B.N.B. and T.H. acknowledge funding by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under Germany&#39;s Excellence Strategy &#8211; EXC-2193/1 &#8211; 390951807, gef&#246;rdert durch die Deutsche Forschungsgemeinschaft (DFG) im Rahmen der Exzellenzstrategie des Bundes und der L&#228;nder &#8211; EXC-2193/1 &#8211; 390951807, and grant HU 997/1-13 (project # 420798410). M.G. acknowledges partial support in the frame of the LOEWE Project iNAPO by the Hessen State Ministry of Higher Education, Research and the Arts. We thank Dr. Wolfgang Bronner and Dr. Agne Zukauskaite from Fraunhofer Institute for Applied Solid State Physics IAF for the donation of high quality gold coated silicon wafers.

NOTE: See Figure 1 for a schematic overview.
1. Reagent setup
NOTE: The polymers used for this protocol are: maleimide-polyethylene glycol-triethoxysilane (silane-PEG-mal, 5 kDa), thiol-polyethylene glycol-thiol (HS-PEG-SH, 35 kDa), thiol terminated poly(N-isopropylacrylamide) (PNiPAM-SH, 637 kDa) and thiol terminated polystyrene (PS-SH, 1.3 mDa).
<ol>
	<li>Prepare the well-defined and high molar mass PNiPAM-SH via atom transfer radical polymerization, followed by conversion and reduction of the functional end group for the introduction of a thiol moiety, as described in the literature18. Please see Figure 1 for the detailed structures.</li>
	<li>For storing of the chemicals, prepare smaller aliquots within a dry glovebox system with nitrogen atmosphere to avoid exposure to atmospheric oxygen and moisture. PEG and PNiPAM are hygroscopic45,46 and the functional end groups of PEG, PNiPAM and PS are known to become easily oxidized when stored at ambient conditions33,47,48. All chemicals have to be stored at -20 &#176;C.</li>
	<li>Use analytical grade solvents or higher. Moreover, use ultrapure water to rinse AFM cantilevers chips and glassware because single molecule experiments are very sensitive to all contamination.</li>
</ol>
2. Equipment Setup
NOTE: Use tweezers and beakers made of stainless steel or glass. Use inverted tweezers for a safe grip (e.g., model R3 SA having a low spring constant).
<ol>
	<li>Prepare RCA (ultrapure water, hydrogen peroxide and ammonia (5:1:1)) solution to clean glassware and tweezers.</li>
	<li>Put the vessels in a beaker and fill it with RCA until glassware or tweezers are fully covered.</li>
	<li>Heat the beaker from step 2.2 for 1 h at 80 &#176;C.</li>
	<li>Rinse the vessels subsequently with ultrapure water until no pungent smell is ascertainable anymore (at least three times).</li>
	<li>Dry glassware and tweezers in an oven (120 &#176;C).</li>
</ol>
3. Tip functionalization
NOTE: All steps should be carried out in a fume hood to avoid inhalation of organic vapors. Additionally, gloves, lab coat and eye protection are required. Use nitrile or latex gloves for every step to avoid contamination. Wear solvent resistant gloves when using toluene. All steps, unless specified otherwise, are done at RT. Use fresh equipment and gloves for every step to avoid possible cross-contamination.
<ol>
	<li>Perform surface activation by applying oxygen plasma to the AFM cantilever chip MLCT-Bio-DC. 
	NOTE: The efficiency of the plasma treatment for further functionalization steps scales with the content of oxygen in the plasma chamber.
	<ol>
		<li>Use freshly cleaned tweezers to place AFM cantilever chips in a plasma chamber (40 kHz, 600 W).</li>
		<li>Use custom-modified activation program: evacuation (0.1 mbar) &#8211; flooding with oxygen to a pressure of: 0.2 mbar (4 min) &#8211; plasma process (power: 40%, duration: 2 min, process pressure: 0.2 mbar).</li>
		<li>Ventilate chamber and carry on with step 3.2.2 immediately in order to prevent any adsorption of contaminants to AFM cantilever chips from air.</li>
	</ol>
	</li>
	<li>Silanization and PEGylation 
	NOTE: Timing is a critical parameter between the steps. Prepare solutions as fresh as possible during the waiting times. Maleimide groups are subject to hydrolysis in aqueous media and thiols easily become oxidized to disulfides in solution33,47 impeding AFM tip functionalization reactions.
	<ol>
		<li>Prepare a silane-PEG-mal solution in toluene (1.25 mg/mL) in solvent resistant plastic or glass tubes and pour 6 mL of the solution in flat Petri dishes, 3 mL each. 
		NOTE: If binding of multiple probe polymers is observed in the SMFS experiment, mixing silane-PEG-mal with non-functional silane-PEG can reduce the number of anchoring points. For the adjustment of the passivation layer PEG with different masses (i.e., contour lengths) can be used27.</li>
		<li>Incubate the AFM cantilever chips immediately after step 3.1.3 in the silane-PEG-mal solution (up to 10 chips per Petri dish) for 3 h at 60 &#176;C35.</li>
		<li>Take Petri dishes out of the oven and let the solution cool down for at least 10 min.</li>
		<li>Rinse each AFM cantilever chip carefully. Reduce the impact of capillary forces on the AFM cantilever when passing the air-solvent interface, for example by tilting these chips slightly when immersing into solution.
		<ol>
			<li>For PEG and PS polymers, rinse three times with toluene.</li>
			<li>For PNiPAM polymer, rinse once with toluene and twice with ethanol.</li>
		</ol>
		</li>
		<li>Choose at least two AFM cantilever chips as control AFM cantilever chips, skipping step 3.3 and rinse them as follows to increase the polarity of the solvent:
		<ol>
			<li>For PEG and PS polymers, rinse twice with ethanol and once with ultrapure water.</li>
			<li>For PNiPAM polymer, rinse twice with ultrapure water. 
			NOTE: Control AFM cantilever chips have gone through all functionalization steps except the polymer attachment (step 3.3). They serve to prove the cleanliness of the functionalization process, the AFM cantilever chip holder system, the surfaces and the solvents used for the SMFS experiment.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Covalent polymer attachment 
	NOTE: Even though the AFM cantilever tip is expected to be completely covered with maleimide groups, there are just a few binding sites for the single probe polymer, because maleimide undergoes hydrolysis in water leading to inactive PEGs47. These inactive PEGs act as a passivation layer, as described above.
	<ol>
		<li>Incubate AFM cantilever chips directly after step 3.2.5 in one of the following polymer solutions in 3 mL Petri dishes. If the respective polymer is not dissolved properly, use a 40 &#176;C water bath and stir the solution well. 
		NOTE: As the use of thiol terminated polymers might lead to the formation of disulfide bonds hampering the reaction with the maleimide groups of silane-PEG-mal, a reducing agent is recommended, in particular if step 3.3 is applied in aqueous buffers for water soluble polymers33.
		<ol>
			<li>For PEG and PS polymers, use a concentration of 1.25 mg/mL in toluene for 1 h at 60 &#176;C.</li>
			<li>For PNiPAM polymers, use a concentration of 1.25 mg/mL in ethanol for 3 h at RT. 
			NOTE: If binding of multiple probe polymers is observed in the SMFS experiment, the concentration of the polymer should be reduced.</li>
		</ol>
		</li>
		<li>Carefully rinse each AFM cantilever chip.
		<ol>
			<li>For PEG and PS polymers, rinse twice with toluene, twice with ethanol and once with ultrapure water after a 10 min cool down.</li>
			<li>For PNiPAM polymers, rinse twice with ethanol and twice with ultrapure water.</li>
		</ol>
		</li>
		<li>Store each AFM cantilever chip separately in a small (1 mL) Petri dish filled with ultrapure water at 4 &#176;C until use in an experiment.</li>
	</ol>
	</li>
</ol>
4. Surface preparation
<ol>
	<li>Silicon oxide wafer 
	NOTE: This surface was used for SMFS with PEG and PNiPAM.
	<ol>
		<li>Cut a silicon oxide wafer in small pieces using a diamond knife.</li>
		<li>Put the silicon oxide pieces separately in microcentrifuge tubes and fill these tubes with ethanol.</li>
		<li>Sonicate the silicon oxide pieces for 10 min.</li>
		<li>Rinse the silicon oxide pieces with ethanol twice and dry them under a nitrogen flow carefully. Use the silicon oxide pieces immediately.</li>
	</ol>
	</li>
	<li>Self-assembled monolayer of hydrophobic alkane thiol on gold (SAM) 
