We would like to thank Oxford Nanoimaging for their constructive feedback and guidance. This work was funded by the 5UM1CA121947-10 to R.P.M. and the 1R01DA040394 to D.P.D.

1 Propagation and maintenance of cell lines
<ol>
	<li>Acquire human osteosarcoma cells (U2OS) and place the cells in the growth medium supplemented with 10% exosome-free Fetal Bovine Serum and 1x Penicillin/Streptomycin solution. 
	NOTE: Exosome-free Fetal Bovine Serum was generated following the protocol presented in McNamara et. al.26.</li>
	<li>Maintain the U2OS cells in a copper-coated incubator at 37 &#176;C in 5% CO2 and passage cells in T175 flasks26,27. The cells must be maintained in mid-logarithmic growth phase to prevent subpopulations from arising, or from the accumulation of apoptotic debris during the stationary phase.</li>
</ol>
2 Exosome isolation and purification
<ol>
	<li>Grow the U2OS cell lines to full confluency in 10 separate T175 flasks with 50 mL of media per flask. Remove the cell supernatant and successively passage it through a 0.45 &#181;m and 0.22 &#181;m vacuum filtration apparatus.</li>
	<li>Subject the supernatant to crossflow filtration (also known as tangential flow filtration) on a filtration system equipped with the 750 kDa hollow-fiber cartridge in order to remove smaller proteins or metabolites28.
	<ol>
		<li>Passage the supernatant through the tangential flow filtration operator at a constant forward pressure of 30 psi, while maintaining a retention pressure of &#60;20 psi to produce a &#916; pressure of 10 or more psi. Place a magnetic stir bar in the retentate tank and set to 150 rpm26.</li>
		<li>Maintain the feed rate at 40 mL/min or higher. Collect the permeate in a reservoir and dispose of it.</li>
	</ol>
	</li>
	<li>Concentrate the supernatant to 30 mL, and then equilibrate with four volumes of 1x PBS. Collect the crossflow filtered and equilibrated solution in a 50 mL conical tube.</li>
	<li>Precipitate the EVs at 4 &#176;C overnight in a final concentration of 40 mg/mL polyethylene glycol with a molecular weight of 8,000 Da. Centrifuge the EVs at 1,200 x g at 4 &#176;C for 1 h and resuspend in 500 &#181;L of 1x PBS.</li>
	<li>Incubate the EVs with 50 &#181;g/mL of photoswitchable red membrane intercalating dye and 50 &#181;g/mL of RNase A in 1x PBS at 4 &#176;C for 1 h.</li>
	<li>Remove excess dye through a column on a protein purification system attached to a fraction collector. Identify EV fractions through UV absorbance and pool them into a microcentrifuge tube26.</li>
	<li>Affinity-select a total of 200 &#181;L of EVs using anti-CD81 magnetic beads equilibrated in 1x PBS.
	<ol>
		<li>Dilute 100 &#181;L of the anti-CD81 magnetic beads in 1x PBS to a total volume of 1 mL and allow them to bind at 4 &#176;C for 2 h in a 2 mL microcentrifuge tube with continuous rocking.</li>
	</ol>
	</li>
	<li>Wash the anti-CD81 beads and EV three times with 1x PBS. Elute CD81+ EV from the solution with 100 mM Glycine at pH 2.0 for 30 min at 37 &#176;C.</li>
	<li>Pipette the purified CD81+ EV into an equal volume of 100 mM Tris-HCl at pH 7.5 in 1x PBS. 
	&#8203;NOTE: An aliquot of purified CD81+ EV solution may be reserved for Nanoparticle Tracking Analysis or electron microscopy in order to confirm the purity of the solution and presence of EVs26.</li>
</ol>
3 Fixation and preparation
<ol>
	<li>Place affinity-purified EVs onto glass-bottom &#181;-slide 8-well plates in a total volume of 200 &#181;L (can dilute EV sample in 100 mM Tris-HCl at pH 7.5 in 1x PBS) and allow to adhere to the surface overnight at 4 &#176;C.</li>
	<li>Without removing the existing solution from the 8-well plate, fix EVs onto the plates by adding 200 &#181;L of 4% paraformaldehyde in 1x PBS to the EV-containing solution in each well and allow to incubate for 30 min at room temperature21.</li>
	<li>Carefully remove paraformaldehyde and excess solution with a micropipette in order to not disturb the EVs. Wash the EV with 1x PBS to remove excess paraformaldehyde. Perform the wash procedure three times. Remove excess 1x PBS.</li>
	<li>Prepare 250 &#181;L of dSTORM B-cubed buffer solution per sample by creating a solution of 5 mM protocatechuate dioxygenase (Part A) diluted in imaging buffer (Part B) per manufacturer&#39;s protocol. 
	NOTE: The concentration of the enzyme in the buffer may be doubled to 10 mM if photobleaching occurs.
	<ol>
		<li>Add 250 &#181;L of the prepared buffer to each well per manufacturer&#39;s protocol for 20 min at room temperature before imaging to scavenge oxidizing molecules. 
		&#8203;NOTE: The EV can then be either viewed immediately or stored at 4 &#176;C for up to a week. Replace the buffer following storage before every visualization.</li>
