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.

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.
<ol>
	<li>Kinesin motor protein cluster preparation
	<ol>
		<li>Express and purify recombinant K401-BIO motor proteins (K401 motors) from Escherichia coli following the protocol by Edgar C. Young20. 
		NOTE: K401-BIO motor proteins are dimeric and consist of two heads connected with a helical stalk. The motors were supplied by the Brandeis Biomaterials Facility and used as previously reported16. For the purposes of forming an active nematic, the kinesin molecules are biotinylated and then connected via a streptavidin linkage to form kinesin clusters of up to four motors13,15,21,22. It is helpful to express the kinesin with a green fluorescent protein (GFP) tag.</li>
		<li>After purification and incubation of streptavidin and motors on ice for 40 min, flash-freeze the K401 motors in liquid nitrogen in 5 &#181;L aliquots at a final concentration of 0.7 mg/mL, and store at -80 &#176;C. 
		NOTE: The experiment can be paused here. Gently thaw the kinesin when needed and do not refreeze.</li>
		<li>Prepare clusters of biotin-labeled kinesin (KSA) by mixing 24 vol% of 0.7 mg/mL K401 motors, 27 vol% of 0.325 mg/mL streptavidin, and 3 vol% of 5 mM dithiothreitol (DTT) (to prevent aggregation) at 4 &#176;C in 46 vol% M2B buffer (80 mM PIPES [1,4-Piperazinediethanesulfonic acid, pH 6.8], 2 mM MgCl2, and 1 mM EGTA [ethylene glycol-bis(&#946;-aminoethyl ether)-N,N,N&#8242;,N&#8242;-tetraacetic acid]). Let the KSA incubate on ice for 40 min.</li>
	</ol>
	</li>
	<li>Microtubule solution preparation 
	NOTE: Guanosine-5&#39;-[(&#945;,&#946;)-methyleno]triphosphate, sodium salt (GMPCPP) is a slowly hydrolysable analog of Guanosine triphosphate (GTP), and microtubules formed in the presence of GMPCPP are three times stiffer than GTP microtubules23 and shorter. The use of short, stiff microtubules is favorable for the formation of the active nematic phase as these factors combine to promote liquid crystalline ordering.
	<ol>
		<li>Polymerize unlabeled cycled tubulin (99%, see Table of Materials) using 0.6 mM GMPCPP at a tubulin concentration of 6 mg/mL. 
		NOTE: High quality tubulin can be also purified from bovine or porcine brain following established protocols or obtained from another reliable source such as the Brandeis Biomaterials Facility where materials can be shipped frozen to prevent damage. For tubulin purification from bovine brain, please refer to the published protocol from Bate et al.24. For tubulin purification from porcine brain, refer to the published protocols from Castoldi et al.25 and Tayar et al.26.</li>
		<li>In preparation for polymerization, prepare a heat bath at 37 &#176;C and pre-cool the centrifuge to 4 &#176;C. Combine the unlabeled tubulin solution in M2B buffer (step 1.1.3) in a 500 &#181;L ultracentrifuge tube with 4 mol% rhodamine-labeled tubulin (see Table of Materials) to produce 4% labeled microtubules after polymerization.</li>
		<li>Verify tubulin concentration using a Bradford assay27. The total tubulin concentration in the centrifuge tube should be 6.5-6.9 mg/mL.</li>
		<li>Incubate the tubulin mixture on ice for 10 min and ultracentrifuge for 10 min at 352,700 x g at 4 &#176;C. This step removes dysfunctional tubulin, which will be in the pellet.</li>
		<li>Using a pipette, carefully extract the supernatant containing functional tubulin into a microcentrifuge tube. Add GMPCPP to a final concentration of 0.6 mM to induce tubulin polymerization and DTT to a final concentration of 1 mM to prevent protein aggregation.</li>
		<li>Incubate the mixture in the heat bath at 37 &#176;C for 30 min, then centrifuge again for 10 min at 14,000 x g at room temperature. Remove the supernatant, then dilute the pellet with M2B buffer to reach a final microtubule concentration of 6 mg/mL. 
		NOTE: This solution can be stored at room temperature for at least 4 h before use.</li>
		<li>To check that the microtubules have polymerized successfully, dilute 1 &#181;L of the microtubule solution 100:1 with M2B buffer and pipette on a microscope slide. Cover with a cover slip for imaging using a fluorescence microscope (see Table of Materials) with a 40x objective lens. 
		NOTE: Excitation and emission wavelengths are chosen according to the fluorescence labeling used on the microtubules. In this protocol, rhodamine labeling is used (see step 1.2.2), so imaging is carried out using an excitation band of 515-560 nm and a 590 nm long pass filter. Figure 1 shows a representative example.</li>