	NOTE: This surface was used for SMFS with PS. See literature39,49 for more information about SAMs.
	<ol>
		<li>Use a gold-coated silicon wafer (A [100], 5 nm titanium, 100 nm gold) to perform steps 4.1.1 &#8211; 4.1.4.</li>
		<li>Incubate the surface pieces in a 1-dodecanthiol solution (2 mM) for 18 h.</li>
		<li>Rinse the freshly prepared SAMs in ethanol twice.</li>
		<li>Dry SAMs with nitrogen flow for direct use or store them in ethanol for up to 4 days for later use.</li>
	</ol>
	</li>
</ol>
5. Data acquisition
NOTE: All measurements shown here were performed in ultrapure water with a Cypher ES AFM using a heating and cooling sample stage for temperature variation. Generally, all AFMs providing the capability to measure in liquids can be used.
<ol>
	<li>Insert the functionalized AFM cantilever chip into the AFM.</li>
	<li>Glue the prepared surface into a sample holder that is suitable for measuring in liquids (e.g., High Resolution Replicating Compound 101RF or an UV curable adhesive). 
	NOTE: These bonding agents are highly inert and resistant to a large number of polar solvents. The resistance of the adhesive to nonpolar solvents (e.g., toluene or hexane) or high temperatures should be checked prior to use.</li>
	<li>Immerse the AFM cantilever chip and the probe sample in the liquid, here: ultrapure water. 
	NOTE: A solvent drop (about 100 &#181;L) can be deposited on the AFM cantilever chip holder. Covering the AFM cantilever chip with solvent reduces capillary forces, which would otherwise act on the AFM cantilever when approaching the sample surface passing through the air-solvent interface.</li>
	<li>If required, adjust the temperature and let the system equilibrate. 
	NOTE: Temperature changes may result in a deflection of the AFM cantilever due to a bimetallic effect for AFM cantilevers with a reflective coating like aluminum or gold. Equilibration should be performed away from the surface (several &#181;m) until no further change of the deflection signal is observed (up to 15 min for MLCT-Bio-DC).</li>
	<li>Vary the temperature randomly to exclude any effects of ageing of the functionalization. Make sure that the temperatures applied do not lead to an irreversible bending of the AFM cantilever. 
	NOTE: Any temperature effects on solvent properties (such as evaporation or changes in viscosity) might hamper your experiments. In the presented examples, the temperature was varied over a range of up to 40 K in steps of 10 K taking water as a solvent (e.g., from 278 K to 318 K).</li>
	<li>Approach the surface to determine the InvOLS (inverse optical lever sensitivity) by taking force-extension curves on a hard surface (such as silicon oxide). For this, take the deflection signal of the photodetector (in V) vs piezo distance and determine the slope of the part representing the indentation of the AFM cantilever tip into the underlying surface (repulsive regime) using a linear function. In order to reduce errors, take the average of at least five values to obtain the final InvOLS value. For further details, see the literature4,39. 
	NOTE: The InvOLS can only be reliably determined on hard surfaces. In the case of experiments on soft surfaces or interfaces make sure you place a hard surface close to your soft surfaces. Then, the InvOLS calibration can be done before or after your soft surface experiments without the need of disassembling the AFM setup.</li>
	<li>For spring constant determination, move the AFM cantilever to a height with neither attractive nor repulsive interactions between AFM cantilever tip and surface (several &#181;m). Then, record a thermal noise spectrum where the power spectral density (PSD) vs frequency is plotted. The following steps are usually performed by automated built-in functions in commercial AFM software: first, the acquired thermal noise spectrum is analyzed by fitting a function to the PSD, e.g., a simple harmonic oscillator (SHO). The fit is done up to the minimum between the first and second resonance. Second, the area under the fitted part of the PSD vs frequency plot is determined representing the mean square displacement of the AFM cantilever in vertical direction. Finally, the equipartition theorem is used to obtain the AFM cantilever force constant28,50. 
	NOTE: An appropriate frequency range should be used comprising the first resonance peak of the AFM cantilever. To get a satisfactory signal-to-noise ratio, at least 10 PSDs should be accumulated with the highest possible frequency resolution.</li>
	<li>Start the experiment. Record force maps by taking force-extension curves in a grid-like fashion (e.g., 10 x 10 points for an area of 20 x 20 &#181;m2) to avoid any local surface effects (e.g., impurities, dislocations) and to average different surface areas. 
	NOTE: Typical parameters are a pulling velocity of 1 &#181;m/s and a sampling rate of 5 kHz to ensure sufficient resolution. The sampling rate should be adapted when the pulling velocity is varied. The retract distance should be adapted to the contour or desorption length of the measured polymer (approx. twice the expected length).</li>
	<li>Use and vary the dwell time towards the surface to allow the single polymer to adhere to the surface (typically 0 &#8211; 5 s).</li>
	<li>Repeat the determination of the InvOLS and the spring constant at the end of the experiment to check the consistency and the stability of the system. 
	NOTE: For strong adhesion between polymer and surface, the calibration can be done after the actual experiment to preserve the functionalization.</li>
</ol>
6. Data evaluation
NOTE: For data evaluation, a custom-written software based on Igor Pro was used for performing the following steps.
<ol>
	<li>Convert the raw deflection signal (in Volts) into force values (in Newtons) by multiplication with the recorded InvOLS and the determined spring constant.</li>
	<li>Subtract the deflection of the AFM cantilever (after multiplication of the raw deflection signal with the InvOLS) from the distance driven by the piezo elements in vertical direction in order to obtain the true extension (tip-surface distance)4.</li>
	<li>Correct the force-extension curves obtained for drift by fitting a linear function to the baseline after the last event and subtracting the same from the force-extension curve. The fitted part should represent a sufficient extension from the surface where neither attractive nor repulsive interactions are observed between AFM cantilever tip and underlying surface. Then, the baseline is set to the zero-axis. 
	NOTE: In the case of measurements on highly reflective surfaces like gold, interferences might appear. These result from partial reflection of the laser beam from the surface and from the backside of the AFM cantilever. So, the obtained force-extension curves might show a sinusoidal force signal artifact along the vertical extension. This is an artifact that hampers the final force values. In order to still take these force-extension curves into account, a correction is possible (Figure 2).</li>
	<li>If the interferences appear in the force-extension curves, select a representative force-extension curve (retraction curve) showing no other events than possibly a peak of unspecific adhesion and the same sinusoidal artifact (i.e., amplitude and phase) (Figure 2A). 
	NOTE: Smooth the representative force-extension curve in order to obtain the low frequency pattern of the interference.</li>
	<li>Select a force-extension curve that is to be corrected (Figure 2B).</li>
	<li>Overlay both force-extension curves from steps 6.4. and 6.5. to make sure that both show the same sinusoidal artifact (i.e., amplitude and phase) (Figure 2C).</li>
	<li>Subtract the (smoothed) representative force-extension curve from the force-extension curve to be corrected leading to a straight rather than a sinusoidal baseline (Figure 2D). 
	NOTE: Take care that the unspecific adhesion peak of the representative curve is distinct from any single molecule events appearing in the curves to be corrected. In fact, the selection of the representative curve is crucial for a proper correction.</li>
</ol>

