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JoVE Journal Biology

Biology

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Published: May 10, 2022 doi: 10.3791/63819
Automatic Translation
*1, *1, 1
1Department of Biological Sciences and Center for Biotechnology and Interdisciplinary Studies, Rensselaer Polytechnic Institute
* These authors contributed equally

Summary

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.

Abstract

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.

Introduction

Cells employ microtubule networks to perform a wide variety of mechanical tasks, ranging from vesicle transport1,2,3 to chromosome segregation during mitosis4,5,6. Many of the proteins that interact with microtubules, such as the molecular motor proteins kinesin and dynein, generate forces and are regulated by mechanical loads. To better understand how these critical molecules function, researchers have employed single-molecule biophysical methods, such as optical trapping and TIRF microscopy, to directly monitor critical parameters such as unloaded stepping rates, processivity, and force-velocity relationships for individual proteins. The most commonly used experimental geometry has been to attach motor proteins directly to trapping beads whose spherical geometry and size mimic vesicles undergoing motor-driven transport. Numerous kinesins, including kinesin-17,8,9, kinesin-210,11,12, kinesin-313,14,15,16 kinesin-517,18, kinesin-819,20, as well as dynein and dynein complexes21,22,23,24,25, have been studied with these methods.

In many cellular processes, however, motor and non-motor proteins use microtubules both as track and cargo26,27. Moreover, in these scenarios where microtubule filaments are crosslinked into higher-order bundles, these proteins function as ensembles rather than single units. For example, within dividing somatic cells, dense filament networks self-organize to build the mitotic spindle apparatus28,29,30. The interpolar spindle microtubule network is highly dynamic and is largely arranged with minus-ends pointing toward the spindle poles and plus-ends overlapping near the spindle equator. Filaments within the spindle are crosslinked by motor proteins such as kinesin-531,32,33, kinesin-1234,35,36, and kinesin-1437,38,39, or by non-motor proteins such as PRC140,41,42,43 or NuMA44,45,46. They frequently move or experience mechanical stress during processes such as poleward flux or while coordinating chromosome centering during metaphase or chromosome segregation during anaphase47,48,49,50,51,52. The integrity of the micron-scale spindle apparatus through mitosis, therefore, relies on a carefully regulated balance of pushing and pulling forces generated and sustained by this network of interacting filaments. However, the tools needed to probe this mechanical regulation and explain how protein ensembles work in concert to coordinate microtubule motions and produce the forces needed to properly assemble the spindle have only recently been developed, and we are just beginning to understand the biophysical rules that define dynamic microtubule networks.

The goal of this manuscript is to demonstrate the steps required to reconstitute crosslinked microtubule pairs in vitro, immobilize these bundles in a microscopy chamber that allows for simultaneous fluorescence visualization of both the microtubules and crosslinking proteins and nanoscale force measurement, and process these data robustly. We detail the steps needed to stably polymerize fluorescence-labeled microtubules, prepare microscope coverslips for attachment, prepare polystyrene beads for optical trapping experiments, and assemble crosslinked filament networks that preserve their in vivo functionality while allowing for direct biophysical manipulation.

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Protocol

1. Preparation of microtubules

NOTE: When employing GFP-labeled crosslinking proteins, red (e.g., rhodamine) and far-red (e.g., biotinylated HiLyte647, referred to as biotinylated far red in the rest of the text) organic fluorophore labeling of the microtubules works well. Minimal crosstalk between all three channels can be achieved during imaging by using a high-quality quad band total internal reflection fluorescence (TIRF) filter.