		<li>After polymerization is complete, drop-freeze the microtubules as 2 &#181;L aliquots in liquid nitrogen and store at -80 &#176;C (if required).</li>
	</ol>
	</li>
	<li>Preparation of MIX 
	NOTE: MIX is an aqueous solution that includes the kinesin-streptavidin clusters (KSA). Prepared MIX should be stored at -80 &#176;C in 4 &#181;L aliquots prior to the experiment. When MIX is combined with ATP and the microtubule solution described in step 1.2 at room temperature, activity is initiated. MIX is prepared as follows13,19.
	<ol>
		<li>Prepare a solution for preventing fluorescence fading (ANTIFADE) by mixing two antioxidant solutions. Combine AO1 (250 mM DTT, 65 mM catalase) and AO2 (750 &#181;M catalase, 3 mM glucose oxidase) in a 1:1 volume ratio. Include 20 mM Trolox, another antioxidant used to reduce damage caused by fluorescence microscopy.</li>
		<li>Prepare MIX by making a solution of KSA (described in step 1.1.3) that includes 6 wt% 20 kDa polyethylene glycol (PEG) to induce bundling of the microtubules, 3 vol% ANTIFADE, and 5 vol% pyruvate kinase/lactic dehydrogenase (PKLDH) at 70 mg/mL for ATP regeneration.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>Flow cell method for active nematic formation
	<ol>
		<li>Prepare hydrophilic coverslips with an acrylamide coating.
		<ol>
			<li>Clean the coverslips thoroughly with soapy water, ethanol, and 0.1 M NaOH with alternate rinses using nanopure water. Once rinsed, coat the coverslips with a silane solution composed of 100 mL of ethanol, 1 mL of acetic acid, and 500 &#181;L of 3-(trimethoxysilyl) propylmethacrylate for 15 min, then rinse with nanopure water.</li>
			<li>Prepare an acrylamide solution from 95 mL of nanopure water and 5 mL of 40 wt% acrylamide, then degas the solution for 30 min in a vacuum oven.</li>
			<li>Add 35 &#956;L of tetramethylethylenediamine (TEMED) for a 2.3 mM final concentration and then 0.07 g of ammonium persulfate.&#160;Pour the acrylamide solution over the coverslips while face up and incubate overnight at room temperature.</li>
		</ol>
		</li>
		<li>Prepare hydrophobic microscope slides. Pipette 100 &#181;L of a water repellent solution (see Table of Materials) onto a clean glass microscope slide, then place another clean glass slide on top. This ensures an even coating of the water repellent solution on the surface where it sits for 2 min. After 2 min, remove the second glass slide and rinse the first slide thoroughly with nano-pure water, then dry with nitrogen gas.&#160;Gently clean the region where the tape will be placed (around the pattern) with acetone to ensure the water repellent solution does not prevent adhesion to the glass.</li>
		<li>Prepare a mixture of engineered oil that includes 1.8% (v/v) 008-FluoroSurfactant (see Table of Materials).</li>
		<li>Assemble the glass slide and coverslip with the hydrophobic glass slide on the bottom of the flow cell and the hydrophilic cover slip as the upper surface in a sandwich geometry using 40 &#181;m double-sided adhesive spacers. Place the spacers 1.5 mm apart on the hydrophobic microscope slide. Then place the acrylamide coated coverslip on top of the spacers with the treated side face down to adhere.&#160;Ensure the adhesion is complete with pressure from a blunt object. 
		NOTE: The goal is to assemble a flow cell with a flat oil/water interface inside (Figure 2A,B). This is where the active layer will form. Alternative spacer tapes and films can also be used to produce a similar cell thickness.</li>
		<li>After the flow cell has been constructed, immediately pipette the oil mixture into the flow cell, filling the enclosed space.</li>
		<li>Using a pipette, in a separate vial gently mix 6 &#181;L of active material with 3.73 &#181;L of MIX, 1 &#181;L of microtubule solution, 0.6 &#181;L of ATP solution (the concentration can be varied to vary microtubule velocity), and 0.67 &#181;L of M2B buffer.</li>
		<li>Pipette freshly mixed active material into one open end of the flow cell, volumes will vary but should exceed the volume of the flow cell (roughly 3-6 &#956;L).&#160;Some oil will be displaced by the aqueous solution as it is injected into the channel; this can be wicked up at the opposite end of the flow channel using a small piece of tissue paper.</li>
		<li>After filling, seal both sides of the flow cell with an epoxy glue (see Table of Materials) that hardens when exposed to UV light for 20 s. 
		NOTE: At this point, the active material forms a 3D network that remains suspended in the water layer.</li>
