Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

This video demonstrates the procedure for photo conversion of the Styro Dye FM 1 43. To study synaptic vesicle pools in drosophila melanogaster first drosophila larvae are dissected to expose the ventral muscles which constitute the neuromuscular junction or NMJ preparation. The preparations are incubated in buffer containing FM D shown here in gray labeling cellular membrane.

They're then electrically or chemically stimulated to induce physical fusion vesicle membrane internalized following. Stimulated exocytosis will be stained with FM D.The extracellular FM dye is then washed, removing it from the plasma membrane, and the tissue is fixed under continuous high intensity illumination. The dye is completely bleached and the black color appears due the di amino Ben and precipitation.

This precipitate can then be visualized by transmission em. Hi, I'm Philippe o Paso from the laboratory of Sylvia Olli at the European Neuroscience Institute in Gaon, Germany. Today we will show you a procedure for FM photo conversion inria neuromuscular junction.

We use this procedure in our laboratory to study synaptic vesical recycling. Okay, so let's get started Working under a dissecting microscope. Begin by pinning a drosophila larvae dorsal side up in a so guard coated dish.

Cover it in drosophila saline using a syringe needle and find dissection scissors section the dorsal side longitudinally. Then remove the internal organs. Next, stretch the preparation and again, pin it to the sigar dish.

Several ventral muscles can then be used to protect the FM dyes used in this procedure from bleaching. This procedure should be performed under low light conditions. Cover the pinched drosophila preparation with drosophila saline containing 90 millimolar potassium chloride, and 10 micromolar FM.1 43 die to chemically stimulate the nerves.

Incubate for one minute at room temperature following stimulation. Remove the stimulating solution and submerge the dish with the pinned drosophila. Preparation in a beaker filled with 100 milliliters of drosophila saline two times to remove the extracellular FM 1 43 die working under a chemical safety hood.

Fix the neuromuscular junction preparation in 2.5%glutaraldehyde in phosphate buffered saline or PBS for 45 minutes at room temperature. Wash the preparation once with PBS, then remove it from the cigar dish and submerge it in 100 millimolar ammonium chloride for 15 minutes. To quench the free aldehyde groups of the remaining glute aldehyde fixative.

Using normal PBS wash the ammonium chloride solution, place the neuromuscular junction preparation in a new cigar dish and submerge it in filtered 1.5 milligram per milliliter. Diamine benzine or DAB in PBS and incubate for 30 minutes at four degrees Celsius. Position the sample under a fluorescent microscope equipped with a mercury lamp light source with a lamp housing containing a back mirror using a relatively low magnification dry objective.

Find the sample. Then using a normal blue excitation green fluorescence filter cube, bring the fluorescent signal into focus. The neuromuscular junctions appear as bright spots on the muscles.

Next, using the same fluorescence filter set, illuminate the sample at maximum intensity until the FM D is completely bleached. Using a 20 x objective lens, 30 to 45 minutes should be sufficient. However, the time for photo conversion varies depending on the intensity of illumination, strength penetration of DAB into the preparation and the objective lens used.

Following illumination, examine the sample under transmission light. If photo conversion has taken place, a dark brown precipitate should be seen. Return to the dissection scope, then use scissors to cut the photo converted spot from the larva.

Return to the dissection scope. Then use scissors to cut the photo converted spot from the larvae. Now place the sample in a micro fuge tube containing PBS working under a chemical safety hood and wearing appropriate safety equipment, including gloves and eye protection.

Prepare 300 microliters of 1%osmium tetroxide per sample. Now place the sample in a plastic topped glass flask and add the prepared osmium tetroxide solution to postfix the preparation. While the sample is incubating, prepare separate solutions containing 30 50 70, 90 95 and 100%ethanol.

Wash the sample four to five times with PBS for five minutes each. After each wash, use a plastic PEs to your pipette to transfer the sample to a new flask. After the last wash, transfer the sample to a clean glass flask.

Next, dehydrate the sample by placing it in the series of ethanol solutions. Begin by incubating in one milliliter of 30%ethanol. Then after five minutes, remove the ethanol using a pasture pipette and replace it with one milliliter of 50%ethanol.

Continue with the dehydration, incubating the neuromuscular junction preparation in the 70, 90 and then 95%solutions. The sample will then be incubated in 95%ethanol solution to more times than incubated in 100%Ethanol three times following dehydration. Add two milliliters of one-to-one EIN resin to ethanol and place the sample on a rotator incubate.

Two to four hours at room temperature following rotation at 100%ein and place the sample in an open flask to allow the remaining ethanol to evaporate for four to six hours. Pour fresh EIN into a mold. Add the preparations and incubate at 60 degrees Celsius for 36 hours.

When the EIN embedded preparations are ready, use an ultra microtome to prepare 50 to 80 nanometer thin sections I using a diamond's knife. The samples are cut in thin sections. The sections flow on the water bath of the knife and can be afterwards picked up on normal EM grids.

The preparations are now ready to image using conventional EM procedures. Isolated neuromuscular junctions from Drosophila Melan gaster were stimulated using high potassium and internalized membranes. Were stained using FM 1 43 as shown here.

The fluorescence of a nerve terminal can be observed using a conventional epi fluorescence microscope. Under continuous illumination, the dye is completely bleached and when illumination continues, a black color appears due to dino benzine precipitation. This nerve terminal contains synaptic vesicles, but none of them are photo converted.

This might be caused by not having enough illumination or poor DIA amino benzine penetration in the tissue. By contrast, too much illumination results in an overall dark terminal where no vesicles or organelles are distinguishable with the appropriate amount of illumination photo conversion can be seen. This micrograph shows a synaptic bhuton in which a pool of dark vesicles are seen indicating FM photo conversion.

We have just shown you how to photo convert synaptic vesicles labeled with fem die. When doing this procedure, it's important to remember to always carefully calibrate the illumination times for the conversion step. So that's it.

Thanks for watching. Good luck with your experiments.