Name: atomic force microscopy imaging and force spectroscopy of supported lipid bilayers
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Description: Watch this Scientific Journal Video about Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers at JoVE.com

This work was supported by the Max Planck Society, the German Cancer Research Center, the University of T&#252;bingen, and the Bundesministerium f&#252;r Bildung und Forschung (grant no. 0312040).
We thank Eduard Hermann for helping us automate the analysis of the force curve data and Dr. Jakob Suckale for careful reading of this manuscript.

1. Preparation of Supported Lipid Bilayers (SLB)11,12,21
<ol>
	<li>Preparation of Lipid Mixture and Multilamellar Vesicle Suspensions
	<ol>
		<li>Prepare the following buffers beforehand.
		<ol>
			<li>Prepare PBS buffer at concentrations of 2.7 mM KCl, 1.5 mM KH2PO4, 8 mM Na2HPO4, and 137 mM NaCl, pH 7.2.</li>
			<li>Prepare SLB (supported lipid bilayer) buffer at concentrations of 150 mM NaCl, 10 mM HEPES, pH 7.4.</li>
			<li>Prepare a solution of 1 M CaCl2.</li>
		</ol>
		</li>
		<li>Dissolve the following lipids in chloroform at a desired concentration: for example, di-oleoyl-phosphatidylcholine (DOPC) at 25 mg/ml, sphingomyelin (SM) at 25 mg/ml and cholesterol (Chol) at 10 mg/ml.</li>
		<li>Take 18.38 &#181;l of DOPC, 17.10 &#181;l of SM and 11.30 &#181;l of Chol to make a solution with 1 mg of total lipids in a 2:2:1 DOPC:SM:Chol molar ratio. Vary the volumes accordingly based on the concentrations of the lipids prepared in 1.1.2. However, maintain the molar ratio at 2:2:1 DOPC:SM:Chol, as changing the molar ratio will change the phase characteristics of the membrane25.</li>
		<li>Mix the solution completely by vortexing. Add a fluorescent lipidic dye at 0.05 mol % if one wishes to visualize the bilayer by a fluorescence microscope. 
		Note: The family of long-chain dialkylcarbocyanines, like DiD, DiI and DiO span a wide range of wavelengths and can be ideally used to label lipid membranes. Alternatively, fluorescently labeled lipids, like rhodamine-phosphatidylethanolamine or BODIPY-cholesterol can be used. In any case, the concentration of the dye should be optimized according to the microscopy setting, keeping in mind that high concentrations of the fluorophore bound to the lipid can change the lipid behavior19.</li>
		<li>Dry off the chloroform using N2 gas (or any available inert gas such as Ar and He).</li>
		<li>Further dry the lipid mixture under vacuum for at least 1 hr. 
		Note: If storing this lipid mixture, purge air with inert gas (N2, Ar or He) and store at -20 &#176;C. Aliquot excess lipids-in-chloroform into small brown vials. Dry them in a similar manner as the lipid mixture, displace air with inert gas (N2, Ar or He) and store at -20 &#176;C.</li>
		<li>Dissolve the dried lipids in PBS buffer to a concentration of 10 mg/ml.</li>
		<li>Vortex the mixture vigorously for about 20 sec to 1 min to get a cloudy suspension, containing multilamellar vesicles. Make sure that the there are no lipid films sticking to the sides of the vial.</li>
		<li>Distribute this suspension into 10 &#181;l aliquots (1 mg of lipid makes 10 aliquots).</li>
		<li>Store at -20 &#176;C until further use.</li>
	</ol>
	</li>
	<li>Preparation of Unilamellar Vesicles 
	Note: Various sizes of unilamellar vesicles can be prepared using the sonication method or the extrusion method. Sonication uses sound energy to agitate the mixture and separate the layers of the multilamellar suspension. This is indicated by clearing up of the hazy suspension. Extrusion forces the multilamellar vesicles to pass through a membrane filter with a definite pore size.
	<ol>
		<li>Sonication Method
		<ol>
			<li>Take the 10 &#181;l of multilamellar lipid suspensions and add 150 &#181;l of SLB buffer.</li>
			<li>Transfer the mixture into small (2 ml) capped glass vials and seal with Parafilm.</li>
			<li>Sonicate the mixture at room temperature in a bath sonicator for 10 min or until it becomes clear to yield small unilamellar vesicles (SUVs, below 50 nm in diameter).</li>
		</ol>
		</li>
		<li>Extrusion Method
		<ol>
			<li>Take the 10 &#181;l of multilamellar lipid suspensions and add 150 &#181;l of SLB buffer.</li>
			<li>Transfer the suspension into a cryogenic tube.</li>
			<li>Freeze the lipid suspension by submerging the cryogenic tube for a few seconds in liquid nitrogen.</li>
			<li>Thaw the lipid suspension by submerging the cryogenic tube in a water bath at room temperature or 37 &#176;C for a few minutes. Repeat the freeze-thaw steps at least 10 times.</li>
			<li>In order to extrude the lipid suspension, use a polycarbonate membrane with 200 nm pore size (other pore sizes are available like, 100 nm or 400 nm) and a commercial extrusion device26. 