  1. Prepare GMPCPP microtubule seed stocks
    1. Suspend 1 mg of unlabeled lyophilized tubulin in 84 µL of ice-cold 1x BRB80 (80 mM PIPES, 1 mM MgCl2, 1 mM EGTA; pH 6.8), with DTT added to a final concentration of 1 mM just prior to use. Suspend 60 µg of fluorophore-labeled lyophilized tubulin in 6 µL of cold 1x BRB80 with 1 mM DTT. Suspend 60 µg of biotin-labeled lyophilized tubulin in 6 µL of cold 1x BRB80 with 1 mM DTT.
    2. To prepare non-biotinylated rhodamine-labeled tubulin seed stock with a labeling ratio of 1:10, add 84 µL of unlabeled tubulin, 6 µL of fluorophore-labeled tubulin, and 10 µL of 10 mM GMPCPP stock solution to a 0.5 mL tube. Mix well by gently pipetting and keep on ice.
      NOTE: GMPCPP polymerization is preferred for these assays as the microtubules are more stable and amenable to direct manipulation with the optical trap than if polymerized with GTP or if taxol is omitted.
    3. To prepare biotinylated far red-labeled tubulin seed stock with a fluorescent labeling ratio of 1:10, add 80 µL of unlabeled tubulin, 6 µL of fluorophore-labeled tubulin, 4 µL of biotin-labeled tubulin, and 10 µL of 10 mM GMPCPP stock solution in a 0.5 mL tube. Mix well by gently pipetting and keep on ice. Incubate the seed stock on ice for 5 min.
    4. Transfer the tubulin solution to a polycarbonate tube appropriate for high-speed centrifugation. Centrifuge the seed stock at ~350,000 x g for 5 min at 2 °C to prevent microtubule polymerization and recover the supernatant for aliquoting. The final estimated concentration of tubulin at this stage is ~50 µM.
    5. Using 0.2 mL PCR tubes, prepare 2 µL aliquots of the seed stock and flash freeze with liquid nitrogen. Aliquots can be stored at -80 °C for up to 6 months.
  2. Polymerization of microtubules
    1. Prepare 500 µL of 1x BRB80 and place on ice. Place one each of the biotinylated far red and non-biotinylated rhodamine seed stock aliquots on ice to thaw.
    2. Incubate the seed stocks on ice for 5 min. At the end of the incubation period, immediately add 26 µL of 1x BRB80 to each tube. If longer microtubules are desired, increase the volume of BRB80 by 5-10 µL, whereas, for shorter microtubules, reduce the volume for polymerization by 5 µL.
    3. Incubate the microtubule seed stocks for 1 h at 37 °C in a water bath. Combine 298.5 µL of room temperature 1x BRB80 and 1.5 µL of 4 mM taxol stock solution to produce a 20 µM taxol solution. Prepare 300 µL of 1x BRB80 and keep both tubes at room temperature.
    4. Warm an appropriate high-speed rotor to 30 °C. This can be done in a tabletop ultracentrifuge set to 30 °C. Remove the microtubules from the water bath and keep at room temperature on a benchtop for 10-15 min.
  3. Clarification of microtubules
    1. Add 100 µL of room temperature 1x BRB80 to the microtubule growth solution and mix by flicking the tube ~10 times or gently pipetting. Transfer the microtubule growth solution into a polycarbonate centrifuge tube.
    2. Mark one side of the polycarbonate tube, which will face outward relative to the rotation of the centrifuge. This will assist in locating the microtubule pellet, which will be barely visible due to the small volume of material.
    3. Centrifuge the microtubule solution at ~350,000 x g for 10 min at 30 °C. After centrifugation, inspect the tube containing the microtubules for a pellet if possible. Remove all but the final few microliters of liquid carefully, without disturbing the pellet. If the pellet is not clearly visible, use the marking made as a guide to avoid disturbing the area where the pellet is located.
    4. Immediately add 100 µL of 1x BRB80 + 20 µM taxol to the microtubule tube. This can be added directly onto the pellet; disruption of the pellet at this stage does not significantly impact the clarification process.
    5. Centrifuge the microtubules again at ~350,000 x g for 10 min, being careful to ensure the marked section of the tube is again oriented outward relative to the rotation of the centrifuge.
    6. Remove all but a few microliters of solution as before, again being careful to avoid disrupting the microtubule pellet. Repeat Steps 1.3.4.-1.3.5. and then resuspend the microtubule pellet in 20-50 µL of 1x BRB80 + 20 µM taxol.
    7. Resuspend by pipetting up most of the resuspension volume and gently ejecting it directly onto the pellet, repeating 20-30x or until the pellet is fully resuspended. This should be done with care so as not to shear the microtubules by overly vigorous pipetting. Cutting the pipette tip to increase the size of the opening can also be helpful to reduce microtubule shearing.
    8. Store the microtubules at room temperature by wrapping the tube in foil to shield the fluorophores from light and keeping on the benchtop. Microtubules are generally usable for 2-3 days after clarification.
  4. Assessment of microtubules through microscopy
    1. Prepare a 1:10 dilution of the clarified microtubule solution by mixing 1 µL of microtubules and 9 µL of room temperature 1x BRB80. Immediately image this solution by pipetting 10 µL onto a microscope slide and then placing a coverslip over the droplet to form a squash.
    2. Ensure that microtubules ranging in lengths from 8-25 µm at a high density (several hundred per full field of view layered in a dense meshwork) are visible when visualized using fluorescence microscopy and a 100x objective focused on the coverslip surface in contact with the microtubule dilution.

2. Preparation of passivated coverslips

  1. Washing coverslips
    1. Place 18 mm x 18 mm coverslips in non-reactive plastic racks, then place each rack into a 100 mL beaker. Place one coverslip per slot in the racks, as cleaning both sides of the coverslips is necessary.
    2. Sonicate using a bath sonicator for 5 min, using 50 mL of the following solutions in the order given for each sonication cycle: (1) deionized water (with 18.3 MΩ-cm resistivity), (2) 2% Micro-90 solution, (3) deionized water, (4) freshly prepared 0.1 M KOH, (5) deionized water (Figure 1A). Make sure that the coverslips are fully covered in liquid. Between sonication cycles, rinse each rack of coverslips in deionized water by using tweezers to dip the rack up and down 10-20x in deionized water, replace the water, and repeat the rinse process 2x.
    3. Rinse the coverslips 3x in water, then refill the beakers with 100% ethanol and return the racks to the beakers. Move the beakers into a fume hood and lay out enough tissue wipes to make room for all the coverslip racks. Then, remove the racks with tweezers and place on the tissue wipes.
    4. Using a dry nitrogen gas stream, blow off the ethanol from the coverslips and their racks until dry. Dispose of the remaining ethanol from the beakers and similarly dry them using the dry nitrogen gas stream (Figure 1A).
  2. Aminosilane surface functionalization on coverslips
    1. Prepare (3-Aminopropyl)tiethoxysilane (APTES) reagent solution by adding 40 mL of acetone and 400 µL of APTES reagent to a 50 mL conical tube, capping tightly, and mixing by inverting 10x.
    2. Move one rack of coverslips into its corresponding dry beaker, and then fully submerge in APTES solution. Incubate for 5 min (Figure 1A). Move the APTES-treated rack to a new dry beaker, and move a new rack into the APTES to incubate for 5 m.
    3. Rinse the treated rack with deionized water by dipping the rack 10-20x as described in Step 2.1.2., replacing the water 5x. Repeat until all the coverslips have been treated and washed.
  3. PEGylation of aminosilane surface
    1. Prepare two PEG solutions, one of ~100 µL with 25% w/v PEG and another of ~10 µL with 10% w/v biotin-PEG, both suspended in freshly prepared 0.1 M NaHCO3, sufficient to coat 24 coverslips. Centrifuge the 25% PEG solution at 3,000 x g for 20 s once 0.1 M NaHCO3 is added to fully dissolve the PEG; the solution may remain cloudy. Mix the biotin-PEG by gentle pipetting (Figure 1B).
    2. Prepare a mixture of the PEG and biotin-PEG solutions, with 33.3 times volume of PEG to 1 volume of biotin-PEG (e.g., 100 µL of PEG solution and 3 µL of biotin-PEG solution).
    3. In a fume hood, lay out several sterile and lint-free wipes. Move the coverslip racks onto the wipes and blow dry with a dry nitrogen gas stream as before. On several new and dry wipes, lay out the now fully dried coverslips arranged in pairs and label each at the bottom right corner with an asymmetric symbol such as "b". This marking will be opposite the sample surface of the coverslip and will not interfere with the experiment (Figure 1B).
    4. Flip one coverslip in each pair with tweezers. Onto each flipped coverslip, place 6 µL of PEG/biotin-PEG mixture and place the second coverslip on top so that both "b" labels face outward.
    5. Align the coverslip edges, and place into a humidified and airtight chamber to prevent drying. Incubate the coverslips in the humidity chamber at room temperature for 3 h.
    6. After incubation, remove the coverslips from the humidity chamber, carefully separate the coverslip pairs, and place them back into their racks. Wash each rack in deionized water as before, dipping the racks using tweezers 10-20x and replacing the water a total of 5x.
    7. Place all PEGylated sides of the coverslips facing the same direction so that no two PEGylated coverslip surfaces face each other. The PEGylated surface will be very sticky until it fully dries, so take extra care to prevent the coverslips from touching. Dry each rack and coverslips as before using a dry nitrogen gas stream (Figure 1B).
    8. Once the coverslips and their racks are fully dried, store in a vacuum desiccator to prevent degradation of the surface coating. Properly stored under vacuum and with desiccant to prevent moisture from disrupting the PEG coating; coverslips can be used for approximately 1 week.

