We thank Thierry Emonet&#39;s lab for sharing cleanroom equipments. We thank Yin-Wei Kuo for preparing the purified eGFP-kinesin used in this study. Y.T. acknowledges the support of the Alexander von Humboldt Foundation through the Feodor Lynen Research Fellowship.&#160;This work was supported by NIH Grant R01 GM139337 (to J.H.).

1. Preparation of flow chambers
NOTE: Microfluidic flow chambers will be constructed by adhering polydimethylsiloxane (PDMS) microchannels to a cleaned and functionalized cover glass. The microchannels will be cast in a master mold.
<ol>
	<li>Preparation of PDMS channels
	<ol>
		<li>Prepare cleaned and/or functionalized cover glasses with surface chemistry appropriate to the specific assay being imaged. 
		NOTE: For kinesin motility assays, piranha-cleaned silanized cover glasses are used. These are&#160;prepared as described in Gell et. al.1&#160;Alternatively, a simpler procedure is to sonicate cover glasses in isopropanol followed by methanol for 3x 20 min cycles.</li>
		<li>To prepare a master mold, cut strips of single-sided stationary tape to the desired flow channel size. Adhere the strips to the bottom of a 10 cm Petri dish, arranging them side-to-side with at least 1 cm spacing between the strips. 
		NOTE: If precise channel dimensions are required, the mold can be fabricated on silicon wafers using photolithography14.</li>
		<li>To prepare the PDMS polymer, combine the curing agent and the base elastomer in a 1:10 mass ratio. Mix for 2 min. 
		NOTE: This mixing ratio can be varied to tune the stiffness of the polymer. A higher ratio of base to curing agent results in a softer polymer.</li>
		<li>Degas the mixture in a vacuum chamber until all bubbles disappear.</li>
		<li>Pour the mixture onto the master mold in a ~0.5 cm thick layer, taking care to avoid creating bubbles.</li>
		<li>Bake the mixture in a preheated oven at 70 &#176;C for 40 min. 
		NOTE: A longer curing time may be required if the PDMS block is thick (&#62;1 cm). Continue heating in 5 min increments until it is fully cured.</li>
		<li>Cut out the structured regions of the polymer. Punch holes at each end of the channel using a PDMS puncher. 
		NOTE: In this protocol, the hole diameter is set to 0.75 mm and can be adjusted according to the required flow rates. PDMS channels can be stored in dry environments for extended periods but should be cleaned prior to use.</li>
		<li>Clean the structured side of the PDMS block. Use stationary tape to remove large particles. Rinse with isopropanol and then methanol. Repeat these rinses 3x, rinse with ultrapure water, and blow-dry the surface.</li>
		<li>Plasma clean the PDMS using oxygen or air plasma. 
		NOTE: In this protocol, 18 W air plasma is used.</li>
		<li>Place the plasma cleaned PDMS on an appropriately cleaned cover glass and heat on a hot plate at 80 &#176;C for 15 min. 
		NOTE: Epoxy resin can be applied to the sides of the PDMS block to better adhere it to the cover glass.</li>
		<li>Insert appropriately sized low-density polyethylene (LDPE) tubing into the holes. Connect the outlet tubing to a 0.5 mL syringe. 
		NOTE: In this protocol, tubes with an inner diameter of 0.023 in are connected to the PDMS via a 0.025 in outer diameter metal adaptor.</li>
		<li>Flow solutions into the microchannels by immersing the inlet tube in the solution and drawing the required volume with the syringe.</li>
	</ol>
	</li>
</ol>
2. Optical setup
<ol>
	<li>Modification of the microscope (Figure 1)
	<ol>
		<li>Modify a TIRF microscope to enable IRM imaging13. Replace the 50/50 (Reflectance (R)/Transmission (T)) beam splitter used in the filter wheel of the microscope with a 10/90 (R/T) beam splitter.</li>
		<li>Insert an image splitter between the camera and the microscope.</li>
		<li>In the filter cube of the image splitter, insert a 10/90 (R/T) beam splitter. Place a 600 nm long-pass filter in front of the reflected beam and an appropriate fluorescence emission filter in front of the transmitted beam. 
		NOTE: Beam splitters with different R/T ratios can be used to tune the fractions of the IRM and TIRF emission light that are collected.</li>
		<li>Align the image splitter according to the manufacturer&#39;s specifications to spatially separate the TIRF and IRM signals on the camera chip. 
		&#8203;NOTE: Spatial separation of the signals is required because the TIRF signal of typical fluorophores is much weaker than the IRM signal of a microtubule and would not be detectable if the two signals were overlayed.</li>
	</ol>
	</li>
</ol>
3. Imaging dynamic microtubules and single Kinesin molecules
<ol>
	<li>Surface functionalization and passivation
	<ol>
		<li>Flow BRB80 buffer (80 mM Na-PIPES, 1 mM EGTA, 1 mM MgCl2, titrated to pH 7.8 with NaOH) into the reaction chamber.</li>
		<li>Flow antibiotin solution diluted to 0.025 mg/mL in BRB80 and incubate at room temperature for 10 min.</li>
		<li>Wash the channel with BRB80.</li>
		<li>Flow in F-127 solution (1% Pluronic F-127 (w/v) dissolved in BRB80 overnight) and incubate for at least 20 min for surface passivation. 
		NOTE: If easy-cleaned cover glasses are used instead of silanized cover glasses, passivate using 2 mg/mL casein in BRB80 for &#62;20 min at room temperature.</li>
		<li>Wash the channel with BRB80.</li>
	</ol>
	</li>
	<li>Preparation for imaging
	<ol>
		<li>If temperature control is required, use an objective heater and set it to the correct temperature. 
		NOTE: In this protocol, all the experiments are performed at 28 &#176;C.</li>
		<li>Place the sample on the microscope stage and turn on the epiillumination light source (&#62;600 nm) for IRM imaging. 
		NOTE: In this protocol, a white LED light source is used with a 600 nm long-pass filter.</li>
		<li>Focus the microscope on the sample surface. Look for the correct focal plane near the PDMS-solution interface. 
		NOTE: In IRM, the aqueous interior of the channel should appear much brighter than the PDMS polymer.</li>
		<li>Pick a field of view near the center of the channel.</li>
