Name: use of stopped-flow fluorescence and labeled nucleotides to analyze the atp turnover cycle of kinesins
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Description: Watch this Scientific Journal Video about Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins at JoVE.com

This work is supported by the Biotechnology and Biological Sciences Research Council (BBSRC), the Royal Society and the University of Nottingham.

1. Determination of the Fluorescence Intensity Change upon Binding of Mant-labeled Nucleotide.
<ol>
	<li>Add 1 &#956;M mant-labeled nucleotide to 1 &#956;M kinesin (final concentrations) in an appropriate reaction buffer. Typically a buffer that benefits microtubule growth and stability is used (see discussion). A commonly used buffer contains 80mM piperazine-bis-2-ethanesulfonic acid (PIPES) pH6.9, 1mM MgCl2 and 1mM ethylene glycol-bis-2 aminoether-tetraacetic acid (EGTA). 
	Note: The concentration of kinesin stated in all protocols refers to the concentration of nucleotide binding sites (i.e., motor domains). Different kinesins exist as monomers, dimers or tetramers and so can contain more than one nucleotide-binding site per molecule.</li>
	<li>After an incubation period (1-3 min) and using a standard fluorimeter, measure the emission spectrum for the mant-labeled nucleotide-kinesin complex. Use an excitation wavelength of 365nm and measure emitted light from 400-500 nm in 1nm increments.</li>
	<li>Make a solution of 1mM mant-labeled nucleotide in the same reaction buffer used in Step 1.1.</li>
	<li>Measure the emission spectrum for the mant nucleotide only sample, as previously described (see Step 1.2).</li>
	<li>Normalize the spectra to the signal at the wavelength of peak fluorescence intensity of the spectrum of mant nucleotide in solution (i.e., without kinesin) by dividing through by this value.</li>
	<li>Plot the normalized spectra (e.g., Figure 2B) to observe fluorescence intensity differences between the mantATP-bound and/or mantADP-bound kinesin and mant nucleotide in solution.</li>
</ol>
2. Measurement of the Association and Dissociation Rate Constants for mantATP
<ol>
	<li>To a solution of up to 20 &#956;M kinesin (see discussion) in any appropriate Mg2+-free buffer (e.g., 80mM PIPES pH6.9, 1mM EGTA, 0.1% Tween20), add 1 mM (final concentration) ethylenediaminetetraacetic acid (EDTA) and 1 mM (final concentration) dithiothreitol (DTT). &#160; Note: Addition of Tween20 to the buffer is recommended as it reduces/prevents loss of kinesin protein on the column used for buffer exchange (Steps 2.3 and 3.3)</li>
	<li>Incubate the solution at 25 &#176;C for 15 min.</li>
	<li>Separate free nucleotide and EDTA from the kinesin using a gravity flow G-25 sephadex gel filtration column according to manufacturer&#8217;s instructions (see list of materials).
	<ol>
		<li>Equilibrate the column with at least 3 column volumes of an appropriate Mg2+-free buffer (see Step 2.1). Load the kinesin-containing solution onto the column.&#160;Prepare a collection tube by adding an appropriate volume of 100mM magnesium chloride solution to the empty tube, such that the final concentration of magnesium chloride is 1 mM after collection of the kinesin solution. &#160;Immediate addition of magnesium chloride helps stabilise the nucleotide-free kinesin.</li>
		<li>Elute the kinesin using&#160;an appropriate Mg2+-free buffer. Kinesins are unstable when free of nucleotide so it is important to work quickly. Store the nucleotide-free kinesin on ice and use within 1-2 hr.</li>
	</ol>
	</li>