	</ol>
	</li>
</ol>
4 Direct stochastic optical reconstruction microscopy calibration
<ol>
	<li>Prepare the beads required for the calibration of the super-resolution microscope by diluting 100 nm microspheres to a concentration of 0.5% in molecular biology grade water and pipetting 200 &#181;L into each well of a glass-bottom &#181;-slide 8-well plate.
	<ol>
		<li>Allow the beads to settle in the wells for 1 h at room temperature.</li>
	</ol>
	</li>
	<li>Without removing the existing solution, add 200 &#181;L of 4% paraformaldehyde in PBS to each well to the calibration bead solution and allow to incubate for 30 min at room temperature.</li>
	<li>Carefully remove the paraformaldehyde with a micropipette to not disturb the beads and wash the beads three times with 1x PBS. Prepare the buffer according to step 3.4.</li>
	<li>Remove 1x PBS and add 250 &#181;L of the prepared buffer to each well. Allow the buffer to sit for 20 min before visualization.</li>
	<li>Before placing anything on the stage, connect to the 3-D microscope using the Connect the Microscope button. Add 100x oil to the objective and place the center of the well on top of the objective. In the Acquire setting, turn on the 473 and 640 nm excitation lasers and click on View.
	<ol>
		<li>Without the 3-D lens activated, view the beads under the photon saturation setting by clicking on Photon Counts in the Image Display options. Set the initial laser powers to 8.4 mW for the 473 nm laser and to 11.6 mW for the 640 nm laser.</li>
		<li>Decrease the focus of the laser to around -300 nm or the focal plane of the calibration beads to produce a clear resolution of the individual beads. Once the Z-plane is focused, further adjust the laser power levels to account for variation in each field of view.</li>
	</ol>
	</li>
	<li>Under the instrument functions, complete 3-D mapping calibration and channel mapping calibration to obtain the errors on the X-, Y-, and Z-axis. Set the max number of FOVs to 20, the target number of points to 4,000, the max distance between channels to 5.0 pixels, and the exclusion radius between channels to 10.0 pixels during channel mapping calibration.</li>
	<li>Ensure that the calibration produces a point coverage of &#62;90% and mapping quality that is good. Save the given calibration data for future image acquisitions.</li>
</ol>
5 Visualization of EV in three dimensions
<ol>
	<li>Add 100x oil onto the objective and place the prepared EVs into the microscope. Without the 3-D lens activated, turn on the 640 nm excitation laser and initially raise it to between 1.2 mW and 12.5 mW depending on the intensity of the signal and field of view to excite the red membrane intercalating dye stained EVs.
	<ol>
		<li>Under the Image Display options, switch the viewing method from photon saturation to percentiles to better visualize the EVs. Adjust laser power to minimize noise, while maximizing signal. Maintain all the other parameters.</li>
		<li>Adjust the focus of the Z-plane by clicking the up or down icon on the Z-axis. 
		NOTE: The Z-plane should be focused between -200 and -350 nm, but will vary depending on the field of view.</li>
	</ol>
	</li>
	<li>Set the exposure time to 20 ms, the frame capture to 10,000 frames, and the initial laser power to between 1.2 mW and 12.5 mW, or the laser power determined in step 5.1, depending on the intensity of the signal and field of view.
	<ol>
		<li>Activate the 3-D lens using the icon and start the acquisition by clicking on the Acquire button.</li>
	</ol>
	</li>
	<li>Throughout image acquisition, raise the laser power by 3 increments of 10 every 1000 frames, or enough to maintain a high signal-to-noise ratio. Do not adjust the Z-plane during acquisition. 
	&#8203;NOTE: The laser may be raised to a maximum of 90 mW laser power.</li>
</ol>
6 Post-acquisition modification and EV tracing
<ol>
	<li>After the image acquisition, toggle over to the Analyze viewing window. Perform drift correction on the unfiltered image, and then activate filters. Adjust photon count, localization precision, sigmas, and frame index according to Table 1.</li>
	<li>Overlay an XYZ plane view tool along the X-axis of individual EVs from the field of view and export .CSV files of photoswitching events.</li>
	<li>Bisect individual EVs on the XY-axis in an X by Y field of view using a line histogram tool, which bins photoswitching events into set distance groups.</li>
	<li>Take images of single EVs and save them as .tiff files.</li>
	<li>Create 3-D videos of individual EVs using a 3-D visualization tool and color according to placement along the Z-axis.</li>
</ol>