		<li>Prepare a solution of 0.1 &#181;m fluorescent microbeads in BRB80 (density: 109&#160;beads/mL, corresponding to a 200-fold dilution for the beads used in this protocol).</li>
		<li>Flow in at least one channel volume of the microbead solution.</li>
		<li>Monitor the reaction via IRM. Wait for the microbeads to gradually &#34;land&#34; onto the surface. When the desired density of microbeads is achieved, wash out the excess with BRB80.</li>
	</ol>
	</li>
	<li>Growing biotinylated guanylyl 5&#39;-&#945;,&#946;-methylenediphosphonate (GMPCPP)-stabilized microtubule seeds
	<ol>
		<li>In a 0.6 mL centrifuge tube, prepare 50 &#956;L of a solution containing 1 mM GMPCPP, 1 mM MgCl2, and 2 &#956;M biotinylated tubulin (5-10% labeling stoichiometry) in BRB80. 
		NOTE: The correct labeling stoichiometry can be achieved by combining high-density biotinylated tubulin with unlabeled bovine tubulin in the correct ratio.</li>
		<li>Incubate the solution on ice for 5 min, then incubate at 37 &#176;C for 12.5 min. 
		NOTE: The length of the seeds can be controlled by tuning the polymerization time.</li>
		<li>Stop the polymerization by adding 100 &#956;L of room temperature BRB80.</li>
		<li>Spin down the solution in an ultracentrifuge at room temperature (126,000 x g, 5 min). Discard the supernatant by using a pipette to remove unpolymerized tubulin. 
		NOTE: In this protocol, an air-driven ultracentrifuge is used.</li>
		<li>Add 200 &#956;L of room temperature BRB80 to the pellet. Resuspend the pellet by pipetting gently but thoroughly. Use a 200 &#956;L pipette with a cut tip to reduce the shearing of the microtubules.</li>
	</ol>
	</li>
	<li>Growing dynamic guanosine diphosphate (GDP)-tubulin &#34;extensions&#34; 
	NOTE: The seeds will be immobilized on the antibiotin surface of the flow channel. Dynamic GTP/GDP &#34;extensions&#34; will be grown from the ends of the immobilized seeds.
	<ol>
		<li>Dilute the seeds 20x in BRB80. Flow the diluted seeds into the reaction chamber.</li>
		<li>Monitor the reaction via IRM. Wait for the seeds to gradually &#34;land&#34; onto the surface and bind to it. When the desired density of seeds is achieved, wash out the excess with BRB80.</li>
		<li>Prepare a microtubule extension mixture: 12 &#956;M unlabeled tubulin, 1 mM GTP, 5 mM dithiothreitol (DTT) in BRB80 buffer.</li>
		<li>Flow in at least one channel volume of the extension mixture. Ensure that the reaction temperature is 28 &#176;C.</li>
		<li>Wait for the microtubule extensions to grow from the seeds over time. 
		NOTE: The steady-state length is usually achieved in less than 20 min.</li>
		<li>Use the dynamic microtubules for imaging. Add and visualize fluorescent MAPs on the microtubules, as described in step 3.5 for kinesin-1. 
		NOTE: Because the microtubules are surface-bound, they are easily visible by TIRF.</li>
	</ol>
	</li>
	<li>Kinesin motility assay 
	NOTE: This step describes a protocol for visualizing motile green fluorescent protein (GFP)-labeled kinesin-1 on shrinking microtubules. A truncated rat kinesin-1 construct fused to eGFP (rKin430-eGFP) was expressed and purified, as described previously15,16.
	<ol>
		<li>Prepare motility buffer: 1 mM ATP and 0.2 mg/mL casein in BRB80.</li>
		<li>Dilute kinesin-eGFP in motility buffer to 10 nM.</li>
		<li>Prepare a 2x oxygen scavenger solution to counteract oxidative photobleaching (80 mM glucose, 80 mg/mL glucose oxidase, 32 mg/mL catalase, 0.2 mg/mL casein, 20 mM DTT) supplemented with 2 mM ATP.</li>
		<li>Combine 10 parts of oxygen scavenger, 9 parts of BRB80, and 1 part of 10 nM kinesin-eGFP for a final kinesin concentration of 0.5 nM. 
		NOTE: The total volume should be at least 1.5-fold larger than the channel volume.</li>
		<li>Set the imaging settings on the microscope software. 
		NOTE: In this protocol, videos are recorded at 10 fps with 100 ms exposure time. The laser intensity is ~0.05 kW/cm2.</li>
		<li>Begin imaging and flow the kinesin solution into the chamber.</li>
		<li>After the measurement is complete, record a short (~5 s) video in which the stage is slowly translated in a circular or lateral motion. Use the median projection of this video as a background image and subtract it from the raw measurements.</li>
	</ol>
	</li>
</ol>
4. Image processing and analysis
NOTE: Image processing was carried out using NIH ImageJ2 (imagej.nih.gov/ij/). A macro was developed to automate the splitting and alignment of the TIRF and IRM channels. This macro requires the GaussFit_OnSpot plugin to be installed (available on the ImageJ plugins repository).
<ol>
	<li>From the background recording, create a median projection in ImageJ by clicking on Image | Stacks | Z-project.</li>
	<li>Subtract the median background projection from the raw image data by clicking on Process | Image Calculator. 
	NOTE:&#160;Make sure to check the &#8220;32-bit (float) result&#34; option.</li>
	<li>For image registration, pick a collection of microbeads near the microtubule of interest and use them to align the TIRF and IRM images.</li>
	<li>For each bead in this collection, use the multipoint selection tool to mark the approximate location in the TIRF channel and then the corresponding location in the IRM channel. 
	NOTE: For example, if there are two beads (1 and 2), the multipoint selection would have four points in the following order: (1) bead 1 in TIRF, (2) bead 1 in IRM, (3) bead 2 in TIRF, (4) bead 2 in IRM.</li>
	<li>Run the provided ImageJ macro (ImageSplitterRegistration.ijm). 
	NOTE: This automates the following steps: i) estimating the center location of each bead by fitting to a Gaussian; ii) for each bead, computing the displacement vector separating the center in the TIRF channel from the center in the IRM channel; iii) averaging this displacement vector across all beads; iv) splitting the TIRF and IRM channels into separate images; v) translating the TIRF image by the average displacement vector computed in iii; vi) overlaying the TIRF and IRM images in a multichannel .tiff file.</li>
</ol>

