The authors acknowledge financial support from the National Science Foundation (award number IGERT-0903715), the University of Utah (Department of Chemical Engineering Seed Grant and the Graduate Research Fellowship Award), and Skolkovo Institute of Science and Technology (Skoltech Fellowship).&#160;

1. Isolation of EVs from a biofluid 
<ol>
	<li>Isolate EVs by one of the established methods, such as the differential ultracentrifugation8, precipitation, or size-exclusion chromatography9.</li>
	<li>Confirm the presence of expected surface and luminal biomarkers and the absence of biomarkers indicating cross-contamination of the preparation. Confirm the lipid bilayer morphology of the isolated particles by electron microscopy. 
	NOTE: When isolating the exosomes, the hydrodynamic size distribution measured by nanoparticle tracking analysis (NTA) or dynamic light scattering should be in the expected range. The details of EV and exosome isolation are beyond the scope of this protocol. The selected method will depend on experimental questions and the goal of the study10. The following steps provide a concrete illustration of the procedure to enrich the exosomes by precipitation from the growth medium of MCF-7 breast cancer cells using a commercially available precipitation kit (Table of Materials).</li>
	<li>Before cell culture expansion, store MCF-7 breast cancer cells in liquid nitrogen. Thaw cells to subculture.</li>
	<li>Following aseptic practices, perform cell plating on 150 mm plates. Use the growth medium composed of the Eagle&#8217;s minimum essential medium, 0.01 mg/mL human recombinant insulin, and 10% exosome-free fetal bovine serum.</li>
	<li>Aerate the culture by 95% air and 5% CO2 and incubate at 37 &#176;C.</li>
	<li>After the cells are settled (approximately 24 h after plating), change the media. Split the plate at 1:10 ratio and culture ten plates, each containing 20 mL of media.</li>
	<li>Harvest and pool media from 9 of these plates (180 mL) at ~70&#8722;80% confluence when cells are still in the growth phase.</li>
	<li>Divide the media into 60 mL and 120 mL, further split into 30 mL/tube, and centrifuge at 3,000 x g for 15 min.</li>
	<li>Transfer the supernatant from each tube to a new sterile 50 mL tube and perform the exosome isolation.</li>
	<li>Isolate exosomes by precipitation according to published protocols (see, for example, reference11) or follow the manufacturer&#8217;s instructions if a commercial isolation kit (Table of Materials) is used. As a first step in the latter case, centrifuge cell medium at 3,000 x g for 15 min. Withdraw supernatant and discard cells and cell debris.</li>
	<li>Add the precipitation solution (1:5 volume ratio) to the supernatant, mix, and refrigerate overnight.</li>
	<li>Centrifuge at 1,500 x g for 30 min at room temperature. Discard the supernatant after centrifugation.</li>
	<li>Spin the remaining exosome pellet for another 5 min at 1,500 x g. Without disturbing the pellet, remove the remaining precipitation solution by aspiration.</li>
	<li>Resuspend the pellet in 100&#8722;500 &#181;L of 1x phosphate-buffered saline (PBS) buffer and divide into multiple aliquots as needed for the downstream analysis.</li>
	<li>Immediately proceed to the surface immobilization of the isolated exosomes for AFM imaging. If necessary, freeze the aliquots at -80 &#176;C for later use while taking precautions to avoid damage to the sample during the freeze-thaw cycle.</li>
</ol>
2. Surface fixation of extracellular vesicles
<ol>
	<li>Use strong double-sided tape, epoxy, or an alternative adhesive to firmly attach a mica disk to an AFM/scanning tunneling microscope (STM) magnetic stainless-steel specimen disk.</li>
	<li>Cleave mica disc by using a sharp razor or utility knife, or by attaching an adhesive tape to the top surface and then pealing it off to remove a layer of material. 
	NOTE: Either method should reveal a virgin surface by removing a thin layer of mica previously exposed to the environment. After the procedure, the attachment of mica to the AFM/STM metal specimen disk must remain firm.</li>
	<li>At room temperature, treat the top surface of mica for 10 s with 100 &#181;L of 10 mM NiCl2 solution, which modifies the surface charge from negative to positive.</li>
	<li>Blot NiCl2 solution with a lint-free wipe or blotting paper. Wash the mica surface 3x with deionized (DI) water and dry it with a stream of dry nitrogen. 
	NOTE: It is a good practice to scan the modified surface with AFM to confirm it is free from contaminants.</li>
	<li>Place the AFM specimen disk with the attached surface-modified mica in a petri dish.</li>
	<li>Dilute the exosome sample from step 1.14 with 1x PBS to obtain a concentration between 4.0 x 109 and 4.0 x 1010 particles per mL of solution. Validate the diluted particle concentration using NTA.</li>
	<li>Form a sessile drop on the surface of mica by emptying 100 &#956;L of the diluted exosome solution from a pipette.</li>
	<li>Place lid on the petri dish and seal it with a paraffin film to reduce sample evaporation. Incubate the sample for 12&#8722;18 h at 4 &#176;C. 
	NOTE: The surface density of the immobilized exosomes will increase with the incubation time and the concentration of EVs in the liquid. Longer incubation time may be necessary if exosomes are present in the sample at lower concentrations.</li>
	<li>After incubation, aspirate 80&#8722;90% of the sample without disturbing the surface. At this point, the exosomes will be electrostatically immobilized on the mica substrate.</li>
	<li>Before imaging hydrated EVs, rinse the surface with 1x PBS. Repeat 3x. Take care to keep the sample hydrated throughout the rinsing process.</li>
	<li>After washing the mica surface with 1x PBS, remove 80%&#8722;90% of liquid, and pipette ~40 &#956;L of fresh 1x PBS to cover the sample.</li>
	<li>When imaging the desiccated EVs, rinse the substrate with DI water. Repeat 3x. 
	NOTE: Rinsing with DI water will prevent the formation of salt crystals and the deposition of solutes on the surface as the substrate dries.</li>
