Name: surface passivation for single-molecule protein studies
Uploaded: 2014-04-24T13:00:00
Duration: 10 min 35 s
Description: Watch this Scientific Journal Video about Surface Passivation for Single-molecule Protein Studies at JoVE.com

S.D.C., A.C.H., and C.J. were supported by Starting Grants (ERC-StG-2012-309509) through the European Research Council. J.-M.N. was supported by the National Research Foundation (NRF) (2011-0018198) of Korea; and the Pioneer Research Center Program (2012-009586) through the NRF of Korea funded by the Ministry of Science, ICT, and Future Planning (MSIP). This work was also supported by Center for BioNano Health-Guard funded by MSIP of Korea as the Global Frontier Project (H-GUARD_2013-M3A6B2078947). Y.K.L. and J.-H.H. were supported by the Seoul Science Fellowship Program of Seoul City, Korea. The labeled Rep protein was a generous gift from Dr. Sua Myong.

1. Slide Preparation and Cleaning
A microfluidic chamber is composed of a quartz slide and a coverslip. In prism-type total internal reflection fluorescence (TIRF) microscopy, the surface of the quartz slide is imaged. Therefore, it is important to clean a quartz slide thoroughly using H2O, acetone, KOH, and piranha solutions. This multi-step cleaning eliminates fluorescent organic molecules on a surface which interfere with single molecule fluorescence measurements. Additionally, the piranha etching makes the quartz surface hydrophilic by generating hydroxyl groups. The free hydroxyl groups are essential for the amino-silanization reaction in Step 3.
<ol>
	<li>Drilling quartz slides. Drill a pair of holes into a quartz slide with a 3/4 mm diamond drill bit (Figure 2a). If multiple channels are desired, drill more pairs of holes (Figure 2b). These holes are used for injecting solutions into a microfluidic chamber in Step 7.
	<ol>
		<li>Keep the drill bit wet in H2O during the drilling. H2O performs as a lubricant, which increases the lifetime of the drill bit.</li>
		<li>Occasional wiping the tip of the drill bit helps drilling holes.</li>
		<li>After drilling holes, mark one side of the slide for easy identification during the PEGylation process. The markers can be used as reference to avoid confusion later on. For marking, a diamond drill bit can be used. Do not use any pen since the ink might get on the surface.</li>
	</ol>
	</li>
	<li>Cleaning with H2O. Place the slides in a glass staining jar. Typically 5 to 15 slides can be placed in a single jar. Rinse the slides with MilliQ H2O. Repeat it 3 times and sonicate the slides with MilliQ H2O for 5 min to remove dirt. Dispose the water and rinse the slides again 3 times with MilliQ H2O. Note that 130 W is the sonication power used for this protocol. Higher sonication power might decrease the lifetime of the quartz slides.</li>
	<li>Cleaning with acetone. Replace the MilliQ H2O with acetone. Sonicate the slides with acetone for 20 min or longer.</li>
	<li>Rinsing with H2O. Discard the acetone and rinse the slides with MilliQ H2O. Repeat it for 3 times in order to remove any acetone residue.</li>
	<li>Cleaning with KOH. Replace the water with 1 M KOH and sonicate the slides for 20 min or longer. Excessive etching (e.g. overnight KOH treatment) will enhance the quality of the surface, but will introduce scratches, which might interfere with fluorescence imaging.</li>
	<li>Rinsing with H2O. Rinse the slides with MilliQ H2O for 3 times to remove traces of KOH.</li>
	<li>Piranha etching.
	<ol>
		<li>Transfer the slides to a Duran slide holder or a custom-made Teflon holder and place them in a beaker (1 L) that is located in a chemical hood.</li>
		<li>Fill the beaker with 450 ml of H2SO4.</li>
		<li>Add 150 ml of H2O2 for 3:1 ratio between H2SO4 and H2O2. Once the solution starts to boil spontaneously, the temperature increases over 90 &#176;C. Make sure that the H2O2 solution is at room temperature before starting the reaction. Otherwise, the temperature of the boiling solution may be lower than 90 &#176;C.</li>
		<li>Stir the solution for proper mixing and let the beaker undisturbed for 20 min.</li>
		<li>Take the slides out of the piranha solution together with the slide holder, and put them in a slide holder containing MilliQ H2O. Meanwhile, discard the piranha solution into a designated waste bottle once it reaches room temperature.</li>
		<li>Rinse the slides 3 times with MilliQ H2O. Extra care should be taken in further steps to avoid any possible contamination of slides. Continue to Step 3. It is advised to immediately proceed with the following steps. 
		* Caution: The piranha solution is extremely reactive. When handling this solution extra caution should be taken. In addition, when by mistake mixed with organic solvents such as acetone, it may cause an explosion.</li>
	</ol>
	</li>
</ol>
2. Coverslip Cleaning
A microfluidic chamber is composed of a quartz slide and a coverslip. When a prism-type TIRF microscope is used, the surface of a coverslip is not imaged. Therefore, it is enough to clean a coverslip only with H2O and KOH. In case a coverslip needs to be imaged (e.g. via the objective-type TIRF microscopy), it is recommended to treat the coverslips with piranha solution (Step 1.7). Note that if PEGylation of the coverslip surface is not of high quality, it might act as a sink for proteins and give rise to variations in the protein concentration.
<ol>
	<li>Rinsing with H2O. Place coverslips (24 x 30, 24 x 40, or 24 x 50 mm2) in a glass staining jar. Typically 5 to 15 coverslips can be placed in a single jar. Rinse the coverslips 3 times with the MilliQ H2O.</li>
	<li>Cleaning with KOH. Replace the water with 1M KOH and sonicate the coverslips for 20 min or longer.</li>
	<li>Rinsing with H2O. Rinse the coverslips 3 times with MilliQ H2O to remove traces of KOH. Continue to Step 3.</li>
</ol>
3. Amino-silanization of Slides and Coverslips
Functionalizing the surface of the quartz slides and the coverslips with amine group via the amino-silanization chemistry. Methanol is used as a solvent and acetic acid as a catalyst for the amino-silanization reaction.
<ol>
	<li>Rinsing with methanol. Replace the MilliQ H2O in the staining dishes (from Steps 1 and 2) with methanol. Keep the slides and the coverslips in methanol until Step 3.3. Do not store the slides and the coverslips in methanol for an unnecessarily long period of time (e.g. several hours) since impurities in methanol will adsorb to the surface.</li>
	<li>Preparing amino-silanization solution.
	<ol>
		<li>Rinse a Pyrex flask several times with methanol. Sonicate the flask with methanol for 5 min or longer. It is advised to have a dedicated flask that is maintained clean.</li>
		<li>Pour 100 ml of methanol into the flask.</li>
		<li>Add 5 ml of acetic acid.</li>
		<li>Add 3 ml of APTES (3-aminopropyl trimethoxysilane) and gently mix by shaking it.</li>
	</ol>
	</li>
	<li>Amino-silanization. Replace the methanol in the staining dishes that contain the slides and the coverslips with the aminosilanization reaction mixture.
