This research was funded by the Slovenian Research Agency (research core funding no. P3-0108 and project J7-2594) and the MRIC UL IP-0510 Infrastructure program. This work contributes to the COST Action CA17116 International Network for Translating Research on Perinatal Derivatives into Therapeutic Approaches (SPRINT), supported by COST (European Cooperation in Science and Technology). The authors would like to thank Linda &#352;trus, Sanja &#268;abraja, Nada Pavlica Dubari&#269;, and Sabina &#381;eleznik for technical help with cell culturing and preparing the samples and Marko Vogrinc, Ota &#352;irca Ro&#353;, and Nejc Debevec for technical help preparing the video.

1. Culturing cancerous urothelial cells and isolation of EVs
NOTE: A protocol to obtain EVs from a human invasive bladder cancer urothelial (T24) cell line is presented. However, culturing conditions have to be optimized to use other cell types.
<ol>
	<li>Plate T24 cells with a density of 3 &#215; 104 cells/cm2 in three flasks (growing surface of 75 cm2) and start with culturing the cells in a CO2 incubator for 3 days at 37 &#176;C and a 5% CO2 (Figure 2A).
	<ol>
		<li>Use a culture medium for T24 cells in a 1:1 (v/v) mixture of A-DMEM and F12 supplemented with 5% fetal bovine serum (FBS), 4 mM Glutamax, 100 mg/mL streptomycin, and 100 U/mL penicillin. 
		NOTE:&#160;The following isolation yields EVs enriched in MVs. Before starting the isolation, inspect the cells with a light microscope to confirm viability and confluence (Figure 2B). Start collecting the EVs from the conditioned culture media when the T24 cells are at 70% confluence.</li>
	</ol>
	</li>
	<li>Collect the culture medium with MVs with a pipette from flasks (T75) into conical centrifuge tubes (Figure 2C). Centrifuge at 300 &#215; g for 10 min at 4 &#176;C (Figure 2D).</li>
	<li>Carefully collect the supernatant into a new centrifuge tube with the pipette (Figure 2E).</li>
	<li>Centrifuge the supernatant at 2,000 &#215; g&#160; for 20&#160;min&#160;at&#160;4 &#176;C.</li>
	<li>Carefully collect the supernatant with the pipette into a new centrifuge tube.</li>
	<li>Centrifuge at 10,000 &#215; g for 40 min at 4 &#176;C. Look for the presence of a whitish pellet at the bottom of the tube. 
	NOTE: If necessary, change the centrifuge rotor to reach the required g-force. The EV pellet is seen as a white pellet at the bottom of the centrifuge tube (Figure 2F); mark its location to avoid losing the pellet during pipetting (Figure 2G).</li>
	<li>Carefully discard the supernatant with a pipette. To the pellet, add 1.5 mL of PBS and resuspend the pellet containing MVs. Put the suspension into a new centrifuge tube.</li>
	<li>Centrifuge at 10,000 &#215; g for 40 min at 4 &#176;C and look for the presence of a white pellet at the bottom of the tube. 
	NOTE: The EV pellet is seen as a white patch at the bottom of the centrifuge tube; mark its location to avoid losing the pellet during pipetting.</li>
	<li>Carefully discard the supernatant with a pipette. Gently add the fixative, 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.2). Leave for 20 min at 4 &#176;C (Figure 2H). 
	CAUTION: Glutaraldehyde and sodium cacodylate are toxic. Work in a fume hood, wear protective gloves, and discard them appropriately.</li>
	<li>Carefully remove the fixative with a pipette. Add washing buffer (0.1 M sodium cacodylate buffer) to the pellet. Leave for 10 min at 4 &#176;C without resuspending the pellet.</li>
</ol>
2. Freezing of EVs
NOTE: Before freezing, first cryoprotect the samples with glycerol.
<ol>
	<li>Prepare 0.1 M sodium cacodylate buffer. Remove the washing buffer and add 50 &#181;L of 30% glycerol in 0.1 M cacodylate buffer. Gently resuspend the EVs to make the solution homogeneous. Incubate for 30 min at 4 &#176;C.</li>
	<li>Clean the copper carriers with a central pit in the ultrasonic bath with chloroform for 5 min (Figure 3A). Air dry them and mark them with colors if using multiple samples (Figure 3B). Until use, keep the carriers in a dry, clean place (e.g., on lens paper in a Petri dish). Do not touch the carriers with bare hands.
	<ol>
		<li>Prepare pipettes, solutions, filter paper, tweezers, a stereomicroscope, a Dewar flask with freon and liquid nitrogen (LN2), a metal rod, and cryovials. Cool it down with LN2. 
		CAUTION: Wear protective equipment and work accordingly when handling LN2 and cooled freon.</li>
	</ol>
	</li>
	<li>Resuspend the EVs again before freezing. Avoid the formation of air bubbles. 