			Note: Currently, there are two extruder devices commercially available: a manual extruder is available for small volumes (200 &#181;l to a few ml). In this case, the sample is extruded through the membrane by manually pushing the suspension back and forth between two syringes. Depending on the manufacturer, lipid aliquots would need to be combined to reach minimum volume of the set up. For bigger volumes (up to 50 ml) more sophisticated devices are used in which the suspension is forced through the polycarbonate membrane by using compressed gas. Set-up and extrusion conditions can change according to the model. Refer to the instructions from the manufacturer before use. This protocol refers to the manual extruder for small volumes.</li>
			<li>Equilibrate the extruder in water by passing water through the extruder. Immerse the membrane in water.</li>
			<li>Place the membrane between the two pistons, assemble the device and equilibrate in buffer, by passing the buffer through the extruder, from one syringe to the other.&#160;This step allows additionally testing the leakiness of the system. If it leaks, disassemble and reassemble it again changing the membrane.</li>
			<li>Take the lipid suspension using one syringe of the extruder (&#39;dirty side&#39;).</li>
			<li>Attach the syringe containing the lipid suspension on one chamber and the empty syringe on the opposing chamber.</li>
			<li>Push the syringe on one end. Pass the solution through the membrane and into the opposing syringe. This is the 1st pass.</li>
			<li>Pass it through the membrane for a total of 31 times such that the solution ends up on the opposing syringe (&#39;clean side&#39;). 
			Note: Check the membrane after extruding. If it was broken, then extrusion process was not successful and needs to be repeated.</li>
			<li>Empty the syringe into a small tube. Large unilamellar vesicles (LUVs) are stable for 1-2 days if kept at 4 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparation of Mica and Coverslips
	<ol>
		<li>Wash coverslips first in water and then in ethanol. Air dry.</li>
		<li>Take mica discs and peel off the outer layer using adhesive tape.</li>
		<li>Attach the mica disc onto the coverslip. Transparent optical glues are optimal so that one can check the bilayers with a fluorescence microscope.</li>
		<li>Attach plastic cylinders (2.5 cm outer diameter, 2 cm inner diameter, and 0.5 cm height) using optical glue/grease. Make sure that everything is sealed and that there are no leaks. Use grease when wanting to re-use the cylinders as they can be easily detached from the coverslip and washed. The dimensions of the plastic cylinders depend on the size of the AFM cantilever holder and should be adjusted accordingly.</li>
	</ol>
	</li>
	<li>Deposition of Unilamellar Vesicles on Solid Support
	<ol>
		<li>Pipet the whole liposome solution directly onto the mica. Add more SLB buffer to reach a volume of 300 &#181;l.</li>
		<li>Add 0.9 &#181;l of 1 M calcium chloride solution to reach a final concentration of 3 mM Ca2+.</li>
		<li>Incubate at 37 &#176;C for 2 min and then at 65 &#176;C for at least 10 min. Always cover the bilayers with buffer. Contact with air and air bubbles will destroy it. During incubation, add more buffer if necessary, especially if incubating at higher temperatures. 
		&#8203;Note: Incubation temperatures vary according to the lipid mixtures and phases required. Incubation time of at least 10 min is needed to form supported bilayers, but it could be longer, depending on the temperature and composition.</li>
		<li>Wash the bilayer with hot (65 &#176;C) SLB buffer 15-20 times to remove calcium and unfused vesicles. Take extra care during the washing steps to keep the bilayer under buffer and gently pipet the washing solution to avoid destroying the bilayer.</li>
		<li>Cool supported bilayers to room temperature before AFM measurements. This is needed to form the different phases of the membrane as well as minimize thermal drift in the instrument.</li>
	</ol>
	</li>
</ol>
2. Atomic Force Microscopy Measurements11,12
<ol>
	<li>Imaging 
	Note: Perform atomic force microscopy imaging in contact mode or tapping mode. Image supported bilayers in solution; do not allow solution to evaporate completely as this will destroy the bilayers. As one would be working with buffer, please observe safety precautions in making sure that any AFM part is protected from liquid spills.
	<ol>
		<li>Select a soft cantilever (below 0.2 N/m spring constant is advised) and mount it to the AFM. The choice of cantilever stiffness is more critical for force spectroscopy (and this will be discussed later).</li>
		<li>Mount the sample on the AFM stage and make sure that all vibration-isolation modules are turned on. Wait for at least 15 min to equilibrate and reduce thermal drift of the system. Depending on the AFM set-up, tuning and imaging specifications may vary. Consult the manufacturer&#8217;s handbook.</li>
		<li>Approach the sample with the AFM tip until contact.</li>
		<li>Since lipid bilayers are usually 4 nm in height, use the Z-range of the instrument that will give the highest z-resolution (for example 1.5 &#181;m).</li>
		<li>Select a region to image. To get an overview of the lipid bilayer, a 50 &#181;m x 50 &#181;m or 25 &#181;m x 25 &#181;m region will be a good starting point to image. If smaller features appear, one can image in smaller areas (10 &#181;m x 10 &#181;m, or 2 &#181;m x 2 &#181;m). The choice of the imaging area also depends on the working range of the piezoelectric scanner. Please consult the instrument manual for this.</li>