		<li>To confine the active layer between the two immiscible fluids in a quasi-2D layer, place the flow cell in a swinging bucket centrifuge (see Table of Materials) with the aqueous phase on top and the denser oil layer underneath. Centrifuge at 212 x g for 10 min. After this step is complete, the flow cell can be taken to an epi-fluorescence microscope for imaging with a 10x or 20x magnification objective. Figure 2C,D shows typical images before and after this step.</li>
	</ol>
	</li>
	<li>Inverted method for active nematic formation 
	NOTE: An alternate method to that described in step 2.1 is to assemble the active nematic layer underneath a thick oil layer confined to a deep PDMS well28. This method produces similar results, and is somewhat easier to master; however, image quality using this method is usually not as good as the flow cell method.
	<ol>
		<li>Prepare the polydimethylsiloxane (PDMS) using an elastomer curing agent and an elastomer base (see Table of Materials). Mix the two components in a 1:10 ratio using a metal spatula. Tiny bubbles appear during mixing which are difficult to remove, and the mixture appears milky. To remove these bubbles, place the mixture under a vacuum to degas for 1 h, after which the uncured PDMS should appear transparent. 
		NOTE: The PDMS is now ready to form any shape using a mold, or it can cure in the container and then be cut or punched in the desired pattern.</li>
		<li>Pour the PDMS into a suitable mold and leave overnight to cure at 60 &#176;C. 
		NOTE: Untreated, the surface of cured PDMS is hydrophobic, but with surface treatment, it can be made hydrophilic.</li>
		<li>To prepare a hydrophilic PDMS surface, coat the PDMS with an acrylamide polymer brush29. This step also prevents proteins from sticking to the surface.
		<ol>
			<li>Start by cleaning the PDMS for 10 min with both ethanol and isopropanol, then rinse thoroughly with deionized water 3x and dry. Use a plasma cleaner for 5 min to clean the dry, cured PDMS. This step makes the surface more hydrophilic.</li>
			<li>Next, prepare a silane solution (98.5 wt% ethanol, 1 wt% acetic acid, and 0.5 wt% Trimethoxysilyl propyl methacrylate) and immerse the substrate in that solution for 15 min to prepare for the acrylamide coating. Rinse the substrate thoroughly with deionized water and immerse in acrylamide solution (2 wt% acrylamide/bis solution, 2.3 mM TEMED, and 3 mM ammonium persulfate). 
			NOTE: These substrates can be stored at room temperature covered in acrylamide solution in a glass Petri dish, and should be used within 2 weeks.</li>
		</ol>
		</li>
		<li>When ready for use, rinse the surface with deionized water and dry with nitrogen for immediate use. Add the active mixture (described in step 2.1.6) to the wells and immediately add silicone oil on top to a thickness of approximately 2 mm. 
		NOTE: At this stage, the active mixture will be sandwiched between the oil and the hydrophilic coating on the PDMS, but it will still be somewhat three-dimensional.</li>
		<li>To push the material further into a 2D layer, glue the PDMS device onto a glass slide, place it in a swinging bucket centrifuge and spin for 12 min at 212 x g. The device will need to be positioned such that the silicone oil is on the top of the aqueous layer. Representative results are shown in Figure 3.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>First, design a master mold for the PDMS. This can be achieved by 3D printing pillars on a substrate. After 3D printing the resin master mold, clean with isopropanol and then cure the mold under a UV lamp for 45 min and in the oven at 120 &#176;C for 2 h (Figure 4). 
	NOTE: Curing under UV and thermal post-curing improves the quality of replication in PDMS by removing monomers and photo inhibitor residues from the resin30.</li>
	<li>Use the master mold to create wells from PDMS. Prepare the PDMS as described in step 2.2.1. Immerse the master mold in uncured PDMS and cure overnight in an oven at 60 &#176;C.</li>
	<li>After PDMS curing is complete, carefully remove the master mold and cut the PDMS as desired to work with the wells (Figure 4). Treat the PDMS surface as described in step 2.2.3. Before the experiment, the surface can be attached to a glass slide with epoxy glue to make imaging easier.</li>
	<li>Pipette 1 &#181;L of the active mixture described in step 2.1.6 onto the PDMS substrate and immediately add silicone oil with 100-1000 cSt viscosity28 on top of the active network droplet. The active network will move into the well; this process takes up to 60 min (Figure 4). As described in step 2.2.5, the 2D network can be enhanced by spinning down the PDMS well in the swinging bucket centrifuge for 12 min at 212 x g.</li>
</ol>

