The authors thank the DFG (Hu 997/2-2) for financial support. FS acknowledges the Hanns-Seidel-Stiftung (HSS). SKr was supported by the Elitenetzwerk Bayern in the framework of the doctorate program Material Science of Complex Interfaces. SKi thanks the SFB 863 for financial support.

NOTE: See Figure 2 for an overview of the process flow comprising the preparation, the data acquisition and data analysis steps.
1. Reagent Setup
NOTE: All chemicals must be handled with care, and thus a lab coat, gloves and eye protection should be used. All operations must be performed in a laboratory hood. In particular, special gloves should be worn in case of chloroform use.
<ol>
	<li>Use chemicals with low water content such as dry chloroform rapidly and store dry but not longer than a week. Store both chemicals at -20 &#176;C and under nitrogen or argon gas because APTES ((3-aminopropyl)triethoxysilan) (see Table 1) and PEG (polyethylenglycol) are hygroscopic and PEG is subject to oxidation in air.</li>
	<li>To avoid frequent exposure of the stock to atmospheric oxygen and moisture, prepare smaller aliquots, ideally within a glovebox system with a nitrogen atmosphere.</li>
</ol>
2. Equipment Setup
NOTE: To avoid possible cross-contamination, use fresh and clean vessels for each step.
<ol>
	<li>Clean glassware and tweezers in detergent solution for 30 min in an ultrasonic bath at 60 &#176;C.</li>
	<li>Rinse and sonicate equipment from step 2.1 two times thoroughly with ultrapure water.</li>
	<li>Heat equipment from step 2.1 in RCA solution (ultrapure water, hydrogen peroxide and ammonia (5:1:1)) to 75 &#176;C in an oven for 45 min and subsequently rinse them with ultrapure water.</li>
	<li>Finally, dry the glassware and tweezers under a stream of dry nitrogen or in an oven (100 &#176;C, 3 hr).</li>
</ol>
3. Tip Functionalization
NOTE: Use tweezers, vessels, etc. made from stainless steel, PTFE, glass or any other material which is chemically stable in organic solutions if applicable. Unless specified otherwise, carry out all steps at RT. The amount of incubation solution needed depends on the number of cantilever chips. Make sure that the cantilevers are immersed in the respective solutions at all time.
NOTE: Use a hood to avoid inhalation of organic vapors.
<ol>
	<li>Formation of OH-groups on cantilever surface (&#8216;activation&#8217;) (approximately 0.5 hr):
	<ol>
		<li>Use tweezers to place fresh cantilever chips (material: SiN, spring constant: 10-100 pN/nm) on a clean glass slide and put them in a plasma chamber (100 W).</li>
		<li>Evacuate chamber (~0.1 mbar).</li>
		<li>Flood chamber with oxygen gas and evacuate again.</li>
		<li>Activate the plasma process (power: 20%, duration: 15 min, process pressure: 0.25 mbar).</li>
	</ol>
	</li>
	<li>Amino-silanization of cantilevers (approximately 1 hr):
	<ol>
		<li>Prepare the 2.5 ml of APTES solution (see Table 1) in a glass Petri dish &#8211; do this ideally during the plasma process.</li>
		<li>Immediately after the plasma process, dip each cantilever for 1 sec in acetone and place them immediately afterwards in the APTES solution.</li>
		<li>Incubate for 15 min at RT.</li>
		<li>Carefully rinse the cantilever chips twice in 10 ml acetone and once in 10 ml chloroform.</li>
		<li>Optional final step: Place cantilever chips from previous step on a clean glass slide and bake them for 30 min at 70 &#176;C. Note that there are also alternative strategies for surface activation, e.g., UV treatment 12.</li>
	</ol>
	</li>
	<li>PEGylation (approximately 2 hr): 
	NOTE: Carry out steps 3.3.1-3.3.4 during amino-silanization. NHS and maleimide groups are subject to hydrolysis in aqueous environments and PEG itself is subject to oxidation in air. Therefore timing (especially in between the steps) is a critical parameter. Refer to 13 for further information.
	<ol>
		<li>Prepare the chloroform solution (see Table 1).</li>
		<li>To avoid condensation, warm PEG powders up to RT before opening the aliquot and weighing the appropriate amount.</li>
		<li>For the coupling of polymers or lipids with amino groups, separately solve NHS-PEG-NHS (6 kDa) and methyl-PEG-NHS (5 kDa) in the chloroform solution by vortexing them until they are completely solved. Refer to Table 1 for concentrations.
		<ol>
			<li>Alternatively. for the coupling of polymers with thiol groups separately solve Mal-NHS- PEG and methyl-PEG-NHS in the chloroform solution.</li>
		</ol>
		</li>
		<li>Mix the solutions as required to adjust a certain number ratio between the NHS- or the maleimide- and the methyl-terminated PEG molecules (typically 1:500). Note that the ideal ratio has to be determined iteratively in a series of preparation-experiment cycles.</li>
		<li>Incubate cantilever chips in the PEG solution for 1 hr within a chloroform saturated atmosphere to prevent evaporation of the chloroform.</li>
	</ol>
	</li>
	<li>Probe molecule conjugation (&#62; 1 hr): 
