Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

Fluorescent speckle microscopy is used to probe the dynamics of various cytoskeletal proteins in living cells. When used correctly, quantitative measurements of the velocity and turnover kinetics of cytoskeletal proteins can be made. We begin this procedure by micro injecting a small volume of fluorescently labeled acton into living cells.

We then assemble the imaging chamber to reduce the effects of photobleaching. First two pieces of double-sided tape are placed on both sides of the micro slide where our cell cover slip will rest. Then imaging media is pipetted into the chamber by dispensing the media near one of the open slits.

Lastly, all four sides are sealed with valet. Hi, my name is James Lim and the laboratory of Galden's Deucer in the Department of Cell Biology at the Scripps Research Institute. Today we're gonna be showing you a procedure on fluorescent speckle microscopy.

We use this procedure to probe Acton Dynamics in migrating epithelial cells. So let's get started. A key component to acquiring good fluorescent speckle microscopy movies is the proper labeling of acton With the fluoro four of your choice, we recommend using fluoro fours of Alexa and X rodine cin, a middle Esther derivatives that target exposed lysine residues on the surface of Acton.

It is important for the labeling ratio to be high around 0.3 to 0.7 dyes per monomer of acton to ensure that speckles will appear bright and discreet prior to long-term storage. Clarify the labeled Acton solution by centrifuging at 75, 000 to 80, 000 times G for 20 minutes at four degrees Celsius. Labeled Acton should be stored in G buffer solution at negative 80 degrees Celsius.

Prepared two microliter aliquots to avoid repeated freeze thawing, which can lead to insoluble aggregates. To avoid photo bleaching of floral fours, do not expose the labeled actin solution to direct light for extended periods of time and use opaque micro centrifuge tubes for long-term storage plate. The cells for microinjection on acid washed square glass cover slips.

Standard substrate coating can be applied to the cover slip and will not affect microinjection. Store the cells at 37 degrees while the actin is being prepared to select cells that are amenable to microinjection. For FSM, consider these criteria.

The cells should be large, greater than 50 micrometers in diameter should spread flat one plated and be naturally adherent to substrates. It is also helpful if the site isol of the cell occupies an area greater than that of its nucleus. Some suitable cell lines include PTK one, newt lung cells and AIA neurons.

In this demonstration, we will inject PTK one epithelial cells. The correct selection of microinjection needles depends on the size of the tip opening. Typically, hole sizes can vary from 0.5 micrometers to as large as three micrometers.

We recommend an optimal opening size between 0.5 to one micrometer. For the injection of Acton Microinjection needles can either be purchased or made. If a commercial needle puller is available for beginners, we recommend having 10 to 20 pulled needles on hand before starting the injection procedure.

The next step is to dilute the labeled acton with ice cold injection buffer. Keep all working solutions on ice. The ideal injection concentration, which is the final concentration of acton loaded in the needle, is between 0.5 to one microgram per microliter.

Based on a 40%labeling ratio of D to actin monomer load 0.5 to one microliter of the diluted actin solution. Using a micro syringe needle, carefully dispense the acton solution. Try to avoid bubbles.

If there are bubbles, they can be easily removed by taking up excess liquids surrounding the bubble. Ensure that the Acton solution is resting completely at the needle tip. There should not be excess solution anywhere along the capillary of the needle because it can disrupt the injection flow.

The loaded needle is now ready to be attached to the injection holder for microinjection. Retrieve your cells from the incubator and place the cover slip on the microscope stage. To begin the microinjection procedure, take the loaded needle to the microscope.

Carefully attach the needle avoiding contact with the needle tip to the injector holder. Position the holder so that the needle makes a 35 to 50 degree angle to the base of the microscope stage. Lower the tip of the needle until it barely breaks the surface of the media bathing.

Your cells now position the needle tip so that it rests directly above the center of the objective. If the needle tip is aligned with the objective, you should see a bright halo through the eyepiece. This is the end of the needle tip.

Slowly moving the needle side to side may help better locate the halo. When navigating ensure the needle tip is well above your cells. To prevent breaking the needle tip, adjust the Z.Focus on the microscope until your cells are in clear view.

Then slowly lower the needle tip while controlling the focus until the halo gradually constricts and finally converges into the needle tip. You can also look for the shaft of the needle, which will cast linear shadows. Lowering the needle will gradually define the needle shaft.

Slight adjustments of the phase ring may be required to improve contrast both the end of the needle and your cells should now be in clear view. When selecting cells to inject cells with free edges should be facing the needle tip. This will ensure the declining slope of the cell from the thickest point where the nucleus resides to the thinnest point of the leading edge will meet the needle tip rather than run parallel to it.

Set the needle pressure at 0.3 to 0.8 PSI. The constant pressure applied will result in a steady flow of the acton solution. When you have decided on a cell to inject, position the needle so that the tip is next to the cell's nucleus slowly, lower and readjust the position of the needle until the tip is depressing.

The membrane incrementally lower the needle until its tip has pierced the membrane. If the needle has fully pierced the membrane, the cell will appear brighter. As soon as you see this change, raise the needle until it is no longer touching the cell.

The actual piercing and injecting process will take 0.5 to two seconds. When done correctly, the cell shape will appear unchanged. To minimize stress on cells, spend no more than 20 to 45 minutes for each microinjection session.

Sometimes you may not see changes in the cell when injecting and the cell does not get brighter. This indicates that the needle tip may be closed. You can test this by pressing the clean button if available on your trans injector.

