The authors gratefully acknowledge financial support under NSF grant NSF-DMR 1807514. The authors thank G. Bachand and V. Vandelinder for providing the GFP-kinesin protein. This work was performed, in part, at the Center for Integrated Nanotechnologies, an Office of Science User Facility operated for the US Department of Energy (DOE) Office of Science by Los Alamos National Laboratory (contract no. DE- AC52-06NA25396) and Sandia National Laboratories (con- tract no. 97 DE-AC04-94AL85000). The authors thank Dr. Jennifer Neff and AllVivo Vascular for their gift of PEG-PPG-PEG functionalized with NTA.

1. Solution preparation
CAUTION: Three of the reagents used in this protocol (toluene, dimethyldichlorosilane, and dithiothreitol) are highly toxic. Please consult the relevant material safety data sheets (MSDS) before use. Furthermore, parts of this protocol need to be performed under a higher level of protection, wearing safety glasses and two sets of protective gloves as indicated. Unless otherwise advised, the experiments can be performed on lab benches while using all the appropriate personal protective equipment (safety glasses, gloves, lab coat, full length pants, and closed-toe shoes).
NOTE: The concentrations of the ATP, kinesin, and microtubules that are used in this protocol can be modified given the needs of each experiment. However, if modified, please ensure that the final concentrations of the other reagents remain the same as given below. All experiments were performed at room temperature, (~25 &#176;C).
<ol>
	<li>Preparation of stock solutions 
	NOTE: Prepare and aliquot the following solutions beforehand. All aliquots can be prepared at room temperature (~25 &#176;C).
	<ol>
		<li>BRB80 buffer 
		NOTE: The buffer for most of the solutions used in this protocol is BRB80. BRB80 can be prepared in large quantities and stored in a -20 &#176;C freezer.
		<ol>
			<li>Dissolve 24.2 g of piperazine-N,N&#8242;-bis(2-ethanesulfonic acid) (PIPES) and 3.1 g of potassium hydroxide (KOH) in 800 mL in deionized water to make 800 mL of 100 mM PIPES. Add 100 mL of 10 mM magnesium chloride (MgCl2) and 100 mL of 10 mM ethylene glycol tetraacetic acid (EGTA) to get a final volume of 1 L. Raising the pH to 8 with potassium hydroxide (KOH) can help in the dissolution of PIPES and EGTA. 
			NOTE: The final concentrations of chemicals in the BRB80 buffer are 80 mM PIPES, 1 mM MgCl2, and 1 mM EGTA.</li>
			<li>Adjust the pH of the buffer to 6.9 using KOH and hydrochloric acid (HCl).</li>
		</ol>
		</li>
		<li>ATP
		<ol>
			<li>Prepare a 1 mL solution of 100 mM ATP in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -80 &#176;C freezer.</li>
		</ol>
		</li>
		<li>GTP
		<ol>
			<li>Prepare a 500 &#956;L solution of 25 mM GTP in ultrapure water. Pipet the solution into 5 &#956;L aliquots. Store the aliquots in a -80 &#176;C freezer.</li>
		</ol>
		</li>
		<li>MgCl2
		<ol>
			<li>Prepare a 1 mL solution of 100 mM MgCl2 in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Casein
		<ol>
			<li>Weigh 1 g of dry casein and transfer it to a 50 mL conical centrifuge tube. Add 35 mL of BRB80 buffer to the tube to dissolve the casein powder.</li>
			<li>Place the solution in a tumbler in a cold room to dissolve overnight. At this point, the solution will look thick and viscous.</li>
			<li>Leave the tube upright in a 4 &#176;C fridge to allow for any large undissolved clumps to settle. Transfer the supernatant to a new tube.</li>
			<li>Spin the tube in a centrifuge at 1,000 x g to pellet out more precipitates. Once again, transfer the supernatant to a new tube.</li>
			<li>Repeatedly filter the solution using 0.2 &#956;m (7 bar maximum pressure) filters. The solution being thick, many filters will get clogged, so repeat this step until the filter is clear and unclogged.</li>
			<li>Determine the casein concentration in the resulting solution using UV/Vis spectrophotometry, using casein&#8217;s extinction coefficient of 19 mM-1&#8729;cm-1 at 280 nm20. Assuming a molecular weight of 23 kDa for casein, dilute the solution to a concentration of 20 mg&#8729;mL-1 in BRB80.</li>
			<li>Pipet the solution into 20 &#956;L aliquots and store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>D-Glucose
		<ol>
			<li>Prepare a 1 mL solution of 2 M D-glucose in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Glucose oxidase
		<ol>
			<li>Prepare a 1 mL solution of 20 mg&#8729;mL-1 glucose oxidase in BRB80. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Catalase
		<ol>
			<li>Prepare a 1 mL solution of 0.8 mg&#8729;mL-1 catalase in BRB80. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Dithiothreitol (DTT) 
		NOTE: Since DTT is slightly volatile and toxic, please perform the following steps under a fume hood.
		<ol>
			<li>Prepare a 1 mL solution of 1 M DTT diluted in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Paclitaxel
		<ol>
			<li>Prepare a 1 mL solution of 1 mM paclitaxel diluted in DMSO. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Dimethyl sulfoxide (DMSO) 
		NOTE: The DMSO used in these experiments is pure.
		<ol>
			<li>Pipet 1 mL of pure DMSO into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Creatine phosphate
		<ol>
			<li>Prepare a 1 mL solution of 0.2 M creatine phosphate in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Creatine phosphokinase
		<ol>
			<li>Prepare a 1 mL solution of 200 units&#183;L-1 creatine phosphokinase in ultrapure water. Pipet the solution into 10 &#956;L aliquots. Store the aliquots in a -20 &#176;C freezer.</li>
		</ol>
		</li>
		<li>Nickel (II) sulfate
		<ol>
			<li>Prepare 500 mL of a solution of 50 mM nickel (II) sulfate in ultrapure water. Store the solution at room temperature (~25 &#176;C).</li>
		</ol>
		</li>
		<li>Poly(ethylene glycol)-block-poly(propylene glycol)-block-poly(ethylene glycol) (PEG-PPG-PEG) solution
		<ol>
			<li>Weigh out 2 mg of PEG-PPG-PEG (number average molecular weight: 14,600 g&#8729;mol-1)-NTA powder on weighing paper. PEG-PPG-PEG-NTA is the triblock copolymer functionalized with a nitrilotriacetic acid (NTA) group. Refer to the Table of Materials for more details about the PEG-PPG-PEG-NTA.</li>
			<li>Transfer the powder into a 1.5 mL microcentrifuge tube. Add 1 mL of the stock nickel (II) sulfate solution to the tube. Vortex until powder is dissolved and no visible clumps remain.</li>
			<li>Store for up to one month at room temperature.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Before starting an experiment
	<ol>
		<li>Fill a bucket with ice.</li>
		<li>Take one aliquot from each of the 13 reagents described in sections 1.1.1 to 1.1.13 above and add them to the bucket, thus keeping them on ice. Thaw each of the reagents before use.</li>
	</ol>
	</li>
	<li>Microtubule Preparation 
	NOTE: Microtubules were polymerized from a 20 &#956;g aliquot of dye-labeled lyophilized tubulin. The excitation wavelength of the dye is of 647 nm.
	<ol>
		<li>Preparation of the microtubule growth buffer
		<ol>
			<li>Pipet 21.8 &#956;L of BRB80 buffer into a small (0.6 mL) microcentrifuge tube. Add 1 &#956;L of the MgCl2 stock solution, 1 &#956;L of the GTP stock solution, and 1.2 &#956;L of the DMSO stock solution. 
			NOTE: Thus, the final concentrations of the reagents in the BRB80 buffer are 4 mM MgCl2, 1 mM GTP, and 5% (v/v) dimethyl sulfoxide.</li>
		</ol>
		</li>
		<li>Microtubule polymerization
		<ol>
			<li>Add 6.25 &#956;L of microtubule growth buffer directly into a 20 &#956;g aliquot of labeled lyophilized tubulin. Vortex the aliquot for 5 s at 30 rps.</li>
			<li>Cool the aliquot on ice for 5 min before incubating it at 37 &#176;C for 45 min and proceed to section 1.3.3.</li>
		</ol>
		</li>
		<li>Microtubule stabilization
		<ol>
			<li>Add 5 &#956;L of the aliquoted paclitaxel solution to 490 &#956;L of BRB80 buffer. Vortex the solution for 10 s at 30 rps.</li>
			<li>Once the 45 min of incubation for the microtubules are up, add 5 &#956;L of the polymerized microtubule solution to the BRB80/paclitaxel solution. 