The authors declare that they have no competing financial interest.

AFM-based SMFS is one of the major tools for investigating single molecule interactions in polymer physics. For a true single molecule experiment, covalent attachment of the probe polymer to an AFM cantilever tip is essential.
Many previous works are based on nanofishing experiments, in particular for PNiPAM, where polymers are adsorbed onto a surface and then stretched by randomly picking them from the substrate using an AFM cantilever tip30,31. This might alter the results and lead to misinterpretation of the single molecule behavior. There, cooperative effects might dominate the results because interactions with neighboring polymers cannot be excluded. This has a large impact on the results, especially for polymers that show significantly different behavior in bulk compared to single isolated molecules57,58.
The functionalization protocol presented here is reliable and can be easily applied to different polymers, irrespective of their contour length, hydrophobicity or the steric hindrance of the monomers. Additionally, a passivation is provided to prevent unwanted interactions between the single probe polymer and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface. Furthermore, the evaluation of force-extension curves showing stretching events is shown. There, a procedure is proposed for the determination of master force-extension curves. This offers a better means of revealing, e.g., temperature-related effects on the force-extension behavior. Furthermore, the analysis of single molecule desorption events featuring constant force plateaus is provided. Also, a simple way of correcting sinusoidal force signal artifacts in force-extension curves is given which might otherwise impair the results of the experiment.
Compared to Stetter et al.39, the functionalization procedure presented here is reduced to three steps instead of four and the robustness of the procedure is improved. The major benefit of performing PEGylation and silanization in one step is to have a better-controlled reaction and to increase the yield. Furthermore, fewer solutions need to be prepared and fewer rinsing steps are required. This reduces the effort and time for preparation and increases the reproducibility. Furthermore, moving AFM cantilevers is always a critical part of the functionalization process. A transfer from one solution to the other always runs the risk of strongly influencing the functionalization quality due to transfers through the air-water interface or of losing AFM cantilevers by improper use of tweezers.
In order to prove proper covalent attachment of a single polymer to an AFM cantilever tip different conditions have to be met. First, control AFM cantilevers are of significant importance and should be prepared for every functionalization. The functionalization process and the fluid cell for performing the experiments are only considered to be clean, if a small number of force-extension curves show stretches or plateaus in the control experiment (in the presented examples less than 2%).
A clear stretching pattern with no further drops or maxima is essential for having proper single molecule stretching events. Additionally, the dependence of rupture force on the force loading rate at rupture or the complete elastic response of the stretching curve should be analyzed in order to exclude simultaneous desorption of multiple polymers59,60. For PEG and PNiPAM, 19% and 42% of the force-extension curves taken at different positions of the surface showed such a stretching pattern, respectively. In order to obtain stretching events, the physisorption of the polymer to the respective underlying surface must be strong. Otherwise a plateau-like desorption event is observed. This is even more decisive for the detection of stretching events at high forces (up to 500 pN or more). As this strong physisorption is not met for every force-extension curve, the yield of such events is less than for pure plateau-like desorption events. As an alternative, strongly adhering groups such as catechols or chemisorption between polymer and underlying surface can be used. However, this requires the introduction of further functional groups or coupling sites at the polymer61,62.
In fact, the mass (i.e., contour length) of the polymer provides a valuable fingerprint. Although the mass cannot directly be translated into the measured contour length for the following reasons, the length distribution is very valuable to define single-molecule events. In the case of a PNiPAM polymer with a low polydispersity (&#393; = 1.28), we found significant differences in the extension values for the obtained stretching events (and thus in the polymer length) in the experiments. One reason for this could be the determination of the polymer length and its distribution. In size-exclusion chromatography (SEC), a relative weight of the target polymer is determined in comparison with standards like PS or poly (methyl methacrylate) (PMMA)63. The presumed relative weight is expected to deviate from the absolute molecular weight because the hydrodynamic radius of the target polymer and the standard can significantly differ. Additionally, the silane layer might be oligomerized by spurious water in toluene during the functionalization process. The attachment of such oligomers to the AFM cantilever tip leads to a more flexible layer with fewer anchor points64. Also, the attachment point of the polymer to the silicon layer might not necessarily be at the apex leading to a shift of the detected length values29. While a polymer model such as the wormlike chain (WLC) or the freely jointed chain (FJC) model cannot reproduce the respective force-extension behavior for PEG or PNiPAM properly over the entire extension range18,29,41,65,66, such a polymer model might be valuable for other polymeric and protein systems10,15,67,68.
The covalent attachment of a single PS polymer (with a contour length of more than 1 &#181;m) is only considered to be successful, when a considerable number of force-extension curves show a long enough plateau of constant force (Figure 5). A plateau resulting from desorbing a single polymer is defined by a single sharp drop of a constant force to the baseline at a certain extension, as given in Figure 5A. If more polymers are attached to the AFM cantilever tip, a cascade of plateaus is observed56 (Figure 5C). The plateau length (desorption length), correlating with the polymer contour length51, has to be significantly longer than any adhesion peak due to unspecific adhesion of the AFM cantilever tip to the underlying surface (here around 200 nm). Features appearing solely in a single force-extension curve, should not be interpreted. In the presented experiments, at least 80 out of 100 curves showed a plateau longer than 200 nm in at least two force maps at two different spots on the surface. Furthermore, the distribution of desorption lengths, using scatter plots such as given in Figure 5B and 5D, reveal if and how many polymers are bound to the AFM cantilever tip. In the case of PS, a narrow distribution of desorption force and length taken from plateaus of the force-extension curves served as evidence of a successful covalent attachment. This finally proved the success of the functionalization protocol. Thus, we strongly recommend to present such force and length distributions in publications.
Evaluating force-extension curves using built-in algorithms that comprise many pre-set parameters should be done with care. Reasons are for example that a fixed sampling rate is not appropriate for every applied pulling velocity or that an automated smoothing of the force-extension curves might average out important details. Usually a proper understanding of the respective evaluation procedure can prevent errors in the evaluation procedure, which can in turn strongly influence the final findings of an AFM-based SMFS experiment.
In summary, we present a functionalization protocol that is reliable and can be easily applied to a variety of polymers. Furthermore, proper evaluation of single molecule force-extension curves is presented, allowing the determination of physical parameters such as stretching force, desorption force and desorption length. The presented protocols and procedures are valuable for the investigation of stimuli-responsive systems at the single molecule level.

