Name: implementation of interference reflection microscopy for label-free, high-speed imaging of microtubules
Uploaded: 2019-08-08T11:30:00.000+00:00
Duration: 9 min 45 s
Description: Watch this Scientific Journal Video about Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules at JoVE.com

The Authors thank Anna Luchniak and Yin-wei Kuo for critical reading and comments on the protocol.

1. Microscope modification and objective lens
<ol>
	<li>Insert a 50/50 mirror into the filter wheel of the fluorescent microscope using an appropriate filter cube (Figure 1). Handle the 50/50 mirror with care as often they have anti-reflection coating. 
	NOTE: We used a 50/50 mirror in an empty filter cube of the microscope. The 50/50 mirror is inserted where the dichroic mirror is located.</li>
	<li>Use a high magnification/high NA oil objective. 
	NOTE: In this protocol, we used a 100x/1.3 NA objective.</li>
</ol>
2. Chamber preparation to adhere microtubules to the surface
<ol>
	<li>Clean&#160;microscope slides and 22 mm x 22 mm coverslips (for upright microscope) or 22 mm x 22 mm and 18 mm x 18 mm coverslips (for inverted microscopes). Modify the surface as needed. For example, clean the coverslips using an alkaline detergent (see Table of Materials) followed by 100% ethanol with distilled water washes in between and after11.</li>
	<li>Cut 3 mm wide strips of plastic paraffin film (see Table of Materials) using a razor blade and another microscope slide as an edge.</li>
	<li>Place two plastic paraffin film strips 3 mm apart on a clean 22 x 22mm cover slip. Then place an 18 mm x 18 mm coverslip to form a channel. If using an upright microscope, place the strips on top of a clean microscope slide and then place a coverslip on top.</li>
	<li>Place the coverslip (or slide) on a heat block at 100 &#176;C for 10-30 s for the paraffin film to form a sealed channel.</li>
	<li>Flow in 50 &#181;g/mL of antibody (in Brinkley Buffer 1980 (BRB80)) by perfusion using a pipette. Incubate for 10 min. 
	NOTE: We use anti-rhodamine antibodies to bind rhodamine-labeled microtubule seeds to the surface. Alternatively, avidin can be used to bind biotin-labelled microtubule seeds or to biotinylated gold particles. To simply look at microtubules in the absence of tubulin in solution (i.e., a non-dynamic assay), an anti-tubulin antibody can be used.</li>
	<li>Wash five times with BRB80 buffer. It is recommended to filter the solutions using a 0.22 &#181;m filter.</li>
	<li>Flow in 1% poloxamer 407 (a triblock copolymer) in BRB80 to block the surface against non-specific binding. Incubate for 10 min.</li>
	<li>Wash five times with BRB80 buffer.</li>
	<li>The sample is ready to be placed on the microscope. To prevent the sample from drying out, add two droplets of BRB80 buffer at the ends of the channel and replenish when needed.</li>
</ol>
3. Microscope alignment
<ol>
	<li>Place the sample on the microscope stage. Turn on the epi-illumination light source.</li>
	<li>Focus on the sample surface. 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. A good method to find the sample surface is to focus on the paraffin film edge. Once the surface is found, set the field of view to the center of the chamber.</li>
	<li>Center the field diaphragm in the field of view by closing it half way and using the adjustment screws. Once aligned, reopen the diaphragm. 
	NOTE: Screw adjustment only needs to be done sporadically, perhaps every 3-6 months or when troubleshooting.</li>
	<li>Slide in the Bertrand lens to view the exit pupil (back focal plane) of the objective.</li>
	<li>Close the aperture diaphragm (AD) beyond the edges of the exit pupil of the objective.</li>
	<li>Use the adjustment screws to center the AD with the exit pupil. Double check by opening the AD and matching its edges with those of the exit pupil. 
	NOTE: This adjustment only needs to be done sporadically.</li>
	<li>Set the aperture diaphragm to about 2/3 of the NA of the objective. See section 7 for detailed procedures for optimizing the aperture diaphragm.</li>
</ol>
4. Imaging stabilized microtubules or 40 nm gold particle
NOTE: Stabilized microtubules and gold nanoparticles serve as good control samples. It is recommended to image surface attached microtubules or gold nanoparticles as a first step to assess IRM performance and help in setting the optimal aperture diaphragm opening (section 7).
<ol>
	<li>Set the exposure time of the camera to 10 ms and adjust the illumination to nearly saturate the camera dynamic range.</li>
	<li>Flow in 10 &#181;L of guanylyl-(alpha, beta)-methylene-diphosphonate (GMPCCP)- or taxol-stabilized microtubules in BRB804,12,13 (or 40 nm gold particles) by perfusion using a pipette to a fresh sample and monitor binding by imaging the surface. Once 10-20 microtubules are bound within the field of view, wash the sample 2x with BRB80. 
	NOTE: With a well aligned microscope, the microtubules should be visible without background subtraction.</li>
	<li>Acquire 10 images by setting a time lapse with 1 second delay period for 10 seconds (at 10 ms exposure).</li>
	<li>Acquire a background image. To do so, move the stage using the stage controller or computer software along the channel long axis while acquiring 100 images with no delay (i.e. streaming close to 100 fps @ 10 ms exposure). 
	NOTE: The background is the median of the 100 images. By taking the median, the illumination profile and other stationary features like dirt on the optics are preserved while everything else is filtered out. There should be no tilt on the sample as this will lead to change in the axial position as the stage is moved and ultimately degrade the background image. If the tilt cannot be avoided, then an alternative method is to acquire averaged background images before flowing the seeds.</li>