	<li>During or prior to preparation of nucleotide-free kinesin, prepare a mantATP concentration series in the same reaction buffer as the final solution of nucleotide-free kinesin. Determine the range of concentrations of mantATP required according to the available concentration of kinesin with a typical series ranging from 0.1 to 50 &#956;M (see step 2.5 and discussion).</li>
	<li>Prepare a concentration series of nucleotide-free kinesin. For each concentration of mantATP (Step 2.4), a concentration of nucleotide-free kinesin should be prepared such that the mant-labeled nucleotide is in 5 to 10-fold molar excess over kinesin nucleotide binding sites.</li>
	<li>Set the excitation wavelength on the stopped-flow fluorimeter to 365 nm and collect emitted light at &#62;400 nm using a long-pass filter.</li>
	<li>Starting with the lowest concentration of mantATP, mix the mantATP with an appropriate concentration of nucleotide-free kinesin in a 1:1 v/v ratio using a stopped-flow fluorimeter (Figure 3A and 3B Inset). Note: The reactants are diluted by 50% upon mixing. Keep a note of the nucleotide concentration both pre and post mixing (i.e., 3.0/1.5 &#956;M) to avoid confusion.</li>
	<li>Fit the change in fluorescence intensity measured at each concentration of mantATP to an exponential function plus a line of constant negative slope to account for photobleaching of the mant group (i.e., Fluorescence = A0.exp(kobs.t)+(m.t+c), where A0 is the amplitude, kobs is the rate constant and t is time, see Figure 3B and discussion). From these fits determine a rate constant (kobs) at each concentration of mantATP. Each kobs is a convolution of an association and dissociation rate constant (Figure 1, k1 and k-1).</li>
	<li>Deconvolve the association and dissociation rate constants by plotting kobs against mantATP concentration (e.g., Figure 3C). Fit these data to a linear function (i.e., kobs = k1[mantATP] + k-1) to obtain the gradient and y-axis intercept. The gradient represents the rate constant for mantATP association (Figure 1, k1) with the units M-1s-1. The intercept represents the dissociation rate constant (Figure 1, k-1) with units of s-1 (see discussion). Note: This approach can also be applied to determine the ADP association and dissociation rate constants (Figure 1, k-4 and k4) by using mantADP as the nucleotide.</li>
</ol>
3. Measurement of the Dissociation Rate Constant for mantADP
<ol>
	<li>To a solution of 2 &#956;M kinesin in a suitable reaction buffer (e.g., 80 mM PIPES pH6.9, 1mM MgCl2, 1mM EGTA, 0.1% Tween20), add 50 &#956;M mantADP (final concentration).</li>
	<li>Incubate at 25 oC for 30 min.</li>
	<li>Remove excess mantADP and other free nucleotide by buffer exchanging the kinesin into the chosen reaction buffer using a G-25 sephadex column (see list of materials) according to manufacturer&#8217;s instructions. The kinesin motor domain should now be loaded with mantADP.</li>
	<li>Set the excitation wavelength on the stopped-flow fluorimeter to 365 nm and collect emitted light at &#62;400 nm using a long-pass filter.</li>
	<li>Mix the mantADP.kinesin complex (~1&#956;M) with a 50-fold or greater molar excess of unlabeled ATP in a 1:1 v/v ratio using a stopped-flow fluorimeter (Figure 4A and 4B Inset).</li>
	<li>Fit the decrease in fluorescence intensity observed over time to a single exponential function plus a line of constant negative slope to account for photobleaching of the mant group (i.e., Fluorescence = A0.exp(kobs.t)+(m.t+c), where A0 is the amplitude, kobs is the rate constant and t is time, see Figure 4B and discussion). The kobs determined from this fit is the rate constant for dissociation of ADP (Figure 1, k4) with units of s-1 (see discussion).</li>
</ol>