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	<li>Gell, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule+dynamics+reconstituted+in+vitro+and+imaged+by+single-molecule+fluorescence+microscopy.">Microtubule dynamics reconstituted in vitro and imaged by single-molecule fluorescence microscopy.</a> Methods in Cell Biology. 95, 221-245 (2010).</li><li>Axelrod, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+7:+Total+internal+reflection+fluorescence+microscopy.">Chapter 7: Total internal reflection fluorescence microscopy.</a> Methods in Cell Biology. 89, 169-221 (2008).</li><li>Kuo, Y. -. W., Trottier, O., Mahamdeh, M., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spastin+is+a+dual-function+enzyme+that+severs+microtubules+and+promotes+their+regrowth+to+increase+the+number+and+mass+of+microtubules.">Spastin is a dual-function enzyme that severs microtubules and promotes their regrowth to increase the number and mass of microtubules.</a> Proceedings of the National Academy of Sciences of the United States of America. 116 (12), 5533-5541 (2019).</li><li>Hinrichs, M. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tau+protein+diffuses+along+the+microtubule+lattice.">Tau protein diffuses along the microtubule lattice.</a> The Journal of Biological Chemistry. 287 (46), 38559-38568 (2012).</li><li>Al-Hiyasat, A., Tuna, Y., Kuo, Y. -. W., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Herding+of+proteins+by+the+ends+of+shrinking+polymers.">Herding of proteins by the ends of shrinking polymers.</a> arXiv:. , (2021).</li><li>Guo, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanism+and+dynamics+of+breakage+of+fluorescent+microtubules.">Mechanism and dynamics of breakage of fluorescent microtubules.</a> Biophysical Journal. 90 (6), 2093-2098 (2006).</li><li>Moerner, W. E., Fromm, D. 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R., Popescu, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+imaging+of+single+microtubule+dynamics+using+spatial+light+interference+microscopy.">Label-free imaging of single microtubule dynamics using spatial light interference microscopy.</a> ACS Nano. 11 (1), 647-655 (2017).</li><li>Mahamdeh, M., Simmert, S., Luchniak, A., Sch&#228;ffer, E., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+high-speed+wide-field+imaging+of+single+microtubules+using+interference+reflection+microscopy.">Label-free high-speed wide-field imaging of single microtubules using interference reflection microscopy.</a> Journal of Microscopy. 272 (1), 60-66 (2018).</li><li>Mahamdeh, M., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Implementation+of+interference+reflection+microscopy+for+label-free,+high-speed+imaging+of+microtubules.">Implementation of interference reflection microscopy for label-free, high-speed imaging of microtubules.</a> Journal of Visualized Experiments: JoVE. (150), e59520 (2019).</li><li>Voldman, J., Gray, M. L., Schmidt, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfabrication+in+biology+and+medicine.">Microfabrication in biology and medicine.</a> Annual Review of Biomedical Engineering. 1, 401-425 (1999).</li><li>Rogers, K. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KIF1D+is+a+fast+non-processive+kinesin+that+demonstrates+novel+K-loop-dependent+mechanochemistry.">KIF1D is a fast non-processive kinesin that demonstrates novel K-loop-dependent mechanochemistry.</a> The EMBO Journal. 20 (18), 5101-5113 (2001).</li><li>Leduc, C., Ruhnow, F., Howard, J., Diez, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+fractional+steps+in+cargo+movement+by+the+collective+operation+of+kinesin-1+motors.">Detection of fractional steps in cargo movement by the collective operation of kinesin-1 motors.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (26), 10847-10852 (2007).</li><li>Tuna, Y., Al-Hiyasat, A., Howard, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+dynamic+microtubules+and+associated+proteins+by+Simultaneous+Interference-Reflection+and+Total-Internal-Reflection-Fluorescence+Microscopy.">Imaging dynamic microtubules and associated proteins by Simultaneous Interference-Reflection and Total-Internal-Reflection-Fluorescence Microscopy.</a> arXiv. , (2022).</li></ol>