M.G.C. has no conflicts of interest to declare. R.P.M and D.P.D. receive material support from Oxford Nanoimaging (ONI) Inc. and Cytiva Inc. (formerly GE Healthcare). R.P.M. and D.P.D. declare competing interests for the possible commercialization of some of the information presented. These are managed by the University of North Carolina. The funding sources were not involved in the interpretations or writing of this manuscript.

EVs have become a popular area of study due to their important role in many intracellular processes and cell-to-cell signaling1,30. However, their visualization proves to be difficult as their small size falls below the diffraction limit of light microscopy. Direct stochastic optical reconstruction microscopy (dSTORM) is a direct method of visualization that bypasses the diffraction limit by capturing photoswitching events of individual fluorophores over time and...

Extracellular vesicles (EVs) are membrane-bound vesicles released by all cell types. They contain lipids, proteins, metabolites, and nucleic acids and transfer these materials locally between cells and distally between tissues and organs. There are three primary subtypes of EVs: apoptotic bodies, microvesicles, and exosomes1,2. Here, we focus our discussion on exosomes and their associated proteins.
Exosomes are secreted vesicles originating from the inward budding of early endosomes into the multivesicular body (MVB). The MVB then fuses with the plasma membrane, releasing the exosomes into the extracellular space to travel to other cells3,4. Exosomes exist on a spectrum of sizes ranging from 40 to 150 nm and are enriched with endosomal transmembrane proteins known as tetraspanins (CD9, CD63, CD81), membrane-bound endosomal sorting complex required for the transport (ESCRT), and lipid raft-associated proteins1,2, 5,6,7.
Characterizing the biochemical makeup of exosomes has become a popular field for researchers to better understand their functional nature. Many methods exist for visualizing and characterizing exosomes, including nanoscale flow cytometry, nanoparticle tracking analysis (NTA), scanning and transmission electron microscopy (TEM), surface plasmon resonance, resistive pulse sensing, and traditional light microscopy, each of which contains intrinsic pros and cons8,9. TEM and cryo-EM can achieve nanometer-based resolution, but often require dehydrating and freeze-fracture steps, thereby shrinking or lysing EVs10,11. NTA relies on light scattering, allowing for the characterization of hundreds of EVs at a time, but is an indirect measurement of particle size and cannot easily distinguish between EVs, viruses, and protein aggregates12,13,14,15,16. Nanoscale flow cytometry employs light scattering from an excitation path, which can then be translated into size measurements, but is an emerging technology, and there is little consensus on what size of particles are within the linear range of detection for various instruments12,17,18.
Traditional light microscopy using fluorescent proteins or dyes has been one of the most heavily employed techniques for visualizing subcellular compartments, protein complexes, and signaling machinery within a cell. While this technique proves useful in visualizing the localization of complexes, the diffraction limit of traditional light microscopy (around 250-400 nm) prevents the clear resolution of proteins or structures in the typical size range of an exosome (40-150 nm)12,19,20.