	<li>For processing and analyzing the images, go to step 6.</li>
</ol>
5. Imaging microtubule dynamics
<ol>
	<li>For microtubule dynamics using brain tubulin, set the sample heater temperature to 37 &#176;C.</li>
	<li>Flow in 10 &#181;L of GMPCCP-stabilized microtubule seeds in BRB8011 by perfusion using a pipette to a fresh sample and monitor them binding to the surface by imaging the surface live (i.e., while streaming). Once 10-20 seeds are bound with field of view, wash the sample using 2x channel volume with BRB80 (BRB80 should be prewarmed to 37 &#176;C or at least at room temperature). 
	NOTE: With a well aligned microscope, the seeds should be visible without background subtraction.</li>
	<li>Flow in the polymerization mix (7.5 &#181;M unlabeled tubulin + 1 mM guanosine triphosphate (GTP) + 1 mM Dithiothreitol (DTT) in BRB80 buffer). For measuring microtubule growth, set a time-lapse using the acquisition software to acquire an image every 5 seconds (0.2 frames per second (fps)) for 15 minutes.</li>
	<li>To enhance the contrast, acquire 10 images (instead of one) at each time point and average them. For microtubule shrinkage, acquire images at 100 fps by setting the time delay to 0 and an exposure time of 10ms (higher fps is possible with smaller regions of interest depending on the camera used).</li>
	<li>Acquire a background image as in step 4.4.</li>
</ol>
6. Image processing and analysis
NOTE: For analysis, this protocol uses Fiji14 but the reader is free to use any software she/he find suitable.
<ol>
	<li>Open saved background images.</li>
	<li>Calculate the median image (i.e. background) using Image &#62; Stack &#62; Z project &#62; Median.</li>
	<li>Open microtubule dynamics movie as a stack (same for non-dynamic microtubules) using File &#62; Open.</li>
	<li>Subtract the background image from the stack using Process &#62; Image calculator. Make sure to check the &#8220;32bit (float) result&#8221; option.</li>
	<li>For dynamic microtubules, using the line tool draw a line along the microtubule of interest and add it to the region of interest manager by pressing &#8220;t&#8221;. Repeat for all microtubules of interest.</li>
	<li>For dynamic microtubules, run the Multi-Kymo macro (Supplementary File 1). The macro will generate a video and a kymograph for every microtubule in the ROI manager. Every microtubule will get a unique identifier.</li>
	<li>For dynamic microtubules, run the Kymo-Analysis macro (Supplementary File 2) and follow its steps to measure the growth rates and shrinkage rates of the microtubules.</li>
</ol>
7. Aperture diaphragm size
NOTE: An important factor for acquiring high contrast images of microtubules using IRM is setting the illumination numerical aperture (INA) correctly10,15. The INA can be changed by changing the size of the incoming illumination beam at the objective&#8217;s exit pupil which is controlled by the size of the AD (the AD is located at a conjugate plane with the exit pupil (back focal plane) of the objective, Figure 1): 
<img alt="Equation" src="/files/ftp_upload/59520/59520eq1.jpg" /> 
where DAD is the diameter of the aperture diaphragm, fobjective is the focal length of the objective and Dep is the objective&#8217;s exit pupil diameter. Typically, the AD is left fully open for fluorescence imaging, so the INA equals the objective&#8217;s NA. In a fluorescence microscope, the AD scale does not indicate its diameter, thus the INA can&#8217;t be calculated. It&#8217;s possible to calibrate the AD size with the help of an objective. Yet, it&#8217;s not necessary since the AD size would be fixed to the size that produces the highest contrast.
<ol>
	<li>Prepare a sample of fluorescently labelled stabilized microtubules stuck to the surface (steps 4.1-4.2). 
	NOTE: We used tetramethylrhodamine labeled microtubules16 (Ex: 550 nm, Em: 580 nm).</li>
	<li>Bring microtubules into focus using the microscope focusing knob while fluorescently imaging them (if microtubules are not labelled, image them with IRM15 or DIC5).</li>
	<li>Set camera exposure to 10 ms using the camera software. 
	NOTE: This exposure is arbitrary and an exposure of 100 ms would also work.</li>
	<li>Close down the AD to its smallest opening. Adjust the illumination to nearly saturate the camera&#8217;s dynamic range or to the maximum possible. 
	NOTE: As a guide, use the look up table typically provided by the acquisition software.</li>
	<li>Acquire 10 images (by streaming or taking an image every second) of a field of view containing 10+ microtubules.</li>
	<li>Acquire a background image as in step 4.4.</li>
	<li>Change the size of the AD and repeat steps 7.5-7.6 for the whole AD opening range (from closed to the exit pupil size, Figure 2). Every time the AD size is changed, adjust the illumination intensity to match that of step 7.4.</li>
	<li>For every field of view acquired, subtract the corresponding background using process &#62; image calculator and choosing &#8220;subtract&#8221; from the drop-down menu. Make sure the &#8220;32bit (float) result&#8221; option is checked. Then average the resulting images using image &#62; stack &#62; Z project &#62; average.</li>
	<li>Measure the signal-to-background noise ratio (SBR) of the microtubules defined as the average intensity of the microtubule signal (intensity of the microtubule minus the intensity of the background) divided by the standard deviation of the background (Figure 3).</li>
	<li>Determine the optimal AD size (i.e. optimal INA) by calculating the average SBR&#160;of the microtubules for every opening size and set the AD size to the one producing the highest SBR (Figure 2). It is possible that there is a range of AD sizes that produce comparable contrast15.</li>
</ol>