	<li>Before imaging desiccated EVs, aspirate as much liquid as possible without touching the surface and dry the rest with a stream of dry nitrogen.</li>
</ol>
3. AFM imaging 
<ol>
	<li>To image the desiccated EVs, select a cantilever designed for scanning in the air in tapping and non-contact imaging modes and mount it onto the probe holder. 
	NOTE: The characteristics of an example cantilever listed in Table of Materials (123 &#956;m length, 40 &#956;m width, 7 nm tip radius, and 37 N/m spring constant) may be used as a guide when selecting a probe compatible with the available AFM instrumentation.
	<ol>
		<li>Place the preparation from step 2.13 on the AFM stage. The magnetic stainless-steel specimen disk will immobilize the sample on the stage. Allow time for the preparation and the stage to equilibrate thermally.</li>
		<li>Use the tapping mode to scan a sufficiently large area of the mica&#8217;s surface. For example, choose an area of 5 x 5 &#181;m, rastered in 512 lines at a scan rate of ~1 Hz. Acquire both the height and phase images as they provide complementary information on the topography and the surface properties of the sample. 
		NOTE: The scan time will increase with the imaged area and the number of lines selected to form the image but decrease with the scan rate defined as the number of lines scanned per second. Fast scan rates may impact the image quality. Therefore, the speed of rastering should judicially balance the tradeoff between the acquisition time and the image quality.</li>
	</ol>
	</li>
	<li>To image hydrated vesicles, select a cantilever appropriate for scanning soft, hydrated samples and mount the cantilever onto a probe holder designed for scanning in liquids. 
	NOTE: When selecting a probe compatible with the available AFM instrumentation, the specifications of the probe listed in Table of Materials (triangular cantilever with 175 &#181;m nominal length, 22 &#181;m width, 20 nm tip radius, 0.07 N/m spring constant, and optimized for imaging with the drive frequency in the range between 4 to 8&#160;kHz) may be used as a guide.
	<ol>
		<li>Wet the tip of the cantilever with 1x PBS to reduce the likelihood of introducing air bubbles into the liquid during scanning.</li>
		<li>Place the preparation from step 2.11 on the AFM stage. The magnetic stainless-steel specimen disk will immobilize the attached mica containing immobilized EVs on its surface.</li>
		<li>Allow time for the preparation and the AFM stage to equilibrate thermally.</li>
		<li>Image the hydrated mica surface in the tapping mode. Acquire both the height and phase images. 
		NOTE: The imaging quality is influenced by the instrumentation, selected probe, and scan parameters. When optimizing the scanning conditions, the following choices may be used as a starting point: 5 x 5 &#181;m area scanned in 512 lines with ~0.8-1.0 Hz scan rate and&#160;drive frequency between 4 to 8 kHz.</li>
	</ol>
	</li>
</ol>
4. Image analysis
NOTE: The following data processing and analysis steps are applied to the acquired height images. A similar procedure may be adapted to analyze the phase data. The description below is specific to Gwyddion12, a free and open source software available under GNU General Public License. Similar capabilities are available in alternative software tools.
<ol>
	<li>Go to Data Process, SPM modes, Tip and choose Model Tip (Figure 2). Select the geometry and the dimensions of the tip used to scan the sample and click OK.</li>
	<li>Correct the tip erosion artifacts by performing the surface reconstruction. Open the image. From the menu, select Data Process, SPM modes, Tip, then choose Surface Reconstruction and click OK (Figure 3).</li>
	<li>Align the imaging plane to match the laboratory XY plane by removing the tilt in the substrate from the scan data. To accomplish this task, select Data Process, Level and choose Plane Level (Figure 4).</li>
	<li>Align rows of the image by selecting Data Process, Correct Data and then choose Align Rows. Several alignment options are available (Figure 5). For example, Median is an algorithm that finds an average height of each scan line and subtracts it from the data.</li>
	<li>Go to Data Process, Correct Data and choose Remove Scars (Figure 6), which removes common scanning errors known as scars.</li>
	<li>Align the mica surface at the zero height, Z = 0, by selecting Flatten Base in Level drop-down menu accessible from Data Process (Figure 7).</li>
	<li>Identify EVs on the scanned surface by using Mark by Threshold in Grains drop-down menu (Figure 8A). This algorithm identifies surface-immobilized exosomes as particles protruding from the zero-surface substrate by the height above the user-selected threshold. Select a threshold in the range between 1 and 3&#160;nm, which will eliminate most of the background interference. Smaller thresholds are used with cleaner background. 
	NOTE: The threshold in Figure 8A is 1.767 nm. The outcome of the MCF-7 exosome identification with this thresholding is shown in Figure 8B. Gwyddion offers several alternatives to thresholding as the algorithm to automatically identify vesicles in the image, including automated thresholding (Otsu&#39;s method), edge detection, and the watershed algorithm. 
	NOTE:&#160;Particle agglomerates, if present in the AFM image, may be masked and excluded from the analysis.</li>
	<li>Perform geometric and dimensional characterization of the identified EVs using the available Distributions algorithms accessible from Grains menu. 
	NOTE: Gwyddion provides tools to assess the distribution of scalar-valued, areal, volumetric, and other properties of immobilized EVs in a hydrated or dessicated state. An example of a scalar-values property is shown in Figure 9, which gives the distribution of maximum heights within the footprint of each identified exosome.</li>
	<li>Export the AFM data from Gwyddion for specialized analysis by other computational tools and custom computer programs.</li>
</ol>