			NOTE: This 100-fold diluted, stabilized, microtubule solution will hereafter be referred to as MT100. It can be used for up to 5 days, and it can be diluted to obtain the desired microtubule density for each experiment.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Motility solution
	<ol>
		<li>Enzymatic antifade, ATP regenerating system, and ATP
		<ol>
			<li>Add 9.0 &#956;L of the aliquoted casein solution to 291 &#956;L of BRB80 buffer.</li>
			<li>If the desired ATP concentration for the experiment is lower than 1 mM, pipet 83 &#956;L of the BRB80/casein solution into a new 0.6 mL microcentrifuge tube. Otherwise, pipet 85 &#956;L of that solution into a new 0.6 mL microcentrifuge tube.</li>
			<li>Add 1 &#956;L of D-glucose, 1 &#956;L of glucose oxidase, 1 &#956;L of catalase, and 1 &#956;L of DTT to that tube. 
			NOTE: These chemicals constitute an enzymatic antifade cocktail21 that will reduce photobleaching by removing dissolved oxygen and quenching reactive radicals. This reduces photobleaching and microtubule disintegration caused by the excitation illumination during fluorescence imaging22,23.</li>
			<li>For experiments in which the ATP concentration is chosen to be significantly lower than 1 mM, add the following reagents to the solution to create an ATP-regenerating system: 1 &#956;L of the aliquoted creatine phosphate solution and 1 &#956;L of the aliquoted creatine phosphokinase solution.</li>
			<li>Add 1 &#956;L of the stock ATP solution to the motility solution. Flick or vortex the aliquot to homogenously distribute the chemicals. 
			NOTE: Thus, the final concentration of chemicals in the solution are 10 &#956;M paclitaxel, 0.5 mg&#183;mL&#8722;1 casein, 20 mM D-glucose, 200 &#956;g&#183;mL&#8722;1 glucose oxidase, 8 &#956;g&#183;mL&#8722;1 catalase, 10 mM dithiothreitol, 2 mM creatine phosphate (if added), 2 units&#183;L&#8722;1 creatine phosphokinase (if added), and 1 mM ATP. This solution will hereafter be referred to as the motility solution.</li>
		</ol>
		</li>
		<li>Kinesin 
		NOTE: The stock kinesin solution used in these experiments is prepared by G. Bachand at the Center for Integrated Nanotechnologies at Sandia National Laboratories and made available under a user agreement (https://cint.lanl.gov/becoming-user/call-for-proposals.php). The buffer used here consists of 40 mM imidazole, 300 mM NaCl, 0.76 g&#183;L-1 EGTA, 37.2 mg&#183;L-1 EDTA, 50 g&#183;L-1 sucrose, 0.2 mM TCEP, and 50 &#956;M Mg-ATP. For this series of experiments, the rkin430eGFP kinesin construct was used. This is a kinesin consisting of the first 430 amino acids of rat kinesin heavy chain fused to eGFP and a C-terminal His-tag at the tail domain24. It was expressed in Escherichia coli and purified using a Ni&#8722;NTA column. The concentration of the GFP-kinesin stock solution was 1.8 &#177; 0.3 &#956;M as determined by UV/Vis spectrophotometry, using an extinction coefficient for GFP of 55 mM-1&#183;cm-1 at 489 nm25, and taking into account that kinesin is a dimer.
		<ol>
			<li>Determine the desired kinesin concentration or surface density for the experiment. For typical assays, this concentration is 20 nM. If the kinesin concentration needs to be diluted more than a hundred-fold, dilute it in a solution of BRB80 that contains 0.5 mg&#183;mL-1 of casein.</li>
			<li>Add 1 &#181;L of kinesin solution to the motility solution in order to get a final concentration of approximatively 20 nM.</li>
		</ol>
		</li>
		<li>Microtubules
		<ol>
			<li>Add 10 &#956;L of the MT100 solution prepared in section 1.3 to the motility solution. 
			NOTE: This motility solution can be used for up to 3 hours. After that time, the antifade system loses its effectiveness because the glucose in the solution is depleted by the enzymatic reaction.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Assembling the flow cells
<ol>
	<li>Washing the coverslips
	<ol>
		<li>For flow cells, use a large coverslip (dimension: 60 mm x 25 mm) and a small one (dimensions: 22 mm x 22 mm)19.</li>
		<li>Rinse all coverslips twice with ethanol and twice with ultrapure water. Sonicate the coverslips in ultrapure water for 5 min. Dry them in an oven at a temperature of 50&#8211;75 &#176;C.</li>
		<li>Using a UV/ozone cleaner (see Table of Materials and manufacturer&#8217;s instructions), treat one side of each coverslip for 15 min. Perform this step can at room temperature (~25 &#176;C) and in normal atmospheric condition (pressure of 1 atm).</li>
		<li>Carefully turn each coverslip to its other side (using tweezers) and UV/ozone treat that side as well.</li>
		<li>Sonicate the coverslips in ultrapure water again for 5 min before drying them again in the oven at a temperature of 50&#8211;75 &#176;C.</li>
	</ol>
	</li>
	<li>Treating the coverslips to enable PEG-PPG-PEG coating 
	NOTE: The dimethyldichlorosilane and the toluene used in this part of the protocol being highly toxic, perform the following steps under a fume hood, while taking the following precautions: wear two sets of protective gloves and a long-sleeved shirt under their lab coat. Tuck the edges of the gloves inside the sleeves of the shirt so that no skin from the forearms is directly exposed to the chemicals in case of a spillage. Wear protective glasses.
	<ol>
		<li>Dilute 25 mL of pure dimethyldichlorosilane in 475 mL of toluene. Immerse each coverslip in the dimethyldichlorosilane and toluene solution for 15 seconds.</li>
		<li>Wash the coverslips twice in toluene and three times in methanol. Dry the coverslips using pressurized nitrogen.</li>
	</ol>
	</li>
	<li>Assembling coverslips into flow cells
	<ol>
		<li>Once the coverslips are dry, cut a 2 cm x 2.5 cm piece of double-sided tape vertically into two 1 cm x 2.5 cm stripes.</li>
		<li>Put the large coverslip on a delicate task wiper and stick the tape stripes lengthwise along the edges of the coverslip to create a 1 cm x 2.5 cm area between the pieces of tape.</li>
		<li>Stick the small coverslip on top of the tape stripes to finish the flow cell assembly.</li>
	</ol>
	</li>
</ol>
3. Flowing the solutions into a flow cell
<ol>
	<li>Flow approximatively 20 &#956;L of the PEG-PPG-PEG solution into the assembled flow cell. The volume that is flown in the cell has to be large enough to fill the chamber. In the next steps, use the same volume when exchanging the solutions.</li>
	<li>Allow for the PEG-PPG-PEG solution to absorb on the surface for 5 minutes. Exchange the PEG-PPG-PEG solution with BRB80 buffer 3 times by flowing the buffer in. Flow the motility solution into the flow cell.</li>
	<li>Seal the edges of the flow cell with grease to prevent evaporation if the planned experiment is longer than an hour. 
	NOTE: The flow cell is now ready to be imaged.</li>
</ol>
4. Imaging a flow cell
<ol>
	<li>Perform the imaging using an objective-type total internal reflection fluorescence (TIRF) setup for both the microtubules and the kinesin motors (see Table of Materials). 
	NOTE: Here, we used a microscope with a 100x/1.49 numerical aperture objective lens, using two lasers, one with a wavelength of 642 nm and a maximum power of 140 mW, and another with wavelength of 488 nm and a maximum power of 150 mW.</li>
	<li>Place a drop of immersion oil on the objective.</li>
	<li>Place the flow cell on the microscope platform and bring the objective up until there is contact between the oil on the objective and the flow cell.</li>
	<li>Use the microscope&#8217;s interlock cover system to block all laser light from escaping.</li>
	<li>Turn on the laser and focus on the lower surface of the flow cell. The microtubules are fluorescently labelled and excited at a wavelength of 647 nm. They will be imaged using a 642 nm laser while a 488 nm laser is used for the GFP-kinesin motors.</li>
	<li>Record the images or videos of interest. Typically, the laser power is of about 30 mW and an exposure time of 50 ms for both laser channels. Images can be recorded for as long as there is motility in the flow cell. 
	NOTE: The laser illumination is harmful for the naked eye and can cause irreparable damage. Please make sure that the illuminated area is completely covered with an opaque lid.</li>
</ol>