		<li>As bilayers are very fragile, adjust the set point of the AFM tip during imaging to keep the force at minimum and correct for thermal drift, while maintaining contact with the sample. In contact mode, minimize the set point. In tapping mode, maximize the set point.</li>
		<li>Process images by fitting each scan line to polynomial leveling functions. Perform line removal for scans that lose contact with the membrane using the proprietary software of the AFM manufacturing company.</li>
	</ol>
	</li>
	<li>Force Spectroscopy
	<ol>
		<li>Calibrate the cantilever following the instructions according to manufacturer&#8217;s protocol. Calibration allows the conversion of the electrical signal of the photodiodes to force (Figure 1B).
		<ol>
			<li>Calibrate the tip&#8217;s sensitivity to measure deflection as z-piezo movement (in units of length) instead of electrical signal in volts. 
			Note: AFM detects changes in the deflection of the cantilever using a laser reflected at the back of the cantilever. The laser reaches a photodiode that can spatially detect the position of the laser spot. A change in the deflection of the cantilever as it bends upon contact with the sample is detected by this photodiode as &#8220;vertical deflection&#8221;, and this is in the units of voltage. Since the cantilever is moved by the z-piezo, the vertical deflection is related to the movement in the z-piezo. The ratio between vertical deflection and z-movement is called sensitivity and converts voltage to distance.</li>
			<li>Calculate the cantilever spring constant using the thermal noise frequency method, usually incorporated in the AFM software. In this method, plot the amplitude of vibration from noise with respect to frequency of oscillation, creating a power spectral density plot. This plot will show a peak around the resonance frequency. The frequency where the amplitude is the greatest is the resonance frequency. Fit a Lorentzian function around the peak to automatically calculate the spring constant by the software. Some useful resources for the theory of the thermal noise method are given in references27-30.</li>
		</ol>
		</li>
		<li>After imaging an area of the bilayer, select a 5 &#181;m x 5 &#181;m region in the bilayer to perform force spectroscopy.</li>
		<li>Set up the AFM so that it takes force curves in a 16 x 16 grid of that region. This results in 256 force curves per region. The space between two neighboring points should be enough to avoid that the membrane area being probed by one point would coincide with the next point. This is dependent on the tip radius. A 100 nm distance between two points would be ideal. Divide the 5 &#181;m x 5 &#181;m region into 16 x 16 grid. Depending on the size of the phase, one can select a smaller or bigger region, and then adjust the grid size accordingly.</li>
		<li>Set z-piezo displacement to 400 nm. This determines the maximum range of movement of the z-piezo. It should be big enough to make sure that the cantilever has fully retracted away from the sample in between measurements.</li>
		<li>Set approach speed to 800 nm/sec and retract speed to 200 nm/sec. Use an approach speed between 400 to 6,000 nm/sec depending on the bilayer composition and lipid phase to be studied6,16.</li>
		<li>After setting up all the parameters, acquire force curves, by pressing the &#8220;run&#8221; button of the AFM software.</li>
		<li>Save force curves as force vs. height curves (a measure of z-piezo movement). This does not take into account the bending of the cantilever when in contact with the sample. During the data processing step, the AFM software allows correction for cantilever bending to get force vs. tip-sample separation curves (Figure 1C), which will give proper values for bilayer thickness. The correction values are usually incorporated in the calibration, which is saved with each force curve. Calculate tip-sample separation by subtracting the vertical deflection from the piezo movement at a given point. 
		Note: The peak in the approach force curve (the force value goes down even if the cantilever movement is still on the approach cycle) is the breakthrough force of the membrane, while the difference between the position of this peak and the position of the point where the force begins to rise up again provides the thickness of the membrane.</li>