Super-resolution microscopy, namely, direct stochastic optical reconstruction microscopy (dSTORM), distinguishes itself from conventional light microscopy by employing the photoswitchable properties of specific fluorophores and detecting these blinking events to reconstruct images down to nanometer precision21. Photoswitching events are collected using a high-framerate detection camera over the course of tens of thousands of individual exposures, and a point spread function is used to map with high confidence the exact location of the photoswitching fluorophore19,20,22. This allows dSTORM to bypass the diffraction limit of light microscopy. Several groups have reported the use of super-resolution techniques for visualizing and tracking exosomes and their associated proteins22,23,24,25. The final resolution depends on the biophysical properties of the fluorophore, but often ranges from +/-10-100 nm along the XY-axis, allowing single-molecule resolution.
The ability to resolve individual fluorophores at this scale on the XY-axis has revolutionized microscopy. However, there is little data on the three-dimensional (3-D) dSTORM of an exosome. Therefore, we sought to establish a standard operating procedure (SOP) for dSTORM-based visualization and characterization of purified EVs, including exosomes to nanometer precision in 3-D.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 &#181;-Slide 8 well plates</td>
			<td>Ibidi</td>
			<td>80827</td>
			<td></td>
		</tr>
		<tr>
			<td>1X PBS</td>
			<td>Gibco</td>
			<td>14190-144</td>
			<td></td>
		</tr>
		<tr>
			<td>1X Penicillin Streptomycin solution</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL conical tube</td>
			<td>Thermo Fisher</td>
			<td>339652</td>
			<td></td>
		</tr>
		<tr>
			<td>500 mL 0.22 &#181;m vacuum filtration apparatus</td>
			<td>Genesee</td>
			<td>25-227</td>
			<td></td>
		</tr>
		<tr>
			<td>750 kDa hollow-fiber cartridge cutoff filter</td>
			<td>Cytiva</td>
			<td>29-0142-95</td>
			<td></td>
		</tr>
		<tr>
			<td>AKTA Flux S</td>
			<td>Cytiva</td>
			<td>29-0384-37</td>
			<td></td>
		</tr>
		<tr>
			<td>AKTA Start</td>
			<td>Cytiva</td>
			<td>29022094-ECOMINSSW</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-CD81 magnetic beads</td>
			<td>Thermo Fisher</td>
			<td>10616D</td>
			<td></td>
		</tr>
		<tr>
			<td>B-cubed buffer</td>
			<td>ONI</td>
			<td>&#160;BCA0017</td>
			<td></td>
		</tr>
		<tr>
			<td>CellMask Red</td>
			<td>Thermo Fisher</td>
			<td>C10046</td>
			<td></td>
		</tr>
		<tr>
			<td>Dubelco&#39;s Modified Eagle Medium</td>
			<td>Thermo Fisher</td>
			<td>10566016</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>VWR</td>
			<td>97068-085</td>
			<td></td>
		</tr>
		<tr>
			<td>Frac 30 Fraction collector</td>
			<td>Cytiva</td>
			<td>29022094-ECOMINSSW</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine pH=2.0</td>
			<td>Thermo Fisher</td>
			<td>BP381-5</td>
			<td></td>
		</tr>
		<tr>
			<td>HiTrap CaptoCore 700 Column</td>
			<td>Cytiva</td>
			<td>17548151</td>
			<td></td>
		</tr>
		<tr>
			<td>Molecular Biology Grade Water</td>
			<td>Corning</td>
			<td>9820003</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanoimager</td>
			<td>Oxford Nanoimaging</td>
			<td>Custom</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehhyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene glycol</td>
			<td>Thermo Fisher</td>
			<td>BP233-1</td>
			<td></td>
		</tr>
		<tr>
			<td>RNase A</td>
			<td>Promega</td>
			<td>A797C</td>
			<td></td>
		</tr>
		<tr>
			<td>T175 Flasks</td>
			<td>Genesee</td>
			<td>25-211</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetraspek microspheres</td>
			<td>Invitrogen</td>
			<td>T7279</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris- HCl pH=7.5</td>
			<td>Thermo Fisher</td>
			<td>BP153-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Unicorn V</td>
			<td>Cytiva</td>
			<td>29022094-ECOMINSSW</td>
			<td></td>
		</tr>
	</tbody>
</table>

