mice and optical imaging of light stimulus-evoked changes in cone Ca2+ level. The protocol also allows &#8220;in-slice measurement&#8221; of absolute Ca2+ concentrations; as the recordings can be followed by calibration. This protocol enables studies into functional cone properties and is expected to contribute to the understanding of cone Ca2+ signaling as well as the potential involvement of Ca2+ in photoreceptor death and retinal degeneration.

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

We describe a protocol to monitor Ca2+ dynamics in the axon terminals of cone photoreceptors using an ex-vivo&#160;slice preparation of the mouse retina. This protocol allows comprehensive studies of cone Ca2+ signaling in an important mammalian model system, the mouse.

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to ...

Loading a Calcium Dye into Frog Nerve Endings Through the Nerve Stump: Calcium Transient Registration in the Frog Neuromuscular Junction

One of the most feasible methods of measuring presynaptic calcium levels in presynaptic nerve terminals is optical recording. It is based on using calcium-sensitive fluorescent dyes that change their emission intensity or wavelength depending on the concentration of free calcium in the cell. There are several methods used to stain cells with calcium dyes. Most common are the processes of loading the dyes through a micropipette or pre-incubating with the acetoxymethyl ester forms of the dyes. However, these methods are not quite applicable to neuromuscular junctions (NMJs) due to methodological issues that arise. In this article, we present a method for loading a calcium-sensitive dye through the frog nerve stump of the frog nerve into the nerve endings. Since entry of external calcium into nerve terminals and the subsequent binding to the calcium dye occur within the millisecond time-scale, it is necessary to use a fast imaging system to record these interactions. Here, we describe a protocol for recording the calcium transient with a fast CCD camera.

Here, we describe a method for loading a calcium-sensitive dye through the frog nerve stump into the nerve endings. We also present a protocol for the recording and analysis of fast calcium transients in the peripheral nerve endings.

One of the most feasible methods of measuring presynaptic calcium levels in presynaptic nerve terminals is optical recording. It is based on using ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Live Imaging of Synaptic Vesicle Recycling in the Neuromuscular Junction of Dissected Larval Zebrafish

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) in response to an action potential (AP). Since SVs are recycled locally at the presynaptic terminal, coordination of SV exocytosis and endocytosis is important for sustained synaptic transmission. A pH-sensitive green fluorescent protein, called pHluorin, provides a powerful tool to monitor SV exo/endocytosis by targeting it to the SV lumen. However, tracking AP-driven SV recycling with the pHluorin-based probes is still largely limited to in vitro culture preparations because the introduction of genetically encoded probes and subsequent optical imaging is technically challenging in general for in vivo animal models or tissue preparations. Zebrafish is a model system offering valuable features, including ease of genetic manipulation, optical clarity, and rapid external development. We recently generated a transgenic zebrafish that highly expresses a pHluorin-labeled probe at motor neuron terminals and developed a protocol to monitor AP-driven SV exo/endocytosis at the neuromuscular junction (NMJ), a well-established synapse model that forms in vivo. In this article, we show how to prepare larval zebrafish NMJ preparation suitable for pHluorin imaging. We also show that the preparation allows time-lapse imaging under conventional upright epifluorescence microscope, providing a cost-effective platform for analyzing NMJ function.

The zebrafish larval neuromuscular junction is an attractive model for studying synaptic physiology. It is amenable to many experimental techniques, including electrophysiology and optical imaging. Here, we describe a protocol for imaging synaptic transmission using a pHluorin-based probe under an upright epifluorescence microscope.

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) ...

FM Dyes in Vesicle Recycling

FM dyes are a class of fluorescent molecules that has found important use in studying the vesicle recycling process. By virtue of a chemical structure, these molecules can insert themselves into the outer leaflet of phospholipid bilayer membranes. After membrane insertion, they are internalized into the cell via endocytosed vesicles, and released when these vesicles recycle back to the membrane. Since, these dyes fluoresce strongly in the hydrophobic environment within membranes and weakly in the extracellular compartment, FM fluorescence levels can be used to track vesicular activity throughout the recycling process.This video provides an introduction to the use of FM dyes in experiments aimed to examine vesicle recycling. We first review the biochemistry of FM dyes and how their properties permit their use in these experiments. We then go through a general protocol for using FM dyes in such studies, and finally, discuss some recent research that makes use of these unique molecules.

FM dyes are a class of fluorescent molecules that has found important use in studying the vesicle recycling process. By virtue of a chemical structure, ...

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Advanced Biology

Microelectrode-Mediated FM Dye Loading and Unloading in Drosophila Larvae Nerve Cells

Source: Kopke, D. L., et al., <a href="https://app.jove.com/v/57765">FM dye cycling at the synapse: comparing high potassium depolarization, electrical and channelrhodopsin stimulation.</a> J. Vis. Exp. (2018)
In this video, Drosophila larvae nerve fibers are immersed in FM dye, which binds to cell membrane lipids. Suction microelectrode and electrical pulses stimulate the endocytosis process, incorporating the FM dye into newly formed endocytic vesicles. This method, which visualizes presynaptic stages of neurotransmission, is useful for studying synaptic function and neurotransmitter release.

1. Larval Glue Dissection

	Thoroughly mix 10 parts of silicone elastomer base with 1 part of silicone elastomer curing agent from the elastomer kit.
	...

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

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