		<li>Acquire at least 500 force curves for each condition to get good statistics. Plot a histogram of the values using programs like Origin or MatLab, and fit the histograms to a Gaussian distribution to get the peak and width of the distribution.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Binnig, G., Quate, C. F., Gerber, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+Force+Microscope.">Atomic Force Microscope.</a> Phys. Rev. Lett. 56, 930-933 (1986).</li><li>Hansma, P. K., Elings, V. B., Marti, O., Bracker, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+Tunneling+Microscopy+and+Atomic+Force+Microscopy:+Application+to+Biology+and+Technology.">Scanning Tunneling Microscopy and Atomic Force Microscopy: Application to Biology and Technology.</a> Science. 242, 209-216 (1988).</li><li>Gaczynska, M., Osmulski, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AFM+of+biological+complexes:+What+can+we+learn.">AFM of biological complexes: What can we learn.</a> Curr, Opin. Colloid In. 13, 351-367 (2008).</li><li>Goksu, E. I., Vanegas, J. M., Blanchette, C. D., Lin, W. -. C., Longo, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AFM+for+structure+and+dynamics+of+biomembranes.">AFM for structure and dynamics of biomembranes.</a> BBA-Biomembranes. 1788, 254-266 (2009).</li><li>Muller, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AFM:+A+Nanotool+in+Membrane+Biology.">AFM: A Nanotool in Membrane Biology.</a> Biochemistry-US. 47, 7986-7998 (2008).</li><li>Redondo-Morata, L., Giannotti, M. I., Sanz, F., Bar&#243;, A. M., Reifenberger, R. G., Sanz, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Atomic Force Microscopy in Liquid: Biological Applications. , (2012).</li><li>Castellana, E. T., Cremer, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solid+supported+lipid+bilayers:+From+biophysical+studies+to+sensor+design.">Solid supported lipid bilayers: From biophysical studies to sensor design.</a> Surf. Sci. Rep. 61, 429-444 (2006).</li><li>Frederix, P. L. T. M., Bosshart, P. D., Engel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+Force+Microscopy+of+Biological+Membranes.">Atomic Force Microscopy of Biological Membranes.</a> Biophys. J. 96, 329-338 (2009).</li><li>Mennicke, U., Salditt, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Solid-Supported+Lipid+Bilayers+by+Spin-Coating.">Preparation of Solid-Supported Lipid Bilayers by Spin-Coating.</a> Langmuir. 18, 8172-8177 (2002).</li><li>Raviakine, I., Brisson, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+Supported+Phospholipid+Bilayers+from+Unilamellar+Vesicles+Investigated+by+Atomic+Force+Microscopy.">Formation of Supported Phospholipid Bilayers from Unilamellar Vesicles Investigated by Atomic Force Microscopy.</a> Langmuir. 16, 1806-1815 (2000).</li><li>Chiantia, S., Ries, J., Kahya, N., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combined+AFM+and+Two-Focus+SFCS+Study+of+Raft-Exhibiting+Model+Membranes.+.">Combined AFM and Two-Focus SFCS Study of Raft-Exhibiting Model Membranes. .</a> ChemPhysChem. 7, 2409-2418 (2006).</li><li>Unsay, J., Cosentino, K., Subburaj, Y., Garcia-Saez, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiolipin+effects+on+membrane+structure+and+dynamics.">Cardiolipin effects on membrane structure and dynamics.</a> Langmuir. 29, 15878-15887 (2013).</li><li>Dom&#232;nech, &. #. 2. 1. 0. ;., Sanz, F., Montero, M. T., Hern&#225;ndez-Borrell, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermodynamic+and+structural+study+of+the+main+phospholipid+components+comprising+the+mitochondrial+inner+membrane.">Thermodynamic and structural study of the main phospholipid components comprising the mitochondrial inner membrane.</a> BBA-Biomembranes. 1758, 213-221 (2006).</li><li>Dom&#232;nech, &. #. 2. 1. 0. ;., Morros, A., Caba&#241;as, M. E., Teresa Montero, M., Hern&#225;ndez-Borrell, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Supported+planar+bilayers+from+hexagonal+phases.">Supported planar bilayers from hexagonal phases.</a> BBA-Biomembranes. 1768, 100-106 (2007).</li><li>Garcia-Saez, A. J., Chiantia, S., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+line+tension+on+the+lateral+organization+of+lipid+membranes.">Effect of line tension on the lateral organization of lipid membranes.</a> J Biol Chem. 282, 33537-33544 (2007).</li><li>Alessandrini, A., Seeger, H. M., Caramaschi, T., Facci, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+Force+Spectroscopy+on+Supported+Lipid+Bilayers:+Effect+of+Temperature+and+Sample+Preparation.">Dynamic Force Spectroscopy on Supported Lipid Bilayers: Effect of Temperature and Sample Preparation.</a> Biophys. J. 103, 38-47 (2012).</li><li>Butt, H. -. J., Franz, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rupture+of+molecular+thin+films+observed+in+atomic+force+microscopy+I.+Theory.">Rupture of molecular thin films observed in atomic force microscopy I. Theory.</a> Physical Review E. 66, 031601 (2002).</li><li>Garcia-Manyes, S., Oncins, G., Sanz, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+Temperature+on+the+Nanomechanics+of+Lipid+Bilayers+Studied+by+Force+Spectroscopy.">Effect of Temperature on the Nanomechanics of Lipid Bilayers Studied by Force Spectroscopy.</a> Biophys. J. 89, 4261-4274 (2005).</li><li>Chiantia, S., Kahya, N., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raft+domain+reorganization+driven+by+short-+and+long-chain+ceramide:+a+combined+AFM+and+FCS+study.">Raft domain reorganization driven by short- and long-chain ceramide: a combined AFM and FCS study.</a> Langmuir. 23, 7659-7665 (2007).</li><li>Canale, C., Jacono, M., Diaspro, A., Dante, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Force+spectroscopy+as+a+tool+to+investigate+the+properties+of+supported+lipid+membranes.">Force spectroscopy as a tool to investigate the properties of supported lipid membranes.