3. Preparation of kinesin-coated beads

  1. Selection and preparation of rigor kinesin construct
    1. Express and purify rigor kinesin constructs according to published protocols53,54. Flash freeze 5 µL aliquots of 0.02 mg/mL kinesin solutions and store at -80 °C for up to 1 year.
      NOTE: Rigor kinesin proteins have a G234A point mutation that prevents ATP hydrolysis. When conjugated to microbeads, high affinity interactions with the microtubule allow beads to sustain loads of 10-20 pN for several minutes. Rigor mutated versions of both the commonly employed kinesin-1 homodimer K56053 and a further optimized K439 construct55 have also been employed with success in these assays54. This improved K439 construct contains an EB1 dimerization domain to enhance stability and a C-terminal 6-His tag for specific binding to a 6-His antibody, as described below.
  2. Binding 6-His antibody to streptavidin-coated beads
    1. Add 50 µL of 1 µm diameter streptavidin-coated polystyrene beads into a 0.5 mL centrifuge tube. Sonicate for 30 s.
    2. Add 10 µL of 200 mM DTT to 790 µL of 1x BRB80. Add 10 µL of sonicated beads to 40 µL of this 1x BRB80 + 2.5 mM DTT buffer.
    3. Combine 1 µL of biotinylated 6-His antibody and 49 µL of 1x BRB80 + 2.5 mM DTT; vortex briefly to mix.
    4. Combine the 50 µL of diluted bead mixture and 50 µL of antibody dilution in a new tube. Place the tube in a tube rotator at 4 °C and rotate for 1 h. Spin down the bead mixture for 10 min at 3,000 x g at 4 °C.
    5. Pipette off the supernatant and resuspend the pellet in 100 µL of 2 mg/mL casein solution in 1x BRB80. Repeat this centrifugation and resuspension 2x.
  3. Addition of rigor kinesin
    1. Resuspend the final pellet in 100 µL of 2 mg/mL casein solution. In a 0.5 mL centrifuge tube, combine 5 µL of 0.02 mg/mL rigor kinesin and 5 µL of beads, mixing gently by lightly flicking the tube ~10x.
    2. Keep both the K439 G234A bead suspension and the remaining 6-His beads on the rotor at 4 °C when not in use. Beads must be prepared fresh and used the same day for the experiments as they tend to clump and lose activity beyond this time.

4. Assembly of the microscopy chamber

  1. Preparation of glass slides and chamber construction
    1. Rinse 75 mm x 25 mm microscope glass slides first with ultrapure water, followed by non-streak ammonia-d glass cleaner, and then once more with ultrapure water. Shake to remove excess water droplets. Dry the slides using a nitrogen stream so that no streaks remain and lay the slides down on a paper towel (Figure 2A).
    2. With scissors, cut double-sided tape into strips of ~2-3 mm x 2.5 cm (2 strips per single chamber and 3 per double chamber).
    3. Using forceps, lay two strips of tape onto one glass slide so that there is ~0.5 cm of space in between them to construct a single chamber. For double chambers, use the same spacing, but with three strips of tape (Figure 2B). The volume of the sample channel using this method is approximately 10 µL.
      NOTE: The wider the chamber(s), the more likely it is that bubbles will form when solutions are flowed in; however, chambers that are less than 0.5 cm wide can be challenging to use due to the reduced area for imaging.
    4. Scrape across the length of each strip of tape using forceps so that the tape is firmly adhered to the glass slide.
    5. Release the pressure from the desiccator and use forceps to remove one biotinylated coverslip from a storage rack. Place one biotinylated coverslip on top of the glass slide using forceps. Make sure that the biotinylated side is facing inwards, toward the glass slide.
    6. Using the flat back end of the forceps, gently press on the glass where the coverslip touches the tape to securely seal the chambers. Light pressure must be used to ensure that the coverslip does not crack (Figure 2C).
    7. Store the finished microscopy chambers in an airtight container, away from direct light, until use (Figure 2D).
      NOTE: It is recommended to use chambers on the same day that they are made, as the biotinylated coverslips degrade when exposed to air.
  2. Flow-in of microscopy reagents
    1. Construct a humidity chamber by placing a folded-up lint-free tissue dampened with ultrapure water at the bottom of an empty pipette tip box. The sample chambers should be kept in this box between flow-in steps to reduce evaporation of solutions from the channel.
    2. Introduce all the reagents by pipetting the liquid at one end of the channel and wicking liquid out at the opposite end. To wick, twist one end of the tissue and gently place it on the exit side of the chamber, allowing for capillary action to homogeneously distribute the reagent (Figure 3A). Bring all reagents to room temperature just before flowing in to prevent microtubule depolymerization.
    3. Using a 20 µL angled pipette tip so that it is touching the entry side of one chamber, slowly flow in 10 µL of 0.5 mg/mL streptavidin or Neutravidin. Use smaller pipette tips and a slow, constant flow-in of reagents to decrease the likelihood of obtaining bubbles in the chamber (Figure 3B).
    4. Incubate the slide in the humidity chamber for 4 min with the coverslip side of the chamber facing down. This orientation will allow for larger objects, such as microtubules or beads, to sink near the PEGylated surface.
    5. Flush out the chamber using 20 µL of 1x BRB80, using a lint-free tissue to draw out liquids. If there is a leak present in the chamber, discard the slide and start with another one; small bubbles that occur in the chamber will not detrimentally impact imaging.
    6. Repeat the flow-in, incubation, and flush steps with 10 µL of 0.5 mg/mL casein blocking solution (Figure 3C). Dilute biotinylated far-red microtubules ~1:20 and flow 10 µL into the chamber. Incubate for 5 min and flush the chamber with 20 µL of 1x BRB80 (Figure 3D). The optimal density of these microtubules within a 100 µm x 100 µm imaging field of view should be 10-20; and the dilution at this stage should be adjusted to achieve this surface-binding ratio.
    7. Repeat the flow-in, incubation, and flush steps with 10 µL of an appropriate concentration (typically 1-10 nM) crosslinking motor or non-motor protein to be studied (Figure 3E).
    8. Flow in 10 µL of rhodamine microtubules diluted to a similar concentration as the previous biotinylated far-red microtubules. Repeat the incubation and flush steps (Figure 3F).
    9. Combine 1 µL of rigor kinesin-coated beads and 19 µL of reaction buffer optimized for crosslinking protein activity. For example, for analysis of kinesin-5 activity, use reaction buffer with 1 mM TCEP bond breaker, 0.2 mg/mL alpha casein, 70 mM KCl, oxygen scavenging system (4.5 mg/mL glucose, 350 units/mL glucose oxidase, 34 units/mL catalase), 1 mM DTT, and ATP at the desired concentration (e.g., 1 mM for maximum kinesin velocity conditions) in 1x BRB80 (Figure 3G). For analysis of non-motor proteins, omit ATP and vary the concentration of KCl to modulate protein-microtubule interaction strength in these assays.
    10. Seal both edges of the chamber with clear nail polish and wait until the polish dries prior to imaging (Figure 3H). A successfully constructed chamber will have no leakage present and minimal bubbles.