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Also, how to avoid it.</a> Analytical Chemistry. 93 (19), 7180-7187 (2021).</li><li>Khaladj, D. A., Hirst, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+curved+fluid+boundaries+to+confine+active+nematic+flows.">Using curved fluid boundaries to confine active nematic flows.</a> Frontiers of Physics. 10, 880941 (2022).</li></ol>

Some materials used in this work were provided free of charge by Cytoskeleton Inc. (Denver, USA).

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 &#181;m 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 &#181;m 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&#160;1 and Movie 2,&#160;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.

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 &#8212; a state in which the fluid produces self-driven chaotic flows19&#160;&#8212; 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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>20 kD PEG (polyethylene glycol))</td>
			<td>Sigma Aldrich</td>
			<td>1419109</td>
			<td>Depletion agent 
			CAS Number: 125061-88-3</td>
		</tr>
		<tr>
			<td>3-(trimethoxysilyl)propyl methacrylate</td>
			<td>Sigma Aldrich</td>
			<td>M6514-50ML</td>
			<td>CAS Number: 2530-85-0</td>
		</tr>
		<tr>
			<td>3D printer &#38; Resin</td>
			<td>Phrozen</td>
			<td></td>
			<td>Phrozen sonic mini 8K 3D printer - Aqua Gray 8K resin</td>
		</tr>
		<tr>
			<td>40% Acrylamide Solution</td>
			<td>BIO-RAD</td>
			<td>1610140</td>
			<td>CAS Number: 7732-18-5, 79-06-1</td>
		</tr>
		<tr>
			<td>Acetic Acid</td>
			<td>Fisher</td>
			<td></td>
			<td>CAS Number: 64-19-7</td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CAS Number: 67-64-1</td>
		</tr>
		<tr>
			<td>Adhesive sheets (NOTE: &#34;Parafilm&#34; is an alternative)</td>
			<td>Grace Bio-Labs</td>
			<td>620001</td>
			<td>SecureSeal</td>
		</tr>
		<tr>
			<td>Ammonium Persulfate</td>
			<td>Sigma Aldrich</td>
			<td>A3678</td>
			<td>CAS Number: 7727-54-0</td>
		</tr>
		<tr>
			<td>Aquapel (NOTE: &#34;RainX&#34; is an alternative)</td>
			<td>Aquapel Glass Treatment</td>
			<td></td>
			<td>hydrophobic glass treatment</td>
		</tr>
		<tr>
			<td>ATP (Adenosine triphosphate)</td>
			<td>Sigma Aldrich</td>
			<td>A1852</td>
			<td>CAS Number: 34369-07-8</td>
		</tr>
		<tr>
			<td>Beakers</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Catalase</td>
			<td>Sigma Aldrich</td>
			<td>C9322</td>
			<td>CAS Number: &#34;9001-05-2&#34;</td>
		</tr>
		<tr>
			<td>Desiccator</td>
			<td>Bel-art</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Digital CMOS camera</td>
			<td>Hamamatsu</td>
			<td>ORCA - Flash4.0 LT+</td>
			<td></td>
		</tr>
		<tr>
			<td>DTT (Dithiothreitol)</td>
			<td>Sigma Aldrich</td>
			<td>D9779</td>
			<td>CAS Number: &#34;3483-12-3&#34;</td>
		</tr>
		<tr>
			<td>EGTA (3,12-bis(carboxymethyl)-6,9-dioxa-3,12-diazatetradecane-1,14-dioic acid)</td>
			<td>Sigma Aldrich</td>
			<td>MFCD00004291</td>
			<td>CAS Number: 67-42-5</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CAS Number: 64-17-5</td>
		</tr>
		<tr>
			<td>Fluorescence microscope</td>
			<td>Leica</td>
			<td>DM 2500P</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Coverslips</td>
			<td>VWR</td>
			<td>48368-040</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Slides</td>
			<td>VWR</td>
			<td>16004-430</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma Aldrich</td>
			<td>G7021</td>
			<td>CAS Number: 50-99-7</td>
		</tr>
		<tr>
			<td>Glucose Oxidase</td>
			<td>Sigma Aldrich</td>
			<td>345386</td>
			<td>CAS Number: 9001-37-0</td>
		</tr>
		<tr>
			<td>GMPCPP (guanylyl 5&#39;-&#945;,&#946;-methylenediphosphonate)</td>
			<td>Jena Bioscience</td>
			<td>NU-405S</td>
			<td>CAS Number: 14997-54-7</td>
		</tr>
		<tr>
			<td>HFE7500 Oil</td>
			<td>3M</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hot Plate</td>
			<td>Fisher Scientific</td>
			<td>Thermix hot plate model 100M</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl Alcohol</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KCl (potassium chloride)</td>
			<td>Sigma Aldrich</td>
			<td>P5405</td>
			<td>CAS Number: 7447-40-7</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CAS Number: 67-56-1</td>
		</tr>
		<tr>
			<td>MgCl2 (Magnesium Chloride)</td>
			<td>Sigma Aldrich</td>
			<td>208337</td>
			<td>CAS Number: 7786-30-3</td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes</td>
			<td>Eppendorf - Thermo Fisher</td>
			<td>1.5 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanopure water purifier</td>
			<td>Sartorius</td>
			<td>arium mini</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH (Sodium hydroxide)</td>
			<td>Sigma Aldrich</td>
			<td>SX0603</td>
			<td>CAS Number: 1310-73-2</td>
		</tr>
		<tr>
			<td>Petri Dishes</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PH Meter</td>
			<td>Thermo Scientist</td>
			<td>Orion 3 STAR</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphoenol-pyruvate (PEP)</td>
			<td>Sigma Aldrich</td>
			<td>MFCD00044476</td>
			<td>CAS Number: 4265-07-0</td>
		</tr>
		<tr>
			<td>PIPES (1,4-Piperazinediethanesulfonic acid)</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CAS Number: 5625-37-6</td>
		</tr>
		<tr>
			<td>Pipettes (0.2 - 1000 &#181;l)</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Sigma Aldrich</td>
			<td>2594628</td>
			<td></td>
		</tr>
		<tr>
			<td>RAN Surfactant (NOTE: &#34;FluoSurf&#34; from Emulso is an alternative)</td>
			<td>Ran Biotechnologies</td>
			<td>008-FluoroSurfactant-2wtH-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon Oil (100mpa s-1000 mpa s)</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td>CAS Number: 63148-52-7</td>
		</tr>
		<tr>
			<td>Streptavidin</td>
			<td>Thermofisher</td>
			<td>S888</td>
			<td></td>
		</tr>
		<tr>
			<td>Swinging Bucket Centrifuge</td>
			<td>Thermo Scientist</td>
			<td>Sorvall legend RT+</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 Elastomer base</td>
			<td>World Precision Instruments</td>
			<td>SYLG184</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 Elastomer Curing agent</td>
			<td>World Precision Instruments</td>
			<td>SYLG184</td>
			<td></td>
		</tr>
		<tr>
			<td>Table top centrifuge</td>
			<td>Eppendorf</td>
			<td>MiniSpin Plus</td>
			<td></td>
		</tr>
		<tr>
			<td>TEMED (Tetramethylethylenediamine)</td>
			<td>BIO-RAD</td>
			<td>1610800</td>
			<td>CAS Number: 110-18-9</td>
		</tr>
		<tr>
			<td>Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid)</td>
			<td>Sigma Aldrich</td>
			<td>MFCD00006846</td>
			<td>CAS Number: 53188-07-1</td>
		</tr>
		<tr>
			<td>Tubulin</td>
			<td>Cytoskeleton</td>
			<td>T240-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubulin (Rhodamine labeled)</td>
			<td>Cytoskeleton</td>
			<td>TL590M-A</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td>Beckman</td>
			<td>Optima Max-TL</td>
			<td></td>
		</tr>
		<tr>
			<td>UV Light</td>
			<td>RapidFix</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>UV-curable glue (NOTE: &#34;Norland NO81&#34; is an alternative)</td>
			<td>RapidFix</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Water Bath</td>
			<td>Thelco</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman Filter paper</td>
			<td>Sigma Aldrich</td>
			<td>WHA1001325</td>
			<td></td>
		</tr>
	</tbody>
</table>