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The authors have no conflicts of interest to disclose.

This protocol demonstrated the successful use of IRM for imaging and measurement of the microtubule dynamics. Care should be given to correctly set the illumination numerical aperture as it has the strongest impact on image contrast. Also, using high numerical aperture (NA) objectives is important for getting high resolution/high contrast images, as higher NA objective have higher light collecting power compared to low NA objectives. The cleaner the surface and solutions used the lower the noise as dirt ends up attaching to the surface and adding (over the course of the experiment) speckle like noise to the images. Acquisition of a background image is important as well as it removes illumination inhomogeneities, static noise and surface irregularities.
A recommended modification is to introduce a long pass filter (&#62;600 nm) in the illumination path. The spectrum of white light sources typically contains wavelengths in the UV which can damage microtubules. In addition, using long wave length for IRM comes in handy when combining IRM with fluorescence (e.g., when studying the effect of microtubule associated proteins (MAPs) on microtubule dynamics). Be aware that when imaging for expended period of times, sample drift (especially along the optical axis) decreases image contrast due to the deviation of the image plane from the background plane. Modern microscopes are often equipped with stabilization mechanisms (e.g., perfect focus (Nikon), Definite focus.2 (Zeiss), IX3-ZDC2 (Olympus)). An alternative solution is to thermally stabilize the setup either passively or actively18 or by correcting for drift19,20,21. Finally, microtubule contrast can be increased by reducing the size of field diaphragm (a 70% opening is good choice as it is a balance between increasing contrast and field of view size)15.
While IRM is suitable for imaging microtubules it is not sensitive enough to detect single proteins. For such application, iSCAT is a more suitable technique. Similarly, fluorescence and iSCAT are better suited if tracking precision of less than 10 nm is needed. For IRM, the measured length tracking precision is ~20 nm as shown in Figure 7.
Use of IRM in surface assays can go beyond microtubules; for example, molecular motors can be labelled with gold nanoparticles and tracked as they interact with microtubules. In addition a more advanced form of IRM known as reflective interference contrast microscopy (RICM)22 can, in principle, be used to further enhance microtubules contrast and obtain higher tracking precision.