5. Imaging microtubule bundles with 3-color TIRF

  1. Employ an inverted microscope instrument that has TIRF imaging capabilities, and into which a single-beam optical trap has been constructed (Figure 4).
    NOTE: For the studies described here, an inverted microscope was used with a 100x/NA1.49 oil objective lens, TIRF module, and three lasers at 488 nm, 561 nm, and 640 nm for the GFP-tagged protein, rhodamine-labeled microtubules, and biotinylated far red-labeled microtubules, respectively.
    1. Add one drop of immersion oil onto the coverslip of the sample chamber. Excessive oil may damage the microscope objective. With the coverslip side of the sample chamber facing downward, place the sample to be imaged on the microscope platform.
    2. Slowly bring the objective up so that there is contact between both the coverslip and the objective through the oil.Adjust the objective height so that the coverslip surface of the sample chamber is in focus.
    3. Record single frame images of the sample in all three channels. For the experiments here,use 100-200 ms of exposure as an optimal imaging duration across all three channels; longer exposure times may result in increased photobleaching.
    4. Adjust the laser power to achieve a good signal-to-noise ratio from the samples while minimizing photobleaching over the 2-5 min when the individual bundle is assayed. For the laser used here (Figure 4 and Figure 5), use a laser power of 5 mW for the GFP channel and 3 mW for both microtubule channels as optimal.
      NOTE: Successfully assembled bundles should consist of one surface-immobilized microtubule in the far-red channel, a coextensive region of several microns with significant GFP-protein signal, and a rhodamine microtubule overlapping these regions and then extending several microns away from the bundle (Figure 5B,C and Figure 6). Live imaging of the rhodamine microtubule should reveal subtle fluctuations at the non-crosslinked microtubule end, indicating this filament is unattached to the surface or other proteins in the chamber.
    5. Observe the frequency of microtubule bundles per field of view. For a successfully prepared sample, there should be approximately 3-4 microtubule bundles present per 100 µL x 100 µL field of view per chamber.