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Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Proteins in Food Processing (Second Edition). , 49-92 (2018).</li><li>Wettermark, G., Borglund, E., Brolin, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+regenerating+system+for+studies+of+phosphoryl+transfer+from+ATP.">A regenerating system for studies of phosphoryl transfer from ATP.</a> Analytical Biochemistry. 22, 211-218 (1968).</li><li>Vigers, G. P. A., Coue, M., McIntosh, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+Microtubules+Break+Up+Under+Illumination.">Fluorescent Microtubules Break Up Under Illumination.</a> Journal of Cell Biology. 107, 1011-1024 (1988).</li><li>Brunner, C., Hess, H., Ernst, K. -. H., Vogel, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lifetime+of+biomolecules+in+hybrid+nanodevices.">Lifetime of biomolecules in hybrid nanodevices.</a> Nanotechnology. 15, S540-S548 (2004).</li><li>Rogers, K. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KIF1D+is+a+fast+non-processive+kinesin+that+demonstrates+novel+K-loop-dependent+mechanochemistry.">KIF1D is a fast non-processive kinesin that demonstrates novel K-loop-dependent mechanochemistry.</a> EMBO Journal. 20, 5101-5113 (2001).</li><li>Patterson, G. H., Knobel, S. M., Sharif, W. D., Kain, S. R., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+the+green+fluorescent+protein+and+its+mutants+in+quantitative+fluorescence+microscopy.">Use of the green fluorescent protein and its mutants in quantitative fluorescence microscopy.</a> Biophysical Journal. 73, 2782-2790 (1997).</li></ol>

