Presented here are methods for preparing active nematics from microtubules and kinesin motors, including protein preparation and construction and the use of wells for active nematic confinement.
The formation of biopolymer-based active phases has become an important technique for researchers interested in exploring the emerging field of active liquid crystals and their possible roles in cell biology. These novel systems consist of self-driven sub-units that consume energy locally, producing an out-of-equilibrium dynamic fluid. To form the active liquid crystal phase described in this report, purified protein components including biopolymers and molecular motors are combined, and the active nematic phase spontaneously forms in the presence of adenosine triphosphate (ATP). To observe the nematic state, the material must be confined in a suitable geometry for microscopy at a high enough density. This article describes two different methods for the formation of an active nematic phase using microtubules and kinesin motors: assembly of a two-dimensional active layer at an oil and water interface and assembly under an oil layer using an elastomeric well. Techniques to insert the active material into small wells of different shapes are also described.
Active fluids are composed of energy-driven particles or elements that draw fuel from their local environment. Under the right conditions, these motile active elements can act collectively to produce emergent fluid dynamics over long length-scales. There are a variety of examples of such out-of-equilibrium phase behavior in the literature and active phases can be found across the spectrum of living systems. Some notable examples are colonies of bacteria1, cell sheets2,3, and the flocking or swarming of organisms4,5. Active phases have also been studied extensively in condensed phases of cytoskeletal filaments, either as part of the cell6 or in synthetic systems designed to make use of biologically extracted components7,8,9. Liquid crystalline ordering and the formation of topological defects in both naturally occurring and synthetic systems assembled from biological extracts are of particular interest to the research community. In recent years, research groups have examined such systems, their fundamental physical properties, and their relevance to biology2,3,10,11.
This paper focuses on the formation of the active nematic state from a combination of microtubules and kinesin motor proteins. The traditional nematic liquid crystal is an equilibrium phase of matter in which the constituent molecules exhibit orientational ordering. For example, a fluid consisting of relatively rigid rod-like molecules may exhibit both the nematic phase and, at higher temperatures, an unoriented isotropic fluid phase12. The first experimental example of an active nematic phase was developed by Sanchez et al.13, adapting an earlier in vitro experiment14 in which clusters of motor proteins were used to produce a shearing motion between neighboring microtubule bundles. When this microtubule system was confined to a thin layer, spontaneous nematic ordering emerged. In recent years, the active nematic state has been studied intensively by several experimental15,16 and theoretical17,18 research groups, focusing on phenomena such as active turbulence — a state in which the fluid produces self-driven chaotic flows19 — and mobile topological defects. This paper describes methods to prepare and form the active nematic state from microtubules and kinesin motors in different experimental geometries. First, preparation methods for the different component solutions are described, followed by methods for forming the active nematic using two different flow chamber geometries. Typical imaging results are shown. Finally, methods for confining the active nematic in wells and channels are described.
1. Preparing the active material
NOTE: The 2D active nematic is assembled in a three-step process. First, two separate solutions are prepared: a) polymerized, stabilized microtubules and b) MIX (a solution containing kinesin motors). These are combined and activity is initiated upon adding adenosine triphosphate (ATP). The material is then confined in a suitable geometry, such that its density is high enough for nematic order to emerge. Protocols are included for the preparation of all necessary components and how to assemble the active phase.
2. Creating the active nematic
NOTE: Activity in the material is initiated by ATP addition. The active network is prepared fresh for each experiment by adding ATP at a concentration high enough to induce motor activity. To form a uniform, fully developed active nematic phase, the microtubules must be at a sufficiently high density. This can be achieved by confining the microtubules in between two immiscible fluids to form a two-dimensional (2D) active nematic layer. This method was originally developed at Brandeis University13 and remains a popular technique for producing a homogeneous, high quality, active nematic phase.
3. Preparing active nematics in confined geometries
NOTE: Active nematics such as this quasi-two-dimensional system can be challenging to confine into small microfluidic geometries such as wells or channels. Here, a reliable method to confine the material into different shaped PDMS wells is described.