Label-free imaging of microtubules is of interest as it circumvents the need for fluorescent labeling of tubulin to generate contrast in the images. Fluorescent labeling has several drawbacks: it not feasible if the protein concentration is low1 and photobleaching and photodamage limit the observation time. Several techniques have been used to image label-free microtubules, including video-enhanced differential interference contrast microscopy (DIC) and darkfield microscopy2,3,4,5. Recently, interferometric scattering microscopy (iSCAT)6, rotating-coherent-scattering microscopy (ROCS)7 and spatial light interference microscopy (SLIM)8, have also been used. All these techniques are capable of imaging microtubules and have proved to be valuable for studying microtubule dynamics. However, each have their own limitations. In DIC the contrast depends on the angle between the microtubule and the Nomarski prism axis. &#160;In darkfield, the microtubule signal is degraded by scattered light from impurities or surfaces defects. Though iSCAT exhibits extraordinary sensitivity (down to single proteins) and ROCS can image microtubules deeper into the sample, both methods are technically demanding, requiring laser scanners.
This protocol demonstrates how interference reflection microscopy (IRM)9, 10 can be set up as an alternative technique for label-free imaging of microtubules. IRM is easy to implement as it requires only the addition of an inexpensive 50/50 mirror to a standard fluorescent microscope. When used in conjunction with the software described here, IRM produces high contrast microtubule images, can image large fields of view at high speed, requires a onetime alignment, and can easily be combined with other techniques such as fluorescence imaging.

<table><tbody><tr><td>Microscope</td><td>Nikon</td><td>Ti-Eclipse</td><td>An inverted microscope used for perfoming the expriments</td></tr><tr><td>50/50 beam splitter</td><td>Chroma</td><td>21000</td><td>When buying make sure to choose the splitter dimensions that fit the cube used in the microscope</td></tr><tr><td>NIKON PLAN FLUOR 100X/0.5-1.3 Iris objective</td><td>Nikon</td><td>MRH02902</td><td>Imaging objective. This objective has a NA adjusting iris that was opened to NA 1.3</td></tr><tr><td>Mucasol universal detergent</td><td>Sigma-aAldrich</td><td>Z637181-2L</td><td>Used for cleaning coverslips and slides</td></tr><tr><td>plastic paraffin film (commerical name Parafilm M)</td><td>Sigma-aAldrich</td><td>P7793</td><td>Used for constructing flow channels</td></tr><tr><td>Anti-TAMRA antibody</td><td>Invitrogen</td><td>A-6397</td><td>Used to bind TAMRA labeled molecules (e.g. microtubules) to the sample surface. RRID (AB_2536196)</td></tr><tr><td>Poloxamer 407 (commercial name Pluronic F-127)</td><td>Sigma-aAldrich</td><td></td><td>Used for blocking the channel surface to prevent nonspecific binding</td></tr><tr><td>40 nm gold nanoparticles</td><td>Sigma-aAldrich</td><td>753637</td><td>Used as a control sample</td></tr><tr><td>20 nm gold nanoparticles</td><td>Sigma-aAldrich</td><td>753610</td><td>Used as a control sample</td></tr><tr><td>Zyla 4.2 Camera</td><td>Andor</td><td>Zyla 4.2</td><td>2048x2048 pixles (6.5&amp;micro;m pixel size) with quantum efficiency of 72% and 16bit dynamic range</td></tr><tr><td>Feista tracking software</td><td></td><td></td><td>https://www.bcube-dresden.de/fiesta/wiki/FIESTA</td></tr><tr><td>Stabilized microtubles</td><td></td><td></td><td>prepared in house (see references in text)</td></tr></tbody></table>

implementation of interference reflection microscopy for label-free, high-speed imaging of microtubules