	NOTE: In the following 3.4.1-3.4.3, three examples for the covalent coupling of different probe molecules to the AFM tip are described. For each molecule the protocol has to be adjusted. Further note, that in example 1 and 3 NHS-chemistry is used whereas in example 2 the thiol functionalized polymer PI-g-PS is coupled to the PEG via maleimide-chemistry. For details see 14.
	<ol>
		<li>For poly(amino acid) poly-D-tyrosine (40-100 kDa)
		<ol>
			<li>Dissolve the polytyrosine conjugated probe molecule in 1 M NaOH. Adjust the concentration to 1 mg/ml.</li>
			<li>Exchange the NaOH for sodium borate buffer (pH 8.1) using spin desalting columns (7 kDa MWCO) immediately before the functionalization</li>
			<li>Rinse the cantilevers first with 5 ml of chloroform, then with 5 ml of ethanol and finally with 5 ml of borate buffer.</li>
			<li>Incubate the cantilever chips for 1 hr in the polytyrosine borate buffer solution and then rinse with borate buffer and ultrapure water. Store in ultrapure water until the measurement.</li>
		</ol>
		</li>
		<li>For polymers with a linear backbone (polyisoprene, 119 kDa) having grafted side chains (polystyrene, 88 kDa) 14
		<ol>
			<li>Dissolve the PI-g-PS in dry chloroform. Adjust the concentration of the conjugation solution to 4 mg/ml.</li>
			<li>Rinse the cantilevers with 5 ml of chloroform and incubate for at least 1 hr in the conjugation solution.</li>
			<li>Rinse again in 5 ml of chloroform and store in chloroform in a chloroform-saturated environment (to prevent chloroform evaporation) until the measurement.</li>
		</ol>
		</li>
		<li>For the lipid POPE
		<ol>
			<li>Dissolve POPE in dry chloroform. Adjust the concentration to 20 mM.</li>
			<li>Rinse the cantilevers with 5 ml of ethanol and 5 ml of ultrapure water before incubation of the cantilevers for 1 hr in the lipid solution.</li>
			<li>After incubation, rinse cantilevers with 5 ml of chloroform, 5 ml of ethanol and 5 ml of warm, ultrapure water (in this order) to remove unbound probe molecules and to get rid of chloroform residues. Use functionalized tips immediately. Alternatively, store them in chloroform until needed.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/52456/52456fig1.jpg" /> 
Figure 1. (A) Schematic showing the tip functionalization process using the example of NHS chemistry. (B)&#160;Chemical bonding employed to attach a probe molecule to the tip via an amino group. <a href="https://www.jove.com/files/ftp_upload/52456/52456fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Surface Preparation
<ol>
	<li>Self-assembled monolayer (SAM) 
	NOTE: As surface for the first example, a self-assembled monolayer was chosen. Refer to literature 15 for details.
	<ol>
		<li>Clean glass slides with detergent solution and then twice in ultrapure water in an ultrasonic bath for 30 min each. Then, as an additional cleaning step, place them in the RCA solution (see step 2.3) at 75 &#176;C for 15 min.</li>
		<li>Coat the slides with 10 nm chromium nickel and 100 nm gold in a vacuum coater. Then store in the fridge and clean again in the RCA solution directly prior to the next step.</li>
		<li>Incubate the slides of the previous step in 2 mM 1-dodecanthiol/ethanol solution for 12 hr. The thiol groups bind to the gold surface and a hydrophobic monolayer self assembles. Rinse the slides with ethanol and ultrapure water and then dry by a stream of nitrogen gas. Confirm the hydrophobicity of the prepared surface with static contact angle measurements in a goniometer with a CCD camera10.</li>
	</ol>
	</li>
	<li>Hydrogen terminated diamond 
	NOTE: As a surface for the second example, a hydrogen terminated diamond was chosen. The surface preparation was performed as described previously16.</li>
	<li>Supported lipid bilayer 
	NOTE: In the last example, a surface supported lipid bilayer (SLB) was used as a surface. To obtain such a surface, a solution (0.1 mg/ml) of large unilamellar vesicles (LUVs) consisting of the phospholipid POPC were formed by the extrusion method. Then 50 &#181;l of the LUV solution were put on a freshly cleaved mica sheet (A = 1 cm&#178;) and incubated for 30 min. Finally the solution was rinsed with 20 ml of ultrapure water. See 17,18 for a detailed description of the preparation of SLBs.</li>
</ol>
5. Data Acquisition
NOTE: For the experiments, use an AFM, which provides the capability to measure in liquids. The data acquisition and data analysis procedures are applicable regardless of the AFM model used. Furthermore, in some experiments it is advantageous to have the possibility to control the temperature within the liquid cell. Cantilever deflection was detected via the laser beam deflection method 19. Spring constants were determined with the thermal noise method 20.
<ol>
	<li>Insert the functionalized cantilever into an AFM cantilever holder which is suitable for measuring in fluids.</li>
