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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</ol>

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><tbody><tr><td colspan="4">&lt;strong&gt;Materials&lt;/strong&gt;</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 (&amp;lt; 50 ppm H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma</td><td></td><td></td></tr><tr><td>Dry chloroform (&gt; 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 (&gt;&amp;nbsp;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 &amp;alpha;-methoxy-&amp;omega;-NHS PEG, 5 kDa, &amp;ldquo;methyl-PEG-NHS&amp;rdquo;</td><td>Rapp, Germany</td><td></td><td></td></tr><tr><td>Heterobifunctional &amp;alpha;,&amp;omega;-bis-NHS PEG, 6&amp;nbsp;kDa, &amp;ldquo;NHS-PEG-NHS&amp;rdquo;</td><td>Rapp, Germany</td><td></td><td></td></tr><tr><td>Heterobifunctional &amp;alpha;-maleimidohexanoic- &amp;omega;-NHS PEG, 5 kDa, &amp;ldquo;Mal-PEG-NHS&amp;rdquo;</td><td>Rapp, Germany</td><td></td><td></td></tr><tr><td>Probe molecule (polymer, lipid, &lt;em&gt;etc.&lt;/em&gt;)</td><td></td><td></td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Equipment&lt;/strong&gt;</td></tr><tr><td>Sufficient amount of glass crystallising dishes with spout (10&amp;nbsp;ml), glass Petri dishes (500 &amp;micro;l) and glass lids</td><td>VWR International GmbH, Germany</td><td></td><td>&lt;br /&gt;&amp;nbsp;&lt;br /&gt;&amp;nbsp;&lt;br /&gt;&amp;nbsp;&lt;br /&gt;&amp;nbsp;&lt;br /&gt;&amp;nbsp;&lt;br /&gt;&amp;nbsp;</td></tr><tr><td colspan="4">[header]</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 &quot;Femto&quot;, 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 &amp;micro;l, 50-200 &amp;micro;l and 100-1,000&amp;nbsp;&amp;micro;l</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>AFM with temperature controlled fluid cell (&lt;em&gt;e.g.&lt;/em&gt; 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, &lt;em&gt;e.g.&lt;/em&gt; 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...

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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

Engineering

10.3K Views.  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.

Video: Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Quantitative and Qualitative Examination of Particle-particle Interactions Using Colloidal Probe Nanoscopy

Colloidal Probe Nanoscopy (CPN), the study of the nano-scale interactive forces between a specifically prepared colloidal probe and any chosen substrate using the Atomic Force Microscope (AFM), can provide key insights into physical interactions present within colloidal systems. Colloidal systems are widely existent in several applications including, pharmaceuticals, foods, paints, paper, soil and minerals, detergents, printing and much more.1-3 Furthermore, colloids can exist in many states such as emulsions, foams and suspensions. Using colloidal probe nanoscopy one can obtain key information on the adhesive properties, binding energies and even gain insight into the physical stability and coagulation kinetics of the colloids present within. Additionally, colloidal probe nanoscopy can be used with biological cells to aid in drug discovery and formulation development. In this paper we describe a method for conducting colloidal probe nanoscopy, discuss key factors that are important to consider during the measurement, and show that both quantitative and qualitative data that can be obtained from such measurements.

Colloidal probe nanoscopy can be used within a variety of fields to gain insight into the physical stability and coagulation kinetics of colloidal systems and aid in drug discovery and formulation sciences using biological systems. The method described within provides a quantitative and qualitative means to study such systems.

Colloidal Probe Nanoscopy (CPN), the study of the nano-scale interactive forces between a specifically prepared colloidal probe and any chosen substrate ...

Chemistry

Adhesion Frequency Assay for In Situ Kinetics Analysis of Cross-Junctional Molecular Interactions at the Cell-Cell Interface