There are several methods for visualizing purified biomolecules near surfaces. Total-internal reflection fluorescence (TIRF) microscopy is a commonly used method, but has the drawback that it requires fluorescent labeling, which can interfere with the activity of the molecules. Also, photobleaching and photodamage are concerns. In the case of microtubules, we have found that images of similar quality to TIRF can be obtained using interference reflection microscopy (IRM). This suggests that IRM might be a general technique for visualizing the dynamics of large biomolecules and oligomers in vitro. In this paper, we show how a fluorescence microscope can be modified simply to obtain IRM images. IRM is easier and considerably cheaper to implement than other contrast techniques such as differential interference contrast microcopy or interferometric scattering microscopy. It is also less susceptible to surface defects and solution impurities than darkfield microscopy. Using IRM, together with the image analysis software described in this paper, the field of view and the frame rate is limited only by the camera; with a sCMOS camera and&#160;wide-field&#160;illumination&#160;microtubule length can be measured with precision up to 20 nm with a bandwidth of 10 Hz.&#160;

As mentioned above, with a well aligned microscope, microtubules should be visible without background subtraction (Figure 4A). Subtracting the background (Figure 4B) enhances the contrast of microtubule (Figure 4C). To further enhance the contrast, averaging or Fourier filtering or a combination of both can be used (Figure 4D,F,E). The line scans in Figure 4G shows the incremental improvement of image quality. Notice the reduction of background noise with each processing step.
Examples of kymographs of microtubule dynamics generated from time-lapse movies are shown in Figure 5. The videos were acquired at two frame rates: 0.2 fps (slow) and 100 fps (fast). The former is suitable for measuring growth rates while the latter is more suitable for measuring shrinkage rate that is an order of magnitude faster than the growth rate.
For the case where gold nanoparticles are used for setting up the microscope, an example image is shown in Figure 6. Gold nanoparticles were passively attached to the surface. While 40 nm particles are recommended, it is also possible to image 20 nm particles, yet at a lower contrast.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59520/59520fig1.jpg" /> 
Figure 1. Schematic representation of IRM. (A) Epi-illumination from the light source passes through the aperture diaphragm before reaching the 50/50 mirror. The aperture diaphragm sets the beam width thus the illumination NA. The 50/50 mirror partially reflects the light up to the objective to illuminate the sample. Light reflected from the sample is collected and then projected onto the camera chip (by the tube lens) where it interferes to generate the image. Image contrast is the result of the interference between the light reflected from the glass/water interface (I1) and the light reflected from the water/microtubule interface (I2). Depending on the microtubule/surface distance (h), the optical path difference between I1 and I2 will result in a constructive (bright signal) or destructive (dark signal) or anything in between. For example, if light with a wavelength of 600 nm is used for imaging, the contrast will switch between dark and bright when the microtubule height changes by about 100 nm. The asterisk indicate conjugate planes &#160;(modified from15). (B) Example of the 50/50 mirror installation. A suitable filter cube was opened and the mirror was inserted where a dichroic mirror usually sits. The mirror was oriented as per manufacturer instructions. Then the cube was inserted in the filter wheel which was inserted back to the microscope (not shown). During the installation, gloves were used, and the mirror was only held by the edges. <a href="https://www.jove.com/files/ftp_upload/59520/59520fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59520/59520fig2.jpg" /> 
Figure 2. Optimal Aperture diaphragm setting. (A) Same field of view was imaged at different aperture diaphragm openings without background subtraction. Visually, the contrast increased as the size of the aperture diaphragm increased till it reached a plateau and started to degrade afterwards. This was confirmed by (B) SBR measurements of background subtracted images. Error bars are standard deviation. &#160;Scale bars are 500 &#181;m (AD) and 3 &#181;m (microtubules). <a href="https://www.jove.com/files/ftp_upload/59520/59520fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59520/59520fig3.jpg" /> 
Figure 3. Measuring signal-to-background noise ratio. Microtubules were isolated in regions of interest. Each region of interest was thresholded to separate the microtubule from the background. The average microtubule signal was obtained from a line scan across the microtubule. The scan line width was set to equal the microtubule length. This way, every point on the scan is an average of the signals of all pixels along the microtubule axis that are parallel to that point. The background noise is the standard deviation of all the pixels below threshold cut off.&#160; <a href="https://www.jove.com/files/ftp_upload/59520/59520fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59520/59520fig4.jpg" /> 
Figure 4. Image processing. After acquiring raw images (A), the background (B) was subtracted (C) to enhance the microtubule contrast. To further improve the contrast the images were either averaged (D) or Fourier filtered (E) or both (F). The line scans (G), whose location is indicated by the dashed red line in (A) are color matched to the various images in (A) to (F). The numbers at the lower corner are average SBRs measured for the whole field of view. The scale bar is 5 &#181;m (modified from15) . <a href="https://www.jove.com/files/ftp_upload/59520/59520fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59520/59520fig5.jpg" /> 
Figure 5. Examples of kymographs. (A) Kymograph examples of microtubule dynamics generated from time-lapse movies acquired at 0.2 fps. (B) Kymograph depicting an example of a shrinkage event generated from a movie acquired at 100 fps. Dashed lines mark the seeds. <a href="https://www.jove.com/files/ftp_upload/59520/59520fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59520/59520fig6.jpg" /> 
Figure 6. Example of gold nanoparticles imaged with IRM. Gold nanoparticles of sizes 20 and 40 nm were passively attached to the surface. 10 images were acquired. After background subtraction, the images were averaged to enhance contrast. <a href="https://www.jove.com/files/ftp_upload/59520/59520fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/59520/59520fig7.jpg" /> 
Figure 7. Microtubule length Tracking precision in IRM images. Stabilized microtubules (i.e., fixed lengths) were imaged 200x at 100 fps then averaged to 10 fps to enhance contrast. Next, the microtubules&#8217; lengths were measured using Fiesta17 tracking software. For every microtubule the mean length and standard deviation were calculated as shown in the figure (dashed line represents the mean and solid red lines represents the standard deviation, length = 3971 &#177; 20 nm. The overall tracking precision was the average of the standard deviation of all tracked &#160;microtubules (n = 6 microtubules x 20 data points = 120 data points).&#160;<a href="https://www.jove.com/files/ftp_upload/59520/59520fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary&#160;File 1.&#160;<a href="https://www.jove.com/files/ftp_upload/59520/Multi_Kymo.ijm" target="_blank">Please click here to download this file.</a>
Supplementary&#160;File 2.&#160;<a href="https://www.jove.com/files/ftp_upload/59520/Kymo_Analysis.ijm" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules at JoVE.com

