Antilichaamkleuring in C. Elegans Een 'Freeze-Cracking "

The overall goal of this procedure is to perform antibody staining on C elegance. To do this, nematodes are rinsed, spread on slides and carefully compressed between two adhesive slides, which are then frozen on dry ice. Next, the slides are separated to crack open the nematodes and immediately placed in fixative.

The nematodes are then stained using standard antibody Staining techniques analysis by fluorescent microscopy reveals successful antibody staining of semi intact nematodes. The main advantage of this technique over existing methods like fixation of intact nematodes is that it minimizes chemical treatment of the sample. Generally, individuals new to this method will struggle because preparing the slides and nematodes is somewhat tricky to demonstrating.

The procedure will be myself and RTO Basu, a graduate student from my laboratory To prepare plates of nematodes for staining begin by placing a flat piece of metal on dry ice to chill it using distilled water and a glass pipette rinse. Worms grown on one 60 millimeter NGM plate with bacteria into a 1.5 milliliter micro fuge tube. The worms can be synchronized or mixed ages, but avoid using plates with many dowers which are difficult to crack or starved worms, which will increase autofluorescence using the same glass pipette.

Rinse the worms three to four times with distilled water to free them of bacteria. Then to collect mostly adults, let them sit on the bench for three to five minutes. After the worms have settled to the bottom of the tube, remove most of the water from the tube, leaving about twice the volume of water as worms, but at least 25 microliters total volume.

Label the frosted side of polylysine slides with indelible ink and place the slides on the benchtop next to container with dry ice label side up. This will serve as the bottom slide. Have an equal number of clean non polylysine treated.

Top slides available. Next pipette about 25 microliters of worms in water onto the bottom adherence slide, and using the side of the pipette tip, spread the worms over the center of the slide. Let them settle for 30 seconds to two minutes.

The ends of the worms will stick as they contact the slide. The most difficult aspect of this procedure is keeping the worms from getting too distorted when you're doing the next few steps. The best way to do this is by being patient, having everything ready to go, and don't drink too much caffeine so that your hands are nice and steady.

Holding the bottom slide in one hand, place the top slide over it so that all but the frosted parts are overlapping. The liquid should hold the slides slightly apart so that the worms will still have some mobility. Do not let the slide slip over one another, which will twist the worms.

Next, using the thumbs and forefingers of each hand carefully press straight down on the top slide With the right amount of pressure, a few of the largest hermaphrodites will rupture on the edge of the slide. While most of the adults and larvae will contact each slide but will stay intact immediately and carefully without allowing the slides to slip, put them on the piece of metal on dry ice. Within one minute, the slides will appear frozen, but keep them there for five minutes to ensure that they are thoroughly frozen.

The slides are now ready for fixation. If not proceeding immediately store the slides in a labeled box at minus 80 degrees Celsius for several days. They should be used within a week or so after they are made.

Do not use the slides after long-term storage. Individual nematodes can also be prepared on slides in a method similar to that just shown as before. Place a flat piece of metal on dry ice to chill it.

Next, use a worm pick to transfer individual nematodes to a drop of water on an NGM plate to free them from bacteria. The nematodes should be well fed to decrease autofluorescence. If possible, repeat the transfer to new spots of water as needed until worm is free of bacteria.

Next, prepare the top and bottom slides as before. Then place a five to 10 microliter drop of water on the polylysine treated area of the bottom slide while observing with a microscope. Use a worm pick to transfer up to 20 worms to the drop of water.

The worms should sink down to touch the sticky slide surface. Then place the top slide as before, but do not compress while ensuring that the slides do not slip immediately. Place the slides on the metal on dry ice.

Keep on dry ice for five to 30 minutes and proceed immediately to fixation. These slides cannot be stored for later use. To lightly fix the worms begin by placing staining jars containing fixation solutions on ice to chill them for 10 minutes.

The solutions needed are 100%methanol, 100%acetone and phosphate buffered saline. Take a pair of bottom and top slides from the dry ice swiftly twist them apart and immediately immerse the bottom slide in ice cold methanol for two minutes, discard the top slide. Next, transfer the slides from methanol to ice cold acetone for four minutes.

If the slides had a lot of worms, most of them will fall off in the fixative even with well prepared slides. If single worm slides were properly prepared, the worms should remain adherent. Finally, to rinse off the fixative, dip the slides in the jar with PBS.