forming, confining, and observing microtubule-based active nematics

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.

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&#160;2 and Figure 3, three supplemental movies (Movie 1, Movie 2,&#160;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.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64287/64287fig01.jpg" /> 
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 &#176;C. Imaging was performed at room temperature. Scale bar = 10 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/64287fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64287/64287fig02.jpg" /> 
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 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/64287fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64287/64287fig03.jpg" /> 
Figure 3: Fluorescence microscope image showing an active nematic prepared using the inverted method. Scale bar = 200 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/64287fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64287/64287fig04.jpg" /> 
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 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/64287fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Movie 1: Representative result for active nematic prepared using the flow cell method. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/MOVIE1_JOVE.avi" target="_blank">Please click here to download this Movie.</a>
Movie 2: Representative result for active nematic prepared using the inverted method. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/MOVIE2_JOVE.avi" target="_blank">Please click here to download this Movie.</a>
Movie 3: Representative result for active nematic prepared using the inverted method confined to an elliptical well. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/64287/MOVIE3_JOVE.avi" target="_blank">Please click here to download this Movie.</a>

Watch this Scientific Journal Video about Forming, Confining, and Observing Microtubule-Based Active Nematics at JoVE.com

Forming, Confining, and Observing Microtubule-Based Active Nematics

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 ...

forming-confining-and-observing-microtubule-based-active-nematics

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

2.5K Views.  University of California. 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.

Video: Forming, Confining, and Observing Microtubule-Based Active Nematics

Induction and Monitoring of Active Delayed Type Hypersensitivity (DTH) in Rats

Delayed type hypersensitivity (DTH) is an inflammatory reaction mediated by CCR7- effector memory T lymphocytes that infiltrate the site of injection of an antigen against which the immune system has been primed. The inflammatory reaction is characterized by redness and swelling of the site of antigenic challenge. It is a convenient model to determine the in vivo efficacy of immunosuppressants. Cutaneous DTH can be induced either by adoptive transfer of antigen-specific T lymphocytes or by active immunization with an antigen, and subsequent intradermal challenge with the antigen to induce the inflammatory reaction in a given skin area. DTH responses can be induced to various antigens, for example ovalbumin, tuberculin, tetanus toxoid, or keyhole limpet hemocyanin (KLH).
Here we demonstrate how to induce an active DTH reaction in Lewis rats. We will first prepare a water-in-oil emulsion of KLH, our antigen of interest, in complete Freund's adjuvant and inject this emulsion subcutaneously to rats. This will prime the immune system to develop memory T cells directed to KLH. Seven days later we will challenge the rats intradermally on the back with KLH on one side and with ovalbumin, an irrelevant antigen, on the other side. The inflammatory reaction will be visible 16-72 hours later and the red and swollen area will be measured as an indication of DTH severity.

Induction and Monitoring of Active Delayed Type Hypersensitivity DTH in Rats

Delayed type hypersensitivity (DTH) is an inflammatory reaction mediated by CCR7- effector memory T lymphocytes. Here we demonstrate how to induce active DTH in Lewis rats and monitor the inflammatory response.

Delayed type hypersensitivity (DTH) is an inflammatory reaction mediated by CCR7- effector memory T lymphocytes that infiltrate the site of injection of ...