	<li>Put the surface (preparation see above) to be sampled into the AFM and cover it with liquid. Both, cantilever and surface should now be immersed in the liquid with the cantilever tip pointing towards the surface. Note that in many cases a drop of liquid is enough. In any case evaporation of the fluid should be avoided, e.g., by using a closed fluid cell.</li>
	<li>If applicable, adjust desired temperature on the controller.</li>
	<li>Let the system equilibrate for at least half an hour.</li>
	<li>Record a thermal noise spectrum of the cantilever with the cantilever far away from the surface in order to exclude any surface damping effects. Note that usually 10 or more spectra have to be accumulated to obtain a sufficient signal-to-noise ratio.</li>
	<li>Approach the surface. Note that in order to conserve tip geometry and tip functionalization the approach process has to be carried out carefully. This can be accomplished by, for example, approaching in intermittent contact mode.</li>
	<li>Determine the inverse optical lever sensitivity (InvOLS) measured in nm/volt. Do this by pushing the cantilever tip against a hard surface.
	<ol>
		<li>Determine the InvOLS by measuring the slope of the piezo travel distance vs. the change in photodiode voltage. Fit a line to the part of the force-distance curve where the cantilever tip is in contact with the surface. In experiments involving a soft surface like lipid bilayers, carry out this step on areas which are not covered with a lipid bilayer (&#8216;defect spots&#8217;).</li>
	</ol>
	</li>
	<li>Determine the spring constant (pN/nm) by fitting a harmonic oscillator to the thermal noise spectrum. Use automated software function for spring constant determination; otherwise consult literature 20.</li>
	<li>Start the experiment.
	<ol>
		<li>Record numerous force-distance curves at each experimental condition. Typically, use the following values for the experimental parameters: Sampling rate of 5&#160;kHz, tip speed of 1 &#181;m/sec, retract distance of 1 &#181;m. 
		NOTE: Sometimes, depending on the dynamics of the studied experiment, it might be necessary to wait a certain amount of time with the tip in contact with the surface (&#8216;dwell time&#8217;). 
		NOTE: The experiment specific parameters can deviate substantially from the values given above.</li>
		<li>During long term measurements, zero the laser position on the photodiode from time to time by repositioning the photodiode.</li>
		<li>Optional: After every force curve, relocate the AFM-tip to another x-y position.</li>
		<li>At the end of the experiment, determine the sensitivity and the spring constant again to check for consistency and stability of the system. 
		NOTE: Comparing the spring constant and sensitivity before and after the experiment is also a means to ensure that the properties of the system and the properties of the tip remained unchanged over the course of the experiment. If the difference of the spring constants is not too large (&#60;=15%), they can be averaged, since the intrinsic uncertainty of spring constant determination is also around 15%. For larger differences, it is advisable to first find and eliminate the cause for the changing apparent spring constant before continuing with further experiments.</li>
	</ol>
	</li>
</ol>
6. Data Preparation
NOTE: In this section general data preparation steps which are typically carried out independent of the specific type of experiment to convert units to newton and nanometer as well as to correct the data are described. The experiment specific data analyses are briefly described further below in the respective representative example section.
<ol>
	<li>Typically, carry out all data preparation and analysis steps automatically by a home-written algorithm, e.g., based on the software IGOR Pro 6.</li>
	<li>Convert the raw deflection signal (Defl) [V] into force by multiplying it with the InvOLS [nm/V] and the spring constant [nN/nm].</li>
	<li>Offset the force in such a way that the force on the cantilever in sufficient distance from the surface (unloaded cantilever) becomes 0 nN.</li>
	<li>Calculate the actual tip position (also called separation when measured relative to the surface) z*, which takes the bending of the cantilever into account: 
	<img alt="Equation 1" fo:content-width="2in" src="/files/ftp_upload/52456/52456eq1.jpg" /></li>
	<li>Offset z* in such a way that z* = 0 corresponds to the z position of the surface. 
	NOTE: Here, F is the force which acts on the tip and hence bends the cantilever (determined in step 6.2.), z is the measured z-sensor position (directly given by the instrument) and k represents the spring constant (determined in step 5.8).</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/52456/52456fig2.jpg" /> 
Figure 2. Process flow diagram showing the sample preparation, the data acquisition and the data analysis. <a href="https://www.jove.com/files/ftp_upload/52456/52456fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors declare that they have no competing financial interests.

