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><tbody><tr><td colspan="4">&lt;strong&gt;1. Slide preparation and cleaning&lt;/strong&gt;</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>24 x 32 mm&lt;sup&gt;2&lt;/sup&gt;, 22 x 40 mm&lt;sup&gt;2&lt;/sup&gt;, 24 x 50 mm&lt;sup&gt;2 &lt;/sup&gt;&amp;nbsp;(No.1&amp;frac12;, 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>H&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;</td><td>Boom</td><td>6233905</td><td>1 L, hydrogen peroxide 30%.&lt;br/&gt;Opened bottles should be stored at 4 &amp;deg;C.</td></tr><tr><td>H&lt;sub&gt;2&lt;/sub&gt;SO&lt;sub&gt;4&lt;/sub&gt;</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&amp;rdquo; x 3&amp;rdquo;, 1 mm thick</td></tr><tr><td colspan="4">&lt;strong&gt;2. Coverslip cleaning&lt;/strong&gt;</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 colspan="4">&lt;strong&gt;3. Amino-silanization of slides and coverslips&lt;/strong&gt;</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>3-aminopropyl trimethoxysilane.&lt;br /&gt;Store at the room temperature under N&lt;sub&gt;2&lt;/sub&gt;. Replace once in a month.</td></tr><tr><td>Flask (200 ml)</td><td>VWR</td><td>Flask (200 ml)</td><td>Pyrex flask.&lt;br/&gt;Have one flask dedicated for aminosilanization.</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 colspan="4">&lt;strong&gt;4. Surface passivation using polymer (the first round)&lt;/strong&gt;</td></tr><tr><td>Biotin-mPEG</td><td>Laysan Bio</td><td>Biotin-PEG-SVA-5000-100mg</td><td>Biotin-PEG-SVA, MW 5,000&amp;nbsp;Da.&lt;br /&gt;Store aliquots of 1-2 mg (enough for 10 slides) at -20 &amp;deg;C under N&lt;sub&gt;2&lt;/sub&gt;.</td></tr><tr><td>mPEG</td><td>Laysan Bio</td><td>MPEG-SVA-5000-1g</td><td>mPEG-succinimidyl valerate (SVA), MW 5,000 Da. &lt;br /&gt;Store aliquots of 80 mg (enough for 10 slides) at -20 &amp;deg;C under N&lt;sub&gt;2&lt;/sub&gt;.</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma</td><td>S6014-500G</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;5. Surface passivation (the second round)&lt;/strong&gt;</td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>D8418-100ML</td><td>Dimethyl sulfoxide BioReagent, for molecular biology, &amp;ge;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>Short NHS-ester PEG, MW 333 Da.&lt;br /&gt;Dissolve 100 mg MS4-PEG in 1.1 ml DMSO. Store 20 &amp;micro;l aliquots at -20 &amp;deg;C.</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma</td><td>S6014-500G</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;6. Assembling a microfluidic chamber&lt;/strong&gt;</td></tr><tr><td>Double sticky tape</td><td>Scotch</td><td></td><td>10 mm 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>ImmunoPure NeutrAvidin Biotin-Binding protein.&lt;br /&gt;Stock in 5 mg/ml in H&lt;sub&gt;2&lt;/sub&gt;O at 4 &amp;deg;C.</td></tr><tr><td>Streptavidin</td><td>Invitrogen</td><td>S-888</td><td>Stock 5 mg/ml in H&lt;sub&gt;2&lt;/sub&gt;O or T50 at 4 &amp;deg;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

41.6K Views.  Delft University of Technology.  We beschrijven een werkwijze voor het passiveren van een glasoppervlak met polyethyleenglycol (PEG). Dit protocol heeft betrekking op oppervlaktereiniging, oppervlakte functionalisering, en PEG-coating. We introduceren een nieuwe strategie voor de behandeling van het oppervlak met PEG-moleculen in twee rondes, die een superieure kwaliteit van passivering oplevert vergeleken met bestaande methoden. 

Video: Surface Passivation for Single-molecule Protein Studies

Monitoring Protein Adsorption with Solid-state Nanopores

Solid-state nanopores have been used to perform measurements at the single-molecule level to examine the local structure and flexibility of nucleic acids 1-6, the unfolding of proteins 7, and binding affinity of different ligands 8. By coupling these nanopores to the resistive-pulse technique 9-12, such measurements can be done under a wide variety of conditions and without the need for labeling 3. In the resistive-pulse technique, an ionic salt solution is introduced on both sides of the nanopore. Therefore, ions are driven from one side of the chamber to the other by an applied transmembrane potential, resulting in a steady current. The partitioning of an analyte into the nanopore causes a well-defined deflection in this current, which can be analyzed to extract single-molecule information. Using this technique, the adsorption of single proteins to the nanopore walls can be monitored under a wide range of conditions 13. Protein adsorption is growing in importance, because as microfluidic devices shrink in size, the interaction of these systems with single proteins becomes a concern. This protocol describes a rapid assay for protein binding to nitride films, which can readily be extended to other thin films amenable to nanopore drilling, or to functionalized nitride surfaces. A variety of proteins may be explored under a wide range of solutions and denaturing conditions. Additionally, this protocol may be used to explore more basic problems using nanopore spectroscopy.