Isolation and Large Scale Expansion of Adult Human Endothelial Colony Forming Progenitor Cells

This paper introduces a novel recovery strategy for endothelial colony forming progenitor cells (ECFCs) from heparinized but otherwise unmanipulated adult human peripheral blood within a mean of 12 days. After large scale expansion &gt;1x108 ECFCs can be obtained for further tests. Advantageously by using pHPL the contact of human cells with bovine serum antigens can be excluded. By flow cytometry and immunohistochemistry the isolated cells can be characterized as ECFC and their in vitro functionality to form vascular like structures can be tested in a matrigel assay. Further these cells can be subcutaneously injected in a mouse model to form functional, perfused vessels in vivo. After long term expansion and cryopreservation proliferation, function and genomic stability appear to be preserved. 3,4
This animal-protein free isolation and expansion method is easily applicable to generate a large quantity of ECFCs.

Endothelial colony forming progenitor cells (ECFCs) are a promising tool to study vascular homeostasis and repair.1,2 This paper introduces a novel animal-serum free method for isolation and expansion of ECFC from heparinised adult human peripheral blood with pooled human platelet lysate (pHPL) diminishing the risk of anti-bovine immunisation.

This paper introduces a novel recovery strategy for endothelial colony forming progenitor cells (ECFCs) from heparinized but otherwise unmanipulated adult ...

Isolation and Animal Serum Free Expansion of Human Umbilical Cord Derived Mesenchymal Stromal Cells (MSCs) and Endothelial Colony Forming Progenitor Cells (ECFCs)

The umbilical cord is a rich source for progenitor cells with high proliferative potential including mesenchymal stromal cells (also termed mesenchymal stem cells, MSCs) and endothelial colony forming progenitor cells (ECFCs). Both cell types are key players in maintaining the integrity of tissue and are probably also involved in regenerative processes and tumor formation.
To study their biology and function in a comparative manner it is important to have both cells types available from the same donor. It may also be beneficial for regenerative purposes to derive MSCs and ECFCs from the same tissue.
Because cellular therapeutics should eventually find their way from bench to bedside we established a new method to isolate and further expand progenitor cells without the use of animal protein. Pooled human platelet lysate (pHPL) replaced fetal bovine serum in all steps of our protocol to completely avoid contact of the cells to xenogeneic proteins.
This video demonstrates a methodology for the isolation and expansion of progenitor cells from one umbilical cord.
All materials and procedures will be described.

Isolation and Animal Serum Free Expansion of Human Umbilical Cord Derived Mesenchymal Stromal Cells MSCs and Endothelial Colony Forming Progenitor Cells ECFCs

This protocol describes the isolation and subsequent expansion of mesenchymal stromal cells and endothelial colony forming cells without the use of animal serum to generate autologous pairs for experimental transplantation purposes.

The umbilical cord is a rich source for progenitor cells with high proliferative potential including mesenchymal stromal cells (also termed mesenchymal ...

Obtaining Hemocytes from the Hawaiian Bobtail Squid Euprymna scolopes and Observing their Adherence to Symbiotic and Non-Symbiotic Bacteria

Studies concerning the role of the immune system in mediating molecular signaling between beneficial bacteria and their hosts have, in recent years, made significant contributions to our understanding of the co-evolution of eukaryotes with their microbiota. The symbiotic association between the Hawaiian bobtail squid, Euprymna scolopes and the bioluminescent bacterium Vibrio fischeri has been utilized as a model system for understanding the effects of beneficial bacteria on animal development. Recent studies have shown that macrophage-like hemocytes, the sole cellular component of the squid host's innate immune system, likely play an important role in mediating the establishment and maintenance of this association. This protocol will demonstrate how to obtain hemocytes from E. scolopes and then use these cells in bacterial binding assays. Adult squid are first anesthetized before hemolymph is collected by syringe from the main cephalic blood vessel. The host hemocytes, contained in the extracted hemolymph, are adhered to chambered glass coverslips and then exposed to green fluorescent protein-labeled symbiotic Vibrio fischeri and non-symbiotic Vibrio harveyi. The hemocytes are counterstained with a fluorescent dye (Cell Tracker Orange, Invitrogen) and then visualized using fluorescent microscopy.

Obtaining Hemocytes from the Hawaiian Bobtail Squid Euprymna scolopes and Observing their Adherence to Symbiotic and Non-Symbiotic Bacteria

This video will demonstrate how to obtain hemocytes (blood cells) from the Hawaiian bobtail squid, Euprymna scolopes for use in cell biological and bacterial adhesion assays. Hemocytes will be stained with a fluorescent dye and exposed to GFP-labeled bacteria.

Studies concerning the role of the immune system in mediating molecular signaling between beneficial bacteria and their hosts have, in recent years, made ...

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.
Telomerase is a specialized reverse transcriptase1 that adds short DNA repeats, called telomeres, to the 3' end of linear chromosomes2 that serve to protect them from end-to-end fusion and degradation. Following DNA replication, a short segment is lost at the end of the chromosome3 and without telomerase, cells continue dividing until eventually reaching their Hayflick Limit4. Additionally, telomerase is dormant in most somatic cells5 in adults, but is active in cancer cells6 where it promotes cell immortality7.
The minimal telomerase enzyme consists of two core components: the protein subunit (TERT), which comprises the catalytic subunit of the enzyme and an integral RNA component (TER), which contains the template TERT uses to synthesize telomeres8,9. Prior to 2008, only structures for individual telomerase domains had been solved10,11. A major breakthrough in this field came from the determination of the crystal structure of the active12, catalytic subunit of T. castaneum telomerase, TcTERT1.
Here, we present methods for producing large quantities of the active, soluble TcTERT for structural and biochemical studies, and the reconstitution of the telomerase RNP complex in vitro for telomerase activity assays. An overview of the experimental methods used is shown in Figure 1.

