This work was supported by a grant from the NIH (no. 1DP2GM126894-01), and by funds from the Pew Charitable Trusts and the Smith Family Foundation to R.S. The authors thank Dr. Shuo Jiang for his contribution toward development and optimization of the protocols.

<ol><li>Subramanian, R., Kapoor, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Building+complexity%3A+insights+into+self-organized+assembly+of+microtubule-based+architectures%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Building complexity: insights into self-organized assembly of microtubule-based architectures.</a> Developmental Cell. 23 (5), 874-885 (2012).</li>
<li>Baas, P. W., Rao, A. N., Matamoros, A. J., Leo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stability+properties+of+neuronal+microtubules%5BTitle%5D)+AND+%22Cytoskeleton+%28Hoboken%29%22%5BJournal%5D)">Stability properties of neuronal microtubules.</a> Cytoskeleton (Hoboken). 73 (9), 442-460 (2016).</li>
<li>Bitan, A., Rosenbaum, I., Abdu, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stable+and+dynamic+microtubules+coordinately+determine+and+maintain+Drosophila+bristle+shape%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Stable and dynamic microtubules coordinately determine and maintain Drosophila bristle shape.</a> Development. 139 (11), 1987-1996 (2012).</li>
<li>Foe, V. E., von Dassow, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stable+and+dynamic+microtubules+coordinately+shape+the+myosin+activation+zone+during+cytokinetic+furrow+formation%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Stable and dynamic microtubules coordinately shape the myosin activation zone during cytokinetic furrow formation.</a> The Journal of Cell Biology. 183 (3), 457-470 (2008).</li>
<li>Pous, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+specialization+of+stable+and+dynamic+microtubules+in+protein+traffic+in+WIF-B+cells%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Functional specialization of stable and dynamic microtubules in protein traffic in WIF-B cells.</a> The Journal of Cell Biology. 142 (1), 153-165 (1998).</li>
<li>Uehara, R., Goshima, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+central+spindle+assembly+requires+de+novo+microtubule+generation+in+the+interchromosomal+region+during+anaphase%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Functional central spindle assembly requires de novo microtubule generation in the interchromosomal region during anaphase.</a> The Journal of Cell Biology. 191 (2), 259-267 (2010).</li>
<li>Mitchison, T., Kirschner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dynamic+instability+of+microtubule+growth%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Dynamic instability of microtubule growth.</a> Nature. 312 (5991), 237-242 (1984).</li>
<li>Bieling, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Reconstitution+of+a+microtubule+plus-end+tracking+system+in+vitro%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Reconstitution of a microtubule plus-end tracking system in vitro.</a> Nature. 450 (7172), 1100-1105 (2007).</li>
<li>Bieling, P., Telley, I. A., Surrey, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+minimal+midzone+protein+module+controls+formation+and+length+of+antiparallel+microtubule+overlaps%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">A minimal midzone protein module controls formation and length of antiparallel microtubule overlaps.</a> Cell. 142 (3), 420-432 (2010).</li>
<li>Shimamoto, Y., Forth, S., Kapoor, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Measuring+pushing+and+braking+forces+generated+by+ensembles+of+Kinesin-5+crosslinking+two+microtubules%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Measuring pushing and braking forces generated by ensembles of Kinesin-5 crosslinking two microtubules.</a> Developmental Cell. 34 (6), 669-681 (2015).</li>
<li>Mani, N., Jiang, S., Neary, A. E., Wijeratne, S. S., Subramanian, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Differential+regulation+of+single+microtubules+and+bundles+by+a+three-protein+module%5BTitle%5D)+AND+%22Nature+Chemical+Biology%22%5BJournal%5D)">Differential regulation of single microtubules and bundles by a three-protein module.</a> Nature Chemical Biology. 17 (9), 964-974 (2021).</li>
<li>Hyman, A. A., Salser, S., Drechsel, D., Unwin, N., Mitchison, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Role+of+GTP+hydrolysis+in+microtubule+dynamics%3A+information+from+a+slowly+hydrolyzable+analogue%2C+GMPCPP%5BTitle%5D)+AND+%22Molecular+Biology+of+the+Cell%22%5BJournal%5D)">Role of GTP hydrolysis in microtubule dynamics: information from a slowly hydrolyzable analogue, GMPCPP.</a> Molecular Biology of the Cell. 3 (10), 1155-1167 (1992).</li>
<li>Subramanian, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Insights+into+antiparallel+microtubule+crosslinking+by+PRC1%2C+a+conserved+nonmotor+microtubule+binding+protein%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Insights into antiparallel microtubule crosslinking by PRC1, a conserved nonmotor microtubule binding protein.</a> Cell. 142 (3), 433-443 (2010).</li>
<li>Rasnik, I., McKinney, S. A., Ha, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Nonblinking+and+long-lasting+single-molecule+fluorescence+imaging%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Nonblinking and long-lasting single-molecule fluorescence imaging.</a> Nature Methods. 3 (11), 891-893 (2006).</li>
<li>Wijeratne, S., Subramanian, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Geometry+of+antiparallel+microtubule+bundles+regulates+relative+sliding+and+stalling+by+PRC1+and+Kif4A%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Geometry of antiparallel microtubule bundles regulates relative sliding and stalling by PRC1 and Kif4A.</a> eLife. 7, 32595(2018).</li>
<li>Mani, N., Wijeratne, S. S., Subramanian, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Micron-scale+geometrical+features+of+microtubules+as+regulators+of+microtubule+organization%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Micron-scale geometrical features of microtubules as regulators of microtubule organization.</a> eLife. 10, 63880(2021).</li>
<li>Freal, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Feedback-driven+assembly+of+the+axon+initial+segment%5BTitle%5D)+AND+%22Neuron%22%5BJournal%5D)">Feedback-driven assembly of the axon initial segment.</a> Neuron. 104 (2), 305-321 (2019).</li>
<li>Ledbetter, M., Porter, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+%26%2334%3Bmicrotubule%26%2334%3B+in+plant+cell+fine+structure%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">A &#34;microtubule&#34; in plant cell fine structure.</a> The Journal of Cell Biology. 19 (1), 239-250 (1963).</li>
<li>Wijeratne, S. S., Marchan, M. F., Tresback, J. S., Subramanian, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Atomic+force+microscopy+reveals+distinct+protofilament-scale+structural+dynamics+in+depolymerizing+microtubule+arrays%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Atomic force microscopy reveals distinct protofilament-scale structural dynamics in depolymerizing microtubule arrays.</a> Proceedings of the National Academy of Sciences of the United States of America. , 119(2022).</li>
</ol>

