Here we demonstrate the protocol for performing single molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified. This protocol begins with growing bacteria, which make a protein tagged to a fluorescent dye molecule. After four to six hours, a small volume of cells are extracted from the culture and their flagella filaments truncated by shearing.
A glass cover slip inside a flow cell is coated to allow cells to stick to the surface. Cells are injected into the flow cell and tethered via a gellar stub, which is viewed in fluorescence using laser excitation microscopy. A high quantum efficiency camera then records the bright brightfield and fluorescence images of the tagged molecular complexes.
Hello, my name's Ian Doy and I'm working with Mark Lee at the Department of Biochemistry at the University of Oxford. I'm Alex Robertson working with Mark Lee in the physics department. I'm Nick De from the leak lab, also based in the physics department.
Today we'll be showing you the procedure for visualizing single molecular complexes in living bacteria using advanced fluorescence microscopy. We use this procedure to Study theto this in a functional state. We're also using to investigate real time dynamics of natural biological machine and nanometer lens scale.
So let's get started. To begin thaw 50 microliters of frozen stalks of e coli bacteria. These have been modified genetically.
To tag a protein with a fluorescent dye molecule, inoculate five milliliters of LB growth media and grow the bacteria aerobically shaking overnight at 37 degrees Celsius. The next morning, take 50 microliters of the saturated culture and subculture it into minimal M 63 Glucose culture media incubate at 30 degrees Celsius for four to six hours. Here two different cell strains are used.
One expresses an electron transporting cytochrome fused to GFP. The other expresses a protein involved in the bacterial flagella motor. Fused to GFP cells may either be harvested directly from the growing subculture if they are to be viewed as immobilized samples, or they may be sheared to truncate bacterial flagella if they're viewed under tethered conditions.
To shear the bacteria, place one to five milliliters of the subculture into a device consisting of two sterile syringes by sterile piping. Alternate pushing in each syringe pump to force the culture through the narrow tubing about 50 to 100 times depending on the extent of shearing acquired Next centrifuge the culture to pellet the cells and then resuspend the cells in minimal media to remove flagella fragments. Once the cells are ready, prepare cleaned BK seven glass cover slips by immersing them in a saturated solution of potassium hydroxide and ethanol for 20 minutes.
Rinse thoroughly in deionized water and ethanol and leave to air dry for at least one hour. Next, construct a simple flow cell to house the cells in the microscope. To do this, draw lines of paraffin grease on a BK seven glass microscope slide, and then create a tunnel sandwich by placing one of the cleaned cover slips on top.
Press down gently with a pair of forceps. This should give a flow cell volume of five to 10 microliters. To observe immobilized cells fill the flow cell by injecting with a 0.1%solution of poly L lysine and allow it to incubate at room temperature for at least one minute.
It next, flush out the flow cell by injecting 100 microliters of minimal media from one end of the flow cell and simultaneously wicking the media through the flow cell with tissue paper from the other afterwards, WIC threw 20 microliters of a one in 500 dilution of 200 nanometer diameter latex microspheres in minimal media to mark the cover slip surface. Place the flow cell in a simple humidity chamber and incubate at room temperature for five minutes. The flow cell is inverted such that the cover slip is facing downwards.
The unbound beads are then washed away by wicking through 100 microliters of minimal media with tissue paper. To observe tethered cells, skip the poly L lysine incubation step instead, fill the flow cell with five micrograms per milliliter Anti-Flag gellan antibody solution, and then place it in a humidity chamber for 10 minutes. After incubation, flush the flow cell through by wicking with tissue paper.
Next, WIC 20 microliters of the cell culture through the flow cell with tissue paper, either using the sheared sample to observe tethered cells or the unshared sample to view immobilized cells, the flow cell is inverted and placed in the humidity chamber for 20 minutes. The unbound cells are then washed out by wicking through 100 microliters of minimal media. Place a drop of immersion oil in the center of the top surface of the cover slip.
Place the flow cell gently onto the sample holder of the custom built fluorescence microscope. This should make optical contact with the high numerical aperture objective lens. Next, switch on the microscope's electron multiplying camera and set the camera to be cooled to minus 70 degrees Celsius.
The software is set to acquire images at a typical frame rate of 25 hertz. In frame transfer mode. Switch on the brightfield illumination and bring the image into focus.
Select a suitable cell or group of cells to be imaged on the basis of their being stuck with their long axis. Parallel to the cover slip surface, adjust the focus to ensure that the 200 nanometer latex beads stuck to the cover slip surface are just in focus. Acquire an image sequence in brightfield to record the outline of the cell body.
The brightfield illumination is then switched off and the camera gain is enabled to maximum for imaging using total internal reflection fluorescence, also known as turf. Start the camera acquisition and open the laser shutter to excite the fluorescent proteins within the bacteria. The parameters for laser intensity and speed of acquisition need to be optimized for the particular biological system.
Under study samples are illuminated until photo bleached, which typically takes about 10 seconds. Once the data has been collected, the images are fed into a custom written analysis program, which automatically detects the positions of fluorescent spots in the cells to a precision of a few nanometers and extracts their size and brightness. The brightness of the photo bleaching trace with respect to time of attract molecular complex is then used to estimate the stoichiometry use.
Using this method, the image of the cells viewed in brightfield is very distinct, such that the perimeters of the cell bodies are dark against a white gray cell body using fluorescence with immobilized cells. Distinct spots of intensity of typically 250 to 300 nanometers in width can be seen displayed here in false color. Healthy tethered cells can be seen rotating around the point of tether attachment in brightfield images under fluorescent excitation.
Some molecular complexes might also be seen at the point of attachment indicating a localization of the tanked protein with the Geller motor. These spots are individual molecular complexes. The number of these that are seen will depend upon the illumination mode used and how many of the complexes are actually present in the cell at any one time.
If the density of spots is initially very high, as is the case with the labeled cytochromes used here, then performing an initial frap bleach can improve the imaging contrast. The mobility of the spots depends upon the specific biological system. Under study, We've just shown you how to visualize single molecular complexes using advanced fluorescence microscopy.
When doing this procedure, it's important not to have a shear cells as this may impair the functionality of the flag motor. It's also important not to leave the cells on the microscope slides for more than an hour are they may become oxygen depleted. Automatization is required to find the best imaging conditions.
It can help using purified GFP alone to ascertain the correct laser power for your particular microscope system. So that's it. Thanks for watching and good luck with your experiments.