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The authors declare that they have no competing financial interests.

Here, we present protocols for observation and analysis of the kinetics of transitions between nucleotide states for a kinesin. These protocols utilize the rapid mixing method stopped-flow in conjunction with fluorescently labeled nucleotides. Methods of this type have been used extensively to study nucleotide binding and dissociation for a variety of kinesins9-11,13-16. The methods described do not allow observation of phosphate release (Figure 1, Step 3). However, stopped-flow fluorescence can be used to observe this transition by using a phosphate sensing protein to couple the production of phosphate to a fluorescence intensity change17. The methods presented use nucleotides labeled with the small fluorophore methylanthraniloyl (mant), which generally exhibit an increase in fluorescence intensity when bound to protein. Furthermore, the mant fluorophore, when conjugated to either the 2&#8217; or 3&#8217; position on the ribose, often has little or no effect on the binding, dissociation or hydrolysis of the nucleotide9-12. Mant-labeled nucleotides are readily available from commercial sources (see list of materials) and are provided as mixtures of the 2&#8217; and 3&#8217; conjugate as they quickly re-equilibrate when purified to a single isomer8. Mant-labeled nucleotides make excellent reporter molecules by which the kinetics of nucleotide binding and dissociation can be observed.
The use of fluorescently labeled nucleotides means that the read out of the assays described is a real time change in fluorescence intensity. This makes these assays ideal for stopped-flow fluorescence, which allows rapid mixing of reagents and real time observation of changes in fluorescence associated with the subsequent reaction. The combination of stopped-flow as the mixing technique and fluorescence as the method of detection produces a system that is relatively sample efficient. For example, to acquire the data shown in Figure 3C requires 0.8&#160;-&#160;1.0 mg of purified kinesin-13, MCAK, and to acquire the data shown in Figure 4B requires ~0.3&#160;mg of purified kinesin-13, MCAK. One drawback of the use of mant as a fluorophore is that it is prone to photobleaching. This can be observed in isolation from kinesin reaction kinetics by mixing a solution of mant-labeled nucleotide with reaction buffer, in the absence of kinesin, in the stopped-flow. A slow decrease in fluorescence intensity is observed as the mant group becomes bleached. Over the range of timescales used in the protocols described photobleaching of the mant group is well described by a linear function. Therefore, a simple way to correct data for photobleaching is to fit to an exponential function with a baseline described by a linear function (i.e., m.t + c) rather than the more common flat baseline, described by a single parameter.
mantATP binding
When measuring rate constants for association and dissociation of mantATP (Section 2) the ADP, which remains associated with the kinesin nucleotide-binding site in the absence of microtubules, must be removed. This allows mantATP association and dissociation (Figure 1, step 1) to be observed in isolation from ADP dissociation. To achieve this the kinesin is incubated with at least a 50-fold excess of EDTA over nucleotide binding sites (Step 2.1). The EDTA sequesters Mg2+ ions causing Mg2+ to dissociate from the nucleotide-binding pocket, which results in release of the nucleotide11. Therefore, when carrying out steps 2.1&#160;-&#160;2.3, it is vital to use a buffer free of Mg2+. The nucleotide-free kinesin protein is buffer exchanged to separate it both from free nucleotide and from EDTA. To accomplish this separation, a disposable gravity-flow gel filtration column is sufficient and is simple and rapid to use (see list of materials). The interaction of mantATP with nucleotide free kinesin is carried out at a range of mantATP concentrations (Step 2.4) to enable deconvolution of the association and dissociation rate constants (Step 2.8). The theory behind experiments of this type is well described in reference 18. In performing these experiments one begins with the lowest concentration of mantATP. This removes the necessity for wash steps between different concentrations. It will likely be necessary to adjust the sensitivity of the stopped-flow fluorimeter as the mantATP concentration is increased.&#160;The mantATP concentration must always be in at least 5-fold excess over the concentration of kinesin nucleotide binding sites to maintain pseudo first order conditions. However, as the concentration of mantATP is increased, the signal change can become swamped as the fraction of nucleotide that binds to kinesin decreases. Therefore, the concentration of the kinesin is also increased for each new concentration of mantATP such that the concentration of mantATP is never greater than 10-fold molar excess over nucleotide binding sites (Step 2.6).
mantADP Dissociation 
Although association and dissociation rate constants for mantADP can be determined by the same method described for mantATP (Section 2). Dissociation of ADP from a kinesin can be directly observed by preloading the nucleotide-binding site with mantADP (Steps 3.1&#8211;3.3) The mantADP.kinesin complex is then mixed with an excess of unlabeled ATP (step 3.5). The excess unlabeled ATP prevents rebinding of mantADP to the kinesin and thereby allows the dissociation reaction (Figure 1, k4) to be observed in isolation from association of mantADP. In this assay the concentration of unlabeled ATP must be at least a 50-fold molar excess of nucleotide binding sites. The theory behind this assay is well described in reference 18. This direct method gives a more accurate value for the dissociation rate constant than the extrapolation to the y-axis described in Step 2.8.
Inclusion of Microtubules
The methods described can also be used to determine the kinetics of nucleotide binding and dissociation in the presence of microtubules. Microtubules are introduced to the nucleotide containing syringe prior to mixing in the stopped-flow:the lower syringe in Figure 3B and 4B (Insets). In this configuration, the kinesin will meet the nucleotide at the same time as it meets the microtubules. Stabilized microtubules are used, where the lifetime of the tubulin polymer is extended by the use of either a stabilizing chemical, such as taxol, or a nonhydrolysable GTP analogue, such as guanosine-5&#8217;-[(&#945;,&#946;)-methyleno]triphosphate (GMPCPP, see list of materials). It is possible to use two cycles of tubulin polymerization with GMPCPP to make microtubules that can be stored for many months in liquid nitrogen9,19. The use of pre-prepared microtubules significantly reduces the practical difficulties in performing these assays. To include microtubules in the assays, it is important to use a reaction buffer suitable for microtubule stability. The buffer of choice is PIPES based, with the most commonly used buffer containing 80&#160;mM PIPES pH 6.9, 1&#160;mM MgCl2 and 1&#160;mM EGTA (known as BRB80)20,21.
The methods described are not restricted to use with kinesins, but can be adapted and applied to any nucleotide binding protein. For example, similar methods have been applied to the ATP turnover cycle of members of the myosin family of molecular motors12,22 and the use of mant labeled guanosine nucleotides further extends methods of this type to GTP hydrolyzing proteins23,24.

