Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

The overall goal of this experiment is to stall a replication fork at a nucleoprotein block and observe the DNA structures at the site of the block for the purpose of studying DNA repair pathways. This method can help answer key questions about DNA replication and repair, such as how the DNA is processed when the cell's replication apparatus encounters a protein roadblock. The main advantage of this technique is that we can reversibly stall a replication fork at a specific location on the chromosome across an entire population of living cells.

Demonstrating this procedure will be the following members of my laboratory, Dr.Karla Mettrick, and graduate student, Georgia Weaver, and Tayla-Ann Corocher. An E.coli strain carrying a tetracycline operator array in the pKM1 plasmid is used for this procedure. pKM1 encodes for TetR-YFP, an inducible yellow fluorescent protein-tagged tetracycline repressor protein.

Dilute a fresh overnight culture of this E.coli strain to an optical density at 600 nanometers of 0.01 in a dilute complex medium with antibiotics as required for selection. Grow the culture at 30 degrees celsius with shaking to an optical density at 600 nanometers of 0.05 to 0.1. Remove a 10 milliliter sample to serve as the uninduced control.

Add 0.01%arabinose to the remaining culture to induce the production of TetR-YFP from pKM1. Continue to grow both the uninduced and induced cultures at 30 degrees celsius with shaking. After one hour, check for the presence of a single focal point within each cell of the induced culture using fluorescence microscopy.

If the induced cells are confirmed blocked, indicated by more than 70%of cells having a single focus, record the optical density and remove 7.5 milliliters of the sample for analysis by two-dimensional gel electrophoresis. Remove an equivalent sample from the uninduced control culture. Prior to fluorescence microscopy, prepare the agarose pads for preventing movement of the cells during visualization.

For each agarose pad, pipette 500 microliters of molten agarose onto a microscope slide and overlay the coverslip immediately before the agarose solidifies. Store the slides between wet tissues at four degrees celsius until needed. When it is time to check the cells, remove the coverslip from the agarose pad and pipette 10 microliters of bacterial culture onto the center of the pad.

After approximately five minutes, when the agarose pad has dried, replace the coverslip. Place a drop of immersion oil onto the coverslip and place the slide under the optics. Visualize the cells using phase microscopy at 100x magnification.

On the Acquire dialog box within the imaging software, set Exposure Time to 100 milliseconds. Select Acquire to capture the image. Do not move the stage.

In the Acquired dialog box, select YFP as the illumination for the external shutter linked to the camera. Set the Exposure Time to 1, 000 milliseconds. Turn off the stage light and select Acquire to capture the fluorescence image.

To begin this procedure, add sodium azide to previously-collected culture samples to a final concentration of 0.1%and incubate on ice for at least five minutes. Centrifuge the cells at 5, 000 times g for ten minutes. Discard the supernatant.

Resuspend each cell pellet in 200 microliters of PIV buffer and transfer the cells to a microfuge tube. Microcentrifuge to pellet the cells and discard the supernatant. Treat a microscope slide with 40 microliters of an appropriate glass water repellent or silicone solution.

Rub the slide with a tissue until it is dry. Resuspend the cells with the lowest cell density in 50 microliters of PIV and place at 50 degrees celsius in a heat block. For all other samples, adjust the volumes of PIV so the final cell density is the same in each tube.

Add an equal volume of freshly-prepared 0.8%Agarose solution in PIV to the samples. Keep the samples in the heat block to ensure they are above 50 degrees celsius to prevent solidification. Place 20 microliter drops of the cell-agarose suspension on the treated slide to produce plugs that are hemispherical.

When the plugs have solidified, gently slide all of the plugs from one sample into a single microfuge tube and add one milliliter of cell lysis buffer. Incubate at 37 degrees celsius for two hours. After two hours, remove the cell lysis buffer and add 1 milliliter of EDTA-Sarkosyl-Proteinase K or ESP solution.

Incubate at 50 degrees celsius overnight or until the plugs are transparent. Once the plugs are transparent, remove the ESP buffer and transfer the plugs to a 15-milliliter tube. Add 12 milliliters of TE buffer and let stand for 30 minutes.

Wash the plugs a total of five times with TE buffer. Store the plugs at four degrees celsius in one milliliter of TE buffer. To analyze the replication intermediates, the DNA in the agarose plugs is digested with an appropriate restriction enzyme for the region of interest.

In this example, EcoRV cuts the DNA immediately before the array, within the array, and after the array, to give 5.5 kb and 6.7 kb fragments in the region of interest. To begin this procedure, transfer an agarose plug to a fresh microfuge tube and add 150 microliters of restriction enzyme buffer. Add 25 to 100 units of restriction enzyme and digest at 37 degrees celsius for six to eight hours.

Prepare a 0.4%agarose gel at four degrees celsius. Pour 300 milliliters of molten agarose into a gel tray of approximately 25 by 25 centimeters. Insert a comb to make the wells approximately the same width as the diameter of the plug.

When the gel is set, remove the comb and move the gel to room temperature. Using an inoculation loop, slide one of the digested agarose plugs into a well, positioning the flat side of the plug against the side of the well where the DNA will enter the gel. Insert a single plug in the same manner into every second well.

Pipette molten 0.4%agarose into the wells to seal the plugs in position. Load a one kb DNA ladder in an empty well, leaving a gap between the ladder and samples. Run the gel overnight in 1xTBE at room temperature.

On the following day, stain the gel in a water bath containing 0.3 micrograms per milliliter of ethidium bromide for 20 minutes. Visualize the DNA with a long-wave UV transilluminator and cut out each lane fragment with a straight even cut minimizing the inclusion of excess agarose on either side of the lane. Place the excised gel lane in a gel tray at 90 degrees to the direction of DNA migration.

The next step is to prepare 300 milliliters of agarose for the second dimension gel. When the agarose is cooled to 50 degrees celsius, pipette the molten agarose around the gel slices to solidify their position. Pour the remaining agarose into the tray to a depth that is at least level with the gel slices.

Once the gel is solidified, electrophorese it in 1xTBE containing 0.3 micrograms per milliliter of ethidium bromide at four degrees celsius until the DNA has migrated approximately 10 centimeters. Excise the blocks where the genomic DNA is present including the gel above it, to include the DNA that is not visible. The DNA within the gel is subsequently visualized by Southern hybridization as described in the text protocol.

In this system, a replication block was confirmed by a majority of cells containing one focus corresponding to one copy of the array within the cell. The addition of anhydrotetracycline reversed the block and subsequent duplication of the array was visualized as the accumulation of cells with multiple foci. Cells with blocked replication also showed growth inhibition that was reversed by subsequent growth in the presence of anhydrotetracycline.

To analyze the replication intermediates, the DNA was electrophoresed in the first dimension. The DNA of interest was then excised and a second dimension electrophoresis resulted in a diagonal of linear DNA. Southern hybridization of the array DNA revealed two spots in the unblocked sample corresponding to the expected 5.5kb and 6.7kb fragments of the array.

The blocked sample showed a decrease in the 5.5kb spot and the addition of an elliptical signal indicating an accumulation of Y-shaped DNA. Addition of anhydrotetracycline eliminated the Y-shaped DNA signal. Another common intermediate is the Holliday junction visualized as a cone signal at the top of the Y-arc and a spike from the linear DNA at the end of the Y-arc.

Once mastered, this technique can be done in eight days if it is preformed properly. While a 2D procedure, it is important to remember that the quality of the DNA plugs is critical for the optimal visualization of the DNA structures.