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>

<ol>
	<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. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+single-molecule+fluorescence+spectroscopy+and+microscopy.">Methods of single-molecule fluorescence spectroscopy and microscopy.</a> Review of Scientific Instruments. 74 (8), 3597-3619 (2003).</li><li>Taylor, R. W., Sandoghdar, V., Astratov, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Interferometric scattering (iSCAT) microscopy and related techniques. in Label-free super-resolution microscopy. , 25-65 (2019).</li><li>Young, G., Kukura, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interferometric+scattering+microscopy).">Interferometric scattering microscopy).</a> Annual Review of Physical Chemistry. 70 (1), 301-322 (2019).</li><li>Koch, M. D., Rohrbach, A. <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+and+bending+analysis+of+microtubules+by+ROCS+microscopy+and+optical+trapping.">Label-free imaging and bending analysis of microtubules by ROCS microscopy and optical trapping.</a> Biophysical Journal. 114 (1), 168-177 (2018).</li><li>Kandel, M. E., Teng, K. W., Selvin, P. 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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#956;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/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/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/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/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. ...

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The Cytoskeleton II: Microtubules and Intermediate Filaments

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3.3K Views.  Yale University. 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.

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 ...

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be employed using a conventional confocal microscope equipped with digital detectors. A protocol for the use of the technique in vitro is shown by means of a use case where number and brightness can be seen to accurately quantify the oligomeric state of mVenus-labelled FKBP12F36V before and after the addition of the dimerizing drug AP20187. The importance of using the correct microscope acquisition parameters and the correct data preprocessing and analysis methods are discussed. In particular, the importance of the choice of photobleaching correction is stressed. This inexpensive method can be employed to study protein-protein interactions in many biological contexts.

Calibration-free In Vitro Quantification of Protein Homo-oligomerization Using Commercial Instrumentation and Free, Open Source Brightness Analysis Software

This protocol describes a calibration-free approach for quantifying protein homo-oligomerization in vitro based on fluorescence fluctuation spectroscopy using commercial light scanning microscopy. The correct acquisition settings and analysis methods are shown.

Number and brightness is a calibration-free fluorescence fluctuation spectroscopy (FFS) technique for detecting protein homo-oligomerization. It can be ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

High-Throughput Total Internal Reflection Fluorescence and Direct Stochastic Optical Reconstruction Microscopy Using a Photonic Chip

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced contrast due to optical sectioning. The conventional approach is to use high numerical aperture microscope TIRF objectives for both excitation and collection, severely limiting the field of view and throughput. We present a novel approach to generating TIRF excitation for imaging with optical waveguides, called chip-based nanoscopy. The aim of this protocol is to demonstrate how chip-based imaging is performed in an already built setup. The main advantage of chip-based nanoscopy is that the excitation and collection pathways are decoupled. Imaging can then be done with a low magnification lens, resulting in large field of view TIRF images, at the price of a small reduction in resolution. Liver sinusoidal endothelial cells (LSECs) were imaged using direct stochastic optical reconstruction microscopy (dSTORM), showing a resolution comparable to traditional super-resolution microscopes. In addition, we demonstrate the high-throughput capabilities by imaging a 500 &#181;m x 500 &#181;m region with a low magnification lens, providing a resolution of 76 nm. Through its compact character, chip-based imaging can be retrofitted into most common microscopes and can be combined with other on-chip optical techniques, such as on-chip sensing, spectroscopy, optical trapping, etc. The technique is thus ideally suited for high throughput 2D super-resolution imaging, but also offers great opportunities for multi-modal analysis.

Chip-based super-resolution optical microscopy is a novel approach to fluorescence microscopy and offers advantages in cost effectiveness and throughput. Here, the protocols for chip preparation and imaging are shown for TIRF microscopy and localization-based super-resolution microscopy.

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced ...

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm phenotyping can be used to classify genotypes based on SG morphotype using scanning electron microscopic (SEM) analysis. SGs can be visualized using SEM by slicing through the kernel (pericarp, aleurone layers, and endosperm) and exposing the organellar contents. Current methods require the rice kernel to be embedded in plastic resin and sectioned using a microtome or embedded in a truncated pipette tip and sectioned by hand using a razor blade. The former method requires specialized equipment and is time-consuming, while the latter introduces a new host of problems depending on rice genotype. Chalky rice varieties, particularly, pose a problem for this type of sectioning due to the friable nature of their endosperm tissue. Presented here is a technique for preparing translucent and chalky rice kernel sections for microscopy, requiring only pipette tips and a scalpel blade. Preparing the sections within the confines of a pipette tip prevents rice kernel endosperm from shattering (for translucent or &#8216;vitreous&#8217; phenotypes) and crumbling (for chalky phenotypes). Using this technique, endosperm cell patterning and the structure of intact SGs can be observed.

Transverse Sectioning of Mature Rice Oryza sativa L. Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

This protocol allows for the preparation of transverse sections of cereal seeds (e.g., rice) for the analysis of endosperm and starch granule morphology using scanning electron microscopy.

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm ...

Imaging of Podocytic Proteins Nephrin, Actin, and Podocin with Expansion Microscopy

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte foot processes contain actin bundles that bind to cytoskeletal adaptor proteins such as podocin. Those adaptor proteins, such as podocin, link the backbone of the glomerular slit diaphragm, such as nephrin, to the actin cytoskeleton. Studying the localization and function of these and other podocytic proteins is essential for the understanding of the glomerular filter&#39;s role in health and disease. The presented protocol enables the user to visualize actin, podocin, and nephrin in cells with super resolution imaging on a conventional microscope. First, cells are stained with a conventional immunofluorescence technique. All proteins within the sample are then covalently anchored to a swellable hydrogel. Through digestion with proteinase K, structural proteins are cleaved allowing isotropical swelling of the gel in the last step. Dialysis of the sample in water results in a 4-4.5-fold expansion of the sample and the sample can be imaged via a conventional fluorescence microscope, rendering a potential resolution of 70 nm.

The presented method enables visualization of fluorescently labeled cellular proteins with expansion microscopy leading to a resolution of 70 nm on a conventional microscope.

Disruption of the glomerular filter composed of the glomerular endothelium, glomerular basement membrane and podocytes, results in albuminuria. Podocyte ...

F&#246;rster Resonance Energy Transfer Measurements in Living Plant Cells

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser scanning microscopes have become a workhorse for biologists. Commercial systems offer high flexibility in laser power adjustment and detector sensitivity and often combine different detectors to obtain the perfect image. However, the comparison of intensity-based data from different experiments and setups is often impossible due to this flexibility. Biologist-friendly procedures are of advantage and allow for simple and reliable adjustment of laser and detector settings. 
Furthermore, as FRET experiments in living cells are affected by the variability in protein expression and donor-acceptor ratios, protein expression levels must be considered for data evaluation. Described here is a simple protocol for reliable and reproducible FRET measurements, including routines for the estimation of protein expression and adjustment of laser intensity and detector settings. Data evaluation will be performed by calibration with a fluorophore fusion of known FRET efficiency. To improve simplicity, correction factors have been compared that have been obtained in cells and by measuring recombinant fluorescent proteins.

A protocol is provided for setting up a standard confocal laser-scanning microscope for in vivo F&#246;rster resonance energy transfer measurements, followed by data evaluation.

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser ...

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 ...

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...