A method of using solid-state nanopores to monitor the non-specific adsorption of proteins onto an inorganic surface is described. The method employs the resistive-pulse principle, allowing for the adsorption to be probed in real-time and at the single-molecule level. Because the process of single protein adsorption is far from equilibrium, we propose the employment of parallel arrays of synthetic nanopores, enabling for the quantitative determination of the apparent first-order reaction rate constant of protein adsorption as well as and the Langmuir adsorption constant.

Solid-state nanopores have been used to perform measurements at the single-molecule level to examine the local structure and flexibility of nucleic acids ...

Bioengineering

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

Biochemistry

Light Enhanced Hydrofluoric Acid Passivation: A Sensitive Technique for Detecting Bulk Silicon Defects

A procedure to measure the bulk lifetime (&#62;100 &#181;sec) of silicon wafers by temporarily attaining a very high level of surface passivation when immersing the wafers in hydrofluoric acid (HF) is presented. By this procedure three critical steps are required to attain the bulk lifetime. Firstly, prior to immersing silicon wafers into HF, they are chemically cleaned and subsequently etched in 25% tetramethylammonium hydroxide. Secondly, the chemically treated wafers are then placed into a large plastic container filled with a mixture of HF and hydrochloric acid, and then centered over an inductive coil for photoconductance (PC) measurements. Thirdly, to inhibit surface recombination and measure the bulk lifetime, the wafers are illuminated at 0.2 suns for 1 min&#160;using a halogen lamp, the illumination is switched off, and a PC measurement is immediately taken. By this procedure, the characteristics of bulk silicon defects can be accurately determined. Furthermore, it is anticipated that a sensitive RT surface passivation technique will be imperative for examining bulk silicon defects when their concentration is low (&#60;1012 cm-3).

A RT liquid surface passivation technique to investigate the recombination activity of bulk silicon defects is described. For the technique to be successful, three critical steps are required: (i) chemical cleaning and etching of silicon, (ii) immersion of silicon in 15% hydrofluoric acid and (iii) illumination for 1 min.

A procedure to measure the bulk lifetime (&#62;100 &#181;sec) of silicon wafers by temporarily attaining a very high level of surface passivation when ...

Engineering

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled protein polymerization process is always a challenge. Here, we have developed an enzymatic methodology for constructing polymerized protein step by step in a rationally-controlled sequence. In this method, the C-terminus of a protein monomer is NGL for protein conjugation using OaAEP1 (Oldenlandia affinis asparaginyl endopeptidases) 1) while the N-terminus was a cleavable TEV (tobacco etch virus) cleavage site plus an L (ENLYFQ/GL) for temporary N-terminal protecting. Consequently, OaAEP1 was able to add only one protein monomer at a time, and then the TEV protease cleaved the N-terminus between Q and G to expose the NH2-Gly-Leu. Then the unit is ready for next OaAEP1 ligation. The engineered polyprotein is examined by unfolding individual protein domain using atomic force microscopy-based single-molecule force spectroscopy (AFM-SMFS). Therefore, this study provides a useful strategy for polyprotein engineering and immobilization.

Here, we present a protocol to conjugate protein monomer by enzymes forming protein polymer with a controlled sequence and immobilize it on the surface for single-molecule force spectroscopy studies.

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Biology

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...

Monitoring Conformational Dynamics of Single Unmodified Proteins using Plasmonic Nanotweezers

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes can alter protein biophysics and reduce the usefulness of the data acquired. Plasmonic nanotweezers is a technique that uses localized surface plasmon resonance (LSPR) on gold nanostructures to enhance the electric field within a confined hotspot region. This field enhancement allows for the use of low laser powers to trap single nanoparticles far smaller than conventional optical tweezers, down to only a few nanometers in diameter, such as single proteins. Trapping of single protein molecules within the hotspot region induces a shift in the local refractive index (nprotein &#62; nwater), altering light scattering as a product of the molecule&#39;s polarisability, which is affected by its volume, shape anisotropy, and refractive index. An avalanche photodiode (APD) collects the subsequent changes in light scattering. These alterations can then be analyzed to determine changes in the trapped molecule, including its size, global conformation, and dynamics of conformational change over time. The incorporation of microfluidics within the system allows for controlled environmental changes and real-time monitoring of their subsequent effects on the molecule. In this protocol, we demonstrate the steps to trap single protein molecules, alter their environmental solution conditions, and monitor their corresponding conformational changes using a plasmonic nanotweezers system.

Plasmonic nanotweezers use localized surface plasmon resonance in gold nanostructures to trap single nanoparticles, including proteins, within a nanometer-scale optical field. Changes in the scattered signal reveal protein presence and conformational dynamics, enabling monitoring without fluorophore modifications or surface tethering.

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes ...

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

Surface Passivation for Single-molecule Protein Studies

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