If the needle tip is open, this will significantly increase the constant pressure flow, which can be seen as ripples and the dispersion of nearby debris. If the needle tip is closed, you can either insert a new microinjection needle or attempt to break the tip of the needle by gently lowering the needle until the tip of the needle presses against the glass cover slip. Thus causing the very end of the tip to break.

The breakage area should be no more than a one fifth of the area of the needle point the portion of the needle that starts to converge to a point when the microinjection is finished. Exchange the cell media in the dish with fresh prewarm media to remove any fluorescent residue. After changing the media, allow the cells to recover for 30 minutes in a 37 degree incubator before imaging.

This will also allow for the actin to integrate into the existing cytoskeleton network while the cells are recovering in the incubator, begin assembling the imaging chamber. First prewarm the imaging media in a 37 degree water bath. Next thaw, A 15 microliter aliquot of oxa erase at room temperature.

The addition of RA to the media later will greatly reduce the effects of photobleaching. Keeping the oxa erase wrapped in aluminum foil will ensure that the photo sensitive compound is protected from light. Prewarm the VAP sealant on a hot plate that is set to a low temperature do not overheat or burn the vap, or it will lose its efficacy as a sealant.

Wipe down a micro slide with a Kim wipe doused with pure ethanol to clean and remove small debris. Stack two equal sized lengths, 2.5 centimeters long of double-sided tape on top of each other using a clean flat surface such as another micro slide. As a platform, press out any trapped air bubbles to create a completely flat plane.

Next, use a razor blade to cut two straight strips along the long axis of the tape so that each strip measures 0.5 centimeters in width. Using forceps, place each strip along the long edges of your micro slide so that the two strips of tape create a gap in the center of your micro slide. Press gently on the tape to make a nice seal with the glass.

These strips of tape will act as spacers between the micro slide and the cover slip. Check on the valve to make sure it has completely liquified. Before you begin the next step, retrieve your prewarm imaging media.

Add 15 microliters of oxy rays to every 500 microliters of media. Fold a chem wipe into a triangle to create three hard edges. This will be used to wipe the cover slip later.

Retrieve the cells from the incubator using forceps. Pick up a corner of the cover slip quickly. Place the cover slip on a kim wipe with the cell side facing up and not touching the tissue, and use the folded Kim wipe to wipe two opposing sides of the cover slip to create a semi-dry surface.

Then pick up a corner of the cover slip with forceps and place the cover slip on the micro slide so that the dry surfaces are aligned to the two strips of tape. Notice now you have two closed sides and two open slits. Slowly pipette 200 microliters of your imaging media RA solution close to one opening.

It is important to not introduce any air bubbles. The solution should be sucked up into the newly assembled chamber by capillary forces, thus fully immersing your cells in solution. Using a cotton swab dab the four corners of your cover slip with melted vap to quickly affix and stabilize the cover slip.

You'll notice that the VAP dries within seconds at room temperature. Then beginning with the closed sides, lay a thin strip of VAP by using the cotton swab and mimicking a brushing stroke. Repeat for the open sides.

Slowly build up the coat of vap, reinforcing the seal by repeating the coating steps. The VAP should be confined to the edges leaving the center of the cover slip clear. After the edges of the cover slip are sealed, gently clean the center portion of the cover slip by using a Q-tip with DD H2O to remove any residual media.

Be careful not to crush the cells. Repeat this cleaning step with pure ethanol. You now have a fully enclosed imaging chamber optimized for fluorescent speckle imaging.

Having a fully sealed enclosure will prevent your floral fours from quickly photobleaching. The cells in the fully enclosed chamber are now ready for imaging. We use an inverted wide field microscope with a mercury lamp, epi illuminator, appropriate filters and oil immersion plan.

Apochromatic objectives images are captured with a cooled CCD camera with 6.7 micron pixels and an image acquisition software. Prewarm the microscope stage to 37 degrees. Place the imaging chamber and wait 10 to 15 minutes for the temperature to stabilize before imaging.

Once the temperature has stabilized, focus your cells in brightfield and then find your injected cells by toggling the correct fluorescent settings. Try to minimize the time of fluorescence exposure to reduce photobleaching. Look for injected cells that have healthy, curved leading edges.

Over injected cells will be characterized by retracted and jagged edges resembling numerous phylo podia. Do not image these cells. Acquisition settings will depend on the cell type, the quality of the labeled acton and microscope components.

However, exposure settings should not be greater than two seconds and time interval between frames should be set between five and 10 seconds. Gain settings on the camera can be turned up to increase the signal to noise. Ideal speckle characteristics should have each speckle spaced by approximately two speckle diameters to the next neighboring speckle.

For epithelial cells, the speckled actin network will appear homogeneous. With speckles evenly spread apart. Exposure settings should be adjusted to get the best speckled images.

The key to obtaining good speckle movies is maintaining focus. This is particularly important for post-analysis involving speckle flow and turnover measurements. Thus, the quality of the quantitative analysis will depend on locking in the focus for the duration of the time lapse.

Various strategies for maintaining focus are detailed in the text for this demonstration will be using the Nikon Perfect focus system. Here are various examples of micro injected cells. The left panel shows an under micro injected cell with a low concentration speckle image.

The middle panel shows an over micro injected cell with a too high concentration speckle image, and the right panel shows an optimally injected cell. For FSM with a good speckle image. We are just shown you how to micro inject fluorescently labeled acton into epithelial cells for fluorescent speckle microscopy.

When doing this procedure, it's extremely important to remember that the quality of your labeled acton, as well as the microinjection technique, will dictate the overall quality of your speckle images. So that's it and good luck with your experiments.