During the last decades, single molecule experiments have provided unprecedented insights into molecular mechanisms and turned out to be an invaluable approach in life science and beyond. To achieve good and meaningful statistics from SMFS experiments, ideally one and the same molecule is used over the whole course of the experiment. In contrast to experiments with ensembles of molecules, SMFS experiments are able to detect rare events and hidden molecular states. Another advantage of single molecule experiments is that ...

Over the past 30 years, atomic force microscopy (AFM) has turned out to be a valuable imaging technique to study biological 1,2 and synthetic 3 materials and surfaces since it provides molecular spatial resolution in all three dimensions and can be operated in various solvent environments. In addition, AFM-single molecule force spectroscopy (SMFS) enables to measure forces ranging from the pN to &#181;N regime and has given unprecedented insight for example into protein folding 4,5, polymer physics 6&#8211;8 , and single molecule-surface interaction9&#8211;12.The rationale behind studying single molecules rather than an ensemble of molecules is to avoid averaging effects which often mask rare events or hidden molecular states. Furthermore, a multitude of molecular parameters such as the contour length, the Kuhn length, the adhesion free energy, etc. can be obtained. This is detailed in the examples below. In a typical AFM-SMFS experiment, the probe molecule is coupled to a very sharp tip via a linker molecule. The tip itself is located at the end of a bendable cantilever. If the tip is brought into contact with the surface the probe molecule will interact with this surface. By observing the deflection of the cantilever upon retraction of the tip, the force, and hence the free energy, to detach the molecule from the surface can be determined. To obtain meaningful statistics, a large number of so called force-distance curves have to be acquired. Furthermore, to have true single molecule experiments (i.e., using one and the same probe molecule over the duration of the whole experiment) the probe molecule should be coupled covalently to the AFM tip. Here, an experimental protocol for cantilever functionalization with a single molecule via a covalent bond is presented. The single molecule can either be coupled via an amino or a thiol group to the AFM tip. The conjugation process can be performed in a broad variety of solvents (organic and aqueous) to account for the solvation properties of the polymers used.
In the first part, a general protocol to covalently attach a single molecule (&#8220;probe molecule&#8221;) via a linker molecule to an AFM tip is described. To this end, organic NHS- or maleimide-chemistry is used13. Along with the protocol for three example molecules, the data acquisition and data analysis processes are described and references for further reading are provided. The example molecules are: the (linear) polymer tyrosine, the graft polymer PI-g-PS and the lipid POPE. This includes slight variations of the protocol, for example to covalently attach cysteines. In addition, a section is dedicated to the preparation of different surfaces such as a diamond surface, a CH3-self-assembled monolayer and lipid bilayers. These interfaces have proven to be good references and examples.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hellmanex III alkaline liquid concentrate (detergent solution)</td>
			<td>Hellma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>RCA (ultrapure water, hydrogen peroxide (35 %), ammonia (32%); 5:1:1(v/v/v))</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vectabond reagent / APTES (3-Aminopropyl)triethoxysilane</td>
			<td>Vectorlabs</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dry acetone (&#60; 50 ppm H2O)</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dry chloroform (&#62; 99.9 %)</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Triethylamine</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrapure water</td>
			<td>Biochrom, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Di-sodium tetraborate (&#62; 99.5 %)</td>
			<td>Biochrom, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Boric Acid</td>
			<td>Biochrom, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Monofunctional &#945;-methoxy-&#969;-NHS PEG, 5kDa, &#8220;methyl-PEG-NHS&#8221;</td>
			<td>Rapp, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heterobifunctional &#945;,&#969;-bis-NHS PEG, 6 kDa, &#8220;NHS-PEG-NHS&#8221;</td>
			<td>Rapp, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heterobifunctional &#945;-maleimidohexanoic- &#969;-NHS PEG, 5 kDa, &#8220;Mal-PEG-NHS&#8221;</td>
			<td>Rapp, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Probe molecule (polymer, lipid, etc.)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sufficient amount of glass crystallising dishes with spout (10 ml), glass petri dishes (500 &#181;l) and glass lids</td>
			<td>VWR International GmbH, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory oven model UF30</td>
			<td>Memmert, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature controlled sonicator</td>
			<td>VWR International GmbH, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plasma system &#34;Femto&#34;, 100 W</td>
			<td>Diener, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>One separate glass syringe for each organic solvent</td>
			<td>VWR International GmbH, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex mixer</td>
			<td>VWR International GmbH, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes (0.5 ml or 1.5 ml)</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes: 10-100 &#181;l, 50-200 &#181;l and 100-1000 &#181;l</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AFM with temperature controlled fluid cell (e.g. MFP-3D with BioHeater)</td>
			<td>Asylulm Research, Santa Barbara</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Soft SiN cantilevers cantilever, typically made from silicon nitride (SiN) (spring constant less than 100 pN/nm, e.g. MLCT)</td>
			<td>Bruker AXS, Santa Barbara</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

investigating single molecule adhesion by atomic force spectroscopy

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

In the following, the results for the above described example molecules, namely the polymers poly(amino acid) polytyrosine, the graft polymer PI-g-PS and the phospholipid POPE, are presented. First for each example, experiment specific details for the data acquisition and data preparation are provided. Then, the exemplary results for experiments where these molecules were desorbed from different surfaces (CH3-SAMs, hydrogen terminated diamond and lipid bilayers) are shown. Determination of the adhesio...