6. Performing optical trap experiments on microtubule bundles

  1. Pre-experimental calibration of conditions
    1. To perform these experiments, use a single-beam optical trap with a stiffness of 0.05-0.1 pN/nm. Incorporate the trapping optics and position-sensitive photodetector into a TIRF-capable inverted microscope by introducing short pass filters just below the objective and above the condenser, respectively (Figure 4).
    2. In addition, add a nanopositioning stage on which the sample sits to allow the user to move the microtubules with nanometer-scale precision in a controlled manner during these assays. The system used here to acquire representative data has been described in published work27,53,54,56,57.
      NOTE: Though beyond the scope of this protocol, multiple resources are available that detail the construction and calibration of a single-beam optical trap58,59,60.
    3. Observe the density of beads in the sample using a brightfield or DIC channel and the 100x objective on the microscope. Use brightfield microscopy only to identify and manipulate beads and not to simultaneously record with fluorescence image acquisition. Space the beads to be a minimum of 10 µm from one another on average, and ensure that they are free from any kind of surface confinement or attachment and are not attached to one another or otherwise clustering.
    4. Ensure that the biotinylated far-red microtubules are firmly attached to the surface of the coverslip, with no visible Brownian motion, such as motions along the microtubule axis or free ends of microtubules fluctuating in solution.
    5. Ensure that the rhodamine microtubules are attached to biotinylated microtubules via the crosslinking MAP and are visibly fluctuating at their free ends. Surface attachment of non-biotinylated microtubules often indicates that PEGylation may have been unsuccessful or that the PEG coating has degraded.
  2. Motor-drive microtubule sliding measurements
    1. Express and purify the motor protein of interest following published protocols. For example, full-length GFP-tagged kinesin-5 proteins have been successfully purified and employed in these assays53,61, as well as unlabeled kinesin-1262 and kinesin-1463. Assemble and visualize motor-protein crosslinked bundles as described above in steps 4 and 5.
    2. Capture with the optical trap a free single bead in solution that is exhibiting Brownian motion. Adjust the trap optics as necessary to move the bead to the middle of the field of view. Ensure the reflected interference pattern from the trap beam is symmetric and adjust the trap optics to resolve any beam asymmetries.
    3. Move the sample stage so that the free end of a rhodamine microtubule within a bundle is directly below the bead and the bead is several microns away from the overlap region containing crosslinking proteins to minimize trap beam-induced photobleaching. Lower the bead in the Z-axis until it collides with the microtubule. Take care to lower the bead gently, and do not push the bead against the coverslip surface, as this can cause sticking to the coverslip surface.
    4. Briefly turn off the trap to observe with the brightfield channel the motility of the bead attached to the sliding microtubule. When performing motor protein assays, the attached bead will move directionally as the microtubules slide apart. Reactivate the trap and capture the bead again.
    5. Begin recording fluorescence data in the three (488 nm, 561 nm, 640 nm) channels at a rate of 1-2 frames per s using the laser power and exposure times that were optimized in step 5 above.
    6. Using the trapping computer software, record voltage data from the position-sensitive photodetector (X,Y, and SUM intensity signals), the nanopositioning stage (X,Y, and Z coordinates), and the TIRF camera shutter state. Start digitizing these trap voltage values using a dedicated voltage input data acquisition board controlled by appropriate software (such as custom-written LabView code) at a rate of 1-10 kHz, depending on the experimental requirements.
    7. Monitor the force signal as data is acquired and pay close attention to any anomalies that could interfere with successful data collection.
      NOTE: The anomalies include but are not limited to (1) other beads moving into the trap, (2) the bead escaping the trap prematurely, and (3) sudden plummets of force and subsequent failure to reinitiate force against the trap, indicating issues with the rigor kinesin molecules on the surface of the bead.
    8. After measurement, deactivate the trap to release the bead and repeat with a new bead and new microtubule until the required number of experimental repeats is achieved.
  3. Passive crosslinking resistance measurements
    1. Express and purify the non-motor crosslinking proteins of interest following published protocols. For example, full-length GFP-labelled PRC143,54,57 and a dimeric NuMA truncated construct56 have been successfully employed in these assays. Assemble and visualize the crosslinked bundles as described above in steps 4 and 5.
    2. Identify a suitable microtubule bundle that contains only two microtubules, one biotinylated with full surface attachment and one non-biotinylated microtubule that is partly free in solution; this gives a free end to attach a bead and guarantees that the bead is not attached to the surface-bound microtubule and is aligned on either the X- or Y-axis, such that the stage motion will be along only one axis.
    3. Use the optical trap to capture a free bead undergoing Brownian motion. Once the bead has been trapped, carefully move the bead over to the selected microtubule bundle, ensuring no other beads are pulled into the trap in the process. Ensure that the bead is several microns away from the overlap region containing crosslinking proteins to minimize photobleaching of the GFP tag.
    4. Carefully lower the bead in the Z-axis until it makes contact with the edge of the free segment of the rhodamine microtubule. Pull up gently and move perpendicular to the microtubule axis to check for attachment by observing the rhodamine microtubule bending and following the bead position. If not attached, repeat the gentle bumping of the bead onto the microtubule until attached.
    5. Carefully realign the rhodamine microtubule along the microtubule axis of the surface-attached microtubule by moving the nanopositioning stage and set up the parameters for automated pulling of the microtubule bundle. Set the direction along the parallel axis of the microtubules, set the desired pull speed (25-200 nm/s is a useful range of speeds), and desired pull time (approximately 30 s to 2 min) depending on overlap length.
    6. Begin recording fluorescence data in the three (488 nm, 561 nm, 640 nm) channels at a rate of 1-2 frames per second. Use laser powers and exposure times optimized in step 5 above.
    7. Initiate automated stage motion and record trapping data from the position-sensitive detector, stage, and TIRF camera state. Monitor for any factors that could interfere with data collection, which include but are not limited to (1) another bead entering the trap, (2) premature loss of microtubule crosslinking, or (3) bead detachment from the rhodamine microtubule.
    8. Deactivate the trap to release the bead and repeat the experiment as desired. A typical sample chamber can be used for approximately 30 min before degradation of the bundles and loss of protein activity occurs.
      ​NOTE: Degradation effects will vary by crosslinker used but can manifest as the formation of static puncta on either the microtubules or surface, the loss of microtubule fluctuations indicating non-specific binding, or the inability to stably attach beads to microtubules.

7. Analysis of data and correlation of fluorescence images with optical trap records

NOTE: To optimize data collection, it is beneficial to employ two separate computer control systems: one for the optical trapping software and another for the fluorescence imaging. This setup allows for high-speed data acquisition in both experimental modalities and eliminates nano- and microsecond delays in operation execution being introduced to the data, which can arise when using a single CPU.

  1. Digitize the optical trapping data at a rate optimized for the employed motor or non-motor crosslinking proteins and their expected rates of stepping (e.g., typically between 1-10 kHz).
  2. Record X,Y, and SUM voltage signals from the position-sensitive photodetector, as well as the X,Y, and Z positional data of the nano-positioning stage using dedicated software for controlling the optical trap and sampling the digital voltage signals from these components.
  3. Record the shutter-open state digital signal from the camera with an additional voltage input channel on the optical trapping data acquisition board. This allows for correlation of the fluorescence imaging data with the optical trapping time traces. Begin data collection with the optical trap prior to the initiation of imaging so that the camera signal from the first frame is properly recorded.
  4. Average the raw time series data acquired at high sampling rates using an appropriate filtering method, such as sliding window, median filter, or Gaussian filter, depending on needs.
  5. Quantify the fluorescence image time series data using methods such as linescan analysis, wherein the pixel intensity value along the length of the microtubule region is calculated. From these linescan trajectories, identify the microtubule edges, and, therefore, the overlap regions. In addition, use the fluorescence intensity from the crosslinker protein channel to calculate protein localization, concentration, and density.
  6. Correlation of force and image data is an essential output of this experimental system. For example, select and average all force data points within a given camera exposure region (indicated by the corresponding digital signal spike in the camera channel) to directly compare with the corresponding fluorescence image frame. Subsequently, extract the relationships between force magnitude and the number of crosslinking proteins, overlap length, or crosslinker distribution.