The authors declare no competing interests.

The experiment described here significantly expands the scope and complexity of conventional microtubule reconstitution assays, which are traditionally performed on single microtubules or on one type of array. The current assay provides a method to simultaneously quantify and compare the regulatory MAP activity on two populations, namely, single microtubules and crosslinked bundles. Further, this assay allows for the examination of two types of bundles: those that are pre-formed from stable seeds before the initiation of...

Microtubules are biopolymers that form structural scaffolds essential for multiple cellular processes, ranging from intracellular transport and organelle positioning to cell division and elongation. To execute these diverse functions, individual microtubules are organized into micron-sized arrays, such as mitotic spindles, ciliary axonemes, neuronal bundles, interphase arrays, and plant cortical arrays. A ubiquitous architectural motif found in these structures is a bundle of microtubules crosslinked along their lengths1. An intriguing feature of several microtubule-based structures is the coexistence of bundled microtubules and non-crosslinked single microtubules in close spatial proximity. These microtubule subpopulations can display starkly different polymerization dynamics from each other, as needed for their proper function2,3,4,5. For instance, within the mitotic spindle, stable crosslinked bundles and dynamic single microtubules are present within a micron-scale region at the cell center6. Studying how the dynamic properties of coexisting microtubule populations are specified is, therefore, central to understanding the assembly and function of microtubule-based structures.
Microtubules are dynamic polymers that cycle between phases of polymerization and depolymerization, switching between the two phases in events known as catastrophe and rescue7. The dynamics of cellular microtubules are regulated by myriad Microtubule Associated Proteins (MAPs) that modulate the rates of microtubule polymerization and depolymerization and the frequencies of catastrophe and rescue events. It is challenging to investigate the activity of MAPs on spatially proximal arrays in cells, owing to the limitations of spatial resolution in light microscopy, especially in regions of high microtubule density. Moreover, the presence of multiple MAPs in the same cellular region hinders interpretations of cell biological studies. In vitro reconstitution assays, performed in conjunction with Total Internal Reflection Fluorescence (TIRF) microscopy, circumvent the challenges of examining mechanisms by which specific subsets of MAPs regulate the dynamics of proximal cellular microtubule arrays. Here, the dynamics of microtubules assembled in vitro are examined in the presence of one or more recombinant MAPs under controlled conditions8,9,10. However, conventional reconstitution assays are typically performed on single microtubules or on one type of array, precluding the visualization of coexisting populations.
Here, we present&#160;in vitro reconstitution assays that enable the simultaneous visualization of two microtubule populations under the same solution conditions11. We describe a&#160;method to simultaneously view the collective activity of multiple MAPs on single microtubules&#160;and on microtubule bundles crosslinked by the mitotic spindle-associated protein PRC1.&#160;The protein PRC1 preferentially binds at the overlap between anti-parallel microtubules, crosslinking them9. Briefly, this protocol consists of the following steps: (i) preparation of stock solutions and reagents, (ii) cleaning and surface treatment of coverslips used to create the imaging chamber for microscopy experiments, (iii) preparation of stable microtubule &#34;seeds&#34; from which polymerization is initiated during the experiment, (iv) specification of TIRF microscope settings to visualize microtubule dynamics, (v) immobilization of microtubule seeds and generation of crosslinked microtubule bundles in the imaging chamber, and (vi) visualization of microtubule dynamics in the imaging chamber through TIRF microscopy, upon addition of soluble tubulin, MAPs, and nucleotides. These assays enable the qualitative evaluation and quantitative examination of MAP localization and their effect on the dynamics of two microtubule populations. Additionally, they facilitate the evaluation of synergistic effects of multiple MAPs on these microtubule populations, across a wide range of experimental conditions.