<ol>
	<li>Chernyshev, V. S., 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=Size+and+shape+characterization+of+hydrated+and+desiccated+exosomes.">Size and shape characterization of hydrated and desiccated exosomes.</a> Analytical and Bioanalytical Chemistry. 407, 3285-3301 (2015).</li><li>Ramirez, M. I., 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=Technical+challenges+of+working+with+extracellular+vesicles.">Technical challenges of working with extracellular vesicles.</a> Nanoscale. 10, 881-906 (2018).</li><li>Skliar, M., 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=Membrane+proteins+significantly+restrict+exosome+mobility.">Membrane proteins significantly restrict exosome mobility.</a> Biochemical and Biophysical Research Communications. 501, 1055-1059 (2018).</li><li>Parisse, P., 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=Atomic+force+microscopy+analysis+of+extracellular+vesicles.">Atomic force microscopy analysis of extracellular vesicles.</a> European Biophysics Journal. 46, 813-820 (2017).</li><li>Ito, K., 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=Host+Cell+Prediction+of+Exosomes+Using+Morphological+Features+on+Solid+Surfaces+Analyzed+by+Machine+Learning.">Host Cell Prediction of Exosomes Using Morphological Features on Solid Surfaces Analyzed by Machine Learning.</a> Journal of Physical Chemistry B. 122, 6224-6235 (2018).</li><li>Sharma, S., LeClaire, M., Gimzewski, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ascent+of+atomic+force+microscopy+as+a+nanoanalytical+tool+for+exosomes+and+other+extracellular+vesicles.">Ascent of atomic force microscopy as a nanoanalytical tool for exosomes and other extracellular vesicles.</a> Nanotechnology. 29, 132001 (2018).</li><li>Meyer, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immobilisation+of+living+bacteria+for+AFM+imaging+under+physiological+conditions.">Immobilisation of living bacteria for AFM imaging under physiological conditions.</a> Ultramicroscopy. 110, 1349-1357 (2010).</li><li>Th&#233;ry, C., Amigorena, S., Raposo, G., Clayton, 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=Isolation+and+characterization+of+exosomes+from+cell+culture+supernatants+and+biological+fluids.">Isolation and characterization of exosomes from cell culture supernatants and biological fluids.</a> Current protocols in cell biology. Chapter 3 (Unit 3.22), (2006).</li><li>Taylor, D. D., Zacharias, W., Gercel-Taylor, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exosome+isolation+for+proteomic+analyses+and+RNA+profiling.">Exosome isolation for proteomic analyses and RNA profiling.</a> Methods in Molecular Biology. 728, 235-246 (2011).</li><li>Th&#233;ry, C., 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=Minimal+information+for+studies+of+extracellular+vesicles+2018+(MISEV2018):+a+position+statement+of+the+International+Society+for+Extracellular+Vesicles+and+update+of+the+MISEV2014+guidelines.">Minimal information for studies of extracellular vesicles 2018 (MISEV2018): a position statement of the International Society for Extracellular Vesicles and update of the MISEV2014 guidelines.</a> Journal of Extracellular Vesicles. 8, 1535750 (2019).</li><li>Weng, Y., 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=Effective+isolation+of+exosomes+with+polyethylene+glycol+from+cell+culture+supernatant+for+in-depth+proteome+profiling.">Effective isolation of exosomes with polyethylene glycol from cell culture supernatant for in-depth proteome profiling.</a> The Analyst. 141, 4640-4646 (2016).</li><li>Ne&#269;as, D., Klapetek, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gwyddion:+an+open-source+software+for+SPM+data+analysis.">Gwyddion: an open-source software for SPM data analysis.</a> Open Physics. 10, 181-188 (2012).</li><li>Hsueh, C., Chen, H., Gimzewski, J. K., Reed, J., Abdel-Fattah, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localized+nanoscopic+surface+measurements+of+nickel-modified+Mica+for+single-molecule+DNA+sequence+sampling.">Localized nanoscopic surface measurements of nickel-modified Mica for single-molecule DNA sequence sampling.</a> ACS Applied Materials and Interfaces. 2, 3249-3256 (2010).</li><li>Conde-Vancells, J., 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=Characterization+and+comprehensive+proteome+profiling+of+exosomes+secreted+by+hepatocytes.">Characterization and comprehensive proteome profiling of exosomes secreted by hepatocytes.</a> Journal of Proteome Research. 7, 5157-5166 (2008).</li><li>Zhou, Y., 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=Exosomes+released+by+human+umbilical+cord+mesenchymal+stem+cells+protect+against+cisplatin-induced+renal+oxidative+stress+and+apoptosis+in+vivo+and+in+vitro.">Exosomes released by human umbilical cord mesenchymal stem cells protect against cisplatin-induced renal oxidative stress and apoptosis in vivo and in vitro.</a> Stem Cell Research &amp; Therapy. 4, 34 (2013).</li><li>Coleman, B. M., Hanssen, E., Lawson, V. A., Hill, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prion-infected+cells+regulate+the+release+of+exosomes+with+distinct+ultrastructural+features.">Prion-infected cells regulate the release of exosomes with distinct ultrastructural features.</a> FASEB Journal. 26, 4160-4173 (2012).</li><li>Briegel, 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=Universal+architecture+of+bacterial+chemoreceptor+arrays.">Universal architecture of bacterial chemoreceptor arrays.</a> Proceedings of the National Academy of Sciences. 106, 17181-17186 (2009).</li><li>Akagi, T., 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=On-Chip+Immunoelectrophoresis+of+Extracellular+Vesicles+Released+from+Human+Breast+Cancer+Cells.">On-Chip Immunoelectrophoresis of Extracellular Vesicles Released from Human Breast Cancer Cells.</a> PLOS ONE. 10, e0123603 (2015).</li><li>Sharma, S., 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=Structural-mechanical+characterization+of+nanoparticle+exosomes+in+human+saliva,+using+correlative+AFM,+FESEM,+and+force+spectroscopy.">Structural-mechanical characterization of nanoparticle exosomes in human saliva, using correlative AFM, FESEM, and force spectroscopy.</a> ACS Nano. 4, 1921-1926 (2010).</li><li>Radeghieri, 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=Cultured+human+amniocytes+express+hTERT,+which+is+distributed+between+nucleus+and+cytoplasm+and+is+secreted+in+extracellular+vesicles.">Cultured human amniocytes express hTERT, which is distributed between nucleus and cytoplasm and is secreted in extracellular vesicles.</a> Biochemical and Biophysical Research Communications. 483, 706-711 (2017).</li><li>Woo, J., Sharma, S., Gimzewski, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Role+of+Isolation+Methods+on+a+Nanoscale+Surface+Structure+and+Its+Effect+on+the+Size+of+Exosomes.">The Role of Isolation Methods on a Nanoscale Surface Structure and Its Effect on the Size of Exosomes.</a> Journal of Circulating Biomarkers. 5, 11 (2016).</li><li>Deegan, R. D., 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=Capillary+flow+as+the+cause+of+ring+stains+from+dried+liquid+drops.">Capillary flow as the cause of ring stains from dried liquid drops.</a> Nature. 389, 827-829 (1997).</li><li>Johnson, C. A., Lenhoff, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adsorption+of+Charged+Latex+Particles+on+Mica+Studied+by+Atomic+Force+Microscopy.">Adsorption of Charged Latex Particles on Mica Studied by Atomic Force Microscopy.</a> Journal of Colloid and Interface Science. 179, 587-599 (1996).</li><li>Pastr&#233;, D., 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=Adsorption+of+DNA+to+Mica+Mediated+by+Divalent+Counterions:+A+Theoretical+and+Experimental+Study.">Adsorption of DNA to Mica Mediated by Divalent Counterions: A Theoretical and Experimental Study.</a> Biophysical Journal. 85, 2507-2518 (2003).</li><li>van der Pol, E., B&#246;ing, A. N., Harrison, P., Sturk, A., Nieuwland, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Classification,+functions,+and+clinical+relevance+of+extracellular+vesicles.">Classification, functions, and clinical relevance of extracellular vesicles.</a> Pharmacological Reviews. 64, 676-705 (2012).</li></ol>