Watch this Scientific Journal Video about Investigating Single Molecule Adhesion by Atomic Force Spectroscopy at JoVE.com

Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

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

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

investigating-single-molecule-adhesion-by-atomic-force-spectroscopy

Research

JoVE Journal

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10.3K ビュー.  Technische Universität München. A protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. Procedures and examples to determine the adhesion force and free energy of these molecules on solid supports and bio-interfaces are provided.

ビデオ: Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Compact Quantum Dots for Single-molecule Imaging

Single-molecule imaging is an important tool for understanding the mechanisms of biomolecular function and for visualizing the spatial and temporal heterogeneity of molecular behaviors that underlie cellular biology 1-4. To image an individual molecule of interest, it is typically conjugated to a fluorescent tag (dye, protein, bead, or quantum dot) and observed with epifluorescence or total internal reflection fluorescence (TIRF) microscopy. While dyes and fluorescent proteins have been the mainstay of fluorescence imaging for decades, their fluorescence is unstable under high photon fluxes necessary to observe individual molecules, yielding only a few seconds of observation before complete loss of signal. Latex beads and dye-labeled beads provide improved signal stability but at the expense of drastically larger hydrodynamic size, which can deleteriously alter the diffusion and behavior of the molecule under study.
Quantum dots (QDs) offer a balance between these two problematic regimes. These nanoparticles are composed of semiconductor materials and can be engineered with a hydrodynamically compact size with exceptional resistance to photodegradation 5. Thus in recent years QDs have been instrumental in enabling long-term observation of complex macromolecular behavior on the single molecule level. However these particles have still been found to exhibit impaired diffusion in crowded molecular environments such as the cellular cytoplasm and the neuronal synaptic cleft, where their sizes are still too large 4,6,7.
Recently we have engineered the cores and surface coatings of QDs for minimized hydrodynamic size, while balancing offsets to colloidal stability, photostability, brightness, and nonspecific binding that have hindered the utility of compact QDs in the past 8,9. The goal of this article is to demonstrate the synthesis, modification, and characterization of these optimized nanocrystals, composed of an alloyed HgxCd1-xSe core coated with an insulating CdyZn1-yS shell, further coated with a multidentate polymer ligand modified with short polyethylene glycol (PEG) chains (Figure 1). Compared with conventional CdSe nanocrystals, HgxCd1-xSe alloys offer greater quantum yields of fluorescence, fluorescence at red and near-infrared wavelengths for enhanced signal-to-noise in cells, and excitation at non-cytotoxic visible wavelengths. Multidentate polymer coatings bind to the nanocrystal surface in a closed and flat conformation to minimize hydrodynamic size, and PEG neutralizes the surface charge to minimize nonspecific binding to cells and biomolecules. The end result is a brightly fluorescent nanocrystal with emission between 550-800 nm and a total hydrodynamic size near 12 nm. This is in the same size range as many soluble globular proteins in cells, and substantially smaller than conventional PEGylated QDs (25-35 nm).

We describe the preparation of colloidal quantum dots with minimized hydrodynamic size for single-molecule fluorescence imaging. Compared to conventional quantum dots, these nanoparticles are similar in size to globular proteins and are optimized for single-molecule brightness, stability against photodegradation, and resistance to nonspecific binding to proteins and cells.

Single-molecule imaging is an important tool for  understanding the mechanisms of biomolecular function and for visualizing the  spatial and temporal ...

Scanning-probe Single-electron Capacitance Spectroscopy

The integration of low-temperature scanning-probe techniques and single-electron capacitance spectroscopy represents a powerful tool to study the electronic quantum structure of small systems - including individual atomic dopants in semiconductors. Here we present a capacitance-based method, known as Subsurface Charge Accumulation (SCA) imaging, which is capable of resolving single-electron charging while achieving sufficient spatial resolution to image individual atomic dopants. The use of a capacitance technique enables observation of subsurface features, such as dopants buried many nanometers beneath the surface of a semiconductor material1,2,3. In principle, this technique can be applied to any system to resolve electron motion below an insulating surface.
As in other electric-field-sensitive scanned-probe techniques4, the lateral spatial resolution of the measurement depends in part on the radius of curvature of the probe tip. Using tips with a small radius of curvature can enable spatial resolution of a few tens of nanometers. This fine spatial resolution allows investigations of small numbers (down to one) of subsurface dopants1,2. The charge resolution depends greatly on the sensitivity of the charge detection circuitry; using high electron mobility transistors (HEMT) in such circuits at cryogenic temperatures enables a sensitivity of approximately 0.01 electrons/Hz&frac12; at 0.3 K 5.

