Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

The overall goal of this microscopy-based approach is to observe a central processes during growth of bacterial cells. This method enables us to address important questions in bacteriology, such as how essential processes are coordinated during bacterial cell growth and division. The main advantage of this technique is we're able to use fluorescent reagents to visualize bacterial cell surfaces and structures in living cells.

Here, we use this method to provide insight into essential processes in Agrobacterium tumefaciens. However, this process could also be adopted to other bacteria to study their growth as well. To begin, use a sterile wooden stick or pipette tip to inoculate three milliliters of ATGN Growth Medium, with a single colony of a wild-type or mutant strain of A.Tumifaciens.

Grow the culture at 28 degrees celsius and 225 RPM overnight. The following day, use a spectrophotometer to measure the OD600. Then, dilute the cell culture to an OD600 of approximately 0.2 and continue to grow the cells to an OD600 of 0.6.

Dispense one milliliter of exponential culture into three microfuge tubes. And pellet the cells in a desktop centrifuge at 7, 000 x g for five minutes. Re-suspend each cell pellet in one milliliter of PBS, and then add either one microliter of DMSO Orange stock solution or DAPI stock solution.

Mix gently by pipetting, and incubate the re-suspended cells in the dark for five minutes. Pellet the cells by centrifugation at 7, 000 x g for five minutes, and re-suspend the pellet in one milliliter of PBS to remove excess reagent. Then, wash the cells two more times, and re-suspend the pellet in 50 microliters of PBS or medium.

For simultaneous time-lapse imaging, combine ten microliters of cells from each treatment in a new microfuge tube. To prepare agarose pads for cell imaging, use a cover slip as a guide, and cut a 22 by 22 millimeter square of laboratory film by running a scalpel around the edges. Cut a square out of the center of the laboratory film, leaving an approximately two to five millimeter border to serve as a gasket for the agarose pad.

Then, discard the center cut-out. Place the film gasket onto a glass slide, cleaned with an ammonia and alcohol-free cleaner. And use a heat block set to 70 degrees celsius or a flame to heat the slide until the film is slightly melted onto the glass.

Next, prepare agarose solution by mixing approximately 0.075 grams of agarose and five milliliters of medium in a small flask. Heat the solution in a microwave with periodic swirling to mix, until the agarose is dissolved and the solution is clear. Keep the agarose solution at 55-70 degrees celsius and use it for construction of multiple agarose pads within 48 hours.

Pipette the warm agarose medium into the center of the gasket. Then, place a cover slip over the gasket to evenly distribute the agarose. Place the slide on a cool level surface to solidify for approximately two minutes.

Now, carefully slide the cover slip off the agarose pad, then allow the agarose pad to air-dry for one to two minutes at room temperature, until the surface of the pad appears dry. Proper construction of the agarose pad is crucial for acquiring high-quality images. Don't try to image on a bad pad.

If the agarose pad dries out, ripples or rips, it's best not to be lazy and instead, make a new pad. Then, using a scalpel, remove a small strip of agarose to create an air-pocket approximately two by seven to ten millimeters. A.Tumefacien cells tend to grow best near the air-pocket.

Spot 0.8 to 1 microliter of cells on the agarose pad. Then gently place a cover slip over the top of the agarose pad to distribute the cells across the surface of the pad. For long-term time-lapse imaging, use melted Valap to seal the edges of the cover slip.

Be sure to seal along all the edges and corners of the cover slip with Valap. Failure to do so can result in drifting during the agarose pad time-lapse imaging. To carry out epifluorescence microscopy, place immersion oil on the desired objective and place the inverted slide into the slide-holder on the stage.

Use the focusing knobs to bring the cells into focus. Acquire images in Phase, or DIC and fluorescence, using the desired fluorescence filter for fluorescence images. Optionally, acquire multiple x, y positions by randomly selecting ten fields of cells in close proximity to the air pocket of the agarose pads.

Set up the time sequence acquisition to image in-Phase, or DIC, at the desired time interval. As shown here, cells were imaged directly from culture after washing by centrifugation and after incubating the cells with 0.1 percent DMSO, the results showed DMSO alone does not impact cell morphology. Both Orange and DAPI labeled DNA within live A.Tumefaciens cells.

In late pre-divisional cells, two distinct nucleoloids are observed with Orange staining, whereas DNA labeling appears more diffuse with DAPI staining. In this time-lapse experiment with an equal proportion of unlabeled, DAPI labeled, and Orange labeled wild-type A.Tumefaciens, all cells grew when imaged with Phase-contrast microscopy, indicating that neither stain impaired cell growth. The cells also grew under Phase contrast in epifluoresence microscopy using the TRITC filter.

When imaged with the DAPI filter, however, cells stopped growing within an hour. Following similar methods, other fluorescent reagents can also be used to visualize the bacterial cell wall or membranes. Furthermore, many of these fluorescent dyes can be combined to provide a comprehensive image of the bacterial cell surface.

Visualization of essential processes in live bacteria is exciting, because it opens up the possibility of studying these processes in conditional mutants or non-model bacteria. This allows us to enhance our studies of diverse bacterial species.