The kinesins are a superfamily of proteins characterized by a highly conserved motor domain1. The kinesin motor domain interacts with microtubules in a nucleotide-dependent manner. Members of the kinesin family display a range of functions from translocating kinesins, which carry cargo in a directed fashion along microtubules, to non-translocating microtubule end-binding kinesins, which regulate microtubule dynamics2,3.
Kinesins use the turnover of adensosine-5&#8217;-triphosphate (ATP) to regulate their interaction with the microtubule cytoskeleton4-6. The conformation of the kinesin motor domain and therefore its affinity for the microtubule depends on its nucleotide state: ATP, ADP, ADP.Pi or no nucleotide can be bound (Figure 1). To understand the molecular mechanism of a kinesin, an understanding of the relationship between ATP turnover and microtubule interaction is required. Therefore, a major goal in the kinesin field is to characterize the functional properties of different nucleotide states and the kinetics of the transitions between them both in the presence and absence of microtubules.
<img alt="Figure 1" fo:content-width="2in" width="200" src="/files/ftp_upload/52142/52142fig1highres.jpg" /> 
Figure 1.&#160;The simplest ATP turnover cycle for a kinesin consists of four possible states: nucleotide-free (&#934;), ATP-bound, ADP.Pi-bound and ADP-bound. Each of the four transitions is characterized by a forward and backward rate constant (kx and k-x respectively).
To understand how the chemical cycle of ATP turnover is coupled to the mechanical cycle of microtubule interaction, one must determine which step in the ATP turnover cycle is rate limiting in the absence of microtubules and then determine how the cycle is altered by the presence of microtubules. The simplest ATP turnover cycle consists of four individual chemicals steps: (1) ATP binding to the empty site (&#934; denotes the empty site, i.e., nucleotide-free kinesin); (2) ATP cleavage; (3) phosphate dissociation and (4) ADP dissociation (Figure 1). The rate-limiting step sets the rate constant for the complete ATP turnover cycle. Therefore, to determine which step is rate limiting each of the individual transitions must be measured and compared to the rate constant for the complete cycle. Methods to measure the overall ATP turnover cycle rate constant are not described here but can be found in reference 7. Here, we describe methods by which rate constants for individual chemical steps can be determined. There are a number of methods available to study individual transitions in the turnover cycle of a nucleotide hydrolyzing protein. The methods presented here take advantage of the properties of nucleotides labeled with the fluorophore methylanthraniloyl, usually referred to as mant, which display a change in fluorescence intensity upon binding to protein8. The mant group is used because its small size means that, when coupled to the ribose (Figure 2A), it generally has little or no effect on the kinetics of nucleotide binding or hydrolysis9-12. Here, we present protocols describing the use of mant-labeled nucleotides, in conjunction with stopped-flow fluorescence, to allow observation of nucleotide binding and dissociation in the ATP turnover cycle of a kinesin.

<table><tbody><tr><td>2&amp;rsquo;/3&amp;rsquo;-(N-methylanthraniloyl) adenosine-5&amp;rsquo;triphosphate&amp;nbsp;</td><td>Jena Biosciences</td><td># NU-202</td><td>mantATP</td></tr><tr><td>2&amp;rsquo;/3&amp;rsquo;-(N-methylanthraniloyl) adenosine-5&amp;rsquo;-diphosphate&amp;nbsp;</td><td>Jena Biosciences</td><td># NU-201</td><td>mantADP</td></tr><tr><td>Mg-ATP</td><td>Roche</td><td># 10519979001</td><td>The disodium salt of ATP is made to a concentration of 100&amp;nbsp;mM in 100&amp;nbsp;mM MgCl&lt;sub&gt;2&lt;/sub&gt;. Concentration should be verified by absorption at 260&amp;nbsp;nm (e = 15,400&amp;nbsp;M-1cm-1). Solution stored in aliquots at -20&amp;nbsp;&lt;sup&gt;o&lt;/sup&gt;C.&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Fisher Scientific</td><td>BP120-500</td><td>0.5&amp;nbsp;M stock solution in water, pH 8.0. HAZARD: powder harmful if inhaled.</td></tr><tr><td>Dithiothreitol (DTT)</td><td>Melford</td><td>MB1015</td><td>1&amp;nbsp;M stock solution in water made fresh for use on the same day. HAZARD: powder harmful if inhaled or in contact with the skin.</td></tr><tr><td>NAP-5 column</td><td>GE Healthcare</td><td># 17-0853</td><td>G-25 sephadex resin gravity-flow gel filtration column</td></tr><tr><td>GMPCPP</td><td>Jena Biosciences</td><td># NU-405</td><td>nonhydrolysable analogue of GTP</td></tr><tr><td>Stopped-flow fluorimeter&amp;nbsp;</td><td>Applied Photophysics</td><td>SX20</td><td>A fluorimeter with some plumbing attached, which allows reagents to be rapidly mixed in the observation cuvette. Thereby allowing the kinetics of reactions on timescales of ~0.1&amp;nbsp;-&amp;nbsp;100&amp;nbsp;sec to be monitored.</td></tr></tbody></table>

use of stopped-flow fluorescence and labeled nucleotides to analyze the atp turnover cycle of kinesins

The kinesin superfamily of microtubule associated motor proteins share a characteristic motor domain which both hydrolyses ATP and binds microtubules. Kinesins display differences across the superfamily both in ATP turnover and in microtubule interaction. These differences tailor specific kinesins to various functions such as cargo transport, microtubule sliding, microtubule depolymerization and microtubule stabilization. To understand the mechanism of action of a kinesin it is important to understand how the chemical cycle of ATP turnover is coupled to the mechanical cycle of microtubule interaction. To dissect the ATP turnover cycle, one approach is to utilize fluorescently labeled nucleotides to visualize individual steps in the cycle. Determining the kinetics of each nucleotide transition in the ATP turnover cycle allows the rate-limiting step or steps for the complete cycle to be identified. For a kinesin, it is important to know the rate-limiting step, in the absence of microtubules, as this step is generally accelerated several thousand fold when the kinesin interacts with microtubules. The cycle in the absence of microtubules is then compared to that in the presence of microtubules to fully understand a kinesin&#8217;s ATP turnover cycle. The kinetics of individual nucleotide transitions are generally too fast to observe by manually mixing reactants, particularly in the presence of microtubules. A rapid mixing device, such as a stopped-flow fluorimeter, which allows kinetics to be observed on timescales of as little as a few milliseconds, can be used to monitor such transitions. Here, we describe protocols in which rapid mixing of reagents by stopped-flow is used in conjunction with fluorescently labeled nucleotides to dissect the ATP turnover cycle of a kinesin.