The authors have nothing to disclose.&#160;

The immobilization of EVs from a biological fluid, surface scanning, and image analysis are the essential steps of the developed protocol for the AFM characterization of EVs in liquid. The number of vesicles amenable to AFM imaging scales with the imaged surface area and the surface concentration of the vesicles immobilized on the substrate. Given a negative zeta potential of EVs and exosomes18, we advocate electrostatic fixation of EVs from liquid samples to the AFM substrate. The immobilization is effective when the surface is positively charged. Prior to EV immobilization, the positive surface charge may need to be imparted to the substrate, as in the case of mica &#8213; a layered silicate mineral with general formula KAl2(AlSi3O10)(OH)2. Freshly cleaved mica&#8217;s surface is close to perfectly flat, which is ideal for imaging nanoparticles by the AFM, but its surface charge is negative and, thus, must be modified. The protocol describes the procedure to impart a positive surface change to the AFM substrate. The representative results show marked improvement in EV fixation from a biofluid to the modified mica substrate. 
&#160; 
When imaging hydrated vesicles, it is important to minimize sample evaporation which causes the surface deposition artifacts and convective flows and increases the liquid concentration of the vesicles with time, leading to higher surface concentration of immobilized EVs than expected, especially during prolonged incubations. Probe holders explicitly designed for liquid samples eliminate or slow evaporation and should be used to image hydrated EVs. Nonspecific bindings to the scanning probe are reduced in the presence of ionic species. Therefore, when imaging hydrated EVs, it is preferable to cover the substrate with a buffered medium, such as PBS, instead of DI water.
Importance of surface immobilization&#160; 
The consistent and predictable immobilization of EVs on the modified substrate removes the primary source of variability in the AFM results. All downstream steps, from scanning to data analysis, are more easily controlled by the selection of instrumentation, probes, scanning parameters, and the data analysis sequence and algorithms. The user should be aware of upstream variability in biological samples and EV isolation protocols, which are important issues beyond the scope of this work.
We recommend performing surface immobilization of EV on the modified mica&#8217;s surface from liquid samples even when the goal is to characterize desiccated vesicles in the air &#8212; the need is less evident since vesicles will unavoidably deposit on any substrate as the liquid evaporates. In fact, the AFM results for desiccated EV obtained without modifying mica&#8217;s surface charges, which is a prerequisite step for electrostatic immobilization of EVs from a liquid, have been reported in the past19,20,21. When EVs are not fixed to the surface from the liquid sample, however, their passive deposition by evaporation will produce artifacts collectively known as a coffee ring effect22. Two such artifacts, occurring as a drying liquid recedes, are illustrated in the SEM image (Figure 13A) of serum exosomes deposited by evaporation on a negatively charged glass surface. Significant variations in the surface concentration of precipitated vesicles are immediately apparent. The second artifact, quantified in Figure 13B, is the considerable variability in vesicle sizes in different areas within the perimeter of the dried sample. Given these artifacts, the AFM characterization of passively deposited vesicles from a drying liquid may produce biased or inconsistent results unless the entire surface area initially occupied by a now-dried liquid sample is scanned.&#160;
Two additional issues should be considered when imaging the desiccated samples obtained without firm immobilization of vesicles on the substrate. Recall that our protocol instructs users to thoroughly wash the surface with DI water after the vesicles are immobilized from a liquid sample. This step intends to prevent ionic and other non-vesicular solutes from forming surface deposits during the evaporation of complex biofluids with considerable osmolarity. If EVs are not fixed, thorough washing will detach a large number of vesicles from the surface, potentially biasing the results and leaving too few particles for analysis. Another common difficulty, reduced by immobilizing EVs on the modified mica surface before the AFM imaging, is the adhesion of particles to the probe23 and the misleading artifacts caused by this phenomenon. 
&#160; 
Control of surface density of immobilized EVs 
The two easily controllable factors identified in the protocol allow the user to customize the surface concentration of the immobilized EVs on the modified mica substrate: the concentration of the vesicles in the liquid sample and the time the sample is incubated on the substrate. A high density of immobilized vesicles, achieved with longer incubation times and higher concentration of EVs in the liquid, increases the number of vesicles analyzed during the scanning and the statistical power of conclusions arrived by the analysis of the AFM data. At the same time, an excessively dense surface concentration, as in the case shown in Figure 10C where particles tightly cover the entire surface with no intervening areas of the substrate, complicates the image analysis and the interpretation of results and may lead to scanning artifacts caused by the interaction between closely spaced particles.
Influence of electrostatic screening and hydrodynamic mobility of EVs 