</a> Microsc. Res. Techniq. 73, 965-972 (2010).</li><li>Garc&#237;a-S&#225;ez, A. J., Chiantia, S., Salgado, J., Schwille, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pore+Formation+by+a+Bax-Derived+Peptide:+Effect+on+the+Line+Tension+of+the+Membrane+Probed+by+AFM.">Pore Formation by a Bax-Derived Peptide: Effect on the Line Tension of the Membrane Probed by AFM.</a> Biophys. J. 93, 103-112 (2007).</li><li>Moreno Flores, S., Toca-Herrera, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+new+future+of+scanning+probe+microscopy:+Combining+atomic+force+microscopy+with+other+surface-sensitive+techniques,+optical+microscopy+and+fluorescence+techniques.">The new future of scanning probe microscopy: Combining atomic force microscopy with other surface-sensitive techniques, optical microscopy and fluorescence techniques.</a> Nanoscale. 1, 40-49 (2009).</li><li>Simons, K., Vaz, W. L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+Systems,+Lipid+Rafts,+and+Cell+Membranes1.">Model Systems, Lipid Rafts, and Cell Membranes1.</a> Annu. Rev. Bioph. Biom. 33, 269-295 (2004).</li><li>Pike, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rafts+defined:+a+report+on+the+Keystone+symposium+on+lipid+rafts+and+cell+function.">Rafts defined: a report on the Keystone symposium on lipid rafts and cell function.</a> The Journal of Lipid Research. 47, 1597-1598 (2006).</li><li>Kahya, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+Lipid+Mobility+of+Raft-exhibiting+Model+Membranes+by+Fluorescence+Correlation+Spectroscopy.">Probing Lipid Mobility of Raft-exhibiting Model Membranes by Fluorescence Correlation Spectroscopy.</a> J. Biol. Chem. 278, 28109-28115 (2003).</li><li>Akbarzadeh, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liposome:+classification,+preparation+and+applications.">Liposome: classification, preparation and applications.</a> Nanoscale Research Letters. 8, 102 (2013).</li><li>Butt, H. -. J., Jaschke, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calculation+of+thermal+noise+in+atomic+force+microscopy.">Calculation of thermal noise in atomic force microscopy.</a> Nanotechnology. 6, 1-7 (1995).</li><li>Chon, J. W. M., Mulvaney, P., Sader, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+validation+of+theoretical+models+for+the+frequency+response+of+atomic+force+microscope+cantilever+beams+immersed+in+fluids.">Experimental validation of theoretical models for the frequency response of atomic force microscope cantilever beams immersed in fluids.</a> Journal of Applied Physics. 87, 3973 (2000).</li><li>Sader, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Frequency+response+of+cantilever+beams+immersed+in+viscous+fluids+with+applications+to+the+atomic+force+microscope.">Frequency response of cantilever beams immersed in viscous fluids with applications to the atomic force microscope.</a> Journal of Applied Physics. 84, 64 (1998).</li><li>Sader, J. E., Pacifico, J., Green, C. P., Mulvaney, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=General+scaling+law+for+stiffness+measurement+of+small+bodies+with+applications+to+the+atomic+force+microscope.">General scaling law for stiffness measurement of small bodies with applications to the atomic force microscope.</a> Journal of Applied Physics. 97, 12490310 (2005).</li><li>Canale, C., Torre, B., Ricci, D., Braga, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recognizing+and+avoiding+artifacts+in+atomic+force+microscopy+imaging.">Recognizing and avoiding artifacts in atomic force microscopy imaging.</a> Methods Mol Biol. 736, 31-43 (2011).</li><li>Lee, M. -. T., Chen, F. -. Y., Huang, H. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Energetics+of+Pore+Formation+Induced+by+Membrane+Active+Peptides.">Energetics of Pore Formation Induced by Membrane Active Peptides.</a> Biochemistry-US. 43, 3590-3599 (2004).</li><li>Henriksen, J. R., Ipsen, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+membrane+elasticity+by+micro-pipette+aspiration.">Measurement of membrane elasticity by micro-pipette aspiration.</a> The European physical journal. E, Soft matter. 14, 149-167 (2004).</li><li>Nichols-Smith, S., Teh, S. -. Y., Kuhl, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermodynamic+and+mechanical+properties+of+model+mitochondrial+membranes.">Thermodynamic and mechanical properties of model mitochondrial membranes.</a> BBA-Biomembranes. 1663, 82-88 (2004).</li><li>Tian, A., Johnson, C., Wang, W., Baumgart, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Line+Tension+at+Fluid+Membrane+Domain+Boundaries+Measured+by+Micropipette+Aspiration.">Line Tension at Fluid Membrane Domain Boundaries Measured by Micropipette Aspiration.</a> Phys. Rev. Lett. 98, (2007).</li><li>Rigaud, J. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+proteins:+functional+and+structural+studies+using+reconstituted+proteoliposomes+and+2-D+crystals.">Membrane proteins: functional and structural studies using reconstituted proteoliposomes and 2-D crystals.</a> Brazilian Journal of Medical and Biological Research. 35, 753-766 (2002).</li></ol>

