Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules

This protocol is a guide for implementing interference reflection microscopy on a standard fluorescence microscope for label-free, high-contrast, high-speed imaging of microtubules using in vitro surfaces assays.

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The significance of this protocol is that it introduces a label-free imaging technique capable of acquiring high contrast images at high frame rates that is simpler and cheaper than other techniques like DIC and dark field. The main advantage of this technique is that it's easy to implement and it's easy to use. It's inexpensive and it can be combined with fluorescence microscopes easily.

The method was developed for visualizing individual microtubules. It has the potential to be used for visualizing large protein complexes and nanoparticles. The main difficulty is the activation energy required to tinker with the filter cube to add the half silver mirror.

My advice to someone who is trying the technique for the first time is just do it. To begin, insert a 50/50 mirror into the filter wheel of a fluorescent microscope using an appropriate filter cube. Handle the mirror with care as often they have an anti-reflection coding.

Turn to a high magnification objective that also has a high numerical aperture. The one shown here is a 100x oil objective with a numerical aperture of 1.3. Next, use a razor blade and a microscope slide as a straight edge to cut three millimeter wide strips of plastic paraffin film.

Place two of the plastic paraffin film strips three millimeters apart on a clean 22 by 22 millimeter cover slip, then place an 18 by 18 millimeter cover slip on top of the strips to form a channel. Transfer the cover slip to a heat block at 100 degrees celsius for 10 to 30 seconds for the paraffin film to form a sealed channel. Using a pipette, flow in 50 micrograms per milliliter of an anti-rhodamine antibody by perfusion and incubate the slide for 10 minutes.

Following incubation, wash the channel five times using filtered BRB80, then flow in 1%Poloxamer 407 in the filtered BRB80 to block the surface against non-specific binding and incubate the slide for 10 minutes. Again, wash the channel five times using filtered BRB80. To prevent the sample from drying out, add two droplets of the filtered BRB80 at the ends of the channel and add more buffer as needed.

Place the sample on the microscope stage and turn on the epi-illumination light source. Focus on the paraffin film edge to find the sample surface and then move the field to set the view to the center of the chamber. You will observe multiple surfaces as the objective is moved up and down due to back reflection of light from optics within the optical path.

Next, center the field diaphragm in the field of view by closing it halfway and using the adjustment screws. Once the diaphragm is properly aligned, re-open it. Then, slide in the Bertrand lens to view the back focal plane, also known as the exit pupil of the objective.

Close the aperture diaphragm beyond the edges of the exit pupil and use the adjustment screws to center the aperture diaphragm with respect to the exit pupil. Double check by opening the aperture diaphragm and matching its edges with those of the exit pupil. Then, set the aperture diaphragm to about two-thirds of the numerical aperture of the objective.

To begin, set the exposure time of the camera to 10 milliseconds and adjust the illumination to nearly saturate the camera dynamic range. Next, use a pipette to flow in 10 microliters of GMPCPP-stabilized microtubules in 0.22 micrometer filtered BRB80. Monitor the microtubule binding on the imaging of the surface.

Once 10 to 20 microtubules are bound within the field of view, wash the sample twice with the filtered BRB80. Acquire 10 images of the microtubules by setting up a time lapse with a 10 millisecond exposure and a one second delay period for a total of 10 seconds. Then, acquire background images by moving the stage using the stage controller along the channel's long axis while acquiring 100 images with no delay.

To image microtubule dynamics using brain tubulin, start by setting the sample heater temperature to 37 degrees celsius. Using a pipette, flow in 10 microliters of GMPCPP-stabilized microtubule seeds and monitor them binding to the surface by imaging the surface live. Once 10 to 20 seeds are bound within the field of view, wash the sample using twice the channel volume of pre-warmed and filtered BRB80.

Next, flow in 10 microliters of the polymerization mix. To measure microtubule growth, set up a time lapse using the acquisition software to acquire an image every five seconds for 15 minutes. Enhance the contrast by acquiring an averaged image of 10 at each time point.

Acquire background images as shown in the previous section. Calculate the median by going to image, selecting stack, then Z project, then median. Subtract the corresponding background by going to process, going to image calculator, and choosing subtract from the dropdown menu.

Make sure the 32-bit float result option is checked. For microtubule shrinkage, acquire images at 100 frames per second by setting the time delay to zero and keeping the exposure time at 10 milliseconds. An important factor for acquiring high contrast images with the interference induction microscopy is to properly set the numerical aperture of the illumination.

This can be done by guiding the size of the illumination beam at the exit pupil of the objective, which is controlled by the aperture diaphragm size. Using a sample of florescently-labeled stabilized microtubules, bring the microtubules into focus using the microscope's focusing knob while fluorescently imaging them. Set camera exposure to 10 milliseconds and close down the aperture diaphragm to its smallest opening.

Also, adjust the illumination to nearly saturate the camera's dynamic range or until the maximum is reached. Acquire 10 images by streaming a field of view containing 10 or more microtubules. Then acquire a background image.

Change the size of the diaphragm and adjust the illumination intensity to match that which was previously determined. Acquire 10 new images and a new background. Repeat this process until the entire range of the diaphragm is completed.

For every field of view acquired, subtract the corresponding background and average the resulting background subtracted images as shown previously. Then, calculate the average signal to background noise ratio of the microtubules for every opening size. Set the diaphragm size to the one producing the highest signal to background noise ratio.

It is possible that there is a range of sizes that produce comparable contrast as shown here. With a well-aligned microscope, microtubules should be visible without background subtraction. Subtracting the background, however, does enhance the contrast of the microtubule.

To further enhance the contrast, averaging, 4a filtering, or a combination of both can be used. The line scans shown here describe the incremental improvement of image quality. These describe a noticeable reduction of background noise with each processing step.

Microtubule dynamics can be reported in kinographs, which are generated from time lapse movies. An example kinograph acquired at a frame rate of 0.2 frames per second is shown here. The dashed lines mark the seeds.

When performing this procedure, it's important to work with high numerical aperture objectives. It's also important to align and set the size of the aperture diameter frame correctly. It is easy to combine IRM with fluorescence imaging to study, for example, microtubule binding proteins and how they modify microtubule dynamics.

The technique reduces photo damage considerably and allows for higher frame rate acquisition, making it easier to study labeled biopolymers such as microtubules for extended periods of time with high temporal resolution and good tracking precision.

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Journal (Bioengineering)

Journal (Bioengineering)

Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules

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