The transparent control over the surface concentration of immobilized EVs as a function of factors influencing it allows a user to customize experimental conditions to meet the specific needs of a study. When performing customization, it is important to recognize that the electrostatic surface immobilization is a transport-limited process influenced by the ionic strength of the biofluid.&#160;
The concentration of ionic and positively charged species inversely impacts the Debye length over which the surface and vesicle charges are screened. Beyond this length, the electrostatic forces are negligible. The boundary layer of the substrate&#8217;s electrostatic attraction will be much smaller in ionically-rich PBS than in DI water. This difference implies that, after a short incubation corresponding to the time needed to deplete the layer of liquid where the electrostatic attractions are felt, a surface density of EVs immobilized from the suspension in DI water will be higher than from PBS suspension, assuming the concentration of EVs is the same in both liquids. Put differently, more vesicles must be immobilized to deplete a thicker attraction layer in DI water than in PBS under otherwise identical conditions.&#160;
After the vesicles are depleted from the boundary layer, the immobilization becomes an entirely transport-limited process. In this regime, the rate of the deposition will not depend on the suspending medium (e.g., DI water or PBS) as long as the viscosity is the same and the transport is entirely diffusive. However, the transport of vesicles into the attraction boundary layer may not be entirely diffusive. For instance, if the sample in a sessile drop on the AFM substrate partially evaporates during the incubation, the fluid inside the drop will be subjected to the evaporation-driven flow, and the transport of the vesicles towards the substrate will have, both, diffusive and convective contributions. When the evaporation is not adequately controlled, the contribution of a convective transport will be considerable, and the rate of immobilization will be higher than expected. The impact of the convective transport will change with the thickness of the attraction layer, which itself depends on the ionic content of the liquid. Furthermore, the evaporation will enhance the vesicle immobilization on the substrate by concentrating EVs in the solution. At higher EV concentrations, the concentration gradient between the attraction layer and the adjacent liquid will increase, creating a larger thermodynamic driving force to the migration of vesicles towards the substrate.
Immobilized vesicles may represent a liquid sample with a bias. For the case when the rate of immobilization is limited by diffusion, vesicles with smaller hydrodynamic sizes, determined by the combination of the vesicle size and the thickness of the coronal layer surrounding it (Figure 1), are more likely to enter the attraction layer because of their higher mobility. Consequently, after the initial depletion period, the hydrodynamically small EVs will be overrepresented on the substrate compared to their contribution to the EV population in the liquid sample. Note that a smaller hydrodynamic size does not automatically point at EVs with smaller vesicle sizes because of the heterogeneity in the thickness of the coronal layer3. The biased representation is avoided with long incubations that deplete the entire population of EVs in the liquid by its immobilization on the substrate. When a user aims at immobilizing all EVs from the biofluid, to avoid excessively dense coverage of the surface with the immobilized vesicles, it may be necessary to reduce the EV concentration in the liquid below the range suggested in the protocol.&#160;
Deformation of EVs on the substrate 
Extracellular vesicles in their native hydrated state and after desiccation can be characterized by the AFM, as described in the protocol. The electrostatic forces24 that immobilize EVs on the mica surface also distort their shape from globular geometry in which they exist in the solution. The impact of the desiccation on the size and morphology of immobilized EVs may be analyzed by rescanning the same surface area before and after the sample is allowed to dry.&#160;
It is instructive to examine the impact of the sample preparation on the shape of the desiccated EVs. The electrostatically immobilized EVs maintain the highly oblate geometry after drying but are further flattened by the desiccation. The height above the surface of the desiccated vesicles becomes smaller than in Figure 12A, while their footprint area increases (data not shown). On the other hand, when vesicles are deposited passively during the liquid evaporation and without prior immobilization on the surface, they tend to attain a cup-shape geometry upon desiccation, as has long been observed in the SEM images and, more recently, in AFM scans. This cupped shape is now recognized as a sample preparation artifact25 caused by non-uniformity in capillary forces during surface desiccation, as mechanistically explained in Figure 141. 
&#160; 
Image analysis and interpretation of AFM data 
The responses to electrostatic and capillary forces acting to distort the shape of EVs provide valuable information on structural and compositional properties of EVs. For example, a multidimensional set of biophysical characteristics, such as the deformed size and shape extracted from the AFM data, were recently used to demonstrate the feasibility to differentiate between exosomes secreted by different host cells5. The distortions can also be taken into account and compensated. For example, we showed how to use the AFM data to characterize the globular size of vesicles in the solution by estimating the diameters of spheres that encapsulate the same volume as immobilized exososmes3.&#160;