direct stochastic optical reconstruction microscopy of extracellular vesicles in three dimensions

Extracellular vesicles (EVs) are released by all cell types and play an important role in cell signaling and homeostasis. The visualization of EVs often require indirect methods due to their small diameter (40-250 nm), which is beneath the diffraction limit of typical light microscopy. We have developed a super-resolution microscopy-based visualization of EVs to bypass the diffraction limit in both two and three dimensions. Using this approach, we can resolve the three-dimensional shape of EVs to within +/- 20 nm resolution on the XY-axis and +/- 50 nm resolution along the Z-axis. In conclusion, we propose that super-resolution microscopy be considered as a characterization method of EVs, including exosomes, as well as enveloped viruses.

The goal of this study was to evaluate the effectiveness of super-resolution microscopy in visualizing individual EVs with nanometer resolution in three dimensions (3-D). To analyze the shape and size of individual EVs, we employed photoswitchable dye and incubated the EVs with a far-red, membrane intercalating dye, and removed excess dye through chromatography29. The affinity-captured anti-CD81 and red-stained EVs were then viewed in the super-resolution microscope under the 640 nm excitation las...

Watch this Scientific Journal Video about Direct Stochastic Optical Reconstruction Microscopy of Extracellular Vesicles in Three Dimensions at JoVE.com

Direct Stochastic Optical Reconstruction Microscopy of Extracellular Vesicles in Three Dimensions

Direct stochastic optical reconstruction microscopy (dSTORM) is used to bypass the typical diffraction limit of light microscopy and to view exosomes at the nanometer scale. It can be employed in both two and three dimensions to characterize exosomes.

Extracellular vesicles (EVs) are released by all cell types and play an important role in cell signaling and homeostasis. The visualization of EVs often ...

direct-stochastic-optical-reconstruction-microscopy-extracellular

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3.6K Views.  The University of North Carolina at Chapel Hill School of Medicine. Direct stochastic optical reconstruction microscopy (dSTORM) is used to bypass the typical diffraction limit of light microscopy and to view exosomes at the nanometer scale. It can be employed in both two and three dimensions to characterize exosomes.

Video: Direct Stochastic Optical Reconstruction Microscopy of Extracellular Vesicles in Three Dimensions

Phase Contrast and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by Zernike, in 1942, who received the Nobel prize for his achievement. DIC microscopy, introduced in the late 1960s, has been popular in biomedical research because it highlights edges of specimen structural detail, provides high-resolution optical sections of thick specimens including tissue cells, eggs, and embryos and does not suffer from the  phase halos  typical of phase-contrast images. This protocol highlights the principles and practical applications of these microscopy techniques.