Figure 1 shows a representative image of single microtubules prepared from GMPCPP tubulin. The image depicts short microtubules of similar lengths (with some dispersity present). Sufficient dilution of the microtubule solution should produce an image of well separated microtubules for length verification. The individual microtubules may be challenging to image due to their small size. Use of a high sensitivity camera designed for fluorescence microscopy is best for this application. Figure 2 and Figure 3 show example fluorescence microscopy images of successful experiments performed using the flow cell method (section 2.1) and the inverted method (section 2.2), respectively. A well-formed active nematic layer is homogeneous in texture, with no significant void areas and mobile topological defects present. Note however that there may be some acceptable small voids in the defect cores. In addition to the examples shown in Figure 2 and Figure 3, three supplemental movies (Movie 1, Movie 2, and Movie 3) have been included to demonstrate how the active nematic should appear in a successful experiment. All the movies demonstrate the smooth continuous motion of the active nematic phase. No variations in microtubule concentration are apparent after the material has reached its steady state. As long as sufficient ATP is present in the system, the material will continue to move uniformly.
Figure 1: Fluorescence microscope image of GMPCPP microtubules. The GMPCPP microtubules were labeled at 4% with rhodamine tubulin and polymerized for 20 min at 37 °C. Imaging was performed at room temperature. Scale bar = 10 µm. Please click here to view a larger version of this figure.
Figure 2: Microtubule nematics in a flow cell. (A) Cross sectional schematic of the flow cell, 1 mm x 18 mm geometry. (B) Top view schematic of the flow cell. (C) Fluorescence microscopy image demonstrating the typical appearance of the active solution before assembly at the oil/water interface. (D) Fluorescence microscopy image of the active nematic phase assembled at the oil/water interface inside the flow cell. Scale bar = 100 µm. Please click here to view a larger version of this figure.
Figure 3: Fluorescence microscope image showing an active nematic prepared using the inverted method. Scale bar = 200 µm. Please click here to view a larger version of this figure.
Figure 4: Flow diagram illustrating the method for active nematic confinement in a PDMS well, including mold fabrication and surface treatment. Scale bar on the right image (confined active material) = 200 µm. Please click here to view a larger version of this figure.
Movie 1: Representative result for active nematic prepared using the flow cell method. Please click here to download this Movie.
Movie 2: Representative result for active nematic prepared using the inverted method. Please click here to download this Movie.
Movie 3: Representative result for active nematic prepared using the inverted method confined to an elliptical well. Please click here to download this Movie.
There are a few points throughout the protocols at which the experimenter can make some important checks. Before filling either of the devices with active material, fluorescence microscopy (see Figure 1) should be used to check that the microtubules are polymerized and ideally ~2-3 µm in length. If microtubules are not visible under the microscope, they may have depolymerized and the active nematic will not form. Because individual microtubules are very small, it may be challenging to observe them directly through the microscope. In this study, a high-quality fluorescence camera designed for challenging low-light applications was used with the associated software to verify filament growth. Significant fluorescent aggregates should not be present at this stage, as this may indicate depolymerization or the presence of denatured protein. It is also a good idea to make a simple microscope test slide by combining microtubules, MIX, and ATP in the same ratios as described in the protocols. Activity should begin on combining the components and the material should appear similar to that shown in Figure 2C with bundles present and noticeable filament movements visible throughout.
When using the flow cell method, the centrifuge time and orientation of the flow cell are important for the formation of a uniform active layer. This step may require some fine tuning depending on the centrifuge type used. Centrifuging the flow cell with the active plane oriented perpendicular to the plane of rotation gives the best results as material can be pushed onto the fluid interface uniformly. Double check that the flow cell is carefully sealed before centrifuging.
When using the inverted method to produce confined active nematics there are several steps to optimize. First, it is important to use a 3D printing method that produces high resolution structures. Uneven side walls can cause the microtubules to catch, which will disrupt the flows. The wells should not be too deep (150-200 µm deep wells with a 2 mm thick overlying oil layer were used in this study). Experimenters may need to adjust these parameters slightly by trial and error to get the best result.