Extracellular vesicles (EVs) are present in all body fluids, including blood, urine, saliva, milk, and the amniotic fluid. Exosomes form a district class of EVs differentiated from other EVs by endosomal biogenesis, the markers of the endosomal pathway, and the smallest size among all EVs. The size of exosomes is often reported with substantial variability between studies. The sizing results were found to be method dependent, reflecting the difference in physical principles employed by different analytical techniques to estimate EV sizes1,2. For example, the nanoparticle tracking analysis (NTA) &#8213; the most widely used size characterization technique &#8213; estimates the size of EVs as their hydrodynamic diameters, which characterize the resistance to the Brownian mobility of EVs in the solution. A larger hydrodynamic diameter of a vesicle implies its lower mobility in liquid. The coronal layer around vesicles, consisting of surface proteins and other molecules anchored or adsorbed to the membrane surface, substantially impedes the mobility and increases the hydrodynamic size of EVs. In relative terms, this increase is particularly large for the exosomes3, as illustrated in Figure 1.&#160;
The cryogenic transmission electron microscopy (cryo-TEM) imaging is a definitive technique in characterizing vesicle sizes and morphology in their hydrated states. However, the high cost of the instrumentation and the specialized expertise needed to use it correctly motivate the exploration of alternative techniques that can image hydrated EVs. A relatively small number of EVs observed or characterized in the acquired cryo-TEM images is another notable disadvantage of this technique.
Atomic force microscopy (AFM) visualizes the three-dimensional topography of hydrated or desiccated EVs4,5,6 by scanning a probe across the substrate to raster the image of the particles on the surface. The essential steps of the protocol to characterize EVs by AFM are outlined in this study. Before imaging the vesicles in liquid, they must be immobilized on a substrate by either tethering to a functionalized surface, trapping in a filter, or by electrostatic attraction7. The electrostatic fixation on a positively charged substrate is a particularly convenient option for immobilization of exosomes known to have a negative zeta potential. However, the same electrostatic forces that immobilize the extracellular vesicles on the surface also distort their shape, which makes post-imaging data analysis essential. We elaborate this point by describing the algorithm that estimates the size of the globular vesicles in the solution based on the AFM data on the distorted shape of the exosomes immobilized on the surface.
In the developed protocol, the procedure for the robust electrostatic immobilization of vesicles is presented and followed by the steps needed to perform atomic force imaging in the hydrated or desiccated states. The factors that influence the surface concentration of the immobilized vesicles are identified. The guidance is given on how to perform the electrostatic immobilization for samples with different concentrations of EVs in the solution. The selection of experimental conditions permitting the estimation of empirical probability distributions of different biophysical properties based on a sufficiently large number of immobilized vesicles is discussed. Examples of post-imaging analysis of the AFM data are given. Specifically, an algorithm is described for determining the size of vesicles in the solution based on the AFM characterization of immobilized EVs. The representative results show the consistency of the vesicle sizing by AFM with the results of cryo-TEM imaging.&#160;