<table><tbody><tr><td>(&amp;plusmn;)-6-Hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox)</td><td>Sigma Aldrich</td><td>238813</td><td></td></tr><tr><td>1,4-piperazinediethanesulfonic acid (PIPES)</td><td>Sigma Aldrich</td><td>P6757</td><td></td></tr><tr><td>18x18 mm #1.5 coverslips&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>63787</td><td></td></tr><tr><td>2-Mercaptoethanol (BME)</td><td>Sigma Aldrich</td><td>M-6250</td><td></td></tr><tr><td>24x60 mm #1.5 coverslips</td><td>Electron Microscopy Sciences</td><td>63793</td><td></td></tr><tr><td>405/488/560/647 nm Laser Quad Band&amp;nbsp;</td><td>Chroma</td><td>TRF89901-NK</td><td></td></tr><tr><td>Acetone</td><td>Sigma Aldrich</td><td>320110</td><td></td></tr><tr><td>Adenosine 5&#039;-triphosphate disodium salt hydrate (ATP)</td><td>Sigma Aldrich</td><td>A7699-5G</td><td></td></tr><tr><td>Avidin, NeutrAvidin&amp;reg; Biotin-binding Protein (Molecular Probes&amp;reg;)</td><td>Thermo Fischer Scientific</td><td>A2666</td><td></td></tr><tr><td>Bath sonicator: Branson 2800 Cleaner</td><td>Branson</td><td>CPX2800H</td><td></td></tr><tr><td>Beckman Coulter Polycarbonate Thickwall Tubes, 11 x 34 mm</td><td>Beckman-Coulter&amp;nbsp;</td><td>343778</td><td></td></tr><tr><td>Beckman Coulter Polycarbonate Thickwall Tubes, 8 x 34 mm</td><td>Beckman-Coulter&amp;nbsp;</td><td>343776</td><td></td></tr><tr><td>Biotin-PEG-SVA, MW 5,000</td><td>Laysan Bio</td><td>#Biotin-PEG-SVA-5000</td><td></td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma Aldrich</td><td>2905</td><td></td></tr><tr><td>Catalase</td><td>Sigma Aldrich</td><td>C40</td><td></td></tr><tr><td>Corning LSE Mini Microcentrifuge, AC100-240V</td><td>Corning</td><td>6670</td><td></td></tr><tr><td>Delicate Task Wipes</td><td>Kimtech</td><td>34120</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>GoldBio</td><td>DTT10</td><td></td></tr><tr><td>Emission filter</td><td>Chroma</td><td>ET610/75m</td><td></td></tr><tr><td>Ethanol (200-proof)</td><td>Decon Labs</td><td>2705</td><td></td></tr><tr><td>Ethylene glycol tetraacetic acid (EGTA)</td><td>Sigma Aldrich</td><td>3777</td><td></td></tr><tr><td>Glucose Oxidase</td><td>Sigma Aldrich</td><td>G2133</td><td></td></tr><tr><td>GMPCPP</td><td>Jena Bioscience&amp;nbsp;</td><td>NU-405</td><td></td></tr><tr><td>Guanosine 5&#039;-triphosphate sodium salt hydrate (GTP)</td><td>Sigma Aldrich</td><td>G8877</td><td></td></tr><tr><td>Hellmanex III detergent&amp;nbsp;</td><td>Sigma Aldrich</td><td>Z805939</td><td></td></tr><tr><td>Immersion oil, Type A</td><td>Fisher Scientific</td><td>77010</td><td></td></tr><tr><td>Kappa-casein</td><td>Sigma Aldrich</td><td>C0406</td><td></td></tr><tr><td>Lanolin</td><td>Fisher Scientific</td><td>S25376</td><td></td></tr><tr><td>Lens Cleaning