	<ol>
		<li>Incubate for 20-30 min. During incubation, sonicate once for 1 min.</li>
	</ol>
	</li>
	<li>Rinsing with methanol. Replace the aminosilanization reaction with methanol. Discard the methanol and add a fresh methanol solution. Repeat this procedure three times.</li>
</ol>
4. Surface Passivation Using Polymer (The First Round)
Passivating the amine-coated surface of quartz slides and coverslips by conjugating NHS-ester polyethylene glycol (PEG). This reaction is carried out with the saturating concentration of PEG solution at pH 8.5 overnight.
<ol>
	<li>Drying slides and coverslips. Dry the slides and the coverslips using N2 gas and place them in clean pipette boxes in such a way that the side which has to be PEGylated is facing up. The pipette box is partially filled with MilliQ H2O. This moist environment prevents PEGylation solution from drying out during the overnight incubation.</li>
	<li>Preparing reaction buffer. Prepare 0.1 M of fresh sodium bicarbonate buffer (pH 8.5) by dissolving 84 mg of sodium bicarbonate in 10 ml of MilliQ H2O. There is no need of adjusting pH. This solution may be stored frozen for the next use (e.g. Step 6.1).</li>
	<li>Preparing PEGylation solution.
	<ol>
		<li>Prepare a PEG mixture of 0.2 mg biotinylated NHS-ester PEG (5,000 Da) 13 and 8 mg of NHS-ester mPEG (5,000 Da) in a 1.5 ml tube. When preparing N number of slides, prepare N times larger amount of the PEG mixture in a single tube.</li>
		<li>Add 64 &#181;l (or N times 64 &#181;l for N number of slides) of the freshly prepared buffer (Step 4.2). Pipette it up and down in order to dissolve them completely.</li>
		<li>Centrifuge with 16,100 x g for 1 min to remove air bubbles. Do not pipette afterwards. Otherwise, air bubbles will form which interfere with PEGylation at Step 4.4.</li>
	</ol>
	</li>
	<li>PEGylation.
	<ol>
		<li>Drop 70 &#181;l of the PEGylation mixture to a dried quartz slide from Step 4.1. Use 90 &#181;l in case of 24 x 50 mm2 coverslip.</li>
		<li>Gently place a dried coverslip from Step 4.1 over the solution.</li>
		<li>To have a uniform and high quality of PEGylation, take care not to introduce air bubbles. Due to surface tension, most of the air bubbles may spontaneously leave the reaction volume within five minutes due to surface tension.</li>
		<li>Incubate the slides in a dark and humid environment. The half-life of NHS-ester PEG that we use at pH 8 is about one hour. At a minimum, incubate them for 2 hr. Overnight incubation leads to the higher quality of PEGylation (data not shown).</li>
	</ol>
	</li>
</ol>
5. Long-term Storage
Storing the PEGylated slides and coverslips in N2 at -20 &#186;C.
<ol>
	<li>Drying slides and coverslips. Carefully disassemble the slide and the coverslip by sliding the coverslip to a side, rinse them with MilliQ H2O, and dry them with N2.</li>
	<li>Storing slides and coverslips. For immediate use, follow the procedure for the second round of PEGylation (Step 6). In order to store for a longer period of time, follow the next steps.
	<ol>
		<li>Place a pair of the slide and the coverslip in a 50 ml tube such that the PEGylated surfaces are facing away from each other.</li>
		<li>Partially close the tube, vacuum the tube, and then fill it with N2. These steps help preserving the PEGylated surface for a long period of time. Screw the tube tightly and store it at -20 &#176;C. The quality of the slides remains good for up to 3 months (Figure 3a, right).</li>
	</ol>
	</li>
</ol>
6. Surface Passivation Using Polymer (The Second Round)
Additional round of PEGylation to make the PEG layer denser and also to quench any remaining amine groups on the surface. The use of short NHS-ester PEG molecules (333 Da) may be effective in penetrating into an existing PEG layer. It is recommended to do this second round of PEGylation right before using a slide.
<ol>
	<li>Preparing reaction buffer. Prepare 0.1 M of fresh sodium bicarbonate buffer (pH 8.5). The frozen solution from Step 4.2 can also be used.</li>
	<li>Preparing PEGylation solution. Dissolve 7 &#181;l of 250 mM MS4-PEG in 63 &#956;l of the sodium bicarbonate buffer.</li>
	<li>PEGylation. Follow the same procedure as in Step 4.4. Incubate for 30 min up to overnight.</li>
	<li>Drying slides and coverslips. Disassemble the pair of the slide and the coverslip, rinse them with MilliQ H2O, dry them with N2, and keep them in a clean pipette box. Proceed for assembling a microfluidic chamber in Step 7.</li>
</ol>
7. Assembling a Microfluidic Chamber
Assembling a microfluidic chamber using a pair of a PEGylated quartz slide and a coverslip. Double-sided sticky tape is used as a spacer. The chamber is sealed with Epoxy glue and, solutions are introduced through the holes in the quartz slide.
<ol>
	<li>Place a quartz slide on a flat surface with the PEGylated side facing up.</li>
	<li>Make a channel (width 5 to 7 mm) diagonally on the PEGylated surface by putting double-sided sticky tapes over the slide. Make sure that the holes are positioned at the center of the channel. Take care not to spoil the PEGylated surface during the process of making a chamber. It is possible to make a multi-channel slide by placing and sticking the double-sided sticky tapes differently (see Figure 2).</li>
	<li>Gently place a PEGylated coverslip on top to complete the chamber. The PEGylated side should be facing down.</li>
	<li>Seal the chamber by pressing the coverslip over the area where double-sided tapes are placed. Do it gently but thoroughly so that the chamber becomes water-tightly sealed.</li>
	<li>Close the edges of the chamber with Epoxy glue.</li>
	<li>Immobilize Streptavidin or Neutravidin on the biotinylated PEG layer by adding 50 &#181;l of 0.1 mg/ml of Streptavidin or Neutravidin solution in T50 (10 mM Tris-HCl [pH8.0], 50 mM NaCl buffer) using a p200 pipette. After 1 min of incubation, flush with 100 &#181;l of T50 buffer.</li>
	<li>Add biotinylated biological molecules for your single-molecule imaging.</li>
</ol>
8. Slide Recycling
Recycling quartz slides. Used slides are recycled by taking off the coverslips and the double-sided sticky tapes.
<ol>
	<li>After use, store the chambers in tap water. Long-term incubation in water eases disassembly of the chambers.</li>
	<li>Boil the chambers in tap water using a microwave. Use a Pyrex beaker. Boil for 10 min or longer.</li>
	<li>Take off the coverslips and the double-sided sticky tapes using a razor blade. Do not press a quartz slide perpendicular to its plane. Otherwise, it breaks. Keep the razor blade away from the channel. Otherwise, it introduces scratches on the channel.</li>
	<li>Rinse the slides using a household detergent by rubbing them with fingers.</li>
	<li>Place the slides in a glass staining jar. Typically 5 to 15 slides can be placed in a single jar. Add 10% household detergent and sonicate the slides for 20 min or longer. Rinse the slides with a large quantity of tab water.</li>
	<li>Go to Step 1.2.</li>
</ol>