The authors declare that they have no competing financial interests.

SLBs composed of DOPC:SM:Chol (2:2:1) were formed on mica after vesicle adsorption and rupture induced by calcium chloride. This lipid composition separated into Ld and Lo phases. The Lo phase is enriched in sphingomyelin and cholesterol and is less fluid/more viscous (Figure 1A) than the Ld phase11. The separation of Lo from Ld phase manifests as circular structures elevated above the surrounding (Figure 1B, C)...

Atomic Force Microscopy (AFM) generates an image of a surface by scanning across an area of the sample using a cantilever with a very sharp tip1. The movement of the cantilever probes the surface topology of the sample. AFM has been widely applied to biological molecules &#8212; including proteins, DNA, and membranes, owing to its versatility in analyzing fixed samples in air or near-native state in liquid2-5.
Apart from its high-resolution imaging capability in the nanometer range, the AFM cantilever acts as a spring to probe interaction forces (adhesion and repulsion) and mechanical properties of the sample5,6. This is known as force spectroscopy. In this mode, the probe first approaches the sample and exerts a force on it, then is retracted until it loses contact with the sample (Figure 1A). The generated curves show force as a function of distance of the cantilever for both the approach and retraction. Several properties including the elastic modulus to measure the stiffness of a material, and adhesion forces can be derived.
Supported lipid bilayers are biological model membranes lying on top of a solid support &#8212; usually mica, borosilicate glass, fused silica, or oxidized silicon7. They are prepared using various techniques like vesicle deposition, Langmuir-Blodgett method and spin-coating8,9. AFM imaging has been used to follow the formation of these supported bilayers10, and probe different structures formed by membranes of different compositions11-15.
Performing force spectroscopy on supported bilayers results in a peak in the approach curve. This peak indicates the force needed to pierce the bilayer, and is called breakthrough force. The bilayer thickness can also be measured using the force curve6 . The typical breakthrough force of bilayers range between 1-50 nN6. These properties depend on lipid packing (liquid or gel phase) and structure (acyl chain length and degree of unsaturation) and altered by membrane-active agents16. The theory behind the rupture has been explained17 and other experimental parameters such as cantilever softness, tip radius and approach speed also affect the breakthrough force15,16,18. Force spectroscopy has been used to analyze properties of different lipid phases11,19, composition-dependent changes12,20, as well as effects of other biomolecules, like peptides, on the stability of the membrane21.
The flat orientation of supported bilayers is advantageous for combining AFM with other methods such as surface plasmon resonance22 and fluorescence microscopy11,19 to better characterize structure and properties of membranes.
This detailed video protocol is intended to prepare supported lipid bilayers using vesicle deposition and analyze them with AFM and force spectroscopy. While vesicles of various sizes may be used to prepare bilayers, this protocol focuses on small and large unilamellar vesicles. Supported bilayers that phase separate into liquid ordered (Lo) and liquid disordered (Ld) phases were characterized11,15. The membrane is composed of di-oleoyl-phosphatidylcholine (DOPC), sphingomyelin (SM), and cholesterol (Chol) at 2:2:1 ratio. This composition models the lipid rafts, which are proposed to behave as platforms important for protein trafficking and sorting, cell signaling and other cellular processes23,24.