The authors have no conflicts of interest to disclose.

Studying the regulation of microtubule dynamics by microtubule-associated proteins (MAPs) often requires simultaneous imaging of microtubules and MAPs. Fluorescence microscopy techniques such as TIRF are typically employed for this purpose. However, they are limited by the drawbacks of fluorescence imaging, which include photobleaching, photodamage, and the need for fluorophore labeling. Label-free methods, such as IRM, are suitable for visualizing microtubules but are not capable of imaging single fluorophores. This protocol combines label-free IRM imaging and TIRF microscopy for the simultaneous imaging of dynamic microtubules and MAPs.
The IRM setup employs an LED illumination source filtered to &gt;600 nm, while the TIRF setup utilizes a 488 nm laser. We used an inexpensive plate beam splitter to reflect the illumination light onto the sample and transmit the collected signal to the detector (Figure 1). A beam splitter with 10% reflectance and 90% transmission was chosen to minimize the loss of the single-molecule signal. The 90% loss in the illumination light intensity is compensated for by increasing the power of the illumination laser and LED. 
Spectral separation of the signals was achieved using a 90/10 (R/T) beam splitter and two spectral filters (long-pass 600 nm for IRM and band-pass 535/50 nm for TIRF). The spectrally separated IRM and TIRF signals are projected onto two halves of a single camera chip using an image splitter assembly. The use of a 90/10 beam splitter sacrifices 90% of the IRM signal, but this is compensated for by increasing the intensity of the LED illumination source. A dichroic mirror could also be used here to separate the IRM and TIRF signals more efficiently. Fluorescent microbeads included in the assays enable accurate alignment of the TIRF and IRM images and serve as a reference for focusing the objective.
The most critical optical element in this protocol is the high numerical aperture (NA) objective. This is essential not only to achieve total internal reflection but also to maximize the collection efficiency and image contrast. The quality of the obtained images also depends on the cleanliness of the glass surface and the acquisition of a clear background image to correct for nonuniform illumination and remove static features. For IRM imaging, we recommend using long-wavelength illumination (&gt;600 nm) to minimize the photodamage of microtubules and proteins. This is particularly important if a white LED light source is used, in which case a long-pass filter should be included to remove any UV light. 
This protocol allows for label-free, high-speed imaging of dynamic microtubules and simultaneous high-resolution visualization of fluorescent MAPs17. Compared to the filter cube switching technique, which alternates between capturing images of microtubules and MAPs, this setup is capable of much higher frame rates because it does not depend on the physical rotation of a filter cube. Compared to two-color TIRF imaging techniques, this technique employs a less demanding optical setup and circumvents the need for fluorophore labeling of microtubules. The primary limitations of this setup are due to the TIRF imaging of the MAPs; the frame rate is limited by the exposure time of a fluorophore, and photobleaching of fluorophores remains a possibility. Nonetheless, this protocol improves upon existing techniques because it uses TIRF only when necessary (i.e., to visualize MAPs but not microtubules) and achieves the highest speed possible within the limits of TIRF. Further improvements are possible only if both the microtubules and the MAPs are visualized via an interferometric technique, but this requires labeling MAPs with metal nanoparticles, which has its limitations and experimental challenges.
To demonstrate the capabilities of this technique, we simultaneously visualized two fast dynamical processes via IRM and TIRF: the shrinkage of a microtubule and the walking of a fluorescent kinesin molecule. This technique has been previously employed to visualize the fast diffusion of spastin on shrinking microtubules5. Beyond this application to MAPs and microtubules, this protocol can be used to visualize single fluorescent molecules simultaneously with any macromolecular structure massive enough to be visualized via IRM, such as a cell membrane or actin filament.