Since its invention in the 1980s1, the atomic force microscope (AFM) has become one of the most important imaging techniques in natural science featuring sub-nanometer spatial resolution, sub-piconewton force resolution and the possibility of measuring in various solvent and temperature conditions2,3,4,5,6,7.
Apart from imaging8,9, AFM is used to perform single molecule force spectroscopy (SMFS) giving insight into adhesive interactions between a single polymer and surfaces, physical properties of single polymers and unfolding mechanisms of proteins7,10,11,12,13,14,15,16. In a regular SMFS experiment, the functionalized cantilever tip is brought into contact with a surface so that the polymer at the AFM cantilever tip physisorbs to this surface. By retracting the AFM cantilever tip from the surface, a change in the deflection of the AFM cantilever is converted into a force leading to a force-extension curve4. Physical parameters such as stretching force, desorption force and desorption length can be determined as dependent on different parameters such as pulling velocity, dwell time on surface, indentation depth into the surface, temperature, solvent17,18 and different surfaces like solid substrates, polymer films or supported lipid bilayers19,20,21,22. Furthermore, a polymer can be probed in different spatial directions, thus investigating the frictional properties of the polymer23,24,25,26.
A covalent attachment of the investigated polymer to an AFM cantilever tip is essential for such studies. Thus, a high yield of single molecule events with one and the same polymer bound to an AFM cantilever tip prevents any bias of the results due to calibration of the spring constant of the AFM cantilever27,28, varying attachment points29 or varying polymers (with different contour lengths) such as in the case of nanofishing experiments30,31,32. Also, interactions with other polymers as well as averaging effects can be widely prevented18,28. For the covalent attachment of a polymer to the AFM cantilever tip, different types of chemical modifications can be applied, many of which are summarized in the book by Hermanson33. Amine and thiol-based linking reactions as well as click chemistry represent the most commonly used methods in AFM cantilever tip functionalization34,35,36,37,38,39,40,41,42. Becke et al.40 show how to use 1-ethyl-3-(3-dimethylaminopropyl)carbodiimid (EDC)/NHS chemistry to attach a protein to an AFM cantilever tip. However, the said functional groups tend to crosslink, thus leading to a loss of functionality43,44. Also, carbodiimides show a tendency to fast hydrolysis in solution43. Maleimide and thiol groups are generally more stable and do not show crosslinking reactions. The presented protocol is an optimization of the previously published protocols given in references35,39.
Here, a reliable functionalization protocol is presented that can be easily adjusted to a large number of different polymers, irrespective of properties such as contour length or hydrophobicity. Three different polymers were chosen by way of example: hydrophilic polyethylene glycol (PEG) and poly(N-isopropylacrylamide) (PNiPAM) as well as high molar mass hydrophobic polystyrene (PS). In order to provide for a covalent binding capability with an appropriate linker molecule, the three polymers were selected for featuring a telechelic thiol moiety as functional end group. The linker molecule itself is typically a short PEG polymer with two active sites, a silane group at one end and a maleimide group at the other end. The former enables a covalent attachment to the AFM cantilever tip and the latter a binding reaction with the thiol group of the functionalized high molar mass polymer. Furthermore, inactive PEG linker molecules serve as a passivation layer to prevent unwanted interactions between the probe polymer and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1-Dodecanethiol (&#8805;98%)</td>
			<td>Sigma-Aldrich, USA</td>
			<td>417364-500ML</td>
			<td>Used for SAM</td>
		</tr>
		<tr>
			<td>Ammonia solution (30%)</td>
			<td>Roth, Germany</td>
			<td>CP17.2</td>
			<td>Used for cleaning</td>
		</tr>
		<tr>
			<td>Cypher ES</td>
			<td>Asylum Research, an Oxford Instruments company, USA</td>
			<td>-</td>
			<td>AFM</td>
		</tr>
		<tr>
			<td>Ethanol (&#8805;99.9%)</td>
			<td>Roth, Germany</td>
			<td>PO76.1</td>
			<td>Solvent</td>
		</tr>
		<tr>
			<td>Gold coated silicon wafer</td>
			<td>Fraunhofer Institute for Applied Solid State Physics IAF, Germany</td>
			<td>-</td>
			<td>Used for SAM</td>
		</tr>
		<tr>
			<td>High Resolution Replicating Compound</td>
			<td>Microset Products Ltd, UK</td>
			<td>101RF</td>
			<td>Bonding agent</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide solution</td>
			<td>Sigma-Aldrich, USA</td>
			<td>H1009</td>
			<td>Used for cleaning</td>
		</tr>
		<tr>
			<td>Igor Pro</td>
			<td>Wavemetrics, USA</td>
			<td>-</td>
			<td>Software environment</td>
		</tr>
		<tr>
			<td>Tetra-30-LF-PC</td>
			<td>Diener Electronic, Germany</td>
			<td>-</td>
			<td>Plasma chamber</td>
		</tr>
		<tr>
			<td>Maleimide-polyethylene glycol-triethoxysilane</td>
			<td>Creative PEG works, USA</td>
			<td>PHB-1923</td>
			<td>Linker polymer</td>
		</tr>
		<tr>
			<td>MLCT-Bio-DC</td>
			<td>Bruker, USA</td>
			<td>MLCT-Bio-DC</td>
			<td>AFM cantilever</td>
		</tr>
		<tr>
			<td>Prime CZ-Si wafer, n-type (Phosphor) TTV &#60; 10 &#181;m</td>
			<td>MicroChemicals, Germany</td>
			<td>WSA40600250 P1314SNN1</td>
			<td>Silicon wafer</td>
		</tr>
		<tr>
			<td>Purelab Chorus 1, 18.2 M&#937; cm</td>
			<td>Elga LabWater, Germany</td>
			<td>10034-540</td>
			<td>Ultrapure water source</td>
		</tr>
		<tr>
			<td>R3 SA</td>
			<td>Vomm GmbH, Germany</td>
			<td>5803 Blank</td>
			<td>Tweezers</td>
		</tr>
		<tr>
			<td>Thiol terminated poly(N-isopropylacrylamide)</td>
			<td>Gallei Group, Saarland University, Germany</td>
			<td>-</td>
			<td>PNiPAM probe polymer</td>
		</tr>
		<tr>
			<td>Thiol terminated polystyrene</td>
			<td>Polymer Source, Canada</td>
			<td>P40722-SSH</td>
			<td>PS probe polymer</td>
		</tr>
		<tr>
			<td>Thiol-polyethylene glycol-thiol</td>
			<td>Creative PEGWorks, USA</td>
			<td>PSB-615</td>
			<td>PEG probe polymer</td>
		</tr>
		<tr>
			<td>Toluene (99.99%)</td>
			<td>Fisher Chemicals</td>
			<td>T324-500</td>
			<td>Solvent</td>
		</tr>
	</tbody>
</table>

