eoe sub articles

EoE Sub Articles

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 plexiglass 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 &#956;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 neuromuscular junction (NMJ) presynaptic terminal during dissection.</li>
	<li>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.</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 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 synaptic vesicle (SV) cycling.</li>
</ol>
2. 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 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>
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 &#956;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 &#956;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.</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>
4. 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>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>HRP:Alexa Fluor 647</td>
			<td>Jackson ImmunoResearch</td>
			<td>123-605-021</td>
			<td>To label neuronal membranes</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Millipore Sigma</td>
			<td>H3375</td>
			<td>For saline</td>
		</tr>
		<tr>
			<td>Paintbrush</td>
			<td>Winsor &#38; 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</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</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>To excite HRP:647 during imaging</td>
			<td></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>Stimulator</td>
			<td>Grass</td>
			<td>S48</td>
			<td>To control the LED and electrical stimulation</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>
		<tr>
			<td>40X Achroplan Water Immersion Objective</td>
			<td>Zeiss</td>
			<td>Used during electrical stimulation and confocal imaging</td>
			<td></td>
		</tr>
	</tbody>
</table>

analyzing synaptic vesicle recycling and fluorescent dye uptake in drosophila larvae

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).
This video demonstrates a protocol to study synaptic vesicle recycling at neuromuscular junctions in channelrhodopsin-expressing Drosophila larvae using a fluorescent dye. Blue light activates the channelrhodopsins, causing calcium influx and synaptic vesicle exocytosis. The dye integrates into the neuronal membranes and is internalized during endocytosis, resulting in enhanced fluorescence. Further blue light stimulation induces exocytosis and dye release, which is shown by reduced fluorescence.

<img alt="Figure 1" src="/files/ftp_upload/22876/22876_Figure1.jpg" /> 
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 FM1-4...

Analyzing Synaptic Vesicle Recycling and Fluorescent Dye Uptake in Drosophila Larvae

1. Larval Glue Dissection

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

Place a live Drosophila larval neuromuscular preparation in a recording chamber containing calcium-free saline.&#160;
The larval neurons express a light-sensitive cation channel.
Switch to calcium-added saline containing a fluorescent dye.
The dye fluoresces weakly in the non-internalized state but strongly when internalized by the neurons.
Illuminate the larva with blue light.
This activates the channel, inducing cation influx and neuron depolarization, initiating an action potential.
The action potential reaches the synaptic terminal at the neuromuscular junction, allowing calcium influx and vesicle exocytosis.
Subsequently, the dye-bound membrane is internalized during vesicle recycling.
Wash with calcium-free saline to stop vesicle recycling and remove any non-internalized dye.
Visualize the fluorescence signal.
Next, introduce dye-free calcium-added saline and illuminate again.
This induces another action potential, calcium influx, and dye-containing vesicle exocytosis.
Visualize the signal again.
The dye release due to vesicle exocytosis reduces the fluorescence compared to the state during vesicle internalization.

analyzing-synaptic-vesicle-recycling-fluorescent-dye-uptake

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

39 Aufrufe. Quelle: Kopke, D. L., et al., FM Farbstoffkreislauf an der Synapse: Vergleich von hoher Kalium-Depolarisation, elektrischer und Channelrhodopsin-Stimulation. J. Vis. Exp. (2018). Dieses Video demonstriert ein Protokoll zur Untersuchung des synaptischen Vesikelrecyclings an neuromuskulären Verbindungen in Channelrhodopsin-exprimierenden Drosophila-Larven unter Verwendung eines Fluoreszenzfarbstoffs. Blaues Licht aktiviert die Kanalrhodopsine, was zu Kalziumeinstrom und synaptischer Vesikelexozy...

Serie: Analyse des synaptischen Vesikelrecyclings und der Aufnahme von Fluoreszenzfarbstoffen in Drosophila-Larven - Konzept

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.

Ganzzellige Patch-Clamp-Aufnahmen zur elektrophysiologischen Bestimmung der Ionenselektivität in Channelrhodopsinen

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

Forschung

JoVE Journal

Neurowissenschaften

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.

Messung der synaptischen Vesikel Endozytose in kultivierten Hippocampal Neuronen

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, making it difficult to isolate inputs from individual sources with electrical stimulation. However, by using channelrhodopsin assisted circuit mapping (CRACM), this limitation can now be overcome. Here, we report a method to use CRACM to map ascending inputs from lower auditory brainstem nuclei and commissural inputs to an identified class of neurons in the inferior colliculus (IC), the midbrain nucleus of the auditory system. In the IC, local, commissural, ascending, and descending axons are heavily intertwined and therefore indistinguishable with electrical stimulation. By injecting a viral construct to drive expression of a channelrhodopsin in a presynaptic nucleus, followed by patch clamp recording to characterize the presence and physiology of channelrhodopsin-expressing synaptic inputs, projections from a specific source to a specific population of IC neurons can be mapped with cell type-specific accuracy. We show that this approach works with both Chronos, a blue light-activated channelrhodopsin, and ChrimsonR, a red-shifted channelrhodopsin. In contrast to previous reports from the forebrain, we find that ChrimsonR is robustly trafficked down the axons of dorsal cochlear nucleus principal neurons, indicating that ChrimsonR may be a useful tool for CRACM experiments in the brainstem. The protocol presented here includes detailed descriptions of the intracranial virus injection surgery, including stereotaxic coordinates for targeting injections to the dorsal cochlear nucleus and IC of mice, and how to combine whole cell patch clamp recording with channelrhodopsin activation to investigate long-range projections to IC neurons. Although this protocol is tailored to characterizing auditory inputs to the IC, it can be easily adapted to investigate other long-range projections in the auditory brainstem and beyond.

Channelrhodopsin-assisted circuit mapping (CRACM) is a precision technique for functional mapping of long-range neuronal projections between anatomically and/or genetically identified groups of neurons. Here, we describe how to utilize CRACM to map auditory brainstem connections, including the use of a red-shifted opsin, ChrimsonR.

Langstrecken-Channelrhodopsin-unterstützte Schaltungszuordnung von minderwertigen Colliculus-Neuronen mit blauen und rot verschiebten Channelrhodopsinen

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, ...

Analyzing Synaptic Vesicle Recycling and Fluorescent Dye Uptake in Drosophila Larvae

Transkript

Analyzing Synaptic Vesicle Recycling and Fluorescent Dye Uptake in Drosophila Larvae