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We have nothing to disclose.

Critical steps within this protocol
It is essential to make the surface hydrophilic before the amino-silanization reaction. This was achieved through the piranha etching which generates the free hydroxyl groups on a glass/quartz surface. It is recommended not to keep the piranha etched surface exposed to either H2O or air for a long period of time since the hydrophilicity of the surface goes down gradually.
The NHS-ester PEG molecules are reactive. It is advised to make aliquots and store them under nitrogen in -20 &#186;C. The shelf life of the APTES chemical at the room temperature is short. It is advised to replace it with a new one every month.
Modifications of this protocol
When you cannot use the piranha etching for any practical reason, you may etch using KOH for a long period of time (e.g. overnight), which will also make hydroxyl groups exposed. The pitfall of this alternative approach is that the slide becomes unusable after a few recycles due to severe scratches.
It is often practiced to burn a glass/quartz surface using propane torch, which is effective in eliminating fluorescent organic materials 7. This procedure was not included in this protocol because it is redundant to the piranha etching. Note that this procedure will potentially lead to oxidation of the hydroxyl groups. Therefore, this procedure should not be carried out once a surface was etched using KOH or piranha solution.
When a buffer with pH lower than 7 is used for single-molecule imaging, the conformation of PEG changes from &#8220;mushroom&#8221; to &#8220;brush,&#8221; which reduces the degree of passivation. Therefore, when pH lower than 7.0 is used, it is recommended to further passivate the surface using disuccinimidyl tartarate 7.
Perspectives
This work has provided a robust protocol for achieving high quality surface passivation. This protocol will be useful for single-molecule fluorescence studies that are involved with proteins 14 as well as protein complexes within cell extracts and immunoprecipitates 15. It will be widely used for other single-molecule techniques such as force spectroscopy and torque spectroscopy 16. It will be also useful for preventing cells from adsorbing to a surface 17.
The protocol provided in this work is demanding for its multi-step procedures. Surface passivation using lipid-PEG 8 and poly-lysine PEG 9 are available as an alternative approach. Since, they do not require any chemical reactions it is easy to implement. However, the degree of passivation is not as high as that achieved through chemical modification of a surface.