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Representative Results

The preparation of microtubule bundles suitable for biophysical analysis is considered successful if several of the key criteria are met. First, imaging in three colors should reveal two aligned microtubules with a concentration of crosslinking protein preferentially decorating the overlap region (Figure 5B,C and Figure 6B). Ideally, the distance between the overlap edge and the free end of the rhodamine microtubule should be at least 5 µm to provide sufficient physical space between the trapping beam and the fluorescence-labeled proteins. Second, there should be minimal background fluorescence signal in regions that lack biotinylated far-red microtubules, particularly from the same channel that the crosslinking protein is labeled with. High background fluorescence is indicative of poor coverslip passivation and a high concentration of glass-bound proteins, which will likely interact non-specifically with the microtubules or trapping bead. Additionally, the presence of small round puncta or short fragments seen in either of the microtubule imaging channels suggests that microtubule polymerization and clarification were unsuccessful or that the microtubules were aged or damaged.

Third, the beads used for trapping should appear as single beads and not within clumps containing many beads. It is likely that a small fraction of beads will irreversibly adhere to the surface, but a significant fraction should be observed as individual particles diffused in the sample chamber. Excessive clumping can often be alleviated by sonicating the beads for at least 10 min prior to flowing into the chamber. Fourth, attaching a bead to a microtubule should result in good attachment, with the bead remaining bound when the trapping laser light is shuttered. The beads should not stick non-specifically when brought into contact with the surface, and, once a bead is attached to the microtubule, it should be possible to lightly manipulate (e.g., bend or flex) the filament prior to measurement. Finally, it should be possible to observe changes in the overlap length when force is applied, or motor activity is allowed to proceed. For example, if observing motor protein-driven sliding, the overlap length should decrease at a rate consistent with the rate of motor protein stepping. If examining passive crosslinkers, the application of force should result in a change in the overlap length. These outputs indicate that the rhodamine microtubule is attached only to the surface-bound microtubule via crosslinking proteins, rather than non-specifically sticking to the coverslip surface.

When these criteria are all met, it is possible to perform a wide range of experiments to extract the essential biophysical parameters that define ensemble mechanics. This assay or similar assays have been used extensively to examine mitotic crosslinking proteins in previous studies. For example, we have shown that ensembles of the essential mitotic motor protein kinesin-5 can regulate microtubule sliding by generating both pushing and braking forces that scale linearly with overlap length64 (Figure 5). Microtubules in either an antiparallel or parallel geometry were crosslinked by a small ensemble of kinesin-5 molecules. As these motor proteins stepped toward the microtubule plus-ends, the filaments were either slid apart (antiparallel) or rapidly fluctuated back-and-forth (parallel). By monitoring the mechanics within this mini-spindle geometry, we found the magnitude of force scaled both with the length of filament overlap, as well as the number of crosslinking motor proteins. When microtubules were moved at a velocity faster than kinesin-5's unloaded stepping rate, the motor proteins provided a resistive braking force. It was also found that the C-terminal tail domain is required for efficient crosslinking and force generation61 (Figure 5B,C). The full-length protein is able to generate sustained forces that plateau when all motor proteins reach their individual stall force (Figure 5D). However, the kinesin-5 motor lacking its C-terminal tail generates maximum forces that are nearly five-fold smaller in magnitude (Figure 5E,F). Together, these results reveal how kinesin-5 proteins work in ensembles to push microtubules poleward during spindle assembly and elucidate the biophysical function of essential regulatory protein domains within the molecule.

We have also demonstrated that ensembles of the non-motor mitotic protein PRC1 operate as a mechanical dashpot to resist microtubule sliding in anaphase spindle midzones54 (Figure 6A-C). PRC1 generates frictional resistance that scales linearly with pulling speed, just as a mechanical dashpot produces velocity-dependent resistive forces. These resistive forces do not depend on the length of overlap regions between microtubule pairs or the local crosslinker density, but they do strongly depend on the total concentration of engaged PRC1 molecules (Figure 6D,E). From these results, it is proposed that PRC1 ensembles act as a leaky piston, wherein compression of diffusive crosslinkers produces a velocity-dependent resistance, but the loss of crosslinkers at microtubule plus-ends alleviates large compressive forces.

Similar assay geometries were also used to examine the mitotic kinesin-12 protein Kif1562. By measuring the force produced as microtubules were pushed apart, Reinemann et al.62 found that Kif15 ensembles can push apart filaments up to a critical plateau force and require the C-terminal tail tether region of the protein in order to efficiently crosslink microtubules and build sustained loads. Reinemann et al. also elegantly demonstrated that the kinesin-14 HSET, a minus-end directed kinesin, can similarly slide apart antiparallel microtubules63. When mixed with equal amounts of the plus-end directed kinesin-5 Eg5 protein, HSET serves to inhibit Eg5 sliding activity, engaging in a tug-of-war for microtubule sliding motion within the overlap region. Together, these results all demonstrate the power of direct force measurement across active microtubule bundles and make clear the need for characterizing protein function in the biologically appropriate network geometry.