covalent attachment of single molecules for afm-based force spectroscopy

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and surfaces. For a true single molecule experiment, covalent attachment of the probe molecule is essential because only then can hundreds of force-extension traces with one and the same single molecule be obtained. Many traces are in turn necessary to prove that a single molecule alone is probed. Additionally, passivation is crucial for preventing unwanted interactions between the single probe molecule and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface. The functionalization protocol presented here is reliable and can easily be applied to a variety of polymers. Characteristic single molecule events (i.e., stretches and plateaus) are detected in the force-extension traces. From these events, physical parameters such as stretching force, desorption force and desorption length can be obtained. This is particularly important for the precise investigation of stimuli-responsive systems at the single molecule level. As exemplary systems poly(ethylene glycol) (PEG), poly(N-isopropylacrylamide) (PNiPAM) and polystyrene (PS) are stretched and desorbed from SiOx (for PEG and PNiPAM) and from hydrophobic self-assembled monolayer surfaces (for PS) in aqueous environment.

The following examples show results of single molecule stretching and desorption of the polymers PEG, PNiPAM and PS. All AFM cantilever tips were functionalized with the protocol given above. PEG and PNiPAM were measured on SiOx with temperature variation. For a detailed discussion of the resulting temperature-dependent stretching curves for PEG and PNiPAM, see Kolberg et al.18 A different force-extension motif is a plateau of constant force (e.g., when desorbing PS from self-assembled monolayers of methyl terminated alkane thiols on gold (SAM) in water4,27,39,51).
Example 1: Stretching of PEG and PNiPAM in water 
The temperature-dependent stretching behavior in water was measured using single PNiPAM and PEG polymers covalently bound to an AFM cantilever tip at one end and physisorbed on a SiOx surface at the other end. After the calibration and clean control experiments (less than 2% of the force-extension curves show single molecule events), at least two force maps were recorded for each AFM cantilever. The temperature-dependent experiment was performed by recording at least one force map at each temperature. When only few stretching events appeared, the respective AFM cantilever was discarded and the next AFM cantilever of the chip was taken (usually in the order C, B, D and E of MLCT-Bio-DC). For the exemplary data of PEG, a single stretching event was observed in 95 out of 500 measured force-extension curves (19%). For PNiPAM, 252 out of 600 force-extension curves showed a stretching pattern (42%). For a better comparison of the force-extension curves, a single master curve for every temperature was generated. For this purpose, only those curves with a stretching event to at least 500 pN, where conformational fluctuations and solvent effects are negligible, were chosen52. The final number of stretches taken into account was 3 at 278 K, 7 at 298 K and 4 at 318 K for PEG and 4 at 278 K, 3 at 298 K and 3 at 318 K for PNiPAM18.
The procedure for generating master curves is given in Figure 3. The force-extension curves chosen (Figure 3A) are rescaled to a length L0 (extension at a force of 500 pN), see Figure 3B. The adhesion peak shows a large variation of unspecific adhesion between the surface and the AFM cantilever tip, but does not influence the polymer stretching behavior. After merging the rescaled force-extension curves they are averaged by a binominal smoothing as presented in Figure 3C. For this, a Gaussian filter convolves the data with normalized coefficients derived from Pascal&#39;s triangle at a level equal to the smoothing parameter 2053. Finally, a master curve is obtained for every temperature as given in Figure 3D. The zoom-in shows the range where the temperature effect on the force-extension behavior is most pronounced.
A comparison of the temperature behavior of PEG (A) and PNiPAM (B) can be found in Figure 4. For PEG a decrease of the stretching force with increasing temperature was observed. An increase of approximately 5% of rescaled extension at 100 pN was observed when increasing the temperature from 278 to 318 K. For PNiPAM, an opposite temperature-dependent shift could be revealed. A decrease of approximately 1% of rescaled extension at 100 pN was observed when the temperature was increased from 278 to 328 K. Additionally, the stretching free energy could be obtained from the force-extension master curves by determining the area under the curve for any given force value. This could be used for extracting energetic and entropic contributions of the stretching free energy with the help of molecular dynamics (MD) simulations18.
Example 2: Desorption of PS from a SAM surface in water 
The desorption of PS from a SAM surface in water could be used to determine the desorption force and length and thereby quantify the hydrophobic interaction. After calibration, at least two force maps were recorded at two different spots of the surface. When the polymer attachment was successful, the force-extension curves showed plateaus of constant force, as characteristic feature, see Figure 5A and Figure 5C. Plateau-like desorption is observed when the dynamics of the probed bonds are much faster than the pulling rate of the AFM cantilever tip (quasi-equilibrium). Desorption forces of plateau-like force-extension curves directly provide adhesion free energies by integrating the force-extension trace54. They have been used to determine electrostatic, dispersive and hydrophobic interactions as well as friction properties of single polymers on surfaces in liquid environment2,4,23,51,54,55.
Each plateau of constant force was fitted with a sigmoidal curve to determine the desorption force and desorption length, which were then plotted in histograms. The histograms were fitted with a Gaussian to extract the maximum value and standard deviation. For a better overview, the desorption force and length values were displayed together in a scatter plot, as given in Figure 5B and Figure 5D.