When performing a single-molecule protein study it is essential to achieve a high quality of surface passivation so that the experiment is free from any surface-induced protein malfunction or denaturation 1,2. While a glass surface treated with surfactants, such as bovine serum albumin, is commonly used for single-molecule nucleic acid studies 3, the degree of its passivation is not high enough for protein studies. A glass surface coated with polymer (polyethylene glycol, PEG) is superior in the passivation performance 4-6. Thereby, it has become universally used for single-molecule protein studies ever since it was introduced to a single-molecule fluorescence study 7-10. The polymer coating procedure requires multiple surface treatments 7,11,12. Therefore, it is difficult to follow the whole procedure without detailed instructions. Often the degree of surface passivation varies depending on which protocol is followed. Here we present a robust protocol with step-by-step instructions, which will remove one of the major bottlenecks of single-molecule protein studies. See Figure 1 for overview.

<table border="1" cellpadding="0" cellspacing="0" width="662">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>1. Slide preparation and cleaning</td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Sigma</td>
			<td>32201</td>
			<td>1 L</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>VWR</td>
			<td>631-0136 631-0144 631-0147</td>
			<td>24x32mm2, 22x40mm2, 
			24x50mm2 &#160;(No.1&#189;, 
			Rectangular)</td>
		</tr>
		<tr>
			<td>Diamond drill bits</td>
			<td>MTN-Giethoorn, The Netherlands</td>
			<td>LA-0564-00DB</td>
			<td>3/4 mm</td>
		</tr>
		<tr>
			<td>Duran slide holder</td>
			<td>LGS, The Netherlands</td>
			<td>213170003</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass staining dishes</td>
			<td>Fisher Scientific</td>
			<td>300101</td>
			<td></td>
		</tr>
		<tr>
			<td>H2O2</td>
			<td>Boom</td>
			<td>6233905</td>
			<td>Opened bottles should be stored at 4 &#176;C. 
			1 L, hydrogen peroxide 30%</td>
		</tr>
		<tr>
			<td>H2SO4</td>
			<td>Boom</td>
			<td>80900627.9010</td>
			<td>Sulfuric acid 95-98%</td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>Sigma</td>
			<td>30603</td>
			<td>Potassium hydroxide</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Sigma</td>
			<td>32213</td>
			<td>1 L</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Branson</td>
			<td></td>
			<td>Tabletop ultrasonic cleaner, 3510</td>
		</tr>
		<tr>
			<td>Quartz slides</td>
			<td>Finkenbeiner</td>
			<td></td>
			<td>1&#8221; x 3&#8221;, 1 mm thick</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>2. Coverslip cleaning</td>
		</tr>
		<tr>
			<td>Duran slide holder</td>
			<td>LGS, The Netherlands</td>
			<td>213170003</td>
			<td></td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>Sigma</td>
			<td>30603</td>
			<td>Potassium hydroxide</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Branson</td>
			<td></td>
			<td>Tabletop ultrasonic cleaner, 3510</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>3. Amino-silanization of slides and coverslips</td>
		</tr>
		<tr>
			<td>Acetic acid</td>
			<td>Sigma</td>
			<td>320099-500ML</td>
			<td>Acetic acid, glacial</td>
		</tr>
		<tr>
			<td>APTES</td>
			<td>Sigma-Aldrich</td>
			<td>281778-100ML</td>
			<td>Store at the room temperature under N2 . Replace once in a month. 
			3-aminopropyl trimethoxysilane</td>
		</tr>
		<tr>
			<td>Flask (200mL)</td>
			<td>VWR</td>
			<td>Flask (200 mL)</td>
			<td>Have one flask dedicated for aminosilanization. 
			Pyrex flask</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Sigma</td>
			<td>32213</td>
			<td>1 L</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Branson</td>
			<td></td>
			<td>Tabletop ultrasonic cleaner, 3510</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>4. Surface passivation using polymer (the first round)</td>
		</tr>
		<tr>
			<td>Biotin-mPEG</td>
			<td>Laysan Bio</td>
			<td>Biotin-PEG-SVA-5000-100mg</td>
			<td>Store aliquots of 1-2 mg (enough for 10 slides) at -20 &#176;C under N2. 
			Biotin-PEG-SVA, MW 5,000&#160;Da</td>
		</tr>
		<tr>
			<td>mPEG</td>
			<td>Laysan Bio</td>
			<td>MPEG-SVA-5000-1g</td>
			<td>Store aliquots of 80 mg (enough for 10 slides) at -20 &#176;C under N2. 
			mPEG-succinimidyl valerate (SVA), MW 5,000 Da</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma</td>
			<td>S6014-500G</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>6. Surface passivation (the second round)</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>D8418-100ML</td>
			<td>Dimethyl sulfoxide BioReagent, for molecular biology, &#8805;99.9%</td>
		</tr>
		<tr>
			<td>DST</td>
			<td>Pierce</td>
			<td>20589</td>
			<td>Disuccinimidyl tartarate</td>
		</tr>
		<tr>
			<td>MS4-PEG</td>
			<td>Pierce</td>
			<td>22341</td>
			<td>Dissolve 100 mg MS4-PEG in 1.1 mL DMSO. Store 20 mL aliquots at -20 &#176;C. 
			Short NHS-ester PEG, MW 333 Da</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma</td>
			<td>S6014-500G</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>7. Assembling a microfluidic chamber</td>
		</tr>
		<tr>
			<td>Double sticky tape</td>
			<td>Scotch</td>
			<td></td>
			<td>10mm wide</td>
		</tr>
		<tr>
			<td>Epoxy</td>
			<td>Thorlabs</td>
			<td>G14250</td>
			<td>Devcon 5 minute epoxy</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S9888-1 KG</td>
			<td>Sodium chloride</td>
		</tr>
		<tr>
			<td>Neutravidin</td>
			<td>Pierce</td>
			<td>31000</td>
			<td>Stock in 5 mg/mL in H2O at 4 &#176;C. 
			ImmunoPure NeutrAvidin Biotin-Binding protein</td>
		</tr>
		<tr>
			<td>Streptavidin</td>
			<td>Invitrogen</td>
			<td>S-888</td>
			<td>Stock 5 mg/mL in H2O or T50 at 4 &#176;C.</td>
		</tr>
		<tr>
			<td>Trizma base</td>
			<td>Sigma-Aldrich</td>
			<td>T1503-1 KG</td>
			<td>Tris</td>
		</tr>
	</tbody>
</table>