Phase Contrast and Differential Interference Contrast DIC Microscopy

This protocol highlights the principles and practical applications of Phase and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by ...

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we know about C. elegans is limited to laboratory cultivation of the nematodes that may not necessarily reflect the environments they normally inhabit in nature. Cultivation of C. elegans in a 3D habitat that is more similar to the 3D matrix that worms encounter in rotten fruits and vegetative compost in nature could reveal novel phenotypes and behaviors not observed in 2D. In addition, experiments in 3D can address how phenotypes we observe in 2D are relevant for the worm in nature. Here, a new method in which C. elegans grows and reproduces normally in three dimensions is presented. Cultivation of C. elegans in Nematode Growth Tube-3D (NGT-3D) can allow us to measure the reproductive fitness of C. elegans strains or different conditions in a 3D environment. We also present a novel method, termed Nematode Growth Bottle-3D (NGB-3D), to cultivate C. elegans in 3D for microscopic analysis. These methods allow scientists to study C. elegans biology in conditions that are more reflective of the environments they encounter in nature. These can help us to understand the overlying evolutionary relevance of the physiology and behavior of C. elegans we observe in the laboratory.

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

We present a simple method to construct 3D nematode cultivation systems called NGT-3D and NGB-3D. These can be used to study nematode fitness and behaviors in habitats that are more similar to natural Caenorhabditis elegans habitats than the standard 2D laboratory C. elegans culture plates.

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we ...

Hemi-laryngeal Setup for Studying Vocal Fold Vibration in Three Dimensions

The voice of humans and most non-human mammals is generated in the larynx through self-sustaining oscillation of the vocal folds. Direct visual documentation of vocal fold vibration is challenging, particularly in non-human mammals. As an alternative, excised larynx experiments provide the opportunity to investigate vocal fold vibration under controlled physiological and physical conditions. However, the use of a full larynx merely provides a top view of the vocal folds, excluding crucial portions of the oscillating structures from observation during their interaction with aerodynamic forces. This limitation can be overcome by utilizing a hemi-larynx setup where one half of the larynx is mid-sagittally removed, providing both a superior and a lateral view of the remaining vocal fold during self-sustained oscillation.
Here, a step-by-step guide for the anatomical preparation of hemi-laryngeal structures and their mounting on the laboratory bench is given. Exemplary phonation of the hemi-larynx preparation is documented with high-speed video data captured by two synchronized cameras (superior and lateral views), showing three-dimensional vocal fold motion and corresponding time-varying contact area. The documentation of the hemi-larynx setup in this publication will facilitate application and reliable repeatability in experimental research, providing voice scientists with the potential to better understand the biomechanics of voice production.

This paper introduces a protocol for the preparation of hemi-larynx specimens facilitating a multi-dimensional view of vocal fold vibration, in order to investigate various biophysical aspects of voice production in humans and non-human mammals.

The voice of humans and most non-human mammals is generated in the larynx through self-sustaining oscillation of the vocal folds. Direct visual ...

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, ...

Flow Cytometric Analysis of Extracellular Vesicles from Cell-conditioned Media

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for phenotypic analyses of extracellular vesicles (EV) including exosomes (Exo) in the peripheral blood and other body fluids. The small size of EV mandates the use of dedicated instruments having a detection threshold around 50-100 nm. Alternatively, EV can be bound to latex microbeads that can be detected by FC. Microbeads, conjugated with antibodies that recognize EV-associated markers/Cluster of Differentiation CD63, CD9, and CD81 can be used for EV capture. Exo isolated from CM can be analyzed with or without pre-enrichment by ultracentrifugation. This approach is suitable for EV analyses using conventional FC instruments. Our results demonstrate a linear correlation between Mean Fluorescence Intensity (MFI) values and EV concentration. Disrupting EV through sonication dramatically decreased MFI, indicating that the method does not detect membrane debris. We report an accurate and reliable method for the analysis of EV surface antigens, which can be easily implemented in any laboratory.