The authors have nothing to disclose.

In this work, we present an active nanoscale system which self assembles weakly-binding building blocks to construct its own track. As shown in Figure 1, gliding microtubules accumulate kinesin motors from solution and deposit them on the surface. The kinesin motors remain in the wake of the microtubule for a short period of time before returning to solution. Thus, in this experiment, kinesin motors alternate between 3 states:
(1) A microtubule single-bound state: this is when a kinesin first binds to a microtubule. It exists in equilibrium with state (2).
(2) A double-bound state: in this case, a microtubule single-bound kinesin also binds to the surface via its His-tag. This double-bound state allows for microtubule propulsion.
(3) A single surface-bound state: a double-bound kinesin that has walked off the end of the microtubule and has not yet desorbed from the surface is in this state. These motors can be observed in Figure 1 (combined and green channels): they extend behind the tail of the microtubule for several micrometers and form its diminishing trail.
The most critical step of this protocol is the formation of the hydrophobic surface on the slide. Not only does it use dangerous chemicals, but it also allows the PEG-PPG-PEG functionalized with the NTA group to coat the surface, which then allows the kinesin to reversibly bind to the surface. Another important step is sealing the flow cell with grease. This allows for prolonged imaging without the liquid in the flow cell evaporating.
The primary modifications to this technique consist of changing microtubule concentration, kinesin concentration, and ATP concentration. Changing microtubule concentration will change the number of microtubules gliding on the surface. Changing kinesin concentration will change the number of kinesin molecules that can bind to the microtubule. However, increasing the kinesin concentration above the amounts already defined in this experiment could increase background fluorescence, making it more difficult to see the kinesin trails left behind gliding microtubules. Meanwhile, lowering ATP concentrations below 10 &#181;M will significantly decrease microtubule gliding velocity. If this effect is desired, it is necessary to utilize an ATP regenerating system consisting of creatine phosphatase and phosphokinase.
A possible limitation of this technique is that, due to the large active kinesin content of the system, the ATP can be rapidly consumed, and experiments may last less than an hour in certain conditions. This would for example be the case if one used a twofold higher kinesin concentration and five-fold higher microtubule concentration than what is presented in this protocol.
In our previous work18, we studied the spatial distribution of kinesin motors along the microtubules, proving that gliding microtubules accumulate kinesin motors from solution, resulting in an increase of the density of motors along the length of the microtubule. We also found that the microtubules&#39; gliding stability demonstrated a nonlinear dependence on the solution kinesin concentration and microtubule velocity.
The presented protocol paves the way for a more efficient use of protein motors in nanoscale engineered systems and for further investigation in the design of active nanosystems that are in dynamic equilibrium. Furthermore, the dynamic nature of this system allows it to serve as a model system for studying self-healing and dynamic replacement of molecular components, closing part of the gap between engineered and natural structures.