Figure 1
Figure 1: Passivation of glass coverslips. (A) Schematic depicting the sequential wash steps, drying, and conjugation of No. 1.5 glass coverslips for amino-silane conjugation. (B) Schematic depicting the protocol for covalently linking PEG and biotin-PEG groups to the glass coverslip, washing and drying, and sealing the coverslips under vacuum for short-term storage. Some portions of the schematic are modified images from 65. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Assembly of a sample chamber. (A) Schematic depicting the assembly of a sample flow chamber using passivated coverslips and a standard 3 in microscope slide. (B) Assembly and typical dimensions of single- and dual-channel flow chambers that are optimized for optical trapping and fluorescence imaging of microtubule bundles. Some portions of the schematic are modified images from 65. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Generation of surface-immobilized microtubule bundles for optical trapping and TIRF-based imaging. Schematics depicting the stepwise assembly of optically trappable microtubule bundles. (A) Reagents are flowed in from the entry port of the chamber and fluid is wicked out from the exit channel. (B) Streptavidin (blue) first binds to the biotin-PEG sites on the coverslip, and (C) casein (red) is introduced as an additional blocking agent. (D) Biotinylated far red-labeled microtubules (magenta) are introduced and allowed to incubate for ~5 min, followed by the addition of (E) crosslinking protein (green), (F) non-biotinylated rhodamine-labeled microtubules (red), and, finally, (G) rigor kinesin-coated microspheres suspended in the appropriate reaction buffer for attachment to the bundle and optical trap-based manipulation. (H) The chamber is sealed with clear nail polish to prevent evaporation of the sample buffer during experiments. Some portions of the schematic are modified images from 65. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Schematic of the optical tweezers/TIRF microscope system. A single-beam optical trap is introduced into the optical path of an objective-based TIRF imaging system within an inverted microscope. A short pass filter inserted just below the objective allows the trap beam to be directed into the back aperture of the objective, where it will be focused just above the surface of the sample coverslip, forming an optical trap. A position-sensitive photodetector will collect the transmitted light, allowing for position and force detection of the bead. Multiple channels of fluorescence data can be acquired via TIRF imaging and recorded on a high-sensitivity EMCCD or sCMOS camera. A broad-spectrum light source is used to image trapping beads during the setup of the experiments. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Representative example of measuring force production by kinesin-5. (A) Schematic depicting the experimental geometry, showing microtubules crosslinked by kinesin-5 motor proteins that generate force as they slide the filaments apart, which is measured with an optical trap. Representative fluorescence images of bundles composed of surface-immobilized microtubules, GFP-kinesin-5, and rhodamine-labeled transport microtubules. (B) Full-length and (C) C-terminal tail truncated kinesin-5 constructs were employed. Scale bars = 5 µm. Sample records of force production for (D) full-length and (E) tailless are shown, with average force plotted as a function of time. (F) Plots of the maximum plateau force and corresponding overlap length for both constructs are shown, revealing that the full-length kinesin construct generates overlap-length dependent forces that are substantially larger in magnitude than the tailless kinesin construct. This figure has been modified from61. Please click here to view a larger version of this figure.

Figure 6
Figure 6: Representative example of measuring frictional forces by PRC1. (A) Schematic depicting the experimental geometry, showing microtubules crosslinked by GFP-PRC1 (protein regulator of cytokinesis) crosslinking proteins that generate resistance as the filaments are slid apart by moving the coverslip at constant velocity. (B) Representative time series fluorescence images showing the moving surface-immobilized far-red microtubule, GFP-PRC1 molecules condensing within the shrinking overlap, and the free rhodamine microtubule held with the bead. Time interval between frames = 6 s; Scale bar = 5 µm. (C) Example force trace showing frictional force during the sliding event, with disruption event and force drop to zero (~55 sec) once the overlap reached 0 µm. (D) Correlation of mean force and integrated GFP intensity within the overlap at four different sliding velocities. (E) Mean slopes and calculated fit errors of traces in (D) reveal that the force per PRC1 molecule increases linearly as a function of sliding velocity. This figure has been modified from66. Please click here to view a larger version of this figure.

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Discussion

Microtubule networks are employed by myriad cell types to accomplish a wide range of tasks that are fundamentally mechanical in nature. In order to describe how cells function in both healthy and disease states, it is critical to understand how these micron-scale networks are organized and regulated by the nanometer-sized proteins that collectively build them. Biophysical tools such as optical tweezers are well suited to probing the mechanochemistry of key proteins at this scale. Reflecting the diversity of microtubule network function, complex experimental geometries that employ optical tweezers have been explored, including force-dependent analyses of microtubule tip dynamics67,68, the generation of forces by kinetochore components69,70,71, and two-beam dumbbell trap geometries to probe microtubule filament mechanics72,73. In this protocol, we have described novel methods for reconstituting fundamental microtubule network motifs such as bundles and applying the biophysical tools of single-molecule fluorescence microscopy and optical trapping to assess critical information about cellular function.

While most individual steps in this protocol are technically straightforward, taken together there are numerous potential points of failure that can arise, and substantial care must be taken at each point to ensure successful measurements. First, as with many microscope-based single-molecule experimental assays, proper surface passivation is key, as non-specific binding of proteins or beads to the coverslip surface nearly always prevents the successful completion of these experiments. Second, the expression and purification of the crosslinking protein(s) of interest must be optimized to ensure high quality single-species products, as contaminants can interfere with the assay in numerous ways, such as inducing excess bundling of filaments or interfering with bead and microtubule attachment, not to mention introducing complexities in the interpretation of force data. Third, maintaining robust bead-microtubule attachment requires high quality rigor kinesin motors bound at high density to the trapping bead, such that multiple kinesin-filament attachment points can be made. These bonds can withstand 10 s of pN of force for several minutes, which is suitable for a wide range of potential systems of study employing this technique.

We also note that there are many ways in which this protocol can be modified to best suit the end user's instrumentation or biological system needs. While we have described experiments that employ GFP-labeled crosslinking proteins and red/far red-labeled microtubules, it is also possible to swap fluorescent labeling as needed. For example, if the user does not have enough laser lines to perform three-color TIRF microscopy, it is possible to get high quality data by differentially labeling microtubules with the same fluorophore but using different concentrations (e.g., dim vs. bright) for each species of microtubule. Similarly, it may not always be possible or advantageous to add a fluorescent label to the crosslinking protein. In this situation, certain experimental information such as crosslinker concentration or localization would not be accessible, but the determination of parameters such as microtubule overlap length could still be made. We anticipate that this protocol is highly adaptable for many different types and combinations of microtubule crosslinking proteins. The introduction of multiple crosslinking proteins would likely necessitate either the removal of fluorescence tags from one of the proteins or the exclusion of fluorescent labels from one of the two microtubule types to free up imaging capabilities in an appropriate bandwidth (e.g., using a red or far-red protein-encoded label for the crosslinking protein). It is unlikely that more than three fluorescent channels could be employed with this TIRF-based assay, though there is certainly flexibility in how those are distributed, which should be carefully considered when planning an experiment. Finally, it is likely that this assay could be modified to study different network geometries and types of cytoskeletal filament. The analysis of the mechanics of microtubule branching, mediated by augmin and the gamma-tubulin ring complex, could readily be probed and imaged. In addition, other crosslinked cytoskeletal networks, such as those involving actin, septins, or intermediate filaments, would be quite amenable to study with these tools, assuming proper modifications and orthogonal attachment strategies to the coverslip and trapping bead are made.