The micropipette adhesion assay was developed in 1998 to measure two-dimensional (2D) receptor-ligand binding kinetics1. The assay uses a human red blood cell (RBC) as adhesion sensor and presenting cell for one of the interacting molecules. It employs micromanipulation to bring the RBC into contact with another cell that expresses the other interacting molecule with precisely controlled area and time to enable bond formation. The adhesion event is detected as RBC elongation upon pulling the two cells apart. By controlling the density of the ligands immobilized on the RBC surface, the probability of adhesion is kept in mid-range between 0 and 1. The adhesion probability is estimated from the frequency of adhesion events in a sequence of repeated contact cycles between the two cells for a given contact time. Varying the contact time generates a binding curve. Fitting a probabilistic model for receptor-ligand reaction kinetics1 to the binding curve returns the 2D affinity and off-rate.
The assay has been validated using interactions of Fc&gamma; receptors with IgG Fc1-6, selectins with glycoconjugate ligands6-9, integrins with ligands10-13, homotypical cadherin binding14, T cell receptor and coreceptor with peptide-major histocompatibility complexes15-19.
The method has been used to quantify regulations of 2D kinetics by biophysical factors, such as the membrane microtopology5, membrane anchor2, molecular orientation and length6, carrier stiffness9, curvature20, and impingement force20, as well as biochemical factors, such as modulators of the cytoskeleton and membrane microenvironment where the interacting molecules reside and the surface organization of these molecules15,17,19.
The method has also been used to study the concurrent binding of dual receptor-ligand species3,4, and trimolecular interactions19 using a modified model21.
The major advantage of the method is that it allows study of receptors in their native membrane environment. The results could be very different from those obtained using purified receptors17. It also allows study of the receptor-ligand interactions in a sub-second timescale with temporal resolution well beyond the typical biochemical methods.
To illustrate the micropipette adhesion frequency method, we show kinetics measurement of intercellular adhesion molecule 1 (ICAM-1) functionalized on RBCs binding to integrin &alpha;L&beta;2 on neutrophils with dimeric E-selectin in the solution to activate &alpha;L&beta;2.

Adhesion Frequency Assay for In Situ Kinetics Analysis of Cross-Junctional Molecular Interactions at the Cell-Cell Interface

An adhesion frequency assay for measuring receptor-ligand interaction kinetics when both molecules are anchored on the surfaces of the interacting cells is described. This mechanically-based assay is exemplified using a micropipette-pressurized human red blood cell as adhesion sensor and integrin &alpha;L&beta;2 and intercellular adhesion molecule-1 as interacting receptors and ligands.

The micropipette adhesion assay was developed in 1998 to measure two-dimensional (2D) receptor-ligand binding kinetics1. The assay uses a human red blood ...

Bioengineering

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

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

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

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

Biochemistry

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

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

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

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

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 ...

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 ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

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.

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 ...

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 ...

Functionalization of Atomic Force Microscope Cantilevers with Single-T Cells or Single-Particle for Immunological Single-Cell Force Spectroscopy

Atomic force microscopy based single cell force spectroscopy (AFM-SCFS) is a powerful tool for studying biophysical properties of living cells. This technique allows for probing interaction strengths and dynamics on a live cell membrane, including those between cells, receptor and ligands, and alongside many other variations. It also works as a mechanism to deliver a physical or biochemical stimulus on single cells in a spatiotemporally controlled manner, thus allowing specific cell activation and subsequent cellular events to be monitored in real-time when combined with live-cell fluorescence imaging. The key step in those AFM-SCFS measurements is AFM-cantilever functionalization, or in other words, attaching a subject of interest to the cantilever. Here, we present methods to modify AFM cantilevers with a single T cell and a single polystyrene bead respectively for immunological studies. The former involves a biocompatible glue that couples single T cells to the tip of a flat cantilever in a solution, while the latter relies on an epoxy glue for single bead adhesion in the air environment. Two immunological applications associated with each cantilever modification are provided as well. The methods described here can be easily adapted to different cell types and solid particles.

We present a protocol to functionalize atomic force microscope (AFM) cantilevers with a single T cell and bead particle for immunological studies. Procedures to probe single-pair T cell-dendritic cell binding by AFM and to monitor the real-time cellular response of macrophages to a single solid particle by AFM with fluorescence imaging are shown.

Atomic force microscopy based single cell force spectroscopy (AFM-SCFS) is a powerful tool for studying biophysical properties of living cells. This ...

Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Summary

Explore More Videos

Quantitative and Qualitative Examination of Particle-particle Interactions Using Colloidal Probe Nanoscopy

Adhesion Frequency Assay for In Situ Kinetics Analysis of Cross-Junctional Molecular Interactions at the Cell-Cell Interface

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

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

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

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

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

Functionalization of Atomic Force Microscope Cantilevers with Single-T Cells or Single-Particle for Immunological Single-Cell Force Spectroscopy