<table><tbody><tr><td>1,2-dioleoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine</td><td>Avanti Polar Lipids, Inc.</td><td>850375P</td><td>Comes as lyophilized powder in sealed vials. Dissolve all powder in chloroform upon opening. Store extra as dried lipid films, under inert atmosphere, at -20&amp;nbsp;&amp;deg;C. Visit &lt;a href=&quot;http://www.avantilipids.com/index.php?option=com_content&amp;amp;view=article&amp;amp;id=1679&amp;amp;Itemid=398&quot;&gt;here&lt;/a&gt; for more information on storage and handling.</td></tr><tr><td>Sphingomyelin (Brain, Porcine)</td><td>Avanti Polar Lipids, Inc.</td><td>860062P</td><td>Comes as lyophilized powder in large sealed plastic containers. Dissolve a spatula point of powder powder in chloroform upon opening. Store extra as dried lipid films, under inert atmosphere, at -20&amp;nbsp;&amp;deg;C. Visit &lt;a href=&quot;http://www.avantilipids.com/index.php?option=com_content&amp;amp;view=article&amp;amp;id=1679&amp;amp;Itemid=398&quot;&gt;here&lt;/a&gt; for more information on storage and handling.</td></tr><tr><td>Cholesterol</td><td>Avanti Polar Lipids, Inc.</td><td>700000P</td><td>Comes as lyophilized powder in large sealed plastic containers. Dissolve a spatula point of powder powder in chloroform upon opening. Store extra as dried lipid films, under inert atmosphere, at -20&amp;nbsp;&amp;deg;C. Visit &lt;a href=&quot;http://www.avantilipids.com/index.php?option=com_content&amp;amp;view=article&amp;amp;id=1679&amp;amp;Itemid=398&quot;&gt;here&lt;/a&gt; for more information on storage and handling.</td></tr><tr><td>Sodium chloride (NaCl), 99.8%</td><td>Carl Roth GmbH + Co. KG</td><td>9265.1</td><td></td></tr><tr><td>Potassium chloride (KCl), 88%</td><td>Sigma</td><td>P9541</td><td></td></tr><tr><td>Sodium hydrogenphosphate (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;), &gt;99%</td><td>AppliChem GmbH</td><td>A1046</td><td></td></tr><tr><td>Potassium dihydrogenphosphate (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;), 99%</td><td>Carl Roth GmbH + Co. KG</td><td>3904.1</td><td></td></tr><tr><td>Calcium chloride dihydrate (CaCl&lt;sub&gt;2&lt;/sub&gt;), molecular biology grade</td><td>AppliChem GmbH</td><td>A4689</td><td></td></tr><tr><td>HEPES, molecular biology grade</td><td>AppliChem GmbH</td><td>A3724</td><td></td></tr><tr><td>Glass coverslip, 24&amp;nbsp;x&amp;nbsp;60&amp;nbsp;mm, 1&amp;nbsp;mm thickness</td><td>Duran Group</td><td>2355036</td><td></td></tr><tr><td>Punch and Die Set</td><td>Precision Brand Products, Inc</td><td>40105</td><td></td></tr><tr><td>Optical Adhesive</td><td>Norland Products, Inc.</td><td>NOA 88</td><td>Liquid adhesive that hardens when cured under long wavelength UV light.&amp;nbsp;</td></tr><tr><td colspan="4">[header]</td></tr><tr><td>Mica blocks</td><td>NSC Mica Exports Ltd.</td><td></td><td>These are mica pieces at least 1 sq. inches in area and thickness ranging from 0.006 inches to 0.016 inches. They are cut to a specific size by the company for shipping. Small mica discs can be punched from the mica blocks using the punch and die set.&amp;nbsp; Always handle mica with gloves or tweezers.</td></tr><tr><td>Laboratory Equipment Grease</td><td>Borer Chemie AG</td><td>Glisseal N</td><td></td></tr><tr><td>Liposome Extruder</td><td>Avestin</td><td>LiposoFast-Basic</td><td>As an alternative one can also look at offers from Northern Lipids, Inc.</td></tr><tr><td>Adhesive Tape</td><td>3M</td><td>Scotch(R) Magic (TM) Tape 810 (1-inch)</td><td></td></tr><tr><td>Bath Sonicator</td><td>Bandelin Sonorex Digitec</td><td>DT-31</td><td>No heating, Frequency: 35 kHz, Ultrasonic Peak Output: 160 W, HF Power: 40 W. (&lt;a href=&quot;http://www.bandelin.com/datenblaetter/dt/DT_31_H_1798d_DE_GB_FR_BANDELIN.pdf&quot;&gt;Data sheet&lt;/a&gt;)</td></tr><tr><td>Silicon Nitride AFM Cantilever&amp;nbsp;</td><td>Bruker AFM Probes</td><td>DNP-10</td><td>Each cantilever has four tips and their nominal tip radius is 20 nm (with possible maximum at 60&amp;nbsp;nm). Based on the specifications, we use tip D with resonance frequency of 18 kHz, and nominal spring constant of 0.06 N/m.</td></tr><tr><td>AFM</td><td>JPK</td><td>JPK Nanowizard II mounted on Zeiss Axiovert 200</td><td></td></tr></tbody></table>