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more ...

A Quantitative Assay for Insulin-expressing Colony-forming Progenitors

The field of pancreatic stem and progenitor cell biology has been hampered by a lack of in vitro functional and quantitative assays that allow for the analysis of the single cell. Analyses of single progenitors are of critical importance because they provide definitive ways to unequivocally demonstrate the lineage potential of individual progenitors. Although methods have been devised to generate "pancreatospheres" in suspension culture from single cells, several limitations exist. First, it is time-consuming to perform single cell deposition for a large number of cells, which in turn commands large volumes of culture media and space. Second, numeration of the resulting pancreatospheres is labor-intensive, especially when the frequency of the pancreatosphere-initiating progenitors is low. Third, the pancreatosphere assay is not an efficient method to allow both the proliferation and differentiation of pancreatic progenitors in the same culture well, restricting the usefulness of the assay.
To overcome these limitations, a semi-solid media based colony assay for pancreatic progenitors has been developed and is presented in this report. This method takes advantage of an existing concept from the hematopoietic colony assay, in which methylcellulose is used to provide viscosity to the media, allowing the progenitor cells to stay in three-dimensional space as they undergo proliferation as well as differentiation. To enrich insulin-expressing colony-forming progenitors from a heterogeneous population, we utilized cells that express neurogenin (Ngn) 3, a pancreatic endocrine progenitor cell marker. Murine embryonic stem (ES) cell-derived Ngn3 expressing cells tagged with the enhanced green fluorescent protein reporter were sorted and as many as 25,000 cells per well were plated into low-attachment 24-well culture dishes. Each well contained 500 &mu;L of semi-solid media with the following major components: methylcellulose, Matrigel, nicotinamide, exendin-4, activin &beta;B, and conditioned media collected from murine ES cell-derived pancreatic-like cells. After 8 to 12 days of culture, insulin-expressing colonies with distinctive morphology were formed and could be further analyzed for pancreatic gene expression using quantitative RT-PCR and immunoflourescent staining to determine the lineage composition of each colony.
In summary, our colony assay allows easy detection and quantification of functional progenitors within a heterogeneous population of cells. In addition, the semi-solid media format allows uniform presentation of extracellular matrix components and growth factors to cells, enabling progenitors to proliferate and differentiate in vitro. This colony assay provides unique opportunities for mechanistic studies of pancreatic progenitor cells at the single cell level.

A three-dimensional clonogenic assay that allows pancreatic-like progenitors to differentiate into insulin-expressing colonies is described. This method takes advantage of semi-solid media containing methylcellulose, Matrigel and growth factors, in which single progenitors proliferate and differentiate in vitro, permitting quantification of the number of functional progenitors in a population.

The field of pancreatic stem and progenitor cell biology has been hampered by a lack of in vitro functional and quantitative assays that allow for the ...

Chromatographic Purification of Highly Active Yeast Ribosomes

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers. Particularly troublesome is the fact that lysis of cells releases a large number of proteases and nucleases which can degrade ribosomes. Thus, it is important to separate ribosomes from these enzymes as quickly as possible. Unfortunately, conventional differential ultracentrifugation methods leaves ribosomes exposed to these enzymes for unacceptably long periods of time, impacting their structural integrity and functionality. To address this problem, we utilize a chromatographic method using a cysteine charged Sulfolink resin. This simple and rapid application significantly reduces co-purifying proteolytic and nucleolytic activities, producing high yields of intact, highly biochemically active yeast ribosomes. We suggest that this method should also be applicable to mammalian ribosomes. The simplicity of the method, and the enhanced purity and activity of chromatographically purified ribosome represents a significant technical advancement for the study of eukaryotic ribosomes.

Contamination of preparations of eukaryotic ribosomes purified by traditional methods by co-purifying nucleases and proteases negatively impacts on downstream biochemical and structural analyses. A rapid and simple chromatographic purification method is used to solve this problem using yeast ribosomes as a model system.

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers.  ...

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Phenotypic and Functional Characterization of Endothelial Colony Forming Cells Derived from Human Umbilical Cord Blood