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.

There are several methods for visualizing purified biomolecules near surfaces. Total-internal reflection fluorescence (TIRF) microscopy is a commonly used ...

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The Cytoskeleton II: Microtubules and Intermediate Filaments

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9.9K Views.  Yale University. 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.

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

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Biochemistry

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Label-Free Non-Linear Optics for the Study of Tubulin-Dependent Defects in Central Myelin

The satisfactory visualization of cytoskeletal components in the brain is challenging. The ubiquitous distribution of the networks of microtubules, microfilaments, and intermediate filaments in all the neural tissues, together with the variability in the outcomes of fluorescent protein fusion strategies and their limited applicability to dynamic studies of antibodies and drugs as chromophore vehicles, make classical optical approaches not as effective as for other proteins. When tubulin needs to be studied, the label-free generation of second harmonics is a very suitable option due to the non-centrosymmetric organization of the molecule. This technique, when conjugated to microscopy, can qualitatively describe the volumetric distribution of parallel bundles of microtubules in biological samples, with the additional advantage of working with fresh tissues that are unfixed and unpermeabilized. This work describes how to image tubulin with a commercial second harmonic generation microscopy setup to highlight microtubules in the tubulin-enriched structures of the oligodendrocytes, as&#160;in hypomyelination with atrophy of the basal ganglia and cerebellum (H-ABC) tubulinopathy, a recently described myelin disorder.

In this article, we present a protocol to detect microtubule-loaded oligodendrocytes in a model of tubulinopathy through a simple, innovative second harmonic generation microscopy approach.