The flow cell method and the inverted method have been used by different authors to look at a variety of effects that impact the active flows, including different oils12 and submersed structures13. The choice of method depends on the experimental objective. Using the flow cell method, optical imaging from above the active layer is clearer than for the inverted method due to the different overlying fluids. In the flow cell method, imaging is carried out through a glass cover slip and a thin layer of water, whereas the inverted method is designed to have the oil layer on top. This means that a long working distance objective is needed for the inverted method, and image quality is reduced. Image quality differences can be seen by comparing Figure 2D (flow cell method) and Figure 3 (inverted method), and Movie 1 and Movie 2, respectively. A lower magnification lens with a longer working distance was required for Figure 3 than that used for Figure 2. These imaging disadvantages for the inverted method can be avoided if a suitable inverted microscope is available, combined with objectives with an appropriate working distance for the microscope slide substrates. Thinner glass can be used as a substrate to allow the use of standard working distance objectives.
As an advantage, the inverted geometry allows for the use of a wider range of oil viscosities, does not necessarily require swinging bucket centrifugation (if this is not available), and preparation of the system is relatively easier once the mold is prepared. However, for confinement in wells using the inverted method, some centrifugation may be important to get the material into a well-defined 2D layer.
The flow cell method has recently been used very successfully in experiments where a continuous active layer is required. Our recent work has looked at the dynamics of topological defects in the active layer, where high quality imaging and texture analysis is important19. In addition, the flow cell method has been used to investigate the effects of oil-submersed microstructures on active flows16 and pillars to trap defects in the active flows31. This method works very well for the formation of a continuous active layer, and the image quality is excellent. However, the centrifugation step used to produce the final 2D active layer can be difficult to carry out, and the flow cells are prone to leaks and air bubbles. The inverted method is a very useful alternative with a high success rate, is easy to construct, and can be used for any substrate pattern or geometry provided a high-resolution 3D printed master mold can be created. This method is also useful for looking at the effects of geometrical confinement on active nematic dynamics because it makes filling wells relatively straightforward.
In this paper, two ways to form an active nematic from microtubules and kinesin motors are described, plus a technique to confine the materials in wells. The system presented represents the cleanest example of an active nematic phase currently in the literature and has been reproduced by several groups around the world. The significance of this material not only lies in the biological origins of its components, but also because it opens up an entirely new direction in active ordered fluids. By working with this system and elucidating its fundamental properties, scientists can move toward the design of fully synthetic active phases.
The experiments focused on the effects of confinement on active nematics have the potential to answer fundamental questions regarding the behavior of active flows and topological defect dynamics under topological confinement. The method presented here will aid in the performance of a variety of geometry-focused experiments and their analysis, including microfluidics and active mixing.
The authors would like to acknowledge the National Science Foundation (NSF) award DMR-1808926 for generous funding. The project was also supported by the NSF through the Center of Research Excellence in Science and Technology: Center for Cellular and Biomolecular Machines at the University of California Merced (HRD-1547848) and the Brandeis Biomaterials Facility Materials Research Science and Engineering Center (DMR-2011846). We would like to thank Dr. Bin Liu at the University of California Merced for assistance in 3D printing the mold, and Dr. Jordi Ignes for scientific advice during the development of the inverted experimental method.