Scanning-probe single-electron capacitance spectroscopy facilitates the study of single-electron motion in localized subsurface regions. A sensitive charge-detection circuit is incorporated into a cryogenic scanning probe microscope to investigate small systems of dopant atoms beneath the surface of semiconductor samples.

The integration of low-temperature scanning-probe techniques and single-electron capacitance spectroscopy represents a powerful tool to study the ...

Single Wavelength Shadow Imaging of Caenorhabditis elegans Locomotion Including Force Estimates

This study demonstrates an inexpensive and straightforward technique that allows the measurement of physical properties such as position, velocity, acceleration and forces involved in the locomotory behavior of nematodes suspended in a column of water in response to single wavelengths of light. We demonstrate how to evaluate the locomotion of a microscopic organism using Single Wavelength Shadow Imaging (SWSI) using two different examples.
The first example is a systematic and statistically viable study of the average descent of C. elegans in a column of water. For this study, we used living and dead wildtype C. elegans. When we compared the velocity and direction of nematode active movement with the passive descent of dead worms within the gravitational field, this study showed no difference in descent-times. The average descent was 1.5 mm/sec &#177; 0.1 mm/sec for both the live and dead worms using 633 nm coherent light.
The second example is a case study of select individual C. elegans changing direction during the descent in a vertical water column. Acceleration and force are analyzed in this example. This case study demonstrates the scope of other physical properties that can be evaluated using SWSI while evaluating the behavior using single wavelengths in an environment that is not accessible with traditional microscopes. Using this analysis we estimated an individual nematode is capable of thrusting with a force in excess of 28 nN.
Our findings indicate that living nematodes exert 28 nN when turning, or moving against the gravitational field. The findings further suggest that nematodes passively descend in a column of water, but can actively resist the force of gravity primarily by turning direction.

Single Wavelength Shadow Imaging of Caenorhabditis elegans Locomotion Including Force Estimates

The technique presented here measures the path of freely swimming microscopic species using single wavelength exposure. C. elegans are used to demonstrate shadow imaging as an inexpensive alternative to costly microscopes. This technique can be adapted to accommodate various orientations, environments and species to measure direction, speed, acceleration and forces.

This study demonstrates an inexpensive and straightforward technique that allows the measurement of physical properties such as position, velocity, ...

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be measured to ~10 pN by a laser and sectored detector, which can be converted to image topography. Amplitude modulation or &#8220;tapping mode&#8221; AFM involves the probe making intermittent contact with the surface while oscillating at its resonant frequency to produce an image. Used in conjunction with a fluid cell, tapping-mode AFM enables the imaging of biological macromolecules such as proteins in physiologically relevant conditions. Tapping-mode AFM requires manual tuning of the probe and frequent adjustments of a multitude of scanning parameters which can be challenging for inexperienced users. To obtain high-quality images, these adjustments are the most time consuming.
PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) produces an image by measuring a force response curve for every point of contact with the sample. With ScanAsyst software, PF-QNM can be automated. This software adjusts the set-point, drive frequency, scan rate, gains, and other important scanning parameters automatically for a given sample. Not only does this process protect both fragile probes and samples, it significantly reduces the time required to obtain high resolution images. PF-QNM is compatible for AFM imaging in fluid; therefore, it has extensive application for imaging biologically relevant materials.
The method presented in this paper describes the application of PF-QNM to obtain images of a bacterial red-light photoreceptor, RpBphP3 (P3), from photosynthetic R. palustris in its light-adapted state. Using this method, individual protein dimers of P3 and aggregates of dimers have been observed on a mica surface in the presence of an imaging buffer. With appropriate adjustments to surface and/or solution concentration, this method may be generally applied to other biologically relevant macromolecules and soft materials.

A method for investigating the structure of a protein photoreceptor using atomic force microscopy (AFM) is described in this paper. PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) reveals intact protein dimers on a mica surface.

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be ...

Making Record-efficiency SnS Solar Cells by Thermal Evaporation and Atomic Layer Deposition

Tin sulfide (SnS) is a candidate absorber material for Earth-abundant, non-toxic solar cells. SnS offers easy phase control and rapid growth by congruent thermal evaporation, and it absorbs visible light strongly. However, for a long time the record power conversion efficiency of SnS solar cells remained below 2%. Recently we demonstrated new certified record efficiencies of 4.36% using SnS deposited by atomic layer deposition, and 3.88% using thermal evaporation. Here the fabrication procedure for these record solar cells is described, and the statistical distribution of the fabrication process is reported. The standard deviation of efficiency measured on a single substrate is typically over 0.5%. All steps including substrate selection and cleaning, Mo sputtering for the rear contact (cathode), SnS deposition, annealing, surface passivation, Zn(O,S) buffer layer selection and deposition, transparent conductor (anode) deposition, and metallization are described. On each substrate we fabricate 11 individual devices, each with active area 0.25 cm2. Further, a system for high throughput measurements of current-voltage curves under simulated solar light, and external quantum efficiency measurement with variable light bias is described. With this system we are able to measure full data sets on all 11 devices in an automated manner and in minimal time. These results illustrate the value of studying large sample sets, rather than focusing narrowly on the highest performing devices. Large data sets help us to distinguish and remedy individual loss mechanisms affecting our devices.