An increase in fluorescence intensity of the mant group is typically observed upon binding of mant-labeled nucleotide to a kinesin. However, the magnitude of such a signal change is dependent on the particular kinesin in question. Therefore, it is useful to perform equilibrium measurements to confirm both the presence and magnitude of a fluorescence intensity change before progressing to kinetic assays. Representative fluorescence spectra for mantATP and mantADP, both in solution and in complex with the kinesin-13, MCAK, are shown in Figure 2B. These data highlight the fluorescence intensity increase displayed by both mantATP and mantADP upon binding to MCAK. The change in fluorescence intensity upon binding of mant nucleotide to a kinesin is used to report on the association and dissociation of both ATP and ADP (Sections 2 and 3). In the case of MCAK, a fluorescence intensity difference is observed between mantATP.kinesin and mantADP.kinesin9.
Figure 3A depicts the reaction that occurs upon mixing mantATP with nucleotide-free kinesin. Representative data illustrating the increase in fluorescence intensity that occurs upon binding of mantATP to the kinesin-13, MCAK is shown in Figure 3B. Data of the type shown in Figure 3B is collected at a range of mantATP concentrations. A plot of the rate constants (kobs) determined versus the mantATP concentration is shown in Figure 3C. Fitting these data to a linear function (i.e., kobs = k1[mantATP] + k-1) allows deconvolution of the rate constants that contribute to kobs (Step 2.8). Therefore, enabling the determination of both the association and the dissociation rate constant for mantATP (Figure 1, k1 and k-1 respectively).
Figure 4A depicts the reaction which occurs upon mixing mantADP.kinesin with an excess of unlabeled ATP. The data in Figure 4B shows the fluorescence intensity decrease observed for mantADP upon dissociation from the kinesin-13, MCAK. By fitting this data to an exponential function (Step 3.6) the rate constant for dissociation of mantADP is determined directly.
<img alt="Figure 2" src="/files/ftp_upload/52142/52142fig2highres.jpg" /> 
Figure 2.&#160;Nucleotides labeled with the fluorophore methylanthraniloyl (mant) are used to isolate individual steps in the ATP turnover cycle. (A) Structure of mantATP showing the position of the mant fluorophore conjugated to either the 2&#8217; or 3&#8217; position on the ribose. (B) Normalized fluorescence spectra for mantATP and mantADP in solution (mAXP) and mantATP and mantADP in complex with the kinesin MCAK (mATP+MCAK and mADP+MCAK). The mantATP spectra are normalized to the fluorescence at max for mantATP in solution and the mantADP spectra are normalized to fluorescence at max for mantADP in solution. The spectra of the mant nucleotides mixed with MCAK are collected at 1&#160;min post mixing. The concentration of reagents and fluorimeter settings are as described in protocol steps 1.1 and 1.2.
<img alt="Figure 3" src="/files/ftp_upload/52142/52142fig3highres.jpg" /> 
Figure 3.&#160;Measurement of the association rate constant of mantATP. (A) Schematic depicting the reaction which takes place post mixing by stopped-flow: mantATP (mATP) binds to nucleotide-free kinesin. The fluorescence intensity of the mant group increases upon binding to the protein. (B) Fluorescence signal change (black) observed upon mixing mantATP (25&#160;M) with MCAK (5&#160;M nucleotide-free, ). This data is fit (red dashed) to a single exponential function plus a line of constant negative slope, which accounts for photobleaching of the mant fluorophore (see discussion). Inset: Contents of the syringes prior to mixing on the stopped-flow fluorimeter. (C) Rate constants (kobs) determined for the reaction of mantATP with nucleotide-free MCAK plotted against concentration of mantATP. This plot will have a linear region with the gradient being the association rate constant (Figure 1, k1) with units of M-1 s-1 and the y-intercept being the dissociation rate constant (Figure 1, k-1) with units of s-1. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52142/52142fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52142/52142fig4highres.jpg" /> 
Figure 4.&#160;Measurement of the dissociation rate constant for mantADP. (A) Schematic depicting the reaction which takes place post mixing by stopped-flow: kinesin preloaded with mantADP is diluted into an excess of unlabeled ATP. Dissociating mantADP is replaced by unlabeled ATP, thereby preventing the back reaction of mantADP association (see discussion). The fluorescence intensity of the mant group decreases upon dissociation from the protein. (B) Fluorescence decrease observed upon the dissociation of mantADP from the kinesin MCAK. The fluorescence signal (black) is fit (red dashed) to a single exponential plus a line of constant negative slope to account for photobleaching of the mant group (see discussion). The rate constant determined is the rate constant for dissociation of mantADP (Figure 1, k4). Inset: Contents of the syringes prior to mixing on a stopped-flow fluorimeter. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52142/52142fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins at JoVE.com

Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins

Kinesins are characterized by nucleotide-dependent interaction with microtubules: a cycle of ATP turnover coupled to a cycle of microtubule interaction. Here, we describe protocols to analyze the kinetics of individual nucleotide transitions in the ATP turnover cycle of a kinesin using fluorescently labeled nucleotides and stopped-flow fluorescence.