The interactions governing the behavior of active nanosystems have always been characterized by long-lived, nearly irreversible bonds1,2,3,4,5,6,7,8. A well-studied example of this is the microtubule-kinesin system, where gliding microtubules are propelled by irreversibly surface-bound kinesin motors1,2,3,4,5. Systems in which the components are reversibly connected to one another have been studied theoretically9,10 and achieved at the macroscale11,12, but scaling these systems down to the nanoscale has been challenging. One of the major reasons for this is that breaking and reforming bonds between components often requires a large change in the environmental conditions. Even though such changes have been implemented in the past13,14,15, they would rely on modifying the system itself rather than adapting it to its environment. Designing molecular-scale systems in which components continually assemble and reorganize into structures without disturbing the overall environment in which the experiments take place will open the door to the exploration of a wide range of dynamic behaviors16,17.
Here, we describe and demonstrate the detailed protocol for creating a dynamically assembling and disassembling system functioning at the nanoscale. The system and its general behavior has been introduced earlier18: microtubule filaments are propelled by tracks of reversibly surface-bound kinesin-1 motors. These kinesin motor proteins are recruited from the solution to help propel microtubules forward, before desorbing again shortly afterwards. Once back in solution, they can be recruited again to propel a new microtubule. In the past13,14,15, the breaking and reforming of bonds required environmental modifications; in contrast, the environment of our flow cell remains unchanged while the kinesin motors interact with the surface.
This protocol will help interested researchers to (1) visualize all the steps of the protocol, and (2) assist with troubleshooting this type of assay. It has been derived from the procedures described in Howard et al. 199319.

<table>
	<tbody>
		<tr>
			<td>488 nm laser</td>
			<td>Omicron Laserage</td>
			<td>LuxX 488-150</td>
			<td></td>
		</tr>
		<tr>
			<td>642 nm laser</td>
			<td>Omicron Laserage</td>
			<td>LuxX&#160;642</td>
			<td></td>
		</tr>
		<tr>
			<td>Casein</td>
			<td>Sigma</td>
			<td>C7078-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Catalase from bovine liver</td>
			<td>Sigma</td>
			<td>C40-500MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Creatine Phosphate</td>
			<td>Sigma</td>
			<td>P-7936</td>
			<td></td>
		</tr>
		<tr>
			<td>Creatine Phosphokinase</td>
			<td>Sigma</td>
			<td>C3755-500UN</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>Sigma</td>
			<td>G2133-50KU</td>
			<td></td>
		</tr>
		<tr>
			<td>Dichlorodimethylsilane solution</td>
			<td>Sigma</td>
			<td>40140-25ML</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Sigma</td>
			<td>34869-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>Sigma</td>
			<td>D0632-5G</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>Eclipse TI</td>
			<td>Nikon Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>eGFP rkin430</td>
			<td>Provided by George Bachand</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma</td>
			<td>E4378-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon 50 mL Conical Centrifuge Tubes</td>
			<td>Fisher Scientific</td>
			<td>14-959-49A</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose Oxidase</td>
			<td>Sigma</td>
			<td>G0543-10KU</td>
			<td></td>
		</tr>
		<tr>
			<td>Guanosine Triphosphate</td>
			<td>Sigma</td>
			<td>G8877-10MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes Delicate Task Wipers</td>
			<td>Sigma Pharmaceuticals</td>
			<td>8089</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Sigma</td>
			<td>M1028-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Fisher Chemical</td>
			<td>A412</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>Milli-Q Water Purification System</td>
			<td>Millipore Corporation</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nickel Sulfate</td>
			<td>Sigma</td>
			<td>656895-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Paclitaxel</td>
			<td>Sigma</td>
			<td>T1912-5MG</td>
			<td></td>
		</tr>
		<tr>
			<td>PIPES</td>
			<td>Sigma</td>
			<td>P-6757</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F108-NTA</td>
			<td>Provided by Jennifer Neff and AllVivo Vascular</td>
			<td></td>
			<td>PEG-PPG-PEG-NTA</td>
		</tr>
		<tr>
			<td>Pluronic&#160;F-108</td>
			<td>Sigma</td>
			<td>542342-250G</td>
			<td>PEG-PPG-PEG</td>
		</tr>
		<tr>
			<td>Thermo Scientific Snap Cap Low Retention Microcentrifuge Tubes</td>
			<td>Fisher Scientific</td>
			<td>21-403-190</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluene</td>
			<td>Fisher Chemical</td>
			<td>T324</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>Tubulin, HiLyte647-labeled</td>
			<td>Cytoskeleton, Inc.</td>
			<td>TL670M</td>
			<td></td>
		</tr>
		<tr>
			<td>UV Ozone Procleaner</td>
			<td>BioForce Nanosciences</td>
			<td>PC440</td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman&#160;Puradisc syringe filters</td>
			<td>Sigma</td>
			<td>WHA67840402</td>
			<td></td>
		</tr>
		<tr>
			<td>Zyla 4.2 sCMOS Camera</td>
			<td>Andor Technology</td>
			<td>sCMOS 4.2</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

In these experiments, we used a 1,000 times dilution of the microtubules prepared in section 1.3.2. The kinesin concentration was 20 nM, and the ATP concentration was 1 mM. Imaging was performed using TIRF microscopy. Gliding microtubules were separately imaged from the kinesin motors: microtubules were visible upon excitation with a 647 nm laser (Figure 1, red), and the GFP-kinesin was visible when excited with a 488 nm laser (Figure 1, green). The time between excitation with the red and green light was less than 1 s. The time between frames was 10 s. Microtubules displayed stable gliding. The average microtubule gliding velocity was approximatively 800 nm/s. The microtubule surface density was 400 mm-2. Tracks of kinesin (green) appeared to extend beyond the trailing end of the microtubules (red) for several micrometers.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59068/59068fig1.jpg" /> 
Figure 1: Gliding microtubules propelled by weakly surface-bound kinesin motors. As microtubules move forward, they accumulate kinesin motors from solution. These motors alternate between two states: single-bound to the microtubule, and double-bound to both the microtubule and the surface. When a double bound motor reaches the end of the microtubule, the motor is left behind and slowly desorbs from the surface with an off rate of approximately 0.1 s-1. As a result, trails of kinesin motors remain behind the microtubules. Top row: red (microtubule) channel. Middle row: green (kinesin) channel. Bottom row: combined red (microtubule) and green (GFP-kinesin) channel. Scale bar: 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59068/59068fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Assembling Molecular Shuttles Powered by Reversibly Attached Kinesins at JoVE.com

Assembling Molecular Shuttles Powered by Reversibly Attached Kinesins

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 ...