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of circulating endothelial progenitor cells (EPCs) were first identified in adult human peripheral blood by Asahara et al. in 1997 2 bringing an infusion of new hypotheses and strategies for vascular regeneration and repair. EPCs are rare but normal components of circulating blood that home to sites of blood vessel formation or vascular remodeling, and facilitate either postnatal vasculogenesis, angiogenesis, or arteriogenesis largely via paracrine stimulation of existing vessel wall derived cells3. No specific marker to identify an EPC has been identified, and at present the state of the field is to understand that numerous cell types including proangiogenic hematopoietic stem and progenitor cells, circulating angiogenic cells, Tie2+ monocytes, myeloid progenitor cells, tumor associated macrophages, and M2 activated macrophages participate in stimulating the angiogenic process in a variety of preclinical animal model systems and in human subjects in numerous disease states4, 5. Endothelial colony forming cells (ECFCs) are rare circulating viable endothelial cells characterized by robust clonal proliferative potential, secondary and tertiary colony forming ability upon replating, and ability to form intrinsic in vivo vessels upon transplantation into immunodeficient mice6-8. While ECFCs have been successfully isolated from the peripheral blood of healthy adult subjects, umbilical cord blood (CB) of healthy newborn infants, and vessel wall of numerous human arterial and venous vessels 6-9, CB possesses the highest frequency of ECFCs7 that display the most robust clonal proliferative potential and form durable and functional blood vessels in vivo8, 10-13. While the derivation of ECFC from adult peripheral blood has been presented14, 15, here we describe the methodologies for the derivation, cloning, expansion, and in vitro as well as in vivo characterization of ECFCs from the human umbilical CB.

Endothelial colony forming cells (ECFCs) are circulating endothelial cells with robust clonal proliferative potential that display intrinsic in vivo vessel forming ability. Phenotypic and functional characterization of outgrowth endothelial cells derived from CB are important to identify and isolate bona fide ECFCs for potential clinical application in repairing damaged tissues.

Longstanding views of new blood vessel formation via angiogenesis, vasculogenesis, and arteriogenesis have been recently reviewed1. The presence of ...

Affinity Precipitation of Active Rho-GEFs Using a GST-tagged Mutant Rho Protein (GST-RhoA(G17A)) from Epithelial Cell Lysates

Proteins of the Rho family of small GTPases are central regulators of the cytoskeleton, and control a large variety of cellular processes, including cell migration, gene expression, cell cycle progression and cell adhesion 1. Rho proteins are molecular switches that are active in GTP-bound and inactive in GDP-bound state. Their activation is mediated by a family of Guanine-nucleotide Exchange Factor (GEF) proteins. Rho-GEFs constitute a large family, with overlapping specificities 2. Although a lot of progress has been made in identifying the GEFs activated by specific signals, there are still many questions remaining regarding the pathway-specific regulation of these proteins. The number of Rho-GEFs exceeds 70, and each cell expresses more than one GEF protein. In addition, many of these proteins activate not only Rho, but other members of the family, contributing further to the complexity of the regulatory networks. Importantly, exploring how GEFs are regulated requires a method to follow the active pool of individual GEFs in cells activated by different stimuli. Here we provide a step-by-step protocol for a method used to assess and quantify the available active Rho-specific GEFs using an affinity precipitation assay. This assay was developed a few years ago in the Burridge lab 3,4 and we have used it in kidney tubular cell lines 5,6,7. The assay takes advantage of a "nucleotide free" mutant RhoA, with a high affinity for active GEFs. The mutation (G17A) renders the protein unable to bind GDP or GTP and this state mimics the intermediate state that is bound to the GEF. A GST-tagged version of this mutant protein is expressed and purified from E. coli, bound to glutathione sepharose beads and used to precipitate active GEFs from lysates of untreated and stimulated cells. As most GEFs are activated via posttranslational modifications or release from inhibitory bindings, their active state is preserved in cell lysates, and they can be detected by this assay8. Captured proteins can be probed for known GEFs by detection with specific antibodies using Western blotting, or analyzed by Mass Spectrometry to identify unknown GEFs activated by certain stimuli.

Affinity Precipitation of Active Rho-GEFs Using a GST-tagged Mutant Rho Protein GST-RhoAG17A from Epithelial Cell Lysates

The method presented here describes an assay to follow activation of RhoA specific GDP/GTP Exchange Factors (GEFs) in cultured cells by making use of a mutant RhoA GST fusion protein that has high affinity for activated GEFs. GEFs are precipitated from cell lysates, detected by Western blotting and quantified by densitometry.

Proteins of the Rho family of small GTPases are central regulators of the cytoskeleton, and control a large variety of cellular processes, including cell ...

Forming, Confining, and Observing Microtubule-Based Active Nematics

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Chapters in this video

Induction and Monitoring of Active Delayed Type Hypersensitivity (DTH) in Rats

Isolation and Large Scale Expansion of Adult Human Endothelial Colony Forming Progenitor Cells

Isolation and Animal Serum Free Expansion of Human Umbilical Cord Derived Mesenchymal Stromal Cells (MSCs) and Endothelial Colony Forming Progenitor Cells (ECFCs)

Obtaining Hemocytes from the Hawaiian Bobtail Squid Euprymna scolopes and Observing their Adherence to Symbiotic and Non-Symbiotic Bacteria

In vitro Reconstitution of the Active T. castaneum Telomerase

A Quantitative Assay for Insulin-expressing Colony-forming Progenitors

Chromatographic Purification of Highly Active Yeast Ribosomes

Direct Detection of the Acetate-forming Activity of the Enzyme Acetate Kinase

Phenotypic and Functional Characterization of Endothelial Colony Forming Cells Derived from Human Umbilical Cord Blood

Affinity Precipitation of Active Rho-GEFs Using a GST-tagged Mutant Rho Protein (GST-RhoA(G17A)) from Epithelial Cell Lysates