Total-internal-reflection-fluorescence (TIRF) microscopy is commonly employed for the visualization of single fluorescent molecules. Compared to epifluorescence imaging, TIRF achieves superior background suppression, allowing for high-resolution localization and tracking of single fluorophores. For this reason, TIRF is the preferred method for visualizing fluorescently labeled microtubule-associated proteins and is frequently used to image microtubules1,2.
To investigate the regulation of microtubule dynamics by MAPs, it is often necessary to image both microtubules and MAPs simultaneously. Most existing methods for this purpose are expensive or suffer from technical drawbacks. Simultaneous two-color TIRF, for example, requires two excitation lasers and two cameras. In addition to the high cost, the need for separate cameras poses frame-synchronization and image-registration problems. This need can be circumvented if a rotating filter cube is used to physically switch between excitation lasers in consecutive frames3. In such a setup, a single camera chip can be used, and the frames alternate between images of microtubules and MAPs. This technique, however, is limited by the speed of the filter change, which typically restricts the frame rate to less than 0.5 frames per second3 (fps). Such a frame rate is insufficient to resolve fast dynamic processes, such as the shrinkage of a microtubule occurring at a velocity of up to 500 nm/s, the walking of a kinesin at a velocity on the order of 800 nm/s, or the diffusion of a MAP occurring with diffusion coefficients exceeding 0.3 &#956;m2/s4. This is particularly problematic when tracking the relative positions of two moving targets in each channel, such as the position of a MAP relative to the position of a moving microtubule tip5.
In addition to these optical constraints, two-color TIRF microscopy requires MAPs and microtubules to be labeled with different fluorophores whose emission spectra are sufficiently separated. Fluorescent labeling of tubulin can alter microtubule dynamics6, and the photobleaching of fluorophores limits the imaging speed7. Because of these issues, label-free imaging techniques have been developed to visualize microtubules. These include interferometric scattering microscopy (iSCAT)8,9, rotating-coherent-scattering microscopy (ROCS)10, spatial light interference microscopy (SLIM)11, and interference reflection microscopy (IRM)12,13. These techniques allow fast label-free imaging of microtubules without the drawbacks of fluorescence imaging, but they cannot be used to visualize single MAPs.
Of these label-free techniques, IRM stands out for its low cost and its modest demands on instrumentation. We have recently presented a protocol for combining IRM with a commercial TIRF microscope, allowing microtubules and fluorescent MAPs to be imaged in alternating frames3,13. This paper presents a protocol for modifying this setup to simultaneously capture TIRF and IRM images on a single camera chip. This involves the addition of an inexpensive beam splitter in the excitation path to simultaneously illuminate the sample with a TIRF laser and an IRM LED light source. A modified commercial image splitter is used to spectrally separate the TIRF and IRM signals and project them onto separate halves of the same camera chip. We also employ a microfluidic system that allows the rapid exchange of reagents during imaging. This protocol describes how this setup can be used to image dynamic microtubules and MAPs. The capability of the apparatus is demonstrated by presenting the first visualization of kinesin-1 proteins walking on shrinking microtubules, which is captured at a frame rate of 10 s-1.