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6.8K Views.  Columbia University. 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.

Video: Assembling Molecular Shuttles Powered by Reversibly Attached Kinesins

Molecular Beam Mass Spectrometry With Tunable Vacuum Ultraviolet (VUV) Synchrotron Radiation

Tunable soft ionization coupled to mass spectroscopy is a powerful method to investigate isolated molecules, complexes and clusters and their spectroscopy and dynamics1-4. Fundamental studies of photoionization processes of biomolecules provide information about the electronic structure of these systems. Furthermore determinations of ionization energies and other properties of biomolecules in the gas phase are not trivial, and these experiments provide a platform to generate these data. We have developed a thermal vaporization technique coupled with supersonic molecular beams that provides a gentle way to transport these species into the gas phase. Judicious combination of source gas and temperature allows for formation of dimers and higher clusters of the DNA bases. The focus of this particular work is on the effects of non-covalent interactions, i.e., hydrogen bonding, stacking, and electrostatic interactions, on the ionization energies and proton transfer of individual biomolecules, their complexes and upon micro-hydration by water1, 5-9.
We have performed experimental and theoretical characterization of the photoionization dynamics of gas-phase uracil and 1,3-dimethyluracil dimers using molecular beams coupled with synchrotron radiation at the Chemical Dynamics Beamline10 located at the Advanced Light Source and the experimental details are visualized here. This allowed us to observe the proton transfer in 1,3-dimethyluracil dimers, a system with pi stacking geometry and with no hydrogen bonds1. Molecular beams provide a very convenient and efficient way to isolate the sample of interest from environmental perturbations which in return allows accurate comparison with electronic structure calculations11, 12. By tuning the photon energy from the synchrotron, a photoionization efficiency (PIE) curve can be plotted which informs us about the cationic electronic states. These values can then be compared to theoretical models and calculations and in turn, explain in detail the electronic structure and dynamics of the investigated species 1, 3.

Molecular Beam Mass Spectrometry With Tunable Vacuum Ultraviolet VUV Synchrotron Radiation

A molecular beam coupled to tunable vacuum ultraviolet photoionization mass spectrometer at a synchrotron provides a convenient tool to explore the electronic structure of isolated gas phase molecules and clusters. Proton transfer mechanisms in DNA base dimers were elucidated with this technique.

Tunable soft ionization coupled to mass spectroscopy is a powerful method to investigate isolated molecules, complexes and clusters and their spectroscopy ...

Ultrahigh Density Array of Vertically Aligned Small-molecular Organic Nanowires on Arbitrary Substrates

In recent years &pi;-conjugated organic semiconductors have emerged as the active material in a number of diverse applications including large-area, low-cost displays, photovoltaics, printable and flexible electronics and organic spin valves. Organics allow (a) low-cost, low-temperature processing and (b) molecular-level design of electronic, optical and spin transport characteristics. Such features are not readily available for mainstream inorganic semiconductors, which have enabled organics to carve a niche in the silicon-dominated electronics market. The first generation of organic-based devices has focused on thin film geometries, grown by physical vapor deposition or solution processing. However, it has been realized that organic nanostructures can be used to enhance performance of above-mentioned applications and significant effort has been invested in exploring methods for organic nanostructure fabrication.	
A particularly interesting class of organic nanostructures is the one in which vertically oriented organic nanowires, nanorods or nanotubes are organized in a well-regimented, high-density array. Such structures are highly versatile and are ideal morphological architectures for various applications such as chemical sensors, split-dipole nanoantennas, photovoltaic devices with radially heterostructured "core-shell" nanowires, and memory devices with a cross-point geometry. Such architecture is generally realized by a template-directed approach. In the past this method has been used to grow metal and inorganic semiconductor nanowire arrays. More recently &pi;-conjugated polymer nanowires have been grown within nanoporous templates. However, these approaches have had limited success in growing nanowires of technologically important &pi;-conjugated small molecular weight organics, such as tris-8-hydroxyquinoline aluminum (Alq3), rubrene and methanofullerenes, which are commonly used in diverse areas including organic displays, photovoltaics, thin film transistors and spintronics.	
Recently we have been able to address the above-mentioned issue by employing a novel "centrifugation-assisted" approach. This method therefore broadens the spectrum of organic materials that can be patterned in a vertically ordered nanowire array. Due to the technological importance of Alq3, rubrene and methanofullerenes, our method can be used to explore how the nanostructuring of these materials affects the performance of aforementioned organic devices. The purpose of this article is to describe the technical details of the above-mentioned protocol, demonstrate how this process can be extended to grow small-molecular organic nanowires on arbitrary substrates and finally, to discuss the critical steps, limitations, possible modifications, trouble-shooting and future applications.

We report a simple method for fabricating an ultrahigh density array of vertically ordered small-molecular organic nanowires. This method allows for synthesis of complex heterostructured hybrid nanowire geometries, which can be inexpensively grown on arbitrary substrates. These structures have potential applications in organic electronics, optoelectronics, chemical sensing, photovoltaics and spintronics.

In  recent years &pi;-conjugated  organic semiconductors have emerged as the active material in a number of  diverse applications including large-area, ...