	NOTE: Perform steps 2.4-2.7 rapidly.</li>
	<li>Work under a stereomicroscope (Figure 3C). Transfer 1.5-1.7 &#181;L of the sample into the central pit of the copper carrier to make a convex drop reaching higher than the copper carrier edge while not spilling the sample over the edge (Figure 3D). In case of overspill, use filter paper to dry the outer ring of the carrier.</li>
	<li>Mix solidified freon with a metal rod to liquefy it again (Figure 3E).</li>
	<li>Grip the outer ring of the copper carrier with tweezers and immerse it into cooled freon with a gentle shaking of tweezers for 8 s (Figure 3F). After 8 s, quickly transfer the carrier into another Dewar flask filled with LN2.</li>
	<li>Proceed immediately with fracturing or storing the samples in LN2. In the latter case, mark a cryovial and cool the vial and tweezer tips by submerging them into LN2 (Figure 3G). Under LN2, collect frozen copper carriers into the vial, close it, and store them in the LN2 container (Figure 3H) until freeze fracturing.</li>
</ol>
3. Fracturing of EVs and making the replicas
NOTE: Prepare the freeze-fracturing unit according to the manufacturer&#39;s operating instructions. (Figure 4A). Clean the chamber of the unit. Position platinum (Pt/C) and carbon (C) guns at angles of 45&#176; and 90&#176;, respectively (Figure 4B).
<ol>
	<li>Start the unit vacuum system. When pressure of 6.6 &#215; 10&#8722;2 Pa (5 &#215; 10&#8722;4 Torr) is reached, cool down the unit chamber to &#8722;150 &#176;C.</li>
	<li>Transfer the copper carriers with frozen samples into a freeze-fracturing unit using dry tweezers (Figure 4C). Use cryotransfer or work fast to avoid thawing the sample and deposition of the ice crystals. Secure the carriers onto the sample table.</li>
	<li>Wait until the vacuum reaches again 6.6 &#215; 10&#8722;2 Pa (5 &#215; 10&#8722;4 Torr), and then set the knife temperature to &#8722;100 &#176;C (Figure 4D).</li>
	<li>Wait until the vacuum reaches 1.0 &#215; 10&#8722;3 Pa (8 &#215; 10&#8722;6 Torr).</li>
	<li>Observe the sample with binoculars and carefully approach the knife (Figure 4E).</li>
	<li>Begin sectioning the sample by turning on the motorized movement of the knife (Figure 4F) and section until the surface of the sample is smooth (Figure 4G).</li>
	<li>For fracturing, turn off the motorized movement of the knife and proceed with slow manual control of the knife, thereby fracturing the sample (Figure 4H). In order to protect the fractured surface from forming ice crystals and debris until shadowing, move the knife above the fractured sample.</li>
	<li>Proceed with the Pt shadowing at 45&#176; (Figure 4I, J).
	<ol>
		<li>Prepare a stopwatch and/or turn on the quartz crystal monitor on the freeze-fracturing unit, which will allow supervision of the thickness of the Pt on the sample. The recommended thickness of the Pt layer as measured on the quartz crystal monitor is 2.5 nm, which corresponds to 4 s of shadowing at a given high tension and current (i.e., the speed of Pt shadowing is 0.63 nm/s).</li>
		<li>Before starting the shadowing, move the knife away from the sample. Apply high tension on the Pt/C gun (1,600 V), and increase the current to 60 mA. Sparks of Pt should appear and should begin to shadow the sample. At this point, start the 4 s countdown and/or observe and locate the position of Pt on the quartz crystal monitor. 
		NOTE: The recommended thickness of the C layer is 2.5 nm, which corresponds to 10 s of shadowing at a given high tension and current.</li>
		<li>Turn off the current and high tension when the desired thickness of Pt is obtained.</li>
	</ol>
	</li>
	<li>Proceed with the C shadowing at 90&#176; (i.e., the speed of C shadowing is 0.25 nm/s).
	<ol>
		<li>Apply high tension on the C gun (1,900 V), increase the current to 90 mA, and sparks of C should appear and begin to shadow the sample; at that moment, start the 10 s countdown. Turn off the current and high tension when the desired thickness of C is obtained.</li>
	</ol>
	</li>
</ol>
4. Cleaning the replicas and replica analysis
<ol>
	<li>Use the copper carriers from the sample table and transfer them with tweezers to distilled water at room temperature in a porcelain 12-well plate (Figure 4K). The replicas will float on the water surface, while the carriers sink (Figure 4L).</li>
	<li>Transfer the replicas with a wire loop from distilled water into a well with sodium hypochlorite. Cover and leave overnight at room temperature in a fume hood. 
	CAUTION: Sodium hypochlorite is toxic. Work in a fume hood, wear protective gloves, and discard them appropriately.</li>
	<li>Transfer the replicas to ddH2O and wash for 30 min. Repeat 3 times.</li>
	<li>Transfer the replicas with a wire loop to a mesh copper TEM grid (e.g., square or honeycomb 100+ mesh). Air dry the grids for 2 h, and then store them in a grid storage box until microscopy. 
	NOTE: Generally, grids can be stored for months in dark, dry, room temperature conditions.</li>
	<li>Use a TEM microscope to image the replicas and obtain micrographs.