Tin sulfide (SnS) is a candidate material for Earth-abundant, non-toxic solar cells. Here, we demonstrate the fabrication procedure of the SnS solar cells employing atomic layer deposition, which yields 4.36% certified power conversion efficiency, and thermal evaporation which yields 3.88%.

Tin sulfide (SnS) is a candidate absorber material for Earth-abundant, non-toxic solar cells. SnS offers easy phase control and rapid growth by congruent ...

Sub-nanometer Resolution Imaging with Amplitude-modulation Atomic Force Microscopy in Liquid

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown that when operated in amplitude-modulation (tapping) mode, atomic or molecular-level resolution images can be achieved over a wide range of soft and hard samples in liquid. In these situations, small oscillation amplitudes (SAM-AFM) enhance the resolution by exploiting the solvated liquid at the surface of the sample. Although the technique has been successfully applied across fields as diverse as materials science, biology and biophysics and surface chemistry, obtaining high-resolution images in liquid can still remain challenging for novice users. This is partly due to the large number of variables to control and optimize such as the choice of cantilever, the sample preparation, and the correct manipulation of the imaging parameters. Here, we present a protocol for achieving high-resolution images of hard and soft samples in fluid using SAM-AFM on a commercial instrument. Our goal is to provide a step-by-step practical guide to achieving high-resolution images, including the cleaning and preparation of the apparatus and the sample, the choice of cantilever and optimization of the imaging parameters. For each step, we explain the scientific rationale behind our choices to facilitate the adaptation of the methodology to every user's specific system.

We present a method for achieving sub-nanometer resolution images with amplitude-modulation (tapping mode) atomic force microscopy in liquid. The method is demonstrated on commercial atomic force microscopes. We explain the rationale behind our choices of parameters and suggest strategies for resolution optimization.

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown ...

Quantitative Analysis of Vacuum Induction Melting by Laser-induced Breakdown Spectroscopy

Vacuum induction melting is a popular method for refining high purity metal and alloys. Traditionally, standard process control in metallurgy involves several steps, include drawing samples, cooling, cutting, transport to the laboratory, and analysis. The whole analysis process requires more than 30 minutes, which hinders on-line process control. Laser-induced breakdown spectroscopy is an excellent on-line analysis method that can satisfy the requirements of vacuum induction melting because it is fast and noncontact and does not require sample preparation. The experimental facility uses a lamp-pumped Q-switched laser to ablate melted liquid steel with an output energy of 80 mJ, a frequency of 5 Hz, a FWHM pulse width of 20 ns, and a working wavelength of 1,064 nm. A multi-channel linear charge coupled device (CCD) spectrometer is used to measure the emission spectrum in real time, with a spectral range from 190 to 600 nm and a resolution of 0.06 nm at a wavelength of 200 nm. The protocol includes several steps: standard alloy sample preparation and an ingredient test, smelting of standard samples and determination of the laser breakdown spectrum, and construction of the elements concentration quantitative analysis curve of each element. To realize the concentration analysis of unknown samples, the spectrum of a sample also needs to be measured and disposed with the same process. The composition of all main elements in the melted alloy can be quantitatively analyzed with an internal standard method. The calibration curve shows that the limit of detection of most metal elements ranges from 20-250 ppm. The concentration of elements, such as Ti, Mo, Nb, V, and Cu, can be lower than 100 ppm, and the concentrations of Cr, Al, Co, Fe, Mn, C, and Si range from 100-200 ppm. The R2 of some calibration curves can exceed 0.94.

During vacuum induction melting, laser-induced breakdown spectroscopy is used to perform real-time quantitative analysis of the main-ingredient elements of a molten alloy.

Vacuum induction melting is a popular method for refining high purity metal and alloys. Traditionally, standard process control in metallurgy involves ...

An Efficient Method for Selective Desalination of Radioactive Iodine Anions by Using Gold Nanoparticles-Embedded Membrane Filter

Here, we demonstrate a detail protocol for the preparation of nanomaterials-embedded composite membranes and its application to the efficient and ion-selective removal of radioactive iodines. By using citrate-stabilized gold nanoparticles (mean diameter: 13 nm) and cellulose acetate membranes, gold nanoparticle-embedded cellulose acetate membranes (Au-CAM) have easily been fabricated. The nano-adsorbents on Au-CAM were highly stable in the presence of high concentration of inorganic salts and organic molecules. The iodide ions in aqueous solutions could rapidly be captured by this engineered membrane. Through a filtration process using an Au-CAM containing filter unit, excellent removal efficiency (&gt;99%) as well as ion-selective desalination result was achieved in a short time. Moreover, Au-CAM provided good reusability without significant decrease of its performances. These results suggested that the present technology using the engineered hybrid membrane will be a promising process for the large-scale decontamination of radioactive iodine from liquid wastes.