Spatial Separation of Molecular Conformers and Clusters

Gas-phase molecular physics and physical chemistry experiments commonly use supersonic expansions through pulsed valves for the production of cold molecular beams. However, these beams often contain multiple conformers and clusters, even at low rotational temperatures. We present an experimental methodology that allows the spatial separation of these constituent parts of a molecular beam expansion. Using an electric deflector the beam is separated by its mass-to-dipole moment ratio, analogous to a bender or an electric sector mass spectrometer spatially dispersing charged molecules on the basis of their mass-to-charge ratio. This deflector exploits the Stark effect in an inhomogeneous electric field and allows the separation of individual species of polar neutral molecules and clusters. It furthermore allows the selection of the coldest part of a molecular beam, as low-energy rotational quantum states generally experience the largest deflection. Different structural isomers (conformers) of a species can be separated due to the different arrangement of functional groups, which leads to distinct dipole moments. These are exploited by the electrostatic deflector for the production of a conformationally pure sample from a molecular beam. Similarly, specific cluster stoichiometries can be selected, as the mass and dipole moment of a given cluster depends on the degree of solvation around the parent molecule. This allows experiments on specific cluster sizes and structures, enabling the systematic study of solvation of neutral molecules.

We present a technique that allows the spatial separation of different conformers or clusters present in a molecular beam. An electrostatic deflector is used to separate species by their mass-to-dipole moment ratio, leading to the production of gas-phase ensembles of a single conformer or cluster stoichiometry.

Gas-phase molecular physics and physical chemistry experiments commonly use supersonic expansions through pulsed valves for the production of cold ...

Measurement and Analysis of Atomic Hydrogen and Diatomic Molecular AlO, C2, CN, and TiO Spectra Following Laser-induced Optical Breakdown

In this work, we present time-resolved measurements of atomic and diatomic spectra following laser-induced optical breakdown. A typical LIBS arrangement is used. Here we operate a Nd:YAG laser at a frequency of 10 Hz at the fundamental wavelength of 1,064 nm. The 14 nsec pulses with anenergy of 190 mJ/pulse are focused to a 50 &#181;m spot size to generate a plasma from optical breakdown or laser ablation in air. The microplasma is imaged onto the entrance slit of a 0.6 m spectrometer, and spectra are recorded using an 1,800 grooves/mm grating an intensified linear diode array and optical multichannel analyzer (OMA) or an ICCD. Of interest are Stark-broadened atomic lines of the hydrogen Balmer series to infer electron density. We also elaborate on temperature measurements from diatomic emission spectra of aluminum monoxide (AlO), carbon (C2), cyanogen (CN), and titanium monoxide (TiO).
The experimental procedures include wavelength and sensitivity calibrations. Analysis of the recorded molecular spectra is accomplished by the fitting of data with tabulated line strengths. Furthermore, Monte-Carlo type simulations are performed to estimate the error margins. Time-resolved measurements are essential for the transient plasma commonly encountered in LIBS.

Measurement and Analysis of Atomic Hydrogen and Diatomic Molecular AlO, C2, CN, and TiO Spectra Following Laser-induced Optical Breakdown

Time-resolved atomic and diatomic molecular species are measured using LIBS. The spectra are collected at various time delays following the generation of optical breakdown plasma with Nd:YAG laser radiation&#160;and are analyzed to infer electron density and temperature.

In this work, we present time-resolved measurements of atomic and diatomic spectra following laser-induced optical breakdown. A typical LIBS arrangement ...

Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards understanding its mechanism. Protonation and conformational changes affect the vibration pattern of amino acid side chains and of the peptide bond, respectively, both of which can be probed by infrared (IR) difference spectroscopy. For proteins whose function can be repetitively and reproducibly triggered by light, it is possible to obtain infrared difference spectra with (sub)microsecond resolution over a broad spectral range using the step-scan Fourier transform infrared technique. With ~102-103 repetitions of the photoreaction, the minimum number to complete a scan at reasonable spectral resolution and bandwidth, the noise level in the absorption difference spectra can be as low as ~10-4, sufficient to follow the kinetics of protonation changes from a single amino acid. Lower noise levels can be accomplished by more data averaging and/or mathematical processing. The amount of protein required for optimal results is between 5-100 &#181;g, depending on the sampling technique used. Regarding additional requirements, the protein needs to be first concentrated in a low ionic strength buffer and then dried to form a film. The protein film is hydrated prior to the experiment, either with little droplets of water or under controlled atmospheric humidity. The attained hydration level (g of water / g of protein) is gauged from an IR absorption spectrum. To showcase the technique, we studied the photocycle of the light-driven proton-pump bacteriorhodopsin in its native purple membrane environment, and of the light-gated ion channel channelrhodopsin-2 solubilized in detergent.

Key steps of protein function, in particular backbone conformational changes and proton transfer reactions, often take place in the microsecond to millisecond time scale. These dynamical processes can be studied by time-resolved step-scan Fourier-transform infrared spectroscopy, in particular for proteins whose function is triggered by light.

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards ...

Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. At the same time the AFM tip is passivated to prevent unspecific interactions between the tip and the substrate, which is a prerequisite to study single molecules attached to the AFM tip. Analyses to determine the adhesion force, the adhesion length, and the free energy of these molecules on solid surfaces and bio-interfaces are shortly presented and external references for further reading are provided. Example molecules are the poly(amino acid) polytyrosine, the graft polymer PI-g-PS and the phospholipid POPE (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine). These molecules are desorbed from different surfaces like CH3-SAMs, hydrogen terminated diamond and supported lipid bilayers under various solvent conditions. Finally, the advantages of force spectroscopic single molecule experiments are discussed including means to decide if truly a single molecule has been studied in the experiment.

A protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. Procedures and examples to determine the adhesion force and free energy of these molecules on solid supports and bio-interfaces are provided.

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single ...

Probing C84-embedded Si Substrate Using Scanning Probe Microscopy and Molecular Dynamics

This paper reports an array-designed C84-embedded Si substrate fabricated using a controlled self-assembly method in an ultra-high vacuum chamber. The characteristics of the C84-embedded Si surface, such as atomic resolution topography, local electronic density of states, band gap energy, field emission properties, nanomechanical stiffness, and surface magnetism, were examined using a variety of surface analysis techniques under ultra, high vacuum (UHV) conditions as well as in an atmospheric system. Experimental results demonstrate the high uniformity of the C84-embedded Si surface fabricated using a controlled self-assembly nanotechnology mechanism, represents an important development in the application of field emission display (FED), optoelectronic device fabrication, MEMS cutting tools, and in efforts to find a suitable replacement for carbide semiconductors. Molecular dynamics (MD) method with semi-empirical potential can be used to study the nanoindentation of C84-embedded Si substrate. A detailed description for performing MD simulation is presented here. Details for a comprehensive study on mechanical analysis of MD simulation such as indentation force, Young&#39;s modulus, surface stiffness, atomic stress, and atomic strain are included. The atomic stress and von-Mises strain distributions of the indentation model can be calculated to monitor deformation mechanism with time evaluation in atomistic level.

Probing C84-embedded Si Substrate Using Scanning Probe Microscopy and Molecular Dynamics

This paper reports the nanomaterial fabrication of a fullerene Si substrate inspected and verified by nanomeasurements and molecular dynamic simulation.

This paper reports an array-designed C84-embedded Si substrate fabricated using a controlled self-assembly method in an ultra-high vacuum chamber. The ...

Quantitative Hardness Measurement by Instrumented AFM-indentation

In this work, a combination of amplitude-modulated non-contact atomic force microscopy and atomic force spectroscopy is applied for instrumented hardness measurements on an Au(111) surface with atomistic resolution of single plasticity events. A careful experimental procedure is described that includes the force sensor selection, its calibration, the calibration of the cantilever deflection detection system, and the minimization of instrumental drift for accurate and reproducible force-distance measurements. Also, a method for the data analysis is presented that allows the extraction of force-penetration curves from recorded force-distance curves. A typical curve displays a clear elastic deformation regime up to the first plasticity event, or pop-in, with a length in the range of one to two Burger's vectors. Later plasticity events exhibit the same magnitude. The work of plasticity is further extracted from the measurements. Finally, the hardness is determined in combination with the indentation curve using non-contact atomic force microscopy images of the remaining indents.

This experimental protocol describes how atomic force microscopy can be used to measure hardness at the true nanometer scale and to detect single atomistic plasticity events.

In this work, a combination of amplitude-modulated non-contact atomic force microscopy and atomic force spectroscopy is applied for instrumented hardness ...

Plasma-assisted Molecular Beam Epitaxy of N-polar InAlN-barrier High-electron-mobility Transistors

Plasma-assisted molecular beam epitaxy is well suited for the epitaxial growth of III-nitride thin films and heterostructures with smooth, abrupt interfaces required for high-quality high-electron-mobility transistors (HEMTs). A procedure is presented for the growth of N-polar InAlN HEMTs, including wafer preparation and growth of buffer layers, the InAlN barrier layer, AlN and GaN interlayers and the GaN channel. Critical issues at each step of the process are identified, such as avoiding Ga accumulation in the GaN buffer, the role of temperature on InAlN compositional homogeneity, and the use of Ga flux during the AlN interlayer and the interrupt prior to GaN channel growth. Compositionally homogeneous N-polar InAlN thin films are demonstrated with surface root-mean-squared roughness as low as 0.19 nm and InAlN-based HEMT structures are reported having mobility as high as 1,750 cm2/V&#8729;sec for devices with a sheet charge density of 1.7 x 1013 cm-2.

Molecular beam epitaxy is used to grow N-polar InAlN-barrier high-electron-mobility transistors (HEMTs). Control of the wafer preparation, layer growth conditions and epitaxial structure results in smooth, compositionally homogeneous InAlN layers and HEMTs with mobility as high as 1,750 cm2/V&#8729;sec.

Plasma-assisted molecular beam epitaxy is well suited for the epitaxial growth of III-nitride thin films and heterostructures with smooth, abrupt ...

Scanning SQUID Study of Vortex Manipulation by Local Contact

Local, deterministic manipulation of individual vortices in type 2 superconductors is challenging. The ability to control the position of individual vortices is necessary in order to study how vortices interact with each other, with the lattice, and with other magnetic objects. Here, we present a protocol for vortex manipulation in thin superconducting films by local contact, without applying current or magnetic field. Vortices are imaged using a scanning superconducting quantum interference device (SQUID), and vertical stress is applied to the sample by pushing the tip of a silicon chip into the sample, using a piezoelectric element. Vortices are moved by tapping the sample or sweeping it with the silicon tip. Our method allows for effective manipulation of individual vortices, without damaging the film or affecting its topography. We demonstrate how vortices were relocated to distances of up to 0.8 mm. The vortices remained stable at their new location up to five days. With this method, we can control vortices and move them to form complex configurations. This technique for vortex manipulation could also be implemented in applications such as vortex based logic devices.

We present a protocol for manipulation of individual vortices in thin superconducting films, using local mechanical contact. The method does not include applying current, magnetic field or additional fabrication steps.

Local, deterministic manipulation of individual vortices in type 2 superconductors is challenging. The ability to control the position of individual ...