atomic force microscopy imaging and force spectroscopy of supported lipid bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

Supported lipid bilayers composed of DOPC:SM:Chol (2:2:1) were imaged in AFM (Figure 2 A-C). Because of the lipid composition, SM/Chol-rich Lo and DOPC-rich Ld phases were observed. The height profile from the AFM imaging can provide important information on the membrane structure. By looking at the height profile, the bilayer thickness can be measured in presence of defects in the membrane (Figure 2B), or the difference in height between the Lo/Ld</...

Watch this Scientific Journal Video about Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers at JoVE.com

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

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Research

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Bioengineering

14.7K Views.  Interfaculty Institute for Biochemistry. We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Video: Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

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

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.

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 ...

Engineering

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 ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

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

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

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

Visualizing Solution Structure at Solid-Liquid Interfaces using Three-Dimensional Fast Force Mapping

Amongst the challenges for a variety of research fields are the visualization of solid-liquid interfaces and understanding how they are affected by the solution conditions such as ion concentrations, pH, ligands, and trace additives, as well as the underlying crystallography and chemistry. In this context, three-dimensional fast force mapping (3D FFM) has emerged as a promising tool for investigating solution structure at interfaces. This capability is based on atomic force microscopy (AFM) and allows the direct visualization of interfacial regions in three spatial dimensions with sub-nanometer resolution. Here we provide a detailed description of the experimental protocol for acquiring 3D FFM data. The main considerations for optimizing the operating parameters depending on the sample and application are discussed. Moreover, the basic methods for data processing and analysis are discussed, including the transformation of the measured instrument observables into tip-sample force maps that can be linked to the local solution structure. Finally, we shed light on some of the outstanding questions related to 3D FFM data interpretation and how this technique can become a central tool in the repertoire of surface science.

Here, we present a protocol for using three-dimensional fast force mapping - an atomic force microscopy technique - for visualizing solution structure at solid-liquid interfaces with the subnanometer resolution by mapping the tip-sample interactions within the interfacial region.

Amongst the challenges for a variety of research fields are the visualization of solid-liquid interfaces and understanding how they are affected by the ...

Chemistry

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 ...

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...