<table><tbody><tr><td>10/90 (R/T) beam splitter</td><td>Thorlabs</td><td>BSN10R</td><td>Plane beam splitter used in the excitation beam path</td></tr><tr><td>90/10 (R/T) beam splitter</td><td>Thorlabs</td><td>BSX10R</td><td>Plane beam splitter used in the image splitter</td></tr><tr><td>Anti-biotin antibody</td><td>Sigma-Aldrich</td><td>B3640</td><td>Used to functionalize surface for bonding biotinylated microtubules</td></tr><tr><td>ATP</td><td>Sigma-Aldrich</td><td>FLAAS</td><td>Used for preparing the motility buffer</td></tr><tr><td>Band-pass filter</td><td>Newport</td><td>HPM535-50</td><td>Hard-coated band-pas filter is used in image splitter to image GFP signal</td></tr><tr><td>Biotinylated tubulin</td><td>Cytoskeleton, Inc.</td><td>T333P-A</td><td>Used to bind microtubule seeds to the surface of the flow channel</td></tr><tr><td>Casein</td><td>Sigma-Aldrich</td><td>C8654</td><td>Casein is used to block nonspesific interactions</td></tr><tr><td>Catalase</td><td>Sigma-Aldrich</td><td>C9322</td><td>Used for preparing the oxygen scavenger solution</td></tr><tr><td>Desiccator chamber</td><td>Southern Labware</td><td>55207</td><td>Desiccator is used for degasing the resin</td></tr><tr><td>DTT</td><td>Sigma-Aldrich</td><td>D0632</td><td>Used for preparing the oxygen scavenger solution</td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E4378</td><td>Used for preparing the BRB80 buffer</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G7528</td><td>Used for preparing the oxygen scavenger solution</td></tr><tr><td>Glucose oxidase</td><td>Sigma-Aldrich</td><td>G7016</td><td>Used for preparing the oxygen scavenger solution</td></tr><tr><td>GMPCPP nucleotides</td><td>Jena Bioscience</td><td>NU-405L</td><td>Used for the polymerization of stabilized microtubules</td></tr><tr><td>Image splitter</td><td>Teledyne-Photometrics Imaging</td><td>OptoSplit II</td><td>An image splitter is used to split the images spatially. When buying, make sure about the compatibility with the microscope</td></tr><tr><td>ImajeJ2</td><td>NIH</td><td></td><td>ImageJ2 is used for image analysis</td></tr><tr><td>Kinesin</td><td></td><td></td><td>prepared in house (see references in text)</td></tr><tr><td>LDPE tubing</td><td>Thomas Scientific</td><td>9565S22</td><td>Non-toxic, lower density polyethylene micro bore tubing is used for fluid transfers</td></tr><tr><td>LED light source</td><td>Lumencor</td><td>Lumencor sola light engine</td><td>Used for IRM imaging</td></tr><tr><td>Long-pass filters</td><td>Thorlabs</td><td>FELH0600</td><td>Hard-coated long pass filters. One is used as an excitation filter, other is used in image splitter to image IRM signal</td></tr><tr><td>Magnesium chloride</td><td>Sigma-Aldrich</td><td>63068</td><td>Used for preparing the BRB80 buffer</td></tr><tr><td>Micoscope objective heater</td><td>okolab</td><td>H401-T-DUAL-BL</td><td>Used to keep sample temperature constant via heating the objective</td></tr><tr><td>Microscope</td><td>Nikon</td><td>Ti-Eclipse</td><td>An inverted microscope that is used in the experiments</td></tr><tr><td>Na-PIPES</td><td>Sigma-Aldrich</td><td>P2949</td><td>Used for preparing the BRB80 buffer</td></tr><tr><td>Nikon CFI Apochromat TIRF 100XC Oil objective</td><td>Nikon</td><td>MRD01991</td><td>The imaging objective has 1.49 numerical aperture</td></tr><tr><td>PDMS and curing agent</td><td>Electron Microscopy Sciences</td><td>Sylgard 184 (24236-10)</td><td>Used for contructing the flow channels</td></tr><tr><td>PDMS puncher</td><td>World Precision Instruments LLC</td><td>504529</td><td>Used to punch hole into the PDMS</td></tr><tr><td>Plasma cleaner</td><td>Harrick Plasma</td><td>DPC-32G</td><td>The air plasma is used to remove organic contamination from the PDMS surface</td></tr><tr><td>Poloxamer 407 (commercial name Pluronic F-127)</td><td>Sigma-aldrich</td><td>P2443</td><td>Used for channel surface passivation to minimize nonspecific binding</td></tr><tr><td>Sodium hydroxide</td><td>Sigma-Aldrich</td><td>567530</td><td>Used for preparing the BRB80 buffer</td></tr><tr><td>Stabilized microtubules</td><td></td><td></td><td>prepared in house (see references in text)</td></tr><tr><td>Table-top ultracentrifuge</td><td>Beckman Coulter</td><td>340400</td><td>Used to spin down microtubule seeds</td></tr><tr><td>TetraSpeck beads</td><td>ThermoFisher Scientific</td><td>T7279</td><td>Used as a reference for aligning images</td></tr><tr><td>Zyla 4.2 camera</td><td>Andor</td><td>Zyla 4.2</td><td>Scientific CMOS camera with spesifications: 2048 x 2048 pixels (6.5 &amp;mu;m pixel size) with quantum efficiency of 72% and 16 bit dynamic range</td></tr></tbody></table>

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.