surface passivation for single-molecule protein studies

Single-molecule fluorescence spectroscopy has proven to be instrumental in understanding a wide range of biological phenomena at the nanoscale. Important examples of what this technique can yield to biological sciences are the mechanistic insights on protein-protein and protein-nucleic acid interactions. When interactions of proteins are probed at the single-molecule level, the proteins or their substrates are often immobilized on a glass surface, which allows for a long-term observation. This immobilization scheme may introduce unwanted surface artifacts. Therefore, it is essential to passivate the glass surface to make it inert. Surface coating using polyethylene glycol (PEG) stands out for its high performance in preventing proteins from non-specifically interacting with a glass surface. However, the polymer coating procedure is difficult, due to the complication arising from a series of surface treatments and the stringent requirement that a surface needs to be free of any fluorescent molecules at the end of the procedure. Here, we provide a robust protocol with step-by-step instructions. It covers surface cleaning including piranha etching, surface functionalization with amine groups, and finally PEG coating. To obtain a high density of a PEG layer, we introduce a new strategy of treating the surface with PEG molecules over two rounds, which remarkably improves the quality of passivation. We provide representative results as well as practical advice for each critical step so that anyone can achieve the high quality surface passivation.

After the microfluidic chamber assembly (Step 7.1 - 7.6) and before carrying out Step 7.7, it is advised to carry out the quality control of the PEGylated surface.
If the surface passivation has been done successfully, there are less than 10 non-specifically adsorbed proteins per imaging area (25 mm x 25 mm) observed when 1-10 nM fluorescently labeled protein is added into the chamber (Figure 3a, left).
When any of the cleaning or reaction steps has not been properly carried out, the number of non-specifically adsorbed proteins significantly increases, and the CCD screen may be saturated by fluorescence signals. For example, if the piranha etching is skipped, there is 100 times a larger amount of non-specific adsorption observed (Figure 3a; compare left with middle). When the second PEGylation step is skipped, there was approximately 3 times a larger amount of non-specific adsorption observed (Figure 3b). &#160;A low degree of PEGylation is observed when expired chemicals (e.g. APTES stored at room temperature for several months) are used (data not shown). The quality of the surface also drops down when a significant amount of time has passed by since it was PEGylated (Figure 3a; compare left with right).
To our best knowledge, it is the first time that the two rounds of PEGylation are introduced for single-molecule studies. The two rounds of PEGylation guarantee the highest quality of PEG layer formation (Figure 3b). The superior nature of the double PEGylation is prominently shown in the movies (Compare Movie 1a with Movie 1b). In these movies, the background signals from fluorescent molecules in solution are observed to be much weaker when the double PEGylation was used, which indicates that proteins are repelled more effectively by the doubly PEGylated layer. Though this two-step process is strongly advised, the second PEGylation step might be skipped if your experiment is tolerable to non-optimal passivation.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/50549/50549fig1highres.jpg" width="500" /> 
Figure 1: Schematic of the surface treatments. (a) A microscope slide is cleaned with acetone, KOH, and piranha solutions. It is functionalized with APTES and PEGylated with NHS-ester PEG. (b) A coverslip is cleaned with KOH, as well as with the piranha solution if necessary. It is functionalized with APTES and PEGylated with NHS-ester PEG. 
&#160;
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50549/50549fig2highres.jpg" width="500" /> 
Figure 2: Microfluidic chamber. (a) A single-channel chamber. The microscope slide has two holes drilled. It is assembled with a coverslip using two slices of a double-sided sticky tape. (b) A three-channel chamber. The microscope slide has six holes drilled. It is assembled with a coverslip using four slices of a double-sided sticky tape.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/50549/50549fig3highres.jpg" width="500" /> 
Figure 3: CCD images taken with dye-labeled proteins in solution. (a) CCD images were taken by prism-type total internal reflection fluorescence microscopy using a 60X objective lens with 10 nM Cy3-labeld Rep in solution. (Left) A surface was prepared following the protocol in this article. (Middle) A surface was prepared following the protocol in this article, but piranha etching was skipped. (Right) A surface was prepared following the protocol in this article and stored for 3 months at -20 &#186;C under nitrogen. (b) CCD images taken with 10 nM Cy3-labeld Rep in solution. (Left) A surface was prepared following the protocol in this article. (Right) A surface was prepared following the protocol in this article, but the second round of PEGylation was skipped. Scale bar = 5 &#181;m.
Movie 1: CCD movies taken with dye-labeled proteins in solution. CCD movies were taken by prism-type total internal reflection fluorescence microscopy using a 60X objective lens with 10 nM Cy3-labeld Rep in solution. The time resolution is 100 msec. (a) A surface was prepared following the protocol in this article. (b) A surface was prepared following the protocol in this article, but the second round of PEGylation was skipped. <a href="https://www.jove.com/files/ftp_upload/50549/Movie1a.avi" target="_blank">Click here</a> to view Movie 1a and <a href="https://www.jove.com/files/ftp_upload/50549/Movie1b.avi" target="_blank"> click here</a> to view Movie 1b.