The slides are now ready for staining to perform a hard fix using formaldehyde or glutaraldehyde. Begin by diluting reagent grade formaldehyde or glutaraldehyde solution in PBS to a final concentration of 0.5 to 4%Aliquots can be stored at minus 20 to minus 30 degrees and defrosted as needed. Remember, always the formaldehyde and glutaraldehyde are very toxic, so you need to be wearing gloves and be working in a fume hood.

Take a pair of bottom and top slides from the dry ice and swiftly twist them apart Immediately. Place the bottom slide which has the worms in a flat dish with lid, discard the top slide immediately pipette 200 microliters of the fixative over the center of the area of the slide. With worms, the fixative will partially freeze in place, then melt to cover a larger region of the slide.

If the worms are not totally covered with fixative immediately and carefully, add more fixative using a micro pipette. Do not allow the fixative to flow over the edge of the slide. Add folded wet lint free wipes to the dish to keep it humidified.

Do not allow the wipes to touch the slides. Then cover the dish with a lid and leave it for 10 minutes to overnight at room temperature after fixation is complete, use a pipette to carefully transfer the fixative with loose worms into a 1.5 milliliter tube. Then dip the slide in a staining jar containing PBS.

Spin the tube containing the worms at high speed for 30 seconds to pellet them. Rinse the worms twice with PBS. Then proceed to staining them in parallel with the worms on the slide.

The antibody staining protocol is similar to standard protocols for any tissue on slides and is described in detail in the accompanying document. After staining, transfer the slides to PBS and allow them to remain there up to 12 hours at four degrees Celsius until ready to mount. To mount the slides, remove a slide from the PBS.

Use a lint-free wipe to dry the backside and then briefly place it on a paper towel rapidly. Place about 20 microliters of mounting medium over the worms so that there are approximately equal parts of medium and PBS. On the slide.

Gently tilt the slide to mix the two solutions. Caution mounting medium is toxic. Holding the frosted edge.

Tilt the slide so that the solution gathers near the frosting. Then place one edge of a 24 by 60 millimeter cover slip on the solution to minimize air bubbles which increase bleaching and disrupt viewing. Lower the cover slip gently.

If bubbles do form, slowly lift the edge of the cover slip. Then without sliding across the worms, re lower it. Add more PBS if needed to eliminate air bubbles.

Using a lint-free wipe, compress the edges of the cover slip and slide carefully to dry off the edge. The edge of the slide must be visible all around the edge of the cover slip and should not be too wet. Seal the cover slip and apply two to three layers of nail polish to the edge.

Once the slide is well sealed and dry, clean the cover slip and back of the slide and place it in a slide holder wells. Sealed slides can be kept days at four degrees Celsius or weeks at minus 20 degrees Celsius over longer periods. Dappy staining of DNA and most green dyes fade reseal the cover slip with more nail polish as needed, particularly after using any oil immersion lens.

The following representative images were collected using slides made during the first antibody staining attempts of undergraduates in a cell biology laboratory course. This image shows an example of the heads of two adult hermaphrodites that are properly compressed, cracked, and lightly fixed with methanol acetone. They were labeled with multiple antibodies and dyes.

In this case, virtually the entire worm is accessible to antibodies. Staining here, the location of the nuclei as indicated by DPI staining indicates which parts of the worm are intact. When the worms are subjected to a harder fixative such as formaldehyde, the morphology of the worm may become distorted.

This can be seen in these images as a wavy appearance of the muscles shown in red. Another common problem associated with the use of harder fixatives is uneven fixation or penetration of particular tissues. This can be seen here where the muscle shown in red is stained only in a portion of the worm.

Here, the presence of other staining, including DNA, shown in blue indicates that at least part of the body was present. The resulting pattern of partial staining can be readily identified with practice so that the entire distribution of staining can be determined even if any single worm shows uneven staining. Common problems encountered with freeze cracking using any fixation condition are illustrated here.

The most common problem is twisting of the sample due to relative motion of the two slides. During compression Here, the morphology of the stained tissues indicates that the body of the worm is twisted just posterior of the pharynx. This image also shows the problem with background staining due to antibody sticking to the polylysine coating the slide.

This can be avoided by using the confocal microscope. Note that even with practice, many individual worms will be lost or will show some of the problems shown here. Once mastered, the preparation and fixation can be done in an hour and the staining can be done in a day.

Following this procedure, a variety of fixation methods can be performed in order to optimize staining for any given antibody and tissue.