In conclusion, we have described a protocol for the reconstitution of active force-generating microtubule networks, which can be imaged using single-molecule fluorescence microscopy tools and manipulated via a single-beam optical trap. We have demonstrated the usefulness of this method with datasets that reveal the ensemble mechanics of both a mitotic motor and non-motor protein. We anticipate that many types of highly organized microtubule networks, such as those found in neurons, epithelial tissue, or cardiomyocytes, can be analyzed with these tools, revealing new principles of biophysical function for the dynamic cytoskeleton.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

The authors wish to acknowledge support from R21 AG067436 (to JP and SF), T32 AG057464 (to ET), and Rensselaer Polytechnic Institute School of Science Startup Funds (to SF).

Materials

Name Company Catalog Number Comments
10W Ytterbium Fiber Laser, 1064nm IPG Photonics YLR-10-1064-LP
405/488/561/640nm Laser Quad Band Set for TIRF applications Chroma TRF89901v2
6x His Tag Antibody, Biotin Conjugate Invitrogen #MA1-21315-BTIN
Acetone, HPLC grade Fisher Scientific 18-608-395
Alpha casein from bovine milk Sigma 1002484390
ATP Fisher Scientific BP413-25
Benzonase Novagen 70746-3
Biotin-PEG-SVA-5000 Laysan Bio, Inc. NC0479433
BL21 (DE3) Rosetta Cells Millipore Sigma 71-400-3
Catalase MP Biomedicals LLC 190311
CFI Apo 100X/1.49NA oil immersion TIRF objective Nikon N/A
Chloramphenicol ACROS Organics 227920250
Coverslip Mini-Rack, for 8 coverslips Fisher Scientific C14784
Delicate Task Wipers Kimberly-Clark 34120
Dextrose Anhydrous Fisher Scientific BP3501
D-Sucrose Fisher Scientific BP220-1
DTT Fisher Scientific BP172-25
Ecoline Immersion Thermostat E100 with 003 Bath LAUDA-Brinkmann 27709
EDTA Fisher Scientific BP118-500
EGTA Millipore Corporation 32462-25GM
FIJI / Image J https://fiji.sc/ N/A
Frosted Microscope Slides Corning 12-553-10 75mmx25mm, with thickness of 0.9-1.1mm
Glucose Oxidase MP Biomedicals LLC 195196 Type VII, without added oxygen
GMPCPP Jena Biosciences JBS-NU-405S Can be stored for several months at -20 °C and up to a year at -80 °C
Gold Seal-Cover Glass Thermo Scientific 3405
HEPES Fisher Scientific BP310-500
Imidazole Fisher Scientific 03196-500
IPTG Fisher Scientific BP1755-10
Laboratory dessicator Bel-Art 999320237 190mm plate size
Kanamycin Sulfate Fischer Scientific BP906-5
KIF5A K439 (aa:1-439)-6His Gilbert Lab, RPI N/A doi.org/10.1074/jbc.RA118.002182
Kimwipe Kimberley Clark Z188956 lint-free tissue
Immersion Oil, Type B Cargille 16484
Lens Tissue ThorLabs MC-5
LuNA Laser launch (4 channel: 405, 488, 561, 640nm) Nikon N/A
Lysozyme MP Biomedicals LLC 100834
Magnesium Acetate Tetrahydrate Fisher Scientific BP215-500
Microfuge 18 Beckman Coulter 367160
MPEG-SVA MW-5000 Laysan Bio, Inc. NC0107576
Neutravadin Invitrogen PI31000
Nikon Ti-E inverted microscope Nikon N/A Nikon LuN4 Laser
Ni-NTA Resin Thermo Scientific 88221
Oligonucleotide - CACCTATTCTGAGTTTGCGCGA
GAACTTTCAAAGGC		 IDT N/A
Oligonucleotide - GCCTTTGAAAGTTCTCGCGCAA
ACTCAGAATAGGTG		 IDT N/A
Open-top thickwall polycarbonate tube, 0.2 mL, 7 mm x 22 mm Beckman Coulter 343755
Optima-TLX Ultracentrifuge Beckman Coulter 361544
Paclitaxel (Taxol equivalent) Thermo Fisher Scientific P3456
PIPES ACROS Organics 172615000
PMSF Millipore 7110-5GM
Porcine Tubulin, biotin label Cytoskeleton, Inc. T333P
Porcine Tubulin, HiLyte 647 Fluor Cytoskeleton, Inc. TL670M far red labelled
Porcine Tubulin, Rhodamine Cytoskeleton, Inc. TL590M
Porcine Tubulin, Tubulin Protein Cytoskeleton, Inc. T240
Potassium Acetate Fisher Scientific BP364-500
Prime 95B sCMOS camera Photometric N/A
Quadrant Detector Sensor Head ThorLabs PDQ80A
Quikchange Lightning Kit Agilent Technologies 210518
Sodium Bicarbonate Fisher Scientific S233-500
Sodium Phosphate Dibasic Anhydrous Fisher Scientific BP332-500
Square Cover Glasses Corning 12-553-450 18 mm x 18 mm, with thickness of 0.13-0.17 mm
Streptavidin Microspheres Polysciences Inc. 24162-1
Superose-6 Column GE Healthcare 29-0915--96
TCEP Thermo Scientific 77720
TLA-100 Fixed-Angle Rotor Beckman Coulter 343840
Ultrasonic Cleaner (Sonicator) Vevor JPS-08A(DD) 304 stainless steel, 40 kHz frequency, 60 W power
Vectabond APTES solution Vector Laboratories SP-1800-7
Windex Powerized Glass Cleaner with Ammonia-D S.C. Johnson SJN695237

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Reconstituted Active Microtubule Bundles Measure Forces Cytoskeletal Motifs Mechanical Components Healthy And Disease States Quantify Forces Protein Concentration And Density Cytoskeleton Network Mitosis Neural Development Muscle Contraction Cell Migration Single-beam Optical Trap TIRF Microscope Single-molecule Experiments High-quality Samples Humidity Chamber
Directly Measuring Forces Within Reconstituted Active Microtubule Bundles
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Palumbo, J., Tai, E., Forth, S.… More

Palumbo, J., Tai, E., Forth, S. Directly Measuring Forces Within Reconstituted Active Microtubule Bundles. J. Vis. Exp. (183), e63819, doi:10.3791/63819 (2022).

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