For polystyrene on SAM in water, the determined desorption forces correspond to previously obtained values19,23. As the desorption length correlates with the polymer contour length51, the desorption length distribution can be used as a proof of the covalent binding of the respective polymer to the AFM cantilever tip via its functional end group. Thus, the desorption length serves as a fingerprint.
For more than one polymer attached to the AFM cantilever tip, cascades of plateaus (discrete steps) can be observed in the force-extension curves56. Each plateau represents the desorption of a polymer at a different extension. The experiment given in Figure 5C and Figure 5D showed a typical case of two polymers attached to the AFM cantilever tip at the same time. By fitting the final rupture, a bimodal distribution could be found for the desorption length, while the desorption force showed a narrow distribution. In this case, the smaller desorption length could be found in 90% of the force-extension curves, either as a single plateau or as an additional plateau on the longer plateau, as given in Figure 5C. The higher desorption length was found in 37% of the obtained force-extension curves. Thus, the desorption length distribution could be used to determine the number of different polymers attached to the AFM cantilever tip. In general, a narrow distribution of the desorption length values is a good indication that one and the same single polymer was probed in the obtained force-extension curves. At the same time, a superposition of the respective forces-extension can be used to decide whether one and the same single polymer has been measured.
After proving covalent binding of a single PS polymer, further experiments with this PS polymer can be performed varying substrate (solid surface as well as polymer films), solvent conditions, temperature, pulling velocity or dwell time.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60934/60934fig1a.jpg" /> 
Figure 1: Schematic overview of the tip functionalization process. Includes the chemical modification of the AFM cantilever tip after (1) plasma activation (2) silanization/PEGylation and (3) polymer attachment. Additionally, the detailed chemical structures of the polymers used, namely PEG, PNiPAM and PS are shown. <a href="https://www.jove.com/files/ftp_upload/60934/60934fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60934/60934fig2a.jpg" /> 
Figure 2: Elimination of interferences in force-extension curves. (A) Find a force-extension curve showing a sinusoidal force signal artifact along the extension but having no single molecule stretching event. (B) Choose a force-extension curve with a single molecule event, which is to be corrected from the sinusoidal artifact. (C) Superimpose the curves to control if the sinusoidal artifacts of the curves really match. (D) By subtracting the force-extension curve (A) from (B) a force-extension curve with a straight baseline is obtained. Although the adhesion peak cannot be used for further analysis, the force-extension curve is now corrected for the artifact leading to much more accurate force values in the region of the single molecule event (here: &#62; 0.2 &#181;m of extension). <a href="https://www.jove.com/files/ftp_upload/60934/60934fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60934/60934fig3a.jpg" /> 
Figure 3: Determination of master curves from force-extension curves of PEG at 298 K. (A) Experimental data at 298 K, using 7 force-extension curves. After rescaling to a length L0 at a force of 500 pN (B), the force-extension curves can be merged and averaged by binominal smoothing obtaining a master curve (C). The rescaled curves are given as dots while the master curve is shown as a solid line. Finally, the obtained master curves for different temperatures can be compared (D). The zoom-in indicates the range where the temperature effect on the force-extension behavior is most pronounced. <a href="https://www.jove.com/files/ftp_upload/60934/60934fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60934/60934fig4a.jpg" /> 
Figure 4: Comparison of the temperature-dependent master curves of PNiPAM and PEG. For PEG an increase of rescaled extension at 100 pN (mid-force range) is observed when increasing the temperature (A), while for PNiPAM an opposite temperature-dependent shift is revealed (B). <a href="https://www.jove.com/files/ftp_upload/60934/60934fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60934/60934fig5.jpg" /> 
Figure 5: Analysis of force-extension curves of PS on SAM in water. (A) Exemplary force-extension curve (blue) with a sigmoidal fit of the plateau (purple). Additionally, the arrows mark the determined force (red) and length (green) of the plateau. The desorption force and desorption length values obtained by sigmoidal fits are displayed in a scatter plot and the resulting histograms are fitted with a Gaussian. (B) The determined average desorption force and desorption length values are (112 &#177; 6) pN and (659 &#177; 7) nm, wherein 93% of the force-extension curves show such single plateau events. (C) Exemplary force-extension curve (blue) for two polymers attached to the AFM cantilever tip at the same time. Here, the desorption force shows a unimodal distribution with an average force value of (117 &#177; 5) pN, while a bimodal distribution can be found for the desorption length leading to average length values of (656 &#177; 9) nm and (1050 &#177; 16) nm. (D) 90% of the sampled force-extension curves show only single plateau events. <a href="https://www.jove.com/files/ftp_upload/60934/60934fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy at JoVE.com

Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

Covalent attachment of probe molecules to atomic force microscopy (AFM) cantilever tips is an essential technique for the investigation of their physical properties. This allows us to determine the stretching force, desorption force and length of polymers via AFM-based single molecule force spectroscopy with high reproducibility.

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and ...

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9.5K Views.  Albert-Ludwigs-Universität Freiburg. Covalent attachment of probe molecules to atomic force microscopy (AFM) cantilever tips is an essential technique for the investigation of their physical properties. This allows us to determine the stretching force, desorption force and length of polymers via AFM-based single molecule force spectroscopy with high reproducibility.