The protocol describes a reproducible method designed for use with cell culture supernatants to detect surface epitopes on small extracellular vesicles (EV). It utilizes specific EV immunoprecipitation using beads coupled with antibodies that recognize surface antigen CD9, CD63, and CD81. The method is optimized for downstream flow cytometry analysis.

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for ...

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular vesicles (EVs) function outside the cell to regulate global physiological processes by transferring proteins, nucleic acids, metabolites, and lipids between tissues. EVs reflect the physiological state of their cells of origin. EVs are implicated to have fundamental roles in virtually every aspect of human health. Thus, EV protein and genetic cargos are being increasingly analyzed for biomarkers of health and disease. However, the EV field still lacks a tractable invertebrate model system that permits the study of EV cargo composition. C. elegans is well suited for EV research because it actively secretes EVs outside of its body into its external environment, permitting facile isolation. This article provides all the necessary information for generating, purifying, and quantifying these environmentally secreted C. elegans EVs including how to work quantitatively with very large populations of age-synchronized worms, purifying EVs, and a flow cytometry protocol that directly measures the number of intact EVs in the purified sample. Thus, the large library of genetic reagents available for C. elegans research can be tapped into for investigating the impacts of genetic pathways and physiological processes on EV cargo composition.

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

This article presents methods for generating, purifying, and quantifying Caenorhabditis elegans extracellular vesicles.

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular ...

Reconstruction of Single-Cell Innate Fluorescence Signatures by Confocal Microscopy

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing the innate cellular fluorescence signature from each individual live cell in a population distributed in a three-dimensional (3D) space. The innate fluorescence signature of a cell is a collection of fluorescence signals emitted by various biomolecules within the cell. Previous studies established that innate fluorescence signatures reflect various cellular properties and differences in physiological status and are a rich source of information for cell characterization and identification. Innate fluorescence signatures have been traditionally analyzed at the population level, necessitating a clonal culture, but not at the single-cell level. CRIF is particularly suitable for studies that require 3D resolution and/or selective extraction of fluorescence signals from individual cells. Because the fluorescence signature is an innate property of a cell, CRIF is also suitable for tag-free prediction of the type and/or physiological status of intact and single cells. This method may be a powerful tool for streamlined cell analysis, where the phenotype of each single cell in a heterogenous population can be directly assessed by its autofluorescence signature under a microscope without cell tagging.

Here, a protocol is presented for optically extracting and cataloging innate cellular fluorescence signatures (i.e., cellular autofluorescence) from every individual live cell distributed in a three-dimensional space. This method is suitable for studying the innate fluorescence signature of diverse biological systems at a single-cell resolution, including cells from bacteria, fungi, yeasts, plants, and animals.

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing ...

Nanoparticle Tracking Analysis for the Quantification and Size Determination of Extracellular Vesicles

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving area of research. There are many different methods for EV isolation, each with distinct advantages and disadvantages that affect the downstream yield and purity of EVs. Thus, characterizing the EV prep isolated from a given source by a chosen method is important for interpretation of downstream results and comparison of results across laboratories. Various methods exist for determining the size and quantity of EVs, which can be altered by disease states or in response to external conditions. Nanoparticle tracking analysis (NTA) is one of the prominent technologies used for high-throughput analysis of individual EVs. Here, we present a detailed protocol for quantification and size determination of EVs isolated from mouse perigonadal adipose tissue and human plasma using a breakthrough technology for NTA representing major advances in the field. The results demonstrate that this method can deliver reproducible and valid total particle concentration and size distribution data for EVs isolated from different sources using different methods, as confirmed by transmission electron microscopy. The adaptation of this instrument for NTA will address the need for standardization in NTA methods to increase rigor and reproducibility in EV research.

We demonstrate how to use a novel nanoparticle tracking analysis instrument to estimate the size distribution and total particle concentration of extracellular vesicles isolated from mouse perigonadal adipose tissue and human plasma.

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...