<table border="0" cellpadding="0" cellspacing="0" style="width:581px;" width="581">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:223px;">AFM/STM Controller&#160;</td>
			<td style="width:92px;">Bruker</td>
			<td style="width:93px;">Multimode Nanoscope V</td>
			<td style="width:173px;">This AFM controller supports imaging of biological samples in liquid and air.&#160;</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">AFM/STM metal specimen discs (10 mm)</td>
			<td>TedPella</td>
			<td align="right">16207</td>
			<td>Metal specimen disc on which a mica disk is attached by a double-sided tape or other means.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">AFM/STM Mica discs (10 mm)</td>
			<td>TedPella</td>
			<td align="right">50</td>
			<td>Highest quality grade V1 mica, 0.21mm (0.0085&#8221;) thick. Interleaved, in packages of 10. Can be mounted on AFM/STM discs. Available in four diameters</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">AFM probe for imaging in the air</td>
			<td>Bruker</td>
			<td>TESP-V2</td>
			<td>High quality etched silicon probes for tapping mode and non-contact mode for scanning in the air.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">AFM probe for soft sample imaging in liquid</td>
			<td>Bruker</td>
			<td>MLCT</td>
			<td>Soft silicon nitride cantilevers with silicon nitride tips, which are well-suited for liquid operation.&#160; The range in force constants enables users to image extremely soft samples in contact mode as well as high load vs distance spectroscopy.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Double sided tape</td>
			<td>Spectrum</td>
			<td>360-77705</td>
			<td>Used to fix the mica disk on the metal specimen disc.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">ExoQuick-TC</td>
			<td>System Biosciences</td>
			<td>EXOTC50A-1</td>
			<td>ExoQuick-TC is a proprietary polymer-based kit designed for exosome isolation from tissue culture media.&#160;</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Glass probe AFM holder for imaging in liquid</td>
			<td>Bruker</td>
			<td>&#160;MTFML-V2</td>
			<td>This glass probe holder is designed for scanning in fluid with the MultiMode AFM.&#160; The holder can be used in peak force tapping mode, contact mode, tapping mode, and force modulation.&#160; The probe is acoustically driven by a separate piezo oscillator for larger amplitude modulation.&#160; The holder is supplied with two ports, required fittings, and accessories kit for adding and removing fluids.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Gwyddion</td>
			<td>Czech Metrology Institute.</td>
			<td>Version 2.52</td>
			<td>Open Source software for visualization and analysis of data fields obtained by scanning probe microscopy techniques.</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">Lint-free blotting paper</td>
			<td>GE Healthcare Whatman&#160;</td>
			<td>Grade GB003 Blotting Paper</td>
			<td>Use this blotting paper to remove NiCl2 after the modification of the mica&#39;s substrate.&#160;&#160;</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Lint-free cleanroom wipes</td>
			<td>Texwipe</td>
			<td>AlphaWipe TX1004</td>
			<td>Use these polyester wipes for surface cleaning.&#160;</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">Nickel(II) chloride (NiCl2)</td>
			<td>Sigma-Aldrich</td>
			<td align="right">339350</td>
			<td>Powder used to make 10 mM NiCl2 in DI water</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Phosphate Buffered Saline (1x)</td>
			<td>Gibco</td>
			<td align="right">10010023</td>
			<td>PBS, pH 7.4</td>
		</tr>
	</tbody>
</table>

imaging of extracellular vesicles by atomic force microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

Watch this Scientific Journal Video about Imaging of Extracellular Vesicles by Atomic Force Microscopy at JoVE.com

Imaging of Extracellular Vesicles by Atomic Force Microscopy

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

imaging-of-extracellular-vesicles-by-atomic-force-microscopy

Nanyang Technological University

Research

JoVE Journal

Biochemistry

13.1K Views.  University of Utah. A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties. 