Tissue</td><td>ThorLabs</td><td>MC-5</td><td></td></tr><tr><td>Magnesium Chloride (MgCl&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>M9272</td><td></td></tr><tr><td>Methylcellulose</td><td>Sigma Aldrich</td><td>M0512</td><td></td></tr><tr><td>Microfuge 16 Benchtop Centrifuge</td><td>Beckman-Coulter&amp;nbsp;</td><td>A46474</td><td></td></tr><tr><td>Microscope Slides, Diamond White Glass, 25 x 75mm, 90&amp;deg; Ground Edges, WHITE Frosted</td><td>Globe Scientific</td><td>1380-50W</td><td></td></tr><tr><td>mPEG-Succinimidyl Valerate, MW 5,000&amp;nbsp;</td><td>Laysan Bio</td><td>#NH2-PEG-VA-5K</td><td></td></tr><tr><td>Optima&amp;trade; Max-XP Tabletop Ultracentrifuge</td><td>Beckman-Coulter&amp;nbsp;</td><td>393315</td><td></td></tr><tr><td>Paraffin</td><td>Fisher Scientific</td><td>P31-500</td><td></td></tr><tr><td>PELCO Reverse (self-closing), Fine Tweezers</td><td>Ted Pella</td><td>5377-NM</td><td></td></tr><tr><td>Petrolatum, White</td><td>Fisher Scientific</td><td>18-605-050</td><td></td></tr><tr><td>Plasma Cleaner, 115V</td><td>Harrick Plasma</td><td>PDC-001</td><td></td></tr><tr><td>Potassium Hydroxide (KOH)</td><td>Sigma Aldrich</td><td>221473</td><td></td></tr><tr><td>Sodium bicarbonate</td><td>Sigma Aldrich</td><td>S6014</td><td></td></tr><tr><td>Sucrose</td><td>Sigma Aldrich</td><td>S7903</td><td></td></tr><tr><td>Thermal-Lok 1-Position Dry Heat Bath</td><td>USA Scientific</td><td>2510-1101</td><td></td></tr><tr><td>Thermal-Lok Block for 1.5 and 2.0 mL Tubes</td><td>USA Scientific</td><td>2520-0000</td><td></td></tr><tr><td>Thermo Scientific&amp;trade; Pierce&amp;trade; Bond-Breaker&amp;trade; TCEP Solution, Neutral pH; 500mM</td><td>Thermo Fischer Scientific</td><td>PI-77720</td><td></td></tr><tr><td>TIRF 100X NA 1.49 Oil Objective</td><td>Nikon</td><td>CFI Apochromat TIRF 100XC Oil</td><td></td></tr><tr><td>TIRF microscope</td><td>Nikon</td><td>Eclipse Ti</td><td></td></tr><tr><td>TLA 120.1 rotor</td><td>Beckman-Coulter&amp;nbsp;</td><td>362224</td><td></td></tr><tr><td>TLA 120.2 rotor</td><td>Beckman-Coulter&amp;nbsp;</td><td>357656</td><td></td></tr><tr><td>Tubulin protein (&gt;99% pure): porcine brain</td><td>Cytoskeleton</td><td>T240</td><td></td></tr><tr><td>Tubulin Protein (Biotin): Porcine Brain</td><td>Cytoskeleton</td><td>T333P</td><td></td></tr><tr><td>Tubulin protein (fluorescent HiLyte 647): porcine brain</td><td>Cytoskeleton</td><td>TL670M</td><td></td></tr><tr><td>Tubulin protein (X-rhodamine): bovine brain</td><td>Cytoskeleton</td><td>TL620M</td><td></td></tr><tr><td>VECTABOND&amp;reg; Reagent, Tissue Section Adhesion</td><td>Vector Biolabs</td><td>SP-1800-7</td><td></td></tr><tr><td>VWR&amp;reg; Personal-Sized Incubator, 120V, 50/60Hz, 0.6A</td><td>VWR</td><td>97025-630</td><td></td></tr></tbody></table>