Video: Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

Covalent Binding of BMP-2 on Surfaces Using a Self-assembled Monolayer Approach

Bone morphogenetic protein 2 (BMP-2) is a growth factor embedded in the extracellular matrix of bone tissue. BMP-2 acts as trigger of mesenchymal cell differentiation into osteoblasts, thus stimulating healing and de novo bone formation. The clinical use of recombinant human BMP-2 (rhBMP-2) in conjunction with scaffolds has raised recent controversies, based on the mode of presentation and the amount to be delivered. The protocol presented here provides a simple and efficient way to deliver BMP-2 for in vitro studies on cells. We describe how to form a self-assembled monolayer consisting of a heterobifunctional linker, and show the subsequent binding step to obtain covalent immobilization of rhBMP-2. With this approach it is possible to achieve a sustained presentation of BMP-2 while maintaining the biological activity of the protein. In fact, the surface immobilization of BMP-2 allows targeted investigations by preventing unspecific adsorption, while reducing the amount of growth factor and, most notably, hindering uncontrolled release from the surface. Both short- and long-term signaling events triggered by BMP-2 are taking place when cells are exposed to surfaces presenting covalently immobilized rhBMP-2, making this approach suitable for in vitro studies on cell responses to BMP-2 stimulation.

We describe a method for accomplishing efficient immobilization of BMP-2 on surfaces. Our approach is based on the formation of a self-assembled monolayer to achieve the covalent binding of BMP-2 via its free amine residues. This method is a useful tool to study signaling at the cell membrane.

Bone morphogenetic protein 2 (BMP-2) is a growth factor embedded in the extracellular matrix of bone tissue. BMP-2 acts as trigger of mesenchymal cell ...

Preparation of Silica Nanoparticles Through Microwave-assisted Acid-catalysis

Microwave-assisted synthetic techniques were used to quickly and reproducibly produce silica nanoparticle sols using an acid catalyst with nanoparticle diameters ranging from 30-250 nm by varying the reaction conditions. Through the selection of a microwave compatible solvent, silicic acid precursor, catalyst, and microwave irradiation time, these microwave-assisted methods were capable of overcoming the previously reported shortcomings associated with synthesis of silica nanoparticles using microwave reactors. The siloxane precursor was hydrolyzed using the acid catalyst, HCl. Acetone, a low-tan &#948; solvent, mediates the condensation reactions and has minimal interaction with the electromagnetic field. Condensation reactions begin when the silicic acid precursor couples with the microwave radiation, leading to silica nanoparticle sol formation. The silica nanoparticles were characterized by dynamic light scattering data and scanning electron microscopy, which show the materials&#39; morphology and size to be dependent on the reaction conditions. Microwave-assisted reactions produce silica nanoparticles with roughened textured surfaces that are atypical for silica sols produced by St&#246;ber&#39;s methods, which have smooth surfaces.

Silica nanoparticles were prepared using acid-catalysis of a siloxane precursor and microwave-assisted synthetic techniques resulting in the controlled growth of nanomaterials ranging from 30-250 nm in diameter. The growth dynamics can be controlled by varying the initial silicic acid concentration, time of the reaction, and temperature of reaction.

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Activating Molecules, Ions, and Solid Particles with Acoustic Cavitation

The chemical and physical effects of ultrasound arise not from a direct interaction of molecules with sound waves, but rather from the acoustic cavitation: the nucleation, growth, and implosive collapse of microbubbles in liquids submitted to power ultrasound. The violent implosion of bubbles leads to the formation of chemically reactive species and to the emission of light, named sonoluminescence. In this manuscript, we describe the techniques allowing study of extreme intrabubble conditions and chemical reactivity of acoustic cavitation in solutions. The analysis of sonoluminescence spectra of water sparged with noble gases provides evidence for nonequilibrium plasma formation. The photons and the "hot" particles generated by cavitation bubbles enable to excite the non-volatile species in solutions increasing their chemical reactivity. For example the mechanism of ultrabright sonoluminescence of uranyl ions in acidic solutions varies with uranium concentration: sonophotoluminescence dominates in diluted solutions, and collisional excitation contributes at higher uranium concentration. Secondary sonochemical products may arise from chemically active species that are formed inside the bubble, but then diffuse into the liquid phase and react with solution precursors to form a variety of products. For instance, the sonochemical reduction of Pt(IV) in pure water provides an innovative synthetic route for monodispersed nanoparticles of metallic platinum without any templates or capping agents. Many studies reveal the advantages of ultrasound to activate the divided solids. In general, the mechanical effects of ultrasound strongly contribute in heterogeneous systems in addition to chemical effects. In particular, the sonolysis of PuO2 powder in pure water yields stable colloids of plutonium due to both effects.

Acoustic cavitation in liquids submitted to power ultrasound creates transient extreme conditions inside the collapsing bubbles, which are the origin of unusual chemical reactivity and light emission, known as sonoluminescence. In the presence of noble gases, nonequilibrium plasma is formed. The "hot" particles and the photons generated by collapsing bubbles are able to excite species in solution.

The chemical and physical effects of ultrasound arise not from a direct interaction of molecules with sound waves, but rather from the acoustic ...

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 ...

Probing the Structure and Dynamics of Interfacial Water with Scanning Tunneling Microscopy and Spectroscopy

Water/solid interfaces are ubiquitous and play a key role in many environmental, biophysical, and technological processes. Resolving the internal structure and probing the hydrogen-bond (H-bond) dynamics of the water molecules adsorbed on solid surfaces are fundamental issues of water science, which remains a great challenge owing to the light mass and small size of hydrogen. Scanning tunneling microscopy (STM) is a promising tool for attacking these problems, thanks to its capabilities of sub-&#197;ngstr&#246;m spatial resolution, single-bond vibrational sensitivity, and atomic/molecular manipulation. The designed experimental system consists of a Cl-terminated tip and a sample fabricated by dosing water molecules in situ onto the Au(111)-supported NaCl(001) surfaces. The insulating NaCl films electronically decouple the water from the metal substrates, so the intrinsic frontier orbitals of water molecules are preserved. The Cl-tip facilitates the manipulation of the single water molecules, as well as gating the orbitals of water to the proximity of Fermi level (EF) via tip-water coupling. This paper outlines the detailed methods of submolecular resolution imaging, molecular/atomic manipulation, and single-bond vibrational spectroscopy of interfacial water. These studies open up a new route for investigating the H-bonded systems at the atomic scale.

