アクチンおよび誘導放出の枯渇 (STED) 顕微鏡を用いた B 細胞免疫シナプスにおける微小管細胞骨格の可視化

The overall goal of this protocol is to stain actin and microtubule cytoskeletons at the B-cell immune synapse for stimulated emission depletion, or STED microscopy, and imaging their structure, organization in spatial relationships. This method can help answer key questions about cytoskeleton organization during B-cell activation, including the reorganization of the actin and microtubule cytoskeletons during immune synapse formation. The main advantage of multi-color STED microscopy is that it allows simultaneous imaging of the actin cytoskeleton, the microtubule network, and key cytoskeleton associated proteins at nanometer scale resolution.

Although this method can provide insights into B-cell activation, it can also be applied to immune synapse formation of other immune cell types such as natural killer and T-cells. Generally, individuals new to this method will find that optimizing the microscope settings for the simultaneous imaging and multiple fluorophores using STED will require some practice. Demonstrating this procedure along with Jia and Madison will be Kate Choi and Caitlin Pritchard.

Begin by centrifuging 2.5 times 10 to the 6th, A20 B-lymphoma cells per each transfection, and re-suspending the cells in 100 microliters of transfection reagent supplemented with one to 2.5 micrograms of the plasma DNA of interest. Mix the solutions gently and transfer each aliquot of cells into individual wells of a six-well plate. Adjust the volume in each well to two milliliters with 37 degrees Celsius complete medium and place the cells in a 37 degree Celsius incubator for 18 hours.

The next morning, dip 1.5 18 millimeter round glass coverslips into 100%methanol and let the coverslips dry completely for about 10 minutes. Place the dried coverslips in individual wells of a 12-well tissue culture plate and add 400 microliters of goat anti-mouse immunoglobulin G antibody into the center of each coverslip so that a bubble forms at the center and spreads to the edges without dripping into the well. Incubate the coverslips at room temperature for 30 minutes.

Then rinse the coverslips with one milliliter of sterile PBS per well, three times, to remove any unbound antibodies. After the last wash, store each coverslip in one milliliter of fresh, sterile PBS until cell seeding. When the transfected cells are ready, collect them in a 15 milliliter conical tube by centrifugation and re-suspend the pellets in one milliliter of modified HEPES buffered saline supplemented with FBS.

After counting, dilute the cells to a two by 10 to the 5th cells per milliliter concentration and modified HEPES buffered saline plus FBS, and use forceps to transfer each coverslip onto a piece of paraffin film. Add 250 microliters of cells to each coverslip and incubate the coverslips at 37 degrees Celsius for 15 minutes in the dark to allow the cells to spread across the anti-immunoglobulin G coated surfaces. At the end of the incubation, remove the excess saline from each slide and coat each coverslip with 350 microliters of fixative, taking care that the entire surface is covered.

After 10 minutes in the dark at room temperature, use a micropipette to carefully aspirate the fixative from the edge of each coverslip and wash the coverslips with 500 microliters of permeabilization and blocking buffer. Then, permeabilize and block the cells with 250 microliters of fresh permeabilization and blocking buffer for 10 minutes at room temperature in the dark. Next, remove the excess buffer and label the cells with 50 microliters of the primary antibody of interest for 30 minutes in the dark at room temperature.

At the end of the incubation, wash the coverslip three times with 500 microliters of staining buffer per wash and add 50 microliters of the appropriate secondary antibody to each coverslip for 30 minutes at room temperature. After washing the secondary antibody, add 10 microliters of mounting reagent to a microscope slide and mount the coverslips onto individual microscope slides, cell side down. Then allow the slides to dry overnight in the dark for next day imaging.

To image the cells with STED microscopy, first turn on the STED microscope. Activate the lasers in the epifluorescence illumination lamp and open the microscope software. Set the parameters for the STED depletion lasers and load the first sample onto the microscope.

Using the eyepiece, manually focus on the sample to select a cell with a moderate GFP expression. Zoom in to the selected cell and select the region of interest to be imaged. Adjust the focus so that the XY plane of the cell that is closest to the coverslip is in focus.

Then, under the Acquire tab of the software, check the Between Frames option in the Sequential Scan panel to set the image acquisition to sequential frame acquisition. Set the STED laser power for each fluorophore and use the slide bar to adjust the laser powers according to the fluorophores used in the experiment. To increase the resolution and reduce the background signal, open the Acquisition settings and use the dropdown menu to select a value greater than one to increase the line and/or frame averaging.

Check the gaining option for each fluorophore and the specific values for time gaining to acquire the appropriate fluorescent signal for time gated STED and increase the STED laser power to enhance the resolution. It is critical to optimize the acquisition time gaining for the microscope sample and fluorophores being used to improve the resolution without losing too much of the fluorescent signal. Then manually acquire multiple images to ensure reproducibility and deconvolve the images using a deconvolution software package according to the manufacturer's instructions.

For A20 B-lymphoma cells, spread on immobilized NT immunoglobulin antibodies. STED microscopy used in conjunction with deconvolution software provides higher resolution images of cytoskeletal structures than does confocal microscopy alone. Although deconvolution of confocal images yields a substantial improvement in the resolution of the image, deconvolved STED images provide more detailed structural information than deconvolved confocal images.

Single-colored STED can also be used to acquire a three-dimensional super-resolution images of the entire B-cell cytoskeletal network. In these multi-colored STED images, staining of the peripheral ring of the dendritic actin can be observed in conjunction with microtubule staining. When using cells transfected with fluorescent fusion proteins, achieving optimal expression levels and avoiding artifacts due to over-expression are significant considerations, while too low an expression of the fluorescent fusion protein also results in poor quality images.

Careful focusing, to include the entire region of interest, is also essential. For example, in this image, only parts of the microtubule network was within the focal plane closest to the coverslip, preventing a complete analysis of the sample. Once mastered, this technique can be completed in two days.

One day for the sample preparation and staining and one day for the STED imaging. It's important to remember to titrate the amount of plasma to use to transfect the cells so that you get a sufficient expression of fluorescent protein for detecting during imaging while avoiding artifacts from over-expression. Following this procedure, the expression of different fluorescent fusion proteins can be induced to facilitate the localization of other proteins that regulate cytoskeletal dynamics and organization, or to visualize the sub-cellular distribution of proteins involved in other cellular processes.

After watching this video, you should have a good understanding of how to use STED microscopy for the imaging of cytoskeletal structures and proteins that are involved in immune synapse formation. Don't forget that working with fixatives such as glutaraldehyde can be extremely hazardous and that precautions such as working in a chemical fume hood should always be taken when using these kinds of reagents.