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.

The experiment described above was performed using 647 nm fluorophore-labeled biotinylated microtubules, 560 nm fluorophore-labeled non-biotinylated microtubules, and 560 nm fluorophore-labeled soluble tubulin mix. Microtubules were crosslinked by the crosslinker protein PRC1 (GFP-labeled). After surface-immobilized bundles and single microtubules were generated (step 5.11), the imaging chamber was mounted on a TIRF 100X 1.49 NA oil objective and viewed in the 560 nm and 647 nm fluorescence channels. Single microtubules ...

Watch this Scientific Journal Video about Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy at JoVE.com

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.

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 ...

simultaneous-visualization-dynamics-crosslinked-single-microtubules

Research

JoVE Journal

Biochemistry

2.5K Views.  Massachusetts General Hospital. 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.

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

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

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

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 ...

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

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

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

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

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

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

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

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

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

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 ...

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

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

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

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

Biology

TIRF Microscopy to Visualize Actin and Microtubule Coupling Dynamics

Cut 12 strips of double-backed, double-sided tape, to a length of 24 millimeters. Remove one side of the tape backing, and fix pieces of tape adjacent to the six grooves present on a clean imaging chamber. Remove the second piece of tape backing, to expose the sticky side of the tape along each chamber groove, and place the chamber, tape side up, on a clean surface.
Mix epoxy resin and hardener solutions, one to one in a small weigh boat, and use a P1000 tip to place a drop of mixed epoxy between the tape strips at the end of each imaging chamber groove. Then, place chamber tape, or epoxy side up, on a clean surface.
Remove a coated coverslip from 70 degrees Celsius incubator, and rinse the coated and uncoated surfaces of the coverslips with double-distilled water six times. Dry with filtered nitrogen gas, and then affix to the imaging chamber, with the coverslip coating side toward the tape.
Use a P200 or a P1000 pipette tip to apply pressure on the tape-glass interface, to ensure a good seal between the tape and the coverslip. Incubate the assembled chambers at room temperature for at least five to 10 minutes, to allow the epoxy to fully seal the chamber walls before use.
Perfusion chambers expire within 12 to 18 hours of assembly. Use a perfusion pump, and sequentially exchange conditioning solutions in the perfusion chamber by flowing 50 microliters of 1% BSA to prime the imaging chamber.
Remove the excess buffer from the Luer-lock fitting reservoir. Then, flow 50 microliters of 0.005 milligrams per milliliter streptavidin, and incubate for one to two minutes at room temperature. Remove the excess buffer from the reservoir.
Flow 50 microliters of 1% BSA to block the nonspecific binding, and incubate for 10 to 30 seconds. Remove the excess buffer from the reservoir. Next, flow 50 microliters of warm 1x TIRF buffer, and then 50 microliters of stabilized, and 50% biotinylated microtubule seeds, diluted in 1x TIRF buffer.
Set the stage or objective heater device to maintain 35 to 37 degrees Celsius temperature for 30 minutes, prior to imaging the first biochemical reaction. Then, set the acquisition interval to every five seconds, for 15 to 20 minutes. Next, set 488 and 647-laser exposures to 50 to 100 milliseconds at 5% to 10% power, by first adjusting the polymerization reaction to initiate the actin filament assembly.
Acquire the images at 647 nanometers, and make appropriate adjustments. Then, adjust the polymerization reaction in a second conditioned perfusion well, to initiate the microtubule assembly. Visualize at 488 nanometers, and make appropriate adjustments.
Combine three microliters of 10-micromolar 488-tubulin with the 7.2 microliter aliquot of 100 micromolar fluorescently unlabeled tubulin, no more than 15 minutes before use. Then, combine three microliters of diluted biotinylated actin in appropriate volumes of fluorescently unlabeled and labeled actin, such that the final mix will be 12.5 micromolar total actin, with 10% to 30% fluorescent label.
Prepare the cytoskeleton mix by combining two microliters of the 12.5-micromolar actin mix stock with the tubulin stock mix, no more than 15 minutes prior to imaging. Store on ice until use.
Prepare the protein reaction mix by combining all other experimental components and proteins, including 2x TIRF buffer, anti-bleach, nucleotides, buffers, and accessory proteins. Incubate the cytoskeleton mix in the protein reaction mix separately, at 37 degrees Celsius for 30 to 60 seconds.
To initiate the reaction, add the contents of the protein reaction mix to the cytoskeleton mix, and mix it. Flow 50 microliters of the reaction mix containing 1x TIRF buffer supplemented with 15 micromolar free tubulin, one millimolar GTP, 0.5 micromolar actin monomers, and appropriate volumes of buffer controls to the perfusion chamber.
Record a time-lapse movie using microscope software, to acquire every five seconds, for 15 to 20 minutes. Condition a new perfusion well, and replace the buffer volume with the regulatory protein of interest and buffer controls. Acquire to assess the regulatory proteins for emergent actin microtubule functions.

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

Summary

Explore More Videos

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

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

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

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

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

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

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

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

TIRF Microscopy to Visualize Actin and Microtubule Coupling Dynamics