Here, we present a protocol to investigate the structure and dynamics of interfacial water at the atomic scale, in terms of submolecular resolution imaging, molecular manipulation, and single-bond vibrational spectroscopy.

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Synthesis of Information-bearing Peptoids and their Sequence-directed Dynamic Covalent Self-assembly

This protocol presents the use of Lewis acidic multi-role reagents to circumvent kinetic trapping observed during the self-assembly of information-encoded oligomeric strands mediated by paired dynamic covalent interactions in a manner mimicking the thermal cycling commonly employed for the self-assembly of complementary nucleic acid sequences. Primary amine monomers bearing aldehyde and amine pendant moieties are functionalized with orthogonal protecting groups for use as dynamic covalent reactant pairs. Using a modified automated peptide synthesizer, the primary amine monomers are encoded into oligo(peptoid) strands through solid-phase submonomer synthesis. Upon purification by high-performance liquid chromatography (HPLC) and characterization by electrospray ionization mass spectrometry (ESI-MS), sequence-specific oligomers are subjected to high-loading of a Lewis acidic rare-earth metal triflate which both deprotects the aldehyde moieties and affects the reactant pair equilibrium such that strands completely dissociate. Subsequently, a fraction of the Lewis acid is extracted, enabling annealing of complementary sequence-specific strands to form information-encoded molecular ladders characterized by matrix assisted laser desorption/ionization mass spectrometry (MALDI-MS). The simple procedure outlined in this report circumvents kinetic traps commonly experienced in the field of dynamic covalent assembly and serves as a platform for the future design of robust, complex architectures.

A protocol is presented for the synthesis of information-encoded peptoid oligomers and for the sequence-directed self-assembly of these peptoids into molecular ladders using amines and aldehydes as dynamic covalent reactant pairs and Lewis acidic rare-earth metal triflates as multi-role reagents.

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Analysis of Complex Molecules and Their Reactions on Surfaces by Means of Cluster-Induced Desorption/Ionization Mass Spectrometry

Desorption/Ionization Induced by Neutral SO2 Clusters (DINeC) is employed as a very soft and efficient desorption/ionization technique for mass spectrometry (MS) of complex molecules and their reactions on surfaces. DINeC is based on a beam of SO2 clusters impacting on the sample surface at low cluster energy. During cluster-surface impact, some of the surface molecules are desorbed and ionized via dissolvation in the impacting cluster; as a result of this dissolvation-mediated desorption mechanism, low cluster energy is sufficient and the desorption process is extremely soft. Both surface adsorbates and molecules of which the surface is composed of can be analyzed. Clear and fragmentation-free spectra from complex molecules such as peptides and proteins are obtained. DINeC does not require any special sample preparation, in particular no matrix has to be applied. The method yields quantitative information on the composition of the samples; molecules at a surface coverage as low as 0.1 % of a monolayer can be detected. Surface reactions such as H/D exchange or thermal decomposition can be observed in real-time and the kinetics of the reactions can be deduced. Using a pulsed nozzle for cluster beam generation, DINeC can be efficiently combined with ion trap mass spectrometry. The matrix-free and soft nature of the DINeC process in combination with the MSn capabilities of the ion trap allows for very detailed and unambiguous analysis of the chemical composition of complex organic samples and organic adsorbates on surfaces.

Neutral SO2 clusters of low kinetic energy (&#60; 0.8 eV/constituent) are used to desorb complex surface molecules such as peptides or lipids for further analysis by means of mass spectrometry using an ion trap mass spectrometer. No special sample preparation is required, and real-time observation of reactions is possible.

Desorption/Ionization Induced by Neutral SO2 Clusters (DINeC) is employed as a very soft and efficient desorption/ionization technique for mass ...

A Femtoliter Droplet Array for Massively Parallel Protein Synthesis from Single DNA Molecules

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The overall goal of the protocol is to prepare over one million ordered, uniform, stable, and biocompatible femtoliter droplets on a 1 cm2 planar substrate that can be used for cell-free protein synthesis.

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Self-Assembly of Hybrid Lipid Membranes Doped with Hydrophobic Organic Molecules at the Water/Air Interface

Because of their unique properties, including an ultrathin thickness (3-4 nm), ultrahigh resistivity, fluidity and self-assembly ability, lipid bilayers can be readily functionalized and have been used in various applications such as bio-sensors and bio-devices. In this study, we introduced a planar organic molecule: copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) to dope lipid membranes. The CuPc/lipid hybrid membrane forms at the water/air interface by self-assembly. In this membrane, the hydrophobic CuPc molecules are located between the hydrophobic tails of lipid molecules, forming a lipid/CuPc/lipid sandwich structure. Interestingly, an air-stable hybrid lipid bilayer can be readily formed by transferring the hybrid membrane onto a Si substrate. We report a straightforward method for incorporating nanomaterials into a lipid bilayer system, which represents a new methodology for the fabrication of biosensors and biodevices.

We report a protocol for producing a hybrid lipid membrane at the water/air interface by doping the lipid bilayer with copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) molecules. The resulting hybrid lipid membrane has a lipid/CuPc/lipid sandwich structure. This protocol can also be applied to the formation of other functional nanomaterials.

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Atomic Force Microscopy Combined with Infrared Spectroscopy as a Tool to Probe Single Bacterium Chemistry

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) is a novel combinatory technique, enabling simultaneous characterization of physical properties and chemical composition of sample with nanoscale resolution. By combining AFM with IR, the spatial resolution limitation of conventional IR is overcome, enabling a resolution of 20&#8211;100 nm to be achieved. This opens the door for a broad array of new applications of IR toward probing samples smaller than several micrometers, previously unachievable by means of conventional IR microscopy. AFM-IR is eminently suited for bacterial research, providing both spectral and spatial information at the single cell and intracellular level. The increasing global health concerns and unfavorable future prediction regarding bacterial infections, and especially, rapid development of antimicrobial resistance, has created an urgent need for a research tool capable of phenotypic probing at the single cell and subcellular level. AFM-IR offers the potential to address this need, by enabling detail characterization of chemical composition of a single bacterium. Here, we provide a complete protocol for sample preparation and data acquisition of single spectra and mapping modality, for the application of AFM-IR toward bacterial studies.

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) provides a powerful platform for bacterial studies, enabling to achieve nanoscale resolution. Both, mapping of subcellular changes (e.g., upon cell division) as well as comparative studies of chemical composition (e.g., arising from drug resistance) can be conducted at a single cell level in bacteria.

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) is a novel combinatory technique, enabling simultaneous characterization of physical properties and ...