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>

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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 ...

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;

Surface fixation of EVs is a critical step in the imaging sequence. Electrostatic surface immobilization of exosomes, known to have a negative zeta potential, will robustly occur after the mica&#8217;s substrate is modified to have a positive surface charge. Without the treatment with NiCl2 to impart positive surface changes, the immobilization of EVs on the substrate was found to be ineffective. The height image in Figure 10A, acquired in the air after the MCF-7 exosome sample co...

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

Research

JoVE Journal

Biochemistry

13.0K 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

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 ...

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 ...

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 ...

Isolating Small Extracellular Vesicles and Extracting Their Peptidome

Identification of Peptides of Small Extracellular Vesicles from Bone Marrow-Derived Macrophages

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of &lt;200 nm are present in various body fluids. These sEVs regulate various biological processes such as gene transcription and translation, cell proliferation and survival, immunity and inflammation through their cargos, such as proteins, DNA, RNA, and metabolites. Currently, various techniques have been developed for sEVs isolation. Among them, the ultracentrifugation-based method is considered the gold standard and is widely used for sEVs isolation. The peptides are naturally biomacromolecules with less than 50 amino acids in length. These peptides participate in a variety of biological processes with biological activity, such as hormones, neurotransmitters, and cell growth factors. The peptidome is intended to systematically analyze endogenous peptides in specific biological samples by liquid chromatography-tandem mass spectrometry (LC-MS/MS). Here, we introduced a protocol to isolate sEVs by differential ultracentrifugation and extracted peptidome for identification by LC-MS/MS. This method identified hundreds of sEVs-derived peptides from bone marrow-derived macrophages.

Author Spotlight: Peptidome Extraction from Small Extracellular Vesicles Isolated from Bone Marrow-Derived Macrophages 

This protocol describes a procedure to isolate small extracellular vesicles from macrophages by differential ultracentrifugation and extract the peptidome for identification by mass spectrometry.

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of ...