Watch this Scientific Journal Video about Surface Passivation for Single-molecule Protein Studies at JoVE.com

Surface Passivation for Single-molecule Protein Studies

We describe a method for passivating a glass surface using polyethylene glycol (PEG). This protocol covers surface cleaning, surface functionalization, and PEG coating. We introduce a new strategy for treating the surface with PEG molecules over two rounds, which yields superior quality of passivation compared to existing methods.

Single-molecule fluorescence spectroscopy has proven to be instrumental in understanding a wide range of biological phenomena at the nanoscale. Important ...

surface-passivation-for-single-molecule-protein-studies

Research

JoVE Journal

Chemistry

Transport of Surface-modified Carbon Nanotubes through a Soil Column

Carbon nanotubes (CNTs) are widely manufactured nanoparticles, which are being utilized in a number of consumer products, such as sporting goods, electronics and biomedical applications. Due to their accelerating production and use, CNTs constitute a potential environmental risk if they are released to soil and groundwater systems. It is therefore essential to improve the current understanding of environmental fate and transport of CNTs. The transport and retention of CNTs in both natural and artificial media have been reported in literature, but the findings widely vary and are thus not conclusive. There are a number of physical and chemical parameters responsible for variation in retention and transport. In this study, a complete procedure of selected multiwalled carbon nanotubes (MWCNTs) is presented starting from their surface modification to a complete set of laboratory column experiments at critical physical and chemical scenarios. Results indicate that the stability of the commercially available MWCNTs are critical with their attached surface functional group which can also influence the transport and retention of MWCNT through the surrounding medium.

Surface properties of a nanoparticle are important for their interaction with the surrounding medium. Therefore the surface modification of carbon nanotubes can be critical for their transport and retention through porous media. Here, lab scale column experiments are used to understand the possible transport and retention of these nanoparticles.

Carbon nanotubes (CNTs) are widely manufactured nanoparticles, which are being utilized in a number of consumer products, such as sporting goods, ...

A Method to Manipulate Surface Tension of a Liquid Metal via Surface Oxidation and Reduction

Controlling interfacial tension is an effective method for manipulating the shape, position, and flow of fluids at sub-millimeter length scales, where interfacial tension is a dominant force. A variety of methods exist for controlling the interfacial tension of aqueous and organic liquids on this scale; however, these techniques have limited utility for liquid metals due to their large interfacial tension.
Liquid metals can form soft, stretchable, and shape-reconfigurable components in electronic and electromagnetic devices. Although it is possible to manipulate these fluids via mechanical methods (e.g., pumping), electrical methods are easier to miniaturize, control, and implement. However, most electrical techniques have their own constraints: electrowetting-on-dielectric requires large (kV) potentials for modest actuation, electrocapillarity can affect relatively small changes in the interfacial tension, and continuous electrowetting is limited to plugs of the liquid metal in capillaries.
Here, we present a method for actuating gallium and gallium-based liquid metal alloys via an electrochemical surface reaction. Controlling the electrochemical potential on the surface of the liquid metal in electrolyte rapidly and reversibly changes the interfacial tension by over two orders of magnitude ( &#820;500 mN/m to near zero). Furthermore, this method requires only a very modest potential (&#60; 1 V) applied relative to a counter electrode. The resulting change in tension is due primarily to the electrochemical deposition of a surface oxide layer, which acts as a surfactant; removal of the oxide increases the interfacial tension, and vice versa. This technique can be applied in a wide variety of electrolytes and is independent of the substrate on which it rests.

We present a method to control the interfacial energy of a liquid metal in an electrolyte via electrochemical deposition (or removal) of a surface oxide layer. This simple method can control the capillary behavior of gallium-based liquid metals by tuning the interfacial energy rapidly, significantly, and reversibly using modest voltages.

Controlling interfacial tension is an effective method for manipulating the shape, position, and flow of fluids at sub-millimeter length scales, where ...

CN-GELFrEE - Clear Native Gel-eluted Liquid Fraction Entrapment Electrophoresis

Protein complexes perform an array of crucial cellular functions. Elucidating their non-covalent interactions and dynamics is paramount for understanding the role of complexes in biological systems. While the direct characterization of biomolecular assemblies has become increasingly important in recent years, native fractionation techniques that are compatible with downstream analysis techniques, including mass spectrometry, are necessary to further expand these studies. Nevertheless, the field lacks a high-throughput, wide-range, high-recovery separation method for native protein assemblies. Here, we present clear native gel-eluted liquid fraction entrapment electrophoresis (CN-GELFrEE), which is a novel separation modality for non-covalent protein assemblies. CN-GELFrEE separation performance was demonstrated by fractionating complexes extracted from mouse heart. Fractions were collected over 2 hr and displayed discrete bands ranging from ~30 to 500 kDa. A consistent pattern of increasing molecular weight bandwidths was observed, each ranging ~100 kDa. Further, subsequent reanalysis of native fractions via SDS-PAGE showed molecular-weight shifts consistent with the denaturation of protein complexes. Therefore, CN-GELFrEE was proved to offer the ability to perform high-resolution and high-recovery native separations on protein complexes from a large molecular weight range, providing fractions that are compatible with downstream protein analyses.

This protocol describes how to prepare and perform clear native gel-eluted liquid fraction entrapment electrophoresis (CN-GELFrEE), a native separation technique for non-covalent biomolecular assemblies and proteins from heterogeneous samples that is compatible with various downstream protein analysis techniques.