The satisfactory visualization of cytoskeletal components in the brain is challenging. The ubiquitous distribution of the networks of microtubules, ...

Neuroscience

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, ...

Biology

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a genetically encoded marker for the positive ends of microtubules (EB3 labelled with tdTomato fluorescent protein) and high-speed, high-resolution, live-cell imaging using spinning disk confocal microscopy. Cell cycle synchronization and increased density of microtubules are achieved by inhibiting centrosomal separation in mitotic cells, and analysis of growth is performed using open-source U-Track software. The use of a bright and red-shifted fluorescent protein, in combination with the lower laser power and reduced exposure time required for spinning disk microscopy reduce phototoxicity and the probability of light-induced artifacts. This allows for imaging a larger number of cells in the same preparation while maintaining the cells in a growth medium under standard culture conditions. Because the analysis is performed in a supervised automatic fashion, the results are statistically robust and reproducible.

Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a ...

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as specialized arrays during mitosis to regulate chromosome segregation. Proteins that interact with microtubules include motors such as kinesins and dynein, which can generate active forces and directional motion, as well as non-motor proteins that crosslink filaments into higher-order networks or regulate filament dynamics. To date, biophysical studies of microtubule-associated proteins have overwhelmingly focused on the role of single motor proteins needed for vesicle transport, and significant progress has been made in elucidating the force-generating properties and mechanochemical regulation of kinesins and dyneins. However, for processes in which microtubules act both as cargo and track, such as during filament sliding within the mitotic spindle, much less is understood about the biophysical regulation of ensembles of the crosslinking proteins involved. Here, we detail our methodology for directly probing force generation and response within crosslinked microtubule minimal networks reconstituted from purified microtubules and mitotic proteins. Microtubule pairs are crosslinked by proteins of interest, one microtubule is immobilized to a microscope coverslip, and the second microtubule is manipulated by an optical trap. Simultaneous total internal reflection fluorescence microscopy allows for multichannel visualization of all the components of this microtubule network as the filaments slide apart to generate force. We also demonstrate how these techniques can be used to probe pushing forces exerted by kinesin-5 ensembles and how viscous braking forces arise between sliding microtubule pairs crosslinked by the mitotic MAP PRC1. These assays provide insights into the mechanisms of spindle assembly and function and can be more broadly adapted to study dense microtubule network mechanics in diverse contexts, such as the axon and dendrites of neurons and polar epithelial cells.

Here, we present a protocol for reconstituting microtubule bundles in vitro and directly quantifying the forces exerted within them using simultaneous optical trapping and total internal reflection fluorescence microscopy. This assay allows for nanoscale-level measurement of the forces and displacements generated by protein ensembles within active microtubule networks.

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as ...

TIRF Microscopy to Visualize Actin and Microtubule Coupling Dynamics

Source: Henty-Ridilla, J.L., <a href="https://app.jove.com/v/64074">Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy.</a> J. Vis. Exp. (2022)
This video describes TIRF, a total internal reflection microscopy-based technique to visualize actin and microtubule polymerization dynamics. The method allows the high-resolution imaging of actin and microtubule coupling dynamics in real-time, which is essential for understanding cellular crosstalks.

Source: Henty-Ridilla, J.L., Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy. J. ...

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Interference Reflection Microscopy for Label-Free Visualization of Microtubule Dynamics

Source: Mahamdeh, M., et al. <a href="https://app.jove.com/v/59520">Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules.</a> J. Vis. Exp. (2019).
In this video, we describe the interference reflection microscopy (IRM) technique to visualize microtubules growing on a coverslip surface in the presence of a suitable buffer. Upon illumination with incident light, the coverslip-buffer interface and buffer-microtubule interface reflect light, which combines to create an interference pattern, enabling the visualization of microtubules as high-contrast images against a bright background.

Source: Mahamdeh, M., et al. Implementation of Interference Reflection Microscopy for Label-free, High-speed Imaging of Microtubules. J. Vis. Exp. 
...

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

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Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Label-Free Non-Linear Optics for the Study of Tubulin-Dependent Defects in Central Myelin

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

TIRF Microscopy to Visualize Actin and Microtubule Coupling Dynamics

Interference Reflection Microscopy for Label-Free Visualization of Microtubule Dynamics