The kinesin superfamily of microtubule associated motor proteins share a characteristic motor domain which both hydrolyses ATP and binds microtubules. ...

use-stopped-flow-fluorescence-labeled-nucleotides-to-analyze-atp

Universidad San Sebastian

Research

JoVE Journal

Chemistry

15.5K Views.  University of Nottingham. Kinesins are characterized by nucleotide-dependent interaction with microtubules: a cycle of ATP turnover coupled to a cycle of microtubule interaction. Here, we describe protocols to analyze the kinetics of individual nucleotide transitions in the ATP turnover cycle of a kinesin using fluorescently labeled nucleotides and stopped-flow fluorescence.

Video: Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins

A Semi-High-Throughput Adaptation of the NADH-Coupled ATPase Assay for Screening Small Molecule Inhibitors

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would not occur spontaneously. These proteins are crucial for essentially all aspects of cellular life, including metabolism, cell division, responses to environmental changes and movement. The protocol presented here describes a nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay that has been adapted to semi-high throughput screening of small molecule ATPase inhibitors. The assay has been applied to cardiac and skeletal muscle myosin II's, two actin-based molecular motor ATPases, as a proof of principle. The hydrolysis of ATP is coupled to the oxidation of NADH by enzymatic reactions in the assay. First, the ADP generated by the ATPase is regenerated to ATP by pyruvate kinase (PK). PK catalyzes the transition of phosphoenolpyruvate (PEP) to pyruvate in parallel. Subsequently, pyruvate is reduced to lactate by lactate dehydrogenase (LDH), which catalyzes the oxidation of NADH in parallel. Thus, the decrease in ATP concentration is directly correlated to the decrease in NADH concentration, which is followed by change to the intrinsic fluorescence of NADH. As long as PEP is available in the reaction system, the ADP concentration remains very low, avoiding inhibition of the ATPase enzyme by its own product. Moreover, the ATP concentration remains nearly constant, yielding linear time courses. The fluorescence is monitored continuously, which allows for easy estimation of the quality of data and helps to filter out potential artifacts (e.g., arising from compound precipitation or thermal changes).

A nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay has been adapted to semihigh throughput screening of small molecule myosin inhibitors. This kinetic assay is run in a 384-well microplate format with total reaction volumes of only 20 &#181;L per well. The platform should be applicable to virtually any ADP producing enzyme.

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would ...

Biochemistry

Assembling Molecular Shuttles Powered by Reversibly Attached Kinesins

This protocol describes how to create kinesin-powered molecular shuttles with a weak and reversible attachment of the kinesins to the surface. In contrast to previous protocols, in this system, microtubules recruit kinesin motor proteins from solution and place them on a surface. The kinesins will, in turn, facilitate the gliding of the microtubules along the surface before desorbing back into the bulk solution, thus being available to be recruited again. This continuous assembly and disassembly leads to striking dynamic behavior in the system, such as the formation of temporary kinesin trails by gliding microtubules.
Several experimental methods will be described throughout this experiment: UV-Vis spectrophotometry will be used to determine the concentration of stock solutions of reagents, coverslips will first be ozone and ultraviolet (UV) treated and then silanized before being mounted into flow cells, and total internal reflection fluorescence (TIRF) microscopy will be used to simultaneously image kinesin motors and microtubule filaments.

We present a protocol to build molecular shuttles, where surface-adhered kinesin motor proteins propel dye-labelled microtubules. Weak interactions of the kinesins with the surface enables their reversible attachment to it. This creates a nanoscale system which exhibits dynamic assembly and disassembly of its components while retaining its functionality.

This protocol describes how to create kinesin-powered molecular shuttles with a weak and reversible attachment of the kinesins to the surface. In contrast ...

Engineering

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic properties are adapted for the particular function they execute in cells. Myosin 5a processively walks on F-actin to transport melanosomes and vesicles in cells. Conversely, nonmuscle myosin 2b operates as a bipolar filament containing approximately 30 molecules. It moves F-actin of opposite polarity toward the center of the filament, where the myosin molecules work asynchronously to bind actin, impart a power stroke, and dissociate before repeating the cycle. Nonmuscle myosin 2b, along with its other nonmuscle myosin 2 isoforms, has roles that include&#160;cell adhesion, cytokinesis, and tension maintenance. The mechanochemistry of myosins can be studied by performing in vitro motility assays using purified proteins. In the gliding actin filament assay, the myosins are bound to a microscope coverslip surface and translocate fluorescently labeled F-actin, which can be tracked. In the single molecule/ensemble motility assay, however, F-actin is bound to a coverslip and the movement of fluorescently labeled myosin molecules on the F-actin is observed. In this report, the purification of recombinant myosin 5a from Sf9 cells using affinity chromatography is outlined. Following this, we outline two fluorescence microscopy-based assays: the gliding actin filament assay and the inverted motility assay. From these assays, parameters such as actin translocation velocities and single molecule run lengths and velocities can be extracted using the image analysis software. These techniques can also be applied to study the movement of single filaments of the nonmuscle myosin 2 isoforms, discussed herein in the context of nonmuscle myosin 2b. This workflow represents a protocol and a set of quantitative tools that can be used to study the single molecule and ensemble dynamics of nonmuscle myosins.

Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic ...

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

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

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

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Motility of Single Molecules and Clusters of Bi-Directional Kinesin-5 Cin8 Purified from S. cerevisiae Cells

The mitotic bipolar kinesin-5 motors perform essential functions in spindle dynamics. These motors exhibit a homo-tetrameric structure with two pairs of catalytic motor domains, located at opposite ends of the active complex. This unique architecture enables kinesin-5 motors to crosslink and slide apart antiparallel spindle microtubules (MTs), thus providing the outwardly-directed force that separates the spindle poles apart. Previously, kinesin-5 motors were believed to be exclusively plus-end directed. However, recent studies revealed that several fungal kinesin-5 motors are minus-end directed at the single-molecule level and can switch directionality under various experimental conditions. The Saccharomyces cerevisiae kinesin-5 Cin8 is an example of such bi-directional motor protein: in high ionic strength conditions single molecules of Cin8 move in the minus-end direction of the MTs. It was also shown that Cin8 forms motile clusters, predominantly at the minus-end of the MTs, and such clustering allows Cin8 to switch directionality and undergo slow, plus-end directed motility. This article provides a detailed protocol for all steps of working with GFP-tagged kinesin-5 Cin8, from protein overexpression in S. cerevisiae cells and its purification to in vitro single-molecule motility assay. A newly developed method described here helps to differentiate between single molecules and clusters of Cin8, based on their fluorescence intensity. This method enables separate analysis of motility of single molecules and clusters of Cin8, thus providing the characterization of the dependence of Cin8 motility on its cluster size.

Motility of Single Molecules and Clusters of Bi-Directional Kinesin-5 Cin8 Purified from S. cerevisiae Cells

The bi-directional mitotic kinesin-5 Cin8 accumulates in clusters that split and merge during their motility. Accumulation in clusters also changes the velocity and directionality of Cin8. Here, a protocol for motility assays with purified Cin8-GFP and analysis of motile properties of single molecules and clusters of Cin8 is described.

The mitotic bipolar kinesin-5 motors perform essential functions in spindle dynamics. These motors exhibit a homo-tetrameric structure with two pairs of ...

Cargo Loading onto Kinesin Powered Molecular Shuttles

Cells have evolved sophisticated molecular machinery, such as kinesin motor proteins and microtubule filaments, to support active intracellular transport of cargo. While kinesins tail domain binds to a variety of cargoes, kinesins head domains utilize the chemical energy stored in ATP molecules to step along the microtubule lattice. The long, stiff microtubules serve as tracks for long-distance intracellular transport.
These motors and filaments can also be employed in microfabricated synthetic environments as components of molecular shuttles 1. In a frequently used design, kinesin motors are anchored to the track surface through their tails, and functionalized microtubules serve as cargo carrying elements, which are propelled by these motors. These shuttles can be loaded with cargo by utilizing the strong and selective binding between biotin and streptavidin. The key components (biotinylated tubulin, streptavidin, and biotinylated cargo) are commercially available.
Building on the classic inverted motility assay 2, the construction of molecular shuttles is detailed here. Kinesin motor proteins are adsorbed to a surface precoated with casein; microtubules are polymerized from biotinylated tubulin, adhered to the kinesin and subsequently coated with rhodamine-labeled streptavidin. The ATP concentration is maintained at subsaturating concentration to achieve a microtubule gliding velocity optimal for loading cargo 3. Finally, biotinylated fluorescein-labeled nanospheres are added as cargo. Nanospheres attach to microtubules as a result of collisions between gliding microtubules and nanospheres adhering to the surface.
The protocol can be readily modified to load a variety of cargoes such as biotinylated DNA4, quantum dots 5 or a wide variety of antigens via biotinylated antibodies 4-6.

Molecular shuttles consisting of functionalized microtubules gliding on surface-adhered kinesin motor proteins can serve as a nanoscale transport system. Here, the assembly of a typical shuttle system is described.

Cells have evolved sophisticated molecular machinery, such as kinesin motor proteins and microtubule filaments, to support active intracellular transport ...