Protein complexes perform an array of crucial cellular functions. Elucidating their non-covalent interactions and dynamics is paramount for understanding ...

Synthesis of Cyclic Polymers and Characterization of Their Diffusive Motion in the Melt State at the Single Molecule Level

We demonstrate a method for the synthesis of cyclic polymers and a protocol for characterizing their diffusive motion in a melt state at the single molecule level. An electrostatic self-assembly and covalent fixation (ESA-CF) process is used for the synthesis of the cyclic poly(tetrahydrofuran) (poly(THF)). The diffusive motion of individual cyclic polymer chains in a melt state is visualized using single molecule fluorescence imaging by incorporating a fluorophore unit in the cyclic chains. The diffusive motion of the chains is quantitatively characterized by means of a combination of mean-squared displacement (MSD) analysis and a cumulative distribution function (CDF) analysis. The cyclic polymer exhibits multiple-mode diffusion which is distinct from its linear counterpart. The results demonstrate that the diffusional heterogeneity of polymers that is often hidden behind ensemble averaging can be revealed by the efficient synthesis of the cyclic polymers using the ESA-CF process and the quantitative analysis of the diffusive motion at the single molecule level using the MSD and CDF analyses.

A protocol for the synthesis and characterization of diffusive motion of cyclic polymers at the single molecule level is presented.

We demonstrate a method for the synthesis of cyclic polymers and a protocol for characterizing their diffusive motion in a melt state at the single ...

Insights into the Interactions of Amino Acids and Peptides with Inorganic Materials Using Single-Molecule Force Spectroscopy

The interactions between proteins or peptides and inorganic materials lead to several interesting processes. For example, combining proteins with minerals leads to the formation of composite materials with unique properties. In addition, the undesirable process of biofouling is initiated by the adsorption of biomolecules, mainly proteins, on surfaces. This organic layer is an adhesion layer for bacteria and allows them to interact with the surface. Understanding the fundamental forces that govern the interactions at the organic-inorganic interface is therefore important for many areas of research and could lead to the design of new materials for optical, mechanical and biomedical applications. This paper demonstrates a single-molecule force spectroscopy technique that utilizes an AFM to measure the adhesion force between either peptides or amino acids and well-defined inorganic surfaces. This technique involves a protocol for attaching the biomolecule to the AFM tip through a covalent flexible linker and single-molecule force spectroscopy measurements by atomic force microscope. In addition, an analysis of these measurements is included.

Here we present a protocol to measure the force of interactions between a well-defined inorganic surface and either peptides or amino acids by single-molecule force spectroscopy measurements using an atomic force microscope (AFM). The information obtained from the measurement is important to better understand the peptide-inorganic material interphase.

The interactions between proteins or peptides and inorganic materials lead to several interesting processes. For example, combining proteins with minerals ...

Preparation of Stable Bicyclic Aziridinium Ions and Their Ring-Opening for the Synthesis of Azaheterocycles

Bicyclic aziridinium ions were generated by the removal of an appropriate leaving group through internal nucleophilic attack by nitrogen atom in the aziridine ring. The utility of bicyclic aziridinium ions, specifically 1-azoniabicyclo[3.1.0]hexane and 1-azoniabicyclo[4.1.0]heptane tosylate highlighted in the aziridine ring openings by the nucleophile with the release of the ring strain to yield the corresponding ring-expanded azaheterocycles such as pyrrolidine, piperidine and azepane with diverse substituents on the ring in regio- and stereospecific manner. Herein, we report a simple and convenient method for the preparation of the stable 1-azabicyclo[4.1.0]heptane tosylate followed by selective ring opening via a nucleophilic attack either at the bridge or at the bridgehead carbon to yield piperidine and azepane rings, respectively. This synthetic strategy allowed us to prepare biologically active natural products containing piperidine and azepane motif including sedamine, allosedamine, fagomine and balanol in highly efficient manner.

Bicyclic aziridinium ions such as 1-azoniabicyclo[4.1.0]heptane tosylate were generated from 2-[4-tolenesulfonyloxybutyl]aziridine, which was utilized for the preparation of substituted piperidines and azepanes via regio- and stereospecific ring-expansion with various nucleophiles. This highly efficient protocol allowed us to prepare diverse azaheterocycles including natural products such as fagomine, febrifugine analogue and balanol.

Bicyclic aziridinium ions were generated by the removal of an appropriate leaving group through internal nucleophilic attack by nitrogen atom in the ...

Surface Functionalization of Metal-Organic Frameworks for Improved Moisture Resistance

Metal-organic frameworks (MOFs) are a class of porous inorganic materials with promising properties in gas storage and separation, catalysis and sensing. However, the main issue limiting their applicability is their poor stability in humid conditions. The common methods to overcome this problem involve the formation of strong metal-linker bonds by using highly charged metals, which is limited to a number of structures, the introduction of alkylic groups to the framework by post-synthetic modification (PSM) or chemical vapour deposition (CVD) to enhance overall hydrophobicity of the framework. These last two usually provoke a drastic reduction of the porosity of the material. These strategies do not permit to exploit the properties of the MOF already available and it is imperative to find new methods to enhance the stability of MOFs in water while keeping their properties intact. Herein, we report a novel method to enhance the water stability of MOF crystals featuring Cu2(O2C)4&#160;paddle-wheel units, such as HKUST (where HKUST stands for Hong Kong University of Science &#38; Technology), with the catechols functionalized with alkyl and fluoro-alkyl chains. By taking advantage of the unsaturated metal sites and the catalytic catecholase-like activity of CuII ions, we are able to create robust hydrophobic coatings through the oxidation and subsequent polymerization of the catechol units on the surface of the crystals under anaerobic and water-free conditions without disrupting the underlying structure of the framework. This approach not only affords the material with improved water stability but also provides control over the function of the protective coating, which enables the development of functional coatings for the adsorption and separations of volatile organic compounds. We are confident that this approach could also be extended to other unstable MOFs featuring open metal sites.

