Quantitative Localization of a Golgi Protein by Imaging Its Center of Fluorescence Mass

The overall goal of this imaging-based method is to quantitatively localize a golgi protein at nanometer resolution. This method can help to answer the key questions in the study of the golgi apparatus such as the golgi's organization and how it works. The main advantage of this method is that it uses a conventional light microscope and software tools.

To begin, culture HeLa cells in a T25 flask in DMEM supplemented with 10%FBS and incubate the culture at 37 degrees Celsius and 5%carbon dioxide. When cells reach approximately 80%confluency, aspirate the culture medium and add one milliliter of 25%Trypsin EDTA to the flask. Incubate the cells at 37 degrees Celsius for two minutes.

Remove the cells from the incubator and add one milliliter of complete medium to gently flush the cells off the wall of the flask. Transfer the detached cells to a sterile centrifugation tube and spin the tube at 500 g's for two minutes to pellet the cells. After aspirating the supernatant, use one milliliter of complete medium to suspend the cell pellet.

Remove the medium from the well of a 24-well plate containing a sterilized glass cover slip and seed approximately 100, 000 cells into the well. Use complete medium to top up the volume of the well to 5 milliliters. Then, incubate the plate at 37 degrees Celsius with 5%carbon dioxide.

When the cells have reached 80%confluency, use 80 nanograms of GalT-mCherry and 320 nanograms of TPST1-EGFP plasmid DNA and transfection reagent to transfect the cells according to the manufacturer's protocol. After incubating the cells for four to six hours, change the medium and return the cells to the incubator for 12 hours. Aspirate the medium in the well and add 5 milliliters of 33 micromold Nocodazole containing complete medium and then incubate the cells at 37 degrees Celsius and 5%carbon dioxide for three hours.

After cleaning glass cover slips according to the text protocol, use one times PBS containing 1 microgram per microliter of BSA to dilute 110 nanometers multi-color fluorescent beads 80 fold. Briefly vortex the tube to disperse the bead aggregates. Using a pipette tip, spread 60 microliters of diluted beads onto a clean cover slip.

In the dark, place the cover slip in a 35 millimeter dish and cover it with aluminum foil. Dry the cover slip in the dark in a desiccator connected to a vacuum pump. Then use 50 microliters of mounting medium to mount the cover slip onto a glass slide.

Properly clean cover slips are essential to eliminate fluorescent debris. To image multi-color beads in green, red, and far red channels, at the beginning of the imaging session, acquire 3D stacks of images, taking three sections above and below the best focal plane. Save the stacks as TIFF files.

To image golgi mini-stacks, use TPST1-EGFP and GalT-mCherry transfected in fluorescently labeled slides to find cells that express TPST1-EGFP and a low or medium level GalT-mCherry. Acquire 3D image stacks as just demonstrated. To analyze the images, open image J, choose File, Image, Open, to open a set of bead images consisting of three TIFF files.

Choose Image, Stacks, Z Project, to average three consecutive sections around the best focused section in the red channel. Input the section number of start slice and stop slice and among options of projection type, select average intensity. Draw background ROIs that contain no beads in the image.

Then in Analyze, Set Measurements, check only mean gray value and Standard Deviation or SD.Execute Analyze, Measure, to obtain the background mean intensity and standard deviation. Choose Process, Math, Subtract, to subtract the image with the corresponding values of the background mean intensity plus six times the standard deviation. Then input the calculated value corresponding to this channel.

Repeat the averaging and subtraction for green and far red channel image stacks by using the same start and stop slice and the same background ROI. Merge the three processed images using Image, Color, Merge Channels, by selecting channel R as red, channel G as green, and channel B as blue. Launch the ROI Manager by choosing Analyze, Tools, ROI Manager.

Draw square ROIs around single beads. Then add the ROI to the ROI Manager by pressing T on the keyboard. Under Analyze, Set Measurements, check only center of mass.

In the ROI Manager, select channel R and click Measure to obtain the X and Y coordinates of the centers of ROIs in the results window. Copy and paste the two columns into a spreadsheet. Obtain the coordinates of the centers of ROIs for channel G and channel B.Save the spreadsheet as beads.csv.

In image J, use plugins install to install the two macros, macro golgi ROI inspection and macro output three channels data. Clear the ROI Manager and results window. Open a set of golgi mini-stack images consisting of three TIFF files.

Generate background subtracted images as demonstrated earlier in this video for later use. Then duplicate the images. Under Process, Image Calculator, add the background subtracted channel G, channel R, and channel B images.

To the resulting image under Image, Adjust, Threshold, select Set to input one as the lowest threshold level. Under Analyze, Analyze Particles, input the size range for size pixel too. Input 50 to infinity, check excludes on edges, and add to manager.

Next, use Image, Color, Merge Channels, to merge the three background subtracted images generated earlier by selecting channel R as red, channel G as green, and channel B as blue. Then run the macro macro golgi ROI inspection by selecting Plugins, Macro, Golgi ROI Inspection. It is important to select as many ROIs as possible that contains single golgi mini-stacks in a set of images.

Under Analyze, Set Measurements, check only area, mean gray value, and center of mass. Acquire data by launching the macro tool macro output three channels data. Copy coordinates of centers into a spreadsheet in the order listed here.

Then save the spreadsheet as ministacks.csv. Proceed to chromatic shift correction of the centers and Localization Quotient or LQ calculation according to the text protocol. The modern research grade light microscope equipped with the plain apochromatic lens such as the one used here shows minimal chromatic shift.

However, a careful examination of the multi-colored bead image reveals it. Centers of red fluorescence are defined as the true positions of beads. The relative chromatic shifts of green and far red fluorescence can be represented by vectors.

The chromatic shift within the imaging plane is illustrated by the green and blue vector for each bead. As illustrated here, chromatic shift correction significantly reduces the chromatic shift. In mammalian cells, the golgi complex is aggregated at the perinuclear area which is usually not resolvable under a conventional optical microscope.

After Nocodazole treatment, the perinuclear golgi complex disappears and dozens of golgi mini-stacks assemble at the endoplasmic reticulum exit sites throughout the cytoplasm. An example image that was generated using the image analysis demonstrated in this video is shown here. Using the macro golgi ROI inspection tool, 40 selected golgi mini-stacks were listed.

After applying the three criteria, 21 golgi mini-stacks were analyzable for the calculation of LQs of TPST1-EGFP. Once mastered, this technique can be done in one hour after fluorescence labeling. After its development, this method paved the way for researchers who study the golgi apparatus to quantitatively localize a golgi protein with resolution equivalent to that of immunoelectron microscopy.