Preparation of Drosophila S2 cells for Light Microscopy

The overall goal of this procedure is to describe the preparation of Drosophila S two cells for light microscopy. This is accomplished by first preparing glass bottom culture dishes. The second step of the procedure is to coat the glass bottom with the lectin conval.

A to promote adhesion and flattening of the S two cells. The third step of the procedure is to add culture medium and S two cells to the culture dish. The final step of the procedure is to fix and immunostain the S two cells in the culture dish.

Ultimately, results can be obtained that show localization of fluoro four tagged or immuno labeled proteins of interest through fluorescence microscopy of live or fixed S two cells respectively. Hi, I'm Joey Club from the laboratory of Greg Rogers in the Department of Cell Biology Anatomy at the University of Arizona. I'm John Nye, also from the Rogers Lab.

And I'm Dan Buster, also. I'm Greg Rogers lab. Today we'll show you a procedure for preparing Drosophila S two cells for light microscopy.

We use this procedure in our laboratory to study the regulation of central duplication in Drosophila by studying changes in central number and structure following elimination or overexpression of important regulatory proteins. So let's get started. We use standard 35 millimeter Petri dishes to make our glass bottom dishes.

These glass bottom dishes keep cells in a condition very similar to their normal culturing condition, as well as allow live cells to be examined through microscopy grade glass. For assembly of the glass bottom dishes, a taped surface of a styrofoam box with thick walls is used as a drilling surface. Place the bottom half of a 35 millimeter Petri dish on the drilling surface with the bottom side of the dish against the styrofoam box while holding the sides of the dish firmly use a hand drill with a three quarter inch spade drill bit to drill a hole through the dish.

After drilling all of your dishes, use a conically shaped grinding drill bit with a coarse grit and a diameter about the same as the spade bit to smooth the outside rim of the holes. Place the power drill on its side with the bit projecting out over the edge of the counter. Hold the drill in place by placing something heavy but pliable on it, such as a heavy duty plastic bag containing 10 kilograms of sand.

Use the lock on button to keep the drill continuously running. Smooth the rim of your dish by holding and moving it against the grinding bit to smooth away the rough edge of the new hole. After the outside rim of the holes have been smoothened, put the dish bottoms in a large peaker and rinse several times with distilled water.

This will remove the small loose pieces of plastic or styrofoam, which will float afterwards. Lay the dish bottoms out to dry. When the dishes are dry, prepare the Sard 180 4 glue.

Mix the resin and the curing agent at a 10 to one weight ratio and stir thoroughly. Apply a small circle of glue around the drilled hole on the outside surface of each dish. Next place a cover glass over the hole in contact with the glue.

The cover glass must be at least about one millimeter larger than the hole that is in the Petri dish and smaller than the diameter of the dish. The cover glass should sit against the dish smoothly and evenly. Now, set the dishes aside.

Bottom side, up to dry. This can take a day or more at room temperature, but you can increase the curing rate by putting the dishes in a warm incubator. Once the dishes are dry, they can be coated with the lectin conent A or con A.We use con A because S two cells that are seated onto con A coated cover slips will flatten extensively and to become excellent specimens for microscopy.

To con a coat, a glass bottom dish, spread 10 microliters of conne solution onto the upper side of the glass. After drying, sterilize the dish and its cover with the UV light. In a tissue culture hood, turn the pieces so that their inner surfaces are directly facing the UV bulb and turn on the sterilizing UV light for at least 45 minutes.

S two cells can be cultured in various media. For our purposes we use in VITROGEN SF 902 medium. To prepare RNAi treated S two cells for microscopy, gently resuspend the cells and transfer them to a conne coated glass bottom dish containing two milliliters of medium healthy cells that contact the conne coated cover slip will adhere tightly to it and begin to spread.

This process should be apparent by about 15 minutes and should be complete by 45 minutes to an hour. Since S two cells are easily maintained under normal lab temperature and atmosphere conditions, no special conditions or equipment are needed when the cells are on the microscope to prevent dehydration, keep sufficient medium in the dish on the microscope. Live imaging of S two cells for extended times is possible.

The runtime of these experiments is limited only by the usual problems of photobleaching and phototoxicity. The S two cells can be fixed on either a glass bottom dish or a plain cover slip. But in this video we will demonstrate fixing and immunostaining in a glass bottom dish.

Methanol and formaldehyde are two common fixatives for S two cells for cell fixation with methanol first chill about 300 milliliters of anhydrous methanol in a covered one liter glass beaker in an explosion proof minus 20 degrees Celsius freezer. When the methanol is cold, remove it from the freezer. Quickly remove the medium from the glass bottom dish and then rapidly plunge the dish into the methanol.

Quickly return the beaker containing the dish to the freezer and fix the cells for 10 to 15 minutes. After 10 to minutes, remove the dish from the beaker, pour out any remaining methanol and rehydrate the cells. By adding PBST, the cells are now ready for standard staining procedures as will be demonstrated later for cell fixation.

With formaldehyde, we use 10%formaldehyde and buffer. This relatively high concentration of formaldehyde is used because S two cells lack cytoplasmic intermediate filaments, which would otherwise help maintain cell morphology and organization during the fixation process. Fix the cells in 10%formaldehyde for 10 to 15 minutes.

When the fixation is complete, pour off the formaldehyde into a waste beaker and wash the fixed cells with three brief washes of PBST. Begin the immunostaining procedure by adding one milliliter of blocking solution to the fixed cells for 15 minutes. We routinely use 5%normal goat serum in PBST.

Pour off the blocking solution and add the primary antibody directly onto the cells. Place the Petri lid back on the dish to prevent evaporation of the antibody solution and let the antibody incubate with the cells for 30 minutes at room temperature. After 30 minutes, remove the primary antibody solution.

Use a transfer pipette to add two milliliters of PBST to the dish and wait for five minutes before replacing the PBST. Then wash three times with PBST. After the final wash, add 100 microliters of secondary antibody directly onto the cells.

Place the Petri lid back on the dish and let the antibody incubate with the cells for 30 minutes. At room temperature. When the 30 minutes are up, remove the secondary antibody and perform three five minute washes with PBST.

Finally, add a few drops of mounting media directly onto the cells. Replace the Petri lid and if needed, store the dish at four degree Celsius in the dark. Shown here are representative phase contrast images of S two cells after plating on a conval and a coated glass bottom dish where the numbers indicate the minutes after plating.

Self flattening is apparent by 15 minutes and is essentially complete by 60 minutes. Notice that the cells become phased dark as they flatten on the coated cover slip. In this magnified image of attached cells, their extended and flattened margins indicated by the arrowhead are clearly visible.

S two cells can be transfected to express fluorescently tagged proteins that mark structures of interest in fixed or live cells. In this example, S two cells were transiently transfected with a fusion construct consisting of nucleo foin, a marker for the nucleus fused to the fluoro four EGFP. These cells were plated on a conna coated glass bottomed dish and then imaged with DIC and epi fluorescence.

RNAI treated S two cells can also be fixed and analyzed by immunofluorescence. These images show fixed S two cells immuno stain for the central marker PLP shown in green and microtubules shown in red, as well as hooked stained for chromosomes shown in blue. Prior to fixation and immuno staining, the cells were subjected to a four day treatment with either control, RNAi or RNAi to knockdown NCDA KINESIN like protein that promotes spindle pole focusing.

Treatment with NCD RNAi resulted in cells with splayed spindle poles and disorganized spindles. When doing this procedure, it's important to remember to be very cautious when drilling holes in the culture dishes. So that's it.

Thanks for watching and good luck with your experiments.