An efficient method for the rapid and ion-selective desalination of radioactive iodine in several aqueous solutions is described by using gold nanoparticles-immobilized cellulose acetate membrane filters.

Here, we demonstrate a detail protocol for the preparation of nanomaterials-embedded composite membranes and its application to the efficient and ...

Atomic Force Microscopy Cantilever-Based Nanoindentation: Mechanical Property Measurements at the Nanoscale in Air and Fluid

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by the probe tip and its contact area with the sample can be quantified, it is possible to determine the nanoscale mechanical properties (e.g., elastic or Young&#39;s modulus) of the surface being probed. A detailed procedure for performing quantitative AFM cantilever-based nanoindentation experiments is provided here, with representative examples of how the technique can be applied to determine the elastic moduli of a wide variety of sample types, ranging from kPa to GPa. These include live mesenchymal stem cells (MSCs) and nuclei in physiological buffer, resin-embedded dehydrated loblolly pine cross-sections, and Bakken shales of varying composition.
Additionally, AFM cantilever-based nanoindentation is used to probe the rupture strength (i.e., breakthrough force) of phospholipid bilayers. Important practical considerations such as method choice and development, probe selection and calibration, region of interest identification, sample heterogeneity, feature size and aspect ratio, tip wear, surface roughness, and data analysis and measurement statistics are discussed to aid proper implementation of the technique. Finally, co-localization of AFM-derived nanomechanical maps with electron microscopy techniques that provide additional information regarding elemental composition is demonstrated.

Quantifying the contact area and force applied by an atomic force microscope (AFM) probe tip to a sample surface enables nanoscale mechanical property determination. Best practices to implement AFM cantilever-based nanoindentation in air or fluid on soft and hard samples to measure elastic modulus or other nanomechanical properties are discussed.

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by ...

Parallel Active Cantilever Arrays in AFMS to Enable High-Throughput Inspections

Active Probe Atomic Force Microscopy with Quattro-Parallel Cantilever Arrays for High-Throughput Large-Scale Sample Inspection

An Atomic Force Microscope&#160;(AFM) is&#160;a powerful and versatile tool for nanoscale surface studies to capture 3D topography images of samples. However, due to their limited imaging throughput, AFMs have not been widely adopted for large-scale inspection purposes. Researchers have developed high-speed AFM systems to record dynamic process videos in chemical and biological reactions at tens of frames per second, at the cost of a small imaging area of up to several square micrometers. In contrast, inspecting large-scale nanofabricated structures, such as semiconductor wafers, requires nanoscale spatial resolution imaging of a static sample over hundreds of square centimeters with high productivity. Conventional AFMs use a single passive cantilever probe with an optical beam deflection system, which can only collect one pixel at a time during AFM imaging, resulting in low imaging throughput. This work utilizes an array of active cantilevers with embedded piezoresistive sensors and thermomechanical actuators, which allows simultaneous multi-cantilever operation in parallel operation for increased imaging throughput. When combined with large-range nano-positioners and proper control algorithms, each cantilever can be individually controlled to capture multiple AFM images. With data-driven post-processing algorithms, the images can be stitched together, and defect detection can be performed by comparing them to the desired geometry. This paper introduces principles of the custom AFM using the active cantilever arrays, followed by a discussion on practical experiment considerations for inspection applications. Selected example images of silicon calibration grating, highly-oriented pyrolytic graphite, and extreme ultraviolet lithography masks are captured using an array of four active cantilevers (&#34;Quattro&#34;) with a 125 &#181;m tip separation distance. With more engineering integration, this high-throughput, large-scale imaging tool can provide 3D metrological data for extreme ultraviolet (EUV) masks, chemical mechanical planarization (CMP) inspection, failure analysis, displays, thin-film step measurements, roughness measurement dies, and laser-engraved dry gas seal grooves.

Author Spotlight: Introduction to Active Probe Atomic Force Microscopy with Quattro-Parallel Cantilever Arrays

Large-scale sample inspection with nanoscale resolution has a wide range of applications, especially for nanofabricated semiconductor wafers. Atomic force microscopes can be a great tool for this purpose, but are limited by their imaging speed. This work utilizes parallel active cantilever arrays in AFMs to enable high-throughput and large-scale inspections.

An Atomic Force Microscope&#160;(AFM) is&#160;a powerful and versatile tool for nanoscale surface studies to capture 3D topography images of samples. ...