The optical setup is schematized in Figure 1. Both IRM illumination and TIRF excitation light are directed to the back aperture of the objective (100x, NA: 1.49) via a 10/90 (R/T) beam splitter (BS1). The emitted signal passes through the same beam splitter (BS1) and is reflected to the image splitter via a mirror (M1). The components of the image splitter (enclosed with dashed lines in Figure 1) separate the IRM and TIRF signals via a 90/10 (R/T) beam splitter (BS2) together with appropriate spectral filters. Finally, the split images are projected onto the camera chip for visualization. The alignment of the image splitter is such that the TIRF and IRM signals are projected on separate halves of the chip.
In a well-aligned microscope, the camera image should display a halfway split image, as presented in Figure 2. Surface-bound microtubules should be easily visible in the IRM channel13, and fluorescent kinesin should be visible in the TIRF channel.
The microbeads used to align and register the two channels appear as bright spots in the TIRF images and dark spots in the IRM images. Although the beads are visible in the raw data, background subtraction improves the contrast significantly (Figure 2). The background image used for the subtraction is the temporal median of a video recorded with a moving stage. As described in the protocol, image alignment was performed by selecting a collection of beads near the region of interest and executing the provided macro (imageSplitterRegistration.ijm). The macro fits the points to Gaussians and aligns the images by minimizing the average distance between the center points of the fits in each channel. This process is represented in Figure 2, which shows a good alignment of the fluorescent microbeads (green in the TIRF channel, black in the IRM channel).
Finally, the capabilities of this simultaneous imaging setup are demonstrated by observing single kinesin molecules walking towards the shrinking ends of microtubules. Figure 3 shows a kymograph of eGFP-labeled kinesin molecules (green) walking on a shrinking microtubule (gray). Also presented is a series of snapshots from the recording from which the kymograph was generated.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63730/63730fig01.jpg" /> 
Figure 1: Schematic representation of the optical setup for simultaneous IRM and TIRF imaging of kinesin motility. Epiillumination from an LED light source passes through the aperture diaphragm and reaches the 10/90 (R/T) beam splitter (BS1). The beam splitter partially reflects the red IRM illumination light and the 488 nm TIRF excitation light up to the objective to illuminate the sample. The signal from the sample is collected by the same objective and directed to the image splitting assembly where IRM and TIRF images are spatially separated by the 90/10 (R/T) beam splitter (BS2). The signals are then spectrally filtered before reaching the camera chip. Abbreviations: IRM = interference reflection microscopy; TIRF = total-internal-reflection-fluorescence; LED = light-emitting diode; ITIRF = TIRF illumination; IGFP = GFP fluorescence; Iinc = IRM illumination; Iref = scattered light at the glass/water interface; Iscat = scattered light from the microtubule; IIRM = IRM signal (Interference of Iref and Iscat); R/T = reflected/transmitted; LP600: long pass filter (600 nm); DM = dichroic mirror; BS1 and BS2 = beam splitters 1 and 2; M1, M2, M3, M4 = mirrors; BP535/50 = band pass (535/50 nm); GFP = green fluorescent protein; GMPCPP = guanylyl 5&#39;-&#945;,&#946;-methylenediphosphonate; GDP = guanosine diphosphate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63730/63730fig01large.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/63730/63730fig02.jpg" /> 
Figure 2: Background subtraction and image alignment. The TIRF (left half) and IRM (right half) images appear simultaneously on two halves of the same camera chip (camera image). Temporal median background subtraction increases the contrast of the beads (background-subtracted image), which appear dark in IRM and bright in TIRF images. IRM and TIRF images are aligned by translation (right) based on the localization of selected beads (white rectangles). Scale bars = 10 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63730/63730fig02large.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/63730/63730fig03.jpg" /> 
Figure 3: Kymograph and snapshots of Kinesins&#39; movement during microtubule shrinkage. The kymograph (left) shows eGFP-kinesin-1 (green) walking towards the plus end of the microtubule (dark grey). Snapshots from the corresponding time series are shown (right). White arrows show single kinesin-1 molecules. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63730/63730fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
ImageSplitterRegistration File: <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63730/ImageSplitterRegistration.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins at JoVE.co

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

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. ...

simultaneous-interference-reflection-total-internal-reflection

Nanyang Technological University

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Unit 1

The Cytoskeleton II: Microtubules and Intermediate Filaments

SCIENTISTS IN ACTION

3.4K Views.  Yale University. Het belang van deze methode is dat het labelvrije microtubuli tegelijkertijd met fluorescerend gelabelde eiwitten met hoge framesnelheden mogelijk maakt. Het belangrijkste voordeel van deze techniek is de eenvoudige implementatie en minimale eisen aan optische componenten. Het omzeilt ook de noodzaak van fluoro-4-etikettering van microtubuli.Bereid om te beginnen het PDMS-polymeer voor door het uithardingsmiddel en het basiselastomeer in een massaverhouding van 1:10 te combineren. Meng...