Biology

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

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

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

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

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved single-molecule studies and has gained immense popularity in detecting and understanding the dynamic behavior of fluorescent-tagged molecules. Here, we describe a detailed method for&#160;in vitro single-molecule studies of kinesin-3 family motors using Total Internal Reflection Fluorescence (TIRF) microscopy. Kinesin-3 is a large family that plays critical roles in cellular and physiological functions ranging from intracellular cargo transport to cell division to development. We have shown previously that constitutively active dimeric kinesin-3 motors exhibit fast and superprocessive motility with high microtubule affinity at the single-molecule level using cell lysates prepared by expressing motor in mammalian cells. Our lab studies kinesin-3 motors and their regulatory mechanisms using cellular, biochemical and biophysical approaches, and such studies demand purified proteins at a large scale. Expression and purification of these motors using mammalian cells would be expensive and time-consuming, whereas expression in a prokaryotic expression system resulted in significantly aggregated and&#160;inactive protein. To overcome the limitations posed by bacterial purification systems and mammalian cell lysate, we have established a robust Sf9-baculovirus expression system to express and purify these motors. The kinesin-3 motors are C-terminally tagged with 3-tandem fluorescent proteins (3xmCitirine or 3xmCit) that provide enhanced signals and decreased photobleaching. In vitro single-molecule and multi-motor gliding analysis of Sf9 purified proteins demonstrate that kinesin-3 motors are fast and superprocessive akin to our previous studies using mammalian cell lysates. Other applications using these assays include detailed knowledge of oligomer conditions of motors, specific binding partners paralleling biochemical studies, and their kinetic state.

This study details purification of KIF1A(1-393LZ), a member of kinesin-3 family, using Sf9-baculovirus expression system. In vitro single-molecule and multi-motor gliding analysis of these purified motors exhibited robust motility properties comparable to motors from mammalian cell lysate. Thus, Sf9-baculovirus system is amenable to express and purify motor protein of interest.

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved ...

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and DNA accessibility by catalyzing ATP-dependent nucleosome remodeling. Biochemically dissecting the contributions of individual subunits of such complexes to the multi-step ATP-dependent chromatin remodeling reaction requires the use of assays that monitor the production of reaction products and measure the formation of reaction intermediates. This JOVE protocol describes assays that allow one to measure the biochemical activities of chromatin remodeling complexes or subcomplexes containing various combinations of subunits. Chromatin remodeling is measured using an ATP-dependent nucleosome sliding assay, which monitors the movement of a nucleosome on a DNA molecule using an electrophoretic mobility shift assay (EMSA)-based method. Nucleosome binding activity is measured by monitoring the formation of remodeling complex-bound mononucleosomes using a similar EMSA-based method, and DNA- or nucleosome-dependent ATPase activity is assayed using thin layer chromatography (TLC) to measure the rate of conversion of ATP to ADP and phosphate in the presence of either DNA or nucleosomes. Using these assays, one can examine the functions of subunits of a chromatin remodeling complex by comparing the activities of the complete complex to those lacking one or more subunits. The human INO80 chromatin remodeling complex is used as an example; however, the methods described here can be adapted to the study of other chromatin remodeling complexes.

Here we describe biochemical assays that can be used to characterize ATP-dependent chromatin remodeling enzymes for their abilities to 1) catalyze ATP-dependent nucleosome sliding, 2) engage with nucleosome substrates, and 3) hydrolyze ATP in a nucleosome- or DNA-dependent manner.

Members of the SNF2 family of ATPases often function as components of multi-subunit chromatin remodeling complexes that regulate nucleosome dynamics and ...

Identification of Kinesin-1 Cargos Using Fluorescence Microscopy

Fluorescence microscopy is employed to identify Kinesin-1 cargos. Recently, the heavy chain of Kinesin-1 (KIF5B) was shown to transport the nuclear transcription factor c-MYC for proteosomal degradation in the cytoplasm. The method described here involves the study of a motorless KIF5B mutant for fluorescence microscopy. The wild-type and motorless KIF5B proteins are tagged with the fluorescent protein tdTomato. The wild-type tdTomato-KIF5B appears homogenously in the cytoplasm, while the motorless tdTomato-KIF5B mutant forms aggregates in the cytoplasm. Aggregation of the motorless KIF5B mutant induces aggregation of its cargo c-MYC in the cytoplasm. Hence, this method provides a visual means to identify the cargos of Kinesin-1. A similar strategy can be utilized to identify cargos of other motor proteins.

Here, a protocol is presented to identify Kinesin-1 cargos. A motorless mutant of the Kinesin-1 heavy chain (KIF5B) aggregates in the cytoplasm and induces aggregation of its cargos. Both aggregates are detected under fluorescence microscopy. A similar strategy can be employed to identify cargos of other motor proteins.

Fluorescence microscopy is employed to identify Kinesin-1 cargos. Recently, the heavy chain of Kinesin-1 (KIF5B) was shown to transport the nuclear ...

Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins

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A Semi-High-Throughput Adaptation of the NADH-Coupled ATPase Assay for Screening Small Molecule Inhibitors

Assembling Molecular Shuttles Powered by Reversibly Attached Kinesins

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

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

Motility of Single Molecules and Clusters of Bi-Directional Kinesin-5 Cin8 Purified from S. cerevisiae Cells

Cargo Loading onto Kinesin Powered Molecular Shuttles

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

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

Biochemical Assays for Analyzing Activities of ATP-dependent Chromatin Remodeling Enzymes

Identification of Kinesin-1 Cargos Using Fluorescence Microscopy