Video: Imaging of Extracellular Vesicles by Atomic Force Microscopy

Preparation of Giant Vesicles Encapsulating Microspheres by Centrifugation of a Water-in-oil Emulsion

The constructive biology and the synthetic biology approach to creating artificial life involve the bottom-up assembly of biological or nonbiological materials. Such approaches have received considerable attention in research on the boundary between living and nonliving matter and have been used to construct artificial cells over the past two decades. In particular, Giant Vesicles (GVs) have often been used as artificial cell membranes. In this paper, we describe the preparation of GVs encapsulating highly packed microspheres as a model of cells containing highly condensed biomolecules. The GVs were prepared by means of a simple water-in-oil emulsion centrifugation method. Specifically, a homogenizer was used to emulsify an aqueous solution containing the materials to be encapsulated and an oil containing dissolved phospholipids, and the resulting emulsion was layered carefully on the surface of another aqueous solution. The layered system was then centrifuged to generate the GVs. This powerful method was used to encapsulate materials ranging from small molecules to microspheres.

Giant vesicles containing highly packed micrometer-sized components are useful cell models. The water-in-oil emulsion centrifugation method is a simple, powerful tool for the preparation of giant vesicles with encapsulated materials.

The constructive biology and the synthetic biology approach to creating artificial life involve the bottom-up assembly of biological or nonbiological ...

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. ...

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Isolation of Tissue Extracellular Vesicles from the Liver

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in cell-to-cell communication by shuttling bioactive molecules such as RNA or protein from one cell to another. Most studies of EVs have been performed in cell culture models or in EVs isolated from body fluids. There is emerging interest in the isolation of EVs from tissues to study their contribution to physiological processes and how they are altered in disease. The isolation of EVs with sufficient yield from tissues is technically challenging because of the need for tissue dissociation without cellular damage. This method describes a procedure for the isolation of EVs from mouse liver tissue. The method involves a two-step process starting with in situ collagenase digestion followed by differential ultra-centrifugation. Tissue perfusion using collagenase provides an advantage over mechanical cutting or homogenization of liver tissue due to its increased yield of obtained EVs. The use of this two-step process to isolate EVs from the liver will be useful for the study of tissue EVs.

This is a protocol to isolate tissue extracellular vesicles (EVs) from the liver. The protocol describes a two-step process involving collagenase perfusion followed by differential ultracentrifugation to isolate liver tissue EVs.

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in ...

Rapid Fluorescence-based Characterization of Single Extracellular Vesicles in Human Blood with Nanoparticle-tracking Analysis

Extracellular vesicles (EVs), including exosomes, are specialized membranous nano-sized vesicles found in bodily fluids that are constitutively released from many cell types and play a pivotal role in regulating cell-cell communication and a diverse range of biological processes. Many different methods for the characterization of EVs have been described. However, most of these methods have the disadvantage that the preparation and characterization of the samples are very time-consuming, or it is extremely difficult to analyze specific markers of interest due to their small size and due to the lack of discrete populations. While methods for analysis of EVs have been considerably improved over the last decade, there is still no standardized method for characterization of single EVs. Here, we demonstrate a semi-automated method for characterization of single EVs by fluorescence-based nanoparticle-tracking analysis. The protocol that is presented addresses the common problem of many researchers in this field and provides the complete workflow for rapid isolation of EVs and characterization with PKH67, a general cell membrane linker, as well as with specific surface markers such as CD63, CD9, vimentin, and lysosomal-associated membrane protein 1 (LAMP-1). The presented results show a high level of reproducibility, as confirmed by other methods, such as Western blotting. In the conducted experiments, we exclusively used EVs isolated from human serum samples, but this method is also suitable for plasma or other body fluids and can be adjusted for characterization of EVs from cell culture supernatants. Irrespective of the future progress of research on EV biology, the protocol that is presented here provides a rapid and reliable method for rapid characterization of single EVs with specific markers.

In this protocol, we describe the complete workflow for rapid isolation of extracellular vesicles from human whole blood and characterization of specific markers by fluorescence-based nanoparticle-tracking analysis. The presented results show a high level of reproducibility and can be adjusted to cell culture supernatants.

Extracellular vesicles (EVs), including exosomes, are specialized membranous nano-sized vesicles found in bodily fluids that are constitutively released ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with neurodegenerative conditions such as Alzheimer&#8217;s and Parkinson&#8217;s diseases. In particular low molecular weight aggregates, amyloid oligomers, have been shown to possess generic cytotoxic properties and are implicated as neurotoxins in many forms of dementia. We illustrate the use of methods based on atomic force microscopy (AFM) to address the challenging task of characterizing the morphological, structural and chemical properties of these aggregates, which are difficult to study using conventional structural methods or bulk biophysical methods because of their heterogeneity and transient nature. Scanning probe microscopy approaches are now capable of investigating the morphology of amyloid aggregates with sub-nanometer resolution. We show here that infrared (IR) nanospectroscopy (AFM-IR), which simultaneously exploits the high resolution of AFM and the chemical recognition power of IR spectroscopy, can go further and enable the characterization of the structural properties of individual protein aggregates, and thus offer insights into the aggregation mechanisms. Since the approach that we describe can be applied also to the investigations of the interactions of protein assemblies with small molecules and antibodies, it can deliver fundamental information to develop new therapeutic compounds to diagnose or treat neurodegenerative disorders.

We describe the application of infrared nanospectroscopy and high-resolution atomic force microscopy to visualize the process of protein self-assembly into oligomeric aggregates and amyloid fibrils, which is closely associated with the onset and development of a wide range of human neurodegenerative disorders.

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...