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

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Bioengineering

A TIRF Microscopy Technique for Real-time, Simultaneous Imaging of the TCR and its Associated Signaling Proteins

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex (pMHC) proteins expressed on the surface of antigen presenting cells (APCs). The TCR complex is composed of the ligand binding TCR&alpha;&beta; heterodimer that associates non-covalently with CD3 dimers (the &#949;&delta; and &#949;&gamma; heterodimers and the &zeta;&zeta; homodimer)1. Upon engagement of the receptor, the CD3 &zeta; chains are phosphorylated by the Src family kinase, Lck. This leads to the recruitment of the Syk family kinase, Zap70, which is then phosphorylated and activated by Lck. After that, Zap70 phosphorylates the adapter proteins LAT and SLP76, initiating the formation of the proximal signaling complex containing a large number of different signaling molecules2. 
The formation of this complex eventually results in calcium and Ras-dependent transcription factor activation and the consequent initiation of a complex series of gene expression programs that give rise to T cell differentiation2. TCR signals (and the resulting state of differentiation) are modulated by many other factors, including antigen potency and crosstalk with co-stimulatory/co-inhibitory, chemokine, and cytokine receptors 3-4. Studying the spatial and temporal organization of the proximal signaling complex under various stimulation conditions is, therefore, key to understanding the TCR signaling pathway as well as its regulation by other signaling pathways.
One very useful model system to study signaling initiated by the TCR at the plasma membrane in T cells is glass-supported lipid bilayers, as described previously5-6. They can be utilized to present antigenic pMHC complexes, adhesion, and co-stimulatory molecules to T cells-serving as artificial APCs. By imaging the T cells interacting with the lipid bilayer using total internal reflection fluorescence microscopy (TIRFM), we can restrict the excitation to within 100 nm of the space between the glass and the cell surface 7-8. This allows us to image primarily the signaling events occurring at the plasma membrane. As we are interested in imaging the recruitment of signaling proteins to the TCR complex, we describe a two-camera TIRF imaging system wherein the TCR, labeled with fluorescent Fab (fragment antigen binding) fragments of the H57 antibody (purified from hybridoma H57-597, ATCC, ATCC Number:HB-218) which is specific for TCR&beta;, and signaling proteins, tagged with GFP, may be imaged simultaneously and in real time. This strategy is necessary due to the highly dynamic nature of both the T cells and of the signaling events that are occurring at the TCR. This imaging modality has allowed researchers to image single ligands 9-11 as well as recruitment of signaling molecules to activated receptors and is an excellent system to study biochemistry in-situ12-16.

The compartmentalization of proteins either within the plasma membrane or into intracellular locations is one regulatory mechanism that can greatly influence signaling outcomes; hence, to understand signaling it is important to study the spatial and temporal behavior of the proteins involved. We describe here a TIRF microscopy based system to study signal transduction in T cells, but is broadly applicable.

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex ...

Immunology and Infection

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

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 ...

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 ...

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;

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 ...

TIRF Microscopy to Visualize Actin and Microtubule Coupling Dynamics

Source: Henty-Ridilla, J.L., <a href="https://app-jove-com.remotexs.ntu.edu.sg/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. ...

JoVE Encyclopedia of Experiments

Biological Techniques

Microscopy Techniques

Interference Reflection Microscopy for Label-Free Visualization of Microtubule Dynamics

Source: Mahamdeh, M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/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. 
...

Total Internal Reflection Fluorescence Microscopy

Total internal reflection fluorescence microscopy or TIRF is an advanced microscopic technique used to visualize fluorophores in samples close to a solid surface with a higher refractive index, such as a glass coverslip. TIRF only allows fluorophores in proximity to the solid surface to be excited. When light from a medium with a lower refractive index (such as air) hits the glass coverslip at a critical angle, the light undergoes total internal reflection stead of passing through the glass. This happens as the sample has a lower refractive index than the coverslip and does not allow the light to enter within. The light is reflected from the interface and forms an electromagnetic field emitting short-length evanescent waves. These waves only excite the fluorophore near the surface as they can move only about 100 to 200 nm deep within the cell before dying out.
There are two types of TIRF; prism-based and objective-based. In prism-based TIRF microscopy, a prism is placed on the coverslip surface that directs the evanescent wave to the sample. In objective-based TIRF microscopy, there is no prism at the interphase; the objective is the same as the light source that helps create the evanescent wave.
TIRF has several advantages over traditional fluorescence and confocal microscopes; it prevents the illumination of background fluorophores. It helps in studying the structures close to the cell surface. It reduces the blurring effect and does not allow out-of-focus light to interfere with the image. As the samples are not directly exposed to an intense light beam, the photobleaching is minimum, and the cells are less exposed to phototoxicity.

Total internal reflection fluorescence microscopy or TIRF is an advanced microscopic technique used to visualize fluorophores in samples close to a solid ...

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

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