The overall goal of the following experiment is to gain quantitative structural information about e coli cell division proteins in their native environment. This is achieved by expressing a photo activ fluorescent fusion protein in live cells and immobilizing these cells on an acro gel pad. The cells are then imaged via streaming video.
While excitation and activation lasers are applied to visualize single molecule locations, the resulting movies are computationally analyzed to extract the molecule positions with nanometer accuracy and superpose them to generate a super resolution image. This image can then be analyzed by a variety of quantitative measurements that describe the structure of interest, including its dimensions and molecule density. Photo activated localization, microscopy or palm is an easily implemented super resolution fluorescence microscopy technique.
The instrumentation is straightforward, but careful Optimization of data acquisition and analysis is required to generate reliable results. Another crucial step is the extent of characterization of your fusion protein and expression level to ensure that your results are biologically relevant. The following protocol was optimized for the FTSZ protein fused Muse two and should be adapted to your protein of interest.
To begin inoculate LB media with a single colony of the strain of interest containing a fluorescently tacked protein prepared as described in the written protocol. Accompanying this video, grow overnight in a shaker at 37 degrees Celsius the following day. Dilute the culture one to 1000 into M nine plus minimal media.
Grow to mid log phase in the presence of appropriate antibiotic at room temperature, and then induce the culture to produce protein with 20 micromolar IPTG for two hours. Meanwhile, assemble the imaging chamber by making a 3%agro solution in M nine minus media. Melt the agros at 70 degrees Celsius in a benchtop heat block for 40 minutes.
Store the melted agros at 50 degrees Celsius for up to five hours. Place a clean micro aqueduct slide in the upper half of the imaging chamber with the electrodes facing down. Align the lower gasket on the micro aqueduct slide so as to cover the perfusion channels.
Apply approximately 50 microliters of the melted aros to the center of the glass. Slide immediately top the Agros gel droplet with a clean, dry cover slip. Allow the gel to solidify at room temperature for at least 30 minutes following protein induction.
Pellet the culture in a micro centrifuge.Resus. Suspend the pellet in an equal volume of M nine plus repeat centrifugation and resus suspension after the second resus suspension. Continue growth at room temperature for two hours.
Next, fix the induced culture by adding paraform aldehyde in PBS to achieve a final concentration of 4%paraform aldehyde. If imaging live cells see the written protocol accompanying this video for variations in the procedure, allow the culture to incubate at room temperature for 40 minutes. Pellet the culture and resuspend in an equal volume of PBS.
Next, dilute 50 nanometer gold beads, one to 10 with Resuspended culture. To apply the sample to the prepared imaging chamber, carefully remove the cover slip, leaving the gel pad on the micro aqueduct slide. Immediately add one microliter of cell culture sample to the top of the gel pad.
Allow approximately two minutes for the solution to be absorbed by the gel pad and top the gel pad with a new clean, dry cover slip. Continue to assemble the whole imaging chamber according to the manufacturer's instructions for image acquisition. Turn on the microscope camera and lasers.
Open metamorph software by molecular devices. Place appropriate excitation and emission filters in the imaging path as pictured in this microscope schematic, which can be found in the written procedure accompanying this video. Next set Laser powers and acquisition settings.
Appropriately lock the imaging chamber into the stage via the complementary stage adapter. Focus on the surface of cells closest to the cover slip lip. Designate an appropriate imaging region that is homogenously illuminated by all three lasers.
Identify a sample area that contains both cells and fiducial markers, which are the gold beads in closes proximity but not overlapping. Acquire one brightfield image with 50 milliseconds integration time and move the focus 0.5 micrometers into the sample. Then acquire an ensemble green fluorescence image with excitation from the 488 nanometer laser using a 50 millisecond integration time.
Next, acquire streaming video with continuous illumination from 405 nanometer and 561 nanometer lasers. For fixed cells, a 50 millisecond frame rate is used for a total of 20, 000 frames. With the 405 nanometer laser power ramped by approximately 10%every 1000 frames here, the first 500 frames are shown.
Note that the number of frames acquired were optimized for this system and are dependent on the particular cellular structure. Labeling density activation rate and whether or not exhausting entire pool of floor floors is important for analysis To perform image construction. First, identify prospective spots in the image stream as three adjacent pixels with intensities above an empirically determined threshold fit the intensity of each prospective spot to a two dimensional Gaussian function.
To identify the molecule position and total number of emitted photons, calculate the localization position of each spot using the total number of detected photons. Next, discard poorly fit spots by applying a threshold of the localization precision and remove repeat observations of the same molecule by disregarding any spot that is within 45 nanometers and six frames of any proceeding spot correct for sample drift. By subtracting the frame dependent displacement of the gold beads from each molecule position, superimposed all corrected molecule positions on a single image composed of 15 by 15 nanometer pixels, plotting each unique molecule as a unit area, two dimensional Gaussian profile with a standard deviation equal to the localization precision.
The resulting image is a probability density map where a pixel's intensity is proportional to the likelihood of finding a molecule. In that Pixel illustrated. Here is a two dimensional super resolution rendering of the ZR generated from the Palm Imaging Method.
Qualitatively, it was observed that the Z ring is an irregular structure that adopts multiple configurations, such as single bands or helical arcs that are not distinguishable in conventional fluorescence images. Observations such as these can be used to determine the percent of the cell population that displays a particular structural configuration. The dimensions of the ZR can also be measured from this plot.
The ring width was determined to be 113 nanometers. This value is wider than the expected width of a single FDSC proto filament, which is five to 10 nanometers based on in vitro electron microscopy. Here, the ring diameter is measured as 1050 nanometers.
This diameter can be used to quantitatively describe the degree of constriction during cell division. Another quantitative approach is to calculate the molecule density by counting the number of molecules localized within a defined region. Since palm images are 2D projections of 3D objects, total internal reflection or TIR palm must be employed to restrict the detection plane to a single side of the Z ring.
Since the plotted molecules represent a subset of the total FDSC population, the density determined is a lower limit of the actual density. The maximal density observed in the TIR images of the Z ring suggests that some sections contain overlapping proto filaments. While attending this procedure, it's important to monitor cell morphology and growth rate so as to ensure that the expression of your fusion protein is not affecting cell viability.
Once this procedure is established for your protein of interest, it can be used to test for structural changes associated with cell cycle progression or further expanded to two color imaging to compare the facial distributions of multiple proteins.