Name | Company | Catalog Number | Comments |
20 kD PEG (polyethylene glycol)) | Sigma Aldrich | 1419109 | Depletion agent CAS Number: 125061-88-3 |
3-(trimethoxysilyl)propyl methacrylate | Sigma Aldrich | M6514-50ML | CAS Number: 2530-85-0 |
3D printer & Resin | Phrozen | Phrozen sonic mini 8K 3D printer - Aqua Gray 8K resin | |
40% Acrylamide Solution | BIO-RAD | 1610140 | CAS Number: 7732-18-5, 79-06-1 |
Acetic Acid | Fisher | CAS Number: 64-19-7 | |
Acetone | Sigma Aldrich | CAS Number: 67-64-1 | |
Adhesive sheets (NOTE: "Parafilm" is an alternative) | Grace Bio-Labs | 620001 | SecureSeal |
Ammonium Persulfate | Sigma Aldrich | A3678 | CAS Number: 7727-54-0 |
Aquapel (NOTE: "RainX" is an alternative) | Aquapel Glass Treatment | hydrophobic glass treatment | |
ATP (Adenosine triphosphate) | Sigma Aldrich | A1852 | CAS Number: 34369-07-8 |
Beakers | VWR | ||
Catalase | Sigma Aldrich | C9322 | CAS Number: "9001-05-2" |
Desiccator | Bel-art | ||
Digital CMOS camera | Hamamatsu | ORCA - Flash4.0 LT+ | |
DTT (Dithiothreitol) | Sigma Aldrich | D9779 | CAS Number: "3483-12-3" |
EGTA (3,12-bis(carboxymethyl)-6,9-dioxa-3,12-diazatetradecane-1,14-dioic acid) | Sigma Aldrich | MFCD00004291 | CAS Number: 67-42-5 |
Ethanol | Sigma Aldrich | CAS Number: 64-17-5 | |
Fluorescence microscope | Leica | DM 2500P | |
Glass Coverslips | VWR | 48368-040 | |
Glass Slides | VWR | 16004-430 | |
Glucose | Sigma Aldrich | G7021 | CAS Number: 50-99-7 |
Glucose Oxidase | Sigma Aldrich | 345386 | CAS Number: 9001-37-0 |
GMPCPP (guanylyl 5'-α,β-methylenediphosphonate) | Jena Bioscience | NU-405S | CAS Number: 14997-54-7 |
HFE7500 Oil | 3M | ||
Hot Plate | Fisher Scientific | Thermix hot plate model 100M | |
Isopropyl Alcohol | VWR | ||
KCl (potassium chloride) | Sigma Aldrich | P5405 | CAS Number: 7447-40-7 |
Methanol | Sigma Aldrich | CAS Number: 67-56-1 | |
MgCl2 (Magnesium Chloride) | Sigma Aldrich | 208337 | CAS Number: 7786-30-3 |
Microcentrifuge tubes | Eppendorf - Thermo Fisher | 1.5 mL | |
Nanopure water purifier | Sartorius | arium mini | |
NaOH (Sodium hydroxide) | Sigma Aldrich | SX0603 | CAS Number: 1310-73-2 |
Petri Dishes | VWR | ||
PH Meter | Thermo Scientist | Orion 3 STAR | |
Phosphoenol-pyruvate (PEP) | Sigma Aldrich | MFCD00044476 | CAS Number: 4265-07-0 |
PIPES (1,4-Piperazinediethanesulfonic acid) | Sigma Aldrich | CAS Number: 5625-37-6 | |
Pipettes (0.2 - 1000 µl) | VWR | ||
Pluronic F-127 | Sigma Aldrich | 2594628 | |
RAN Surfactant (NOTE: "FluoSurf" from Emulso is an alternative) | Ran Biotechnologies | 008-FluoroSurfactant-2wtH-50G | |
Silicon Oil (100mpa s-1000 mpa s) | Sigma Aldrich | CAS Number: 63148-52-7 | |
Streptavidin | Thermofisher | S888 | |
Swinging Bucket Centrifuge | Thermo Scientist | Sorvall legend RT+ | |
Sylgard 184 Elastomer base | World Precision Instruments | SYLG184 | |
Sylgard 184 Elastomer Curing agent | World Precision Instruments | SYLG184 | |
Table top centrifuge | Eppendorf | MiniSpin Plus | |
TEMED (Tetramethylethylenediamine) | BIO-RAD | 1610800 | CAS Number: 110-18-9 |
Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) | Sigma Aldrich | MFCD00006846 | CAS Number: 53188-07-1 |
Tubulin | Cytoskeleton | T240-B | |
Tubulin (Rhodamine labeled) | Cytoskeleton | TL590M-A | |
Ultracentrifuge | Beckman | Optima Max-TL | |
UV Light | RapidFix | ||
UV-curable glue (NOTE: "Norland NO81" is an alternative) | RapidFix | ||
Water Bath | Thelco | ||
Whatman Filter paper | Sigma Aldrich | WHA1001325 |
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