Robust functional catechol coatings were produced in one step by direct reaction of the material known as HKUST with synthetic catechols under anaerobic conditions. The formation of homogeneous coatings surrounding the entire crystal is ascribed to the biomimetic catalytic activity of Cu(II) dimers on the external surface of the crystals.

Metal-organic frameworks (MOFs) are a class of porous inorganic materials with promising properties in gas storage and separation, catalysis and sensing. ...

Surface Properties of Synthesized Nanoporous Carbon and Silica Matrices

In this work, we report the synthesis and characterization of ordered nanoporous carbon material (also called ordered mesoporous carbon material [OMC]) with a 4.6 nm pore size, and ordered silica porous matrix, SBA-15, with a 5.3 nm pore size. This work describes the surface properties of nanoporous molecular sieves, their wettability, and the melting behavior of D2O confined in the differently ordered porous materials with similar pore sizes. For this purpose, OMC and SBA-15 with highly ordered nanoporous structures are synthesized via impregnation of the silica matrix by applying a carbon precursor and by the sol-gel method, respectively. The porous structure of investigated systems is characterized by an N2 adsorption-desorption analysis at 77 K. To determine the electrochemical character of the surface of synthesized materials, potentiometric titration measurements are conducted; the obtained results for OMC shows a significant pHpzc shift toward the higher values of pH, relative to SBA-15. This suggests that investigated OMC has surface properties related to oxygen-based functional groups. To describe the surface properties of the materials, the contact angles of liquids penetrating the studied porous beds are also determined. The capillary rise method has confirmed the increased wettability of the silica walls relative to the carbon walls and an influence of the pore roughness on the fluid/wall interactions, which is much more pronounced for silica than for carbon mesopores. We have also studied the melting behavior of D2O confined in OMC and SBA-15 by applying the dielectric method. The results show that the depression of the melting temperature of D2O in the pores of OMC is about 15 K higher relative to the depression of the melting temperature in SBA-15 pores with a comparable 5 nm size. This is caused by the influence of adsorbate/adsorbent interactions of the studied matrices.

Here we report the synthesis and characterization of ordered nanoporous carbon (with a 4.6 nm pore size) and SBA-15 (with a 5.3 nm pore size). The work describes the surface and textural properties of nanoporous molecular sieves, their wettability, and the melting behavior of D2O confined in the materials.

In this work, we report the synthesis and characterization of ordered nanoporous carbon material (also called ordered mesoporous carbon material [OMC]) ...

Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery vehicles and enzyme scaffolds. Due to their biocompatibility and tunability on the genetic level, Elastin-like proteins (ELP) are ideal building blocks for biotechnological and biomedical applications. Nevertheless, the assembly of protein based supramolecular structures with distinct physiochemical properties and good encapsulation potential remains challenging.
Here we provide two efficient protocols for guided self-assembly of amphiphilic ELPs into supramolecular protein architectures such as spherical coacervates, fibers and stable vesicles. The presented assembly protocols generate Protein Membrane-Based Compartments (PMBCs) based on ELPs with adaptable physicochemical properties. PMBCs demonstrate phase separation behavior and reveal method dependent membrane fusion and are able to encapsulate chemically diverse fluorescent cargo molecules. The resulting PMBCs have a high application potential as a drug formulation and delivery platform, artificial cell, and compartmentalized reaction space.

At the interface of organic and aqueous solvents, tailored amphiphilic elastin-like proteins assemble into complex supramolecular structures such as vesicles, fibers and coacervates triggered by environmental parameters. The described assembly protocols yield Protein Membrane-Based Compartments (PMBCs) with tunable properties, enabling the encapsulation of various cargo.

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery ...

Preparation of Nanoparticles for ToF-SIMS and XPS Analysis

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, chemistry, and their potential to enable advanced materials. To effectively understand and regulate the physico-chemical properties and potential adverse effects of nanoparticles, validated measurement procedures for the various properties of nanoparticles need to be developed. While procedures for measuring nanoparticle size and size distribution are already established, standardized methods for analysis of their surface chemistry are not yet in place, although the influence of the surface chemistry on nanoparticle properties is undisputed. In particular, storage and preparation of nanoparticles for surface analysis strongly influences the analytical results from various methods, and in order to obtain consistent results, sample preparation must be both optimized and standardized. In this contribution, we present, in detail, some standard procedures for preparing nanoparticles for surface analytics. In principle, nanoparticles can be deposited on a suitable substrate from suspension or as a powder. Silicon (Si) wafers are commonly used as substrate, however, their cleaning is critical to the process. For sample preparation from suspension, we will discuss drop-casting and spin-coating, where not only the cleanliness of the substrate and purity of the suspension but also its concentration play important roles for the success of the preparation methodology. For nanoparticles with sensitive ligand shells or coatings, deposition as powders is more suitable, although this method requires particular care in fixing the sample.

A number of different procedures for preparing nanoparticles for surface analysis are presented (drop casting, spin coating, deposition from powders, and cryofixation). We discuss the challenges, opportunities, and possible applications of each method, particularly regarding the changes in the surface properties caused by the different preparation methods.

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, ...