	<ol>
		<li>To do so, insert a TEM grid with a replica and start the TEM imaging according to the manufacturer&#39;s operating manuals. Work at 80 kV or 100 kV.</li>
	</ol>
	</li>
	<li>Inspect the sample and acquire micrographs at lower and higher magnifications with a camera on TEM.</li>
	<li>Analyze the micrographs of the MVs. For measurements of the MV diameters, use Fiji/ImageJ software16. The diameter of MVs is 4/&#960; times the measurement on the micrographs according to Hallett et al.17.</li>
</ol>

<ol>
	<li>Raposo, G., Stoorvogel, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+vesicles:+Exosomes,+microvesicles,+and+friends.">Extracellular vesicles: Exosomes, microvesicles, and friends.</a> The Journal of Cell Biology. 200 (4), 373-383 (2013).</li><li>Willms, E., Cabanas, C., Mager, I., Wood, M. J. A., Vader, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+vesicle+heterogeneity:+Subpopulations,+isolation+techniques,+and+diverse+functions+in+cancer+progression.">Extracellular vesicle heterogeneity: Subpopulations, isolation techniques, and diverse functions in cancer progression.</a> Frontiers in Immunology. 9, 738 (2018).</li><li>van Niel, G., D'Angelo, G., Raposo, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shedding+light+on+the+cell+biology+of+extracellular+vesicles.">Shedding light on the cell biology of extracellular vesicles.</a> Nature Reviews Molecular Cell Biology. 19, 213-228 (2018).</li><li>Cocucci, E., Racchetti, G., Meldolesi, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shedding+microvesicles:+Artefacts+no+more.">Shedding microvesicles: Artefacts no more.</a> Trends in Cell Biology. 19 (2), 43-51 (2009).</li><li>Urabe, F., 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=Extracellular+vesicles+as+biomarkers+and+therapeutic+targets+for+cancer.">Extracellular vesicles as biomarkers and therapeutic targets for cancer.</a> American Journal of Physiology-Cell Physiology. 318 (1), 29-39 (2020).</li><li>Tricarico, C., Clancy, J., D'Souza-Schorey, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biology+and+biogenesis+of+shed+microvesicles.">Biology and biogenesis of shed microvesicles.</a> Small GTPases. 8 (4), 220-232 (2017).</li><li>Georgantzoglou, N., Pergaris, A., Masaoutis, C., Theocharis, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+vesicles+as+biomarkers+carriers+in+bladder+cancer:+Diagnosis,+surveillance,+and+treatment.">Extracellular vesicles as biomarkers carriers in bladder cancer: Diagnosis, surveillance, and treatment.</a> International Journal of Molecular Sciences. 22 (5), 2744 (2021).</li><li>S&#322;omka, A., Urban, S. K., Lukacs-Kornek, V., &#379;ekanowska, E., Kornek, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+extracellular+vesicles:+Have+we+found+the+holy+grail+of+inflammation.">Large extracellular vesicles: Have we found the holy grail of inflammation.</a> Frontiers in Immunology. 9, 2723 (2018).</li><li>Han, L., Lam, E. W. F., Sun, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+vesicles+in+the+tumor+microenvironment:+Old+stories,+but+new+tales.">Extracellular vesicles in the tumor microenvironment: Old stories, but new tales.</a> Molecular Cancer. 18 (1), 59 (2019).</li><li>Muralidharan-Chari, V., Clancy, J. W., Sedgwick, A., D'Souza-Schorey, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microvesicles:+Mediators+of+extracellular+communication+during+cancer+progression.">Microvesicles: Mediators of extracellular communication during cancer progression.</a> Journal of Cell Science. 123, 1603-1611 (2010).</li><li>Kowal, 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=Proteomic+comparison+defines+novel+markers+to+characterize+heterogeneous+populations+of+extracellular+vesicle+subtypes.">Proteomic comparison defines novel markers to characterize heterogeneous populations of extracellular vesicle subtypes.</a> Proceedings of the National Academy of Sciences of the United States of America. 113 (8), 968-977 (2016).</li><li>Thery, C., Amigorena, S., Raposo, G., Clayton, A. <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. , (2006).</li><li>Thery, 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. 7 (1), 1535750 (2018).</li><li>Severs, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Freeze-fracture+electron+microscopy.">Freeze-fracture electron microscopy.</a> Nature Protocols. 2 (3), 547-576 (2007).</li><li>Severs, N. J., Robenek, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Freeze-fracture+cytochemistry+in+cell+biology.">Freeze-fracture cytochemistry in cell biology.</a> Methods in Cell Biology. 88, 181-186 (2008).</li><li>Rueden, C. 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=ImageJ2:+ImageJ+for+the+next+generation+of+scientific+image+data.">ImageJ2: ImageJ for the next generation of scientific image data.</a> BMC Bioinformatics. 18 (1), 529 (2017).</li><li>Hallett, F. R., Nickel, B., Samuels, C., Krygsman, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+vesicle+size+distributions+by+freeze-fracture+electron+microscopy.">Determination of vesicle size distributions by freeze-fracture electron microscopy.</a> Journal of Electron Microscopy Technique. 17 (4), 459-466 (1991).</li><li>Nigro, 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=Selective+loss+of+microvesicles+is+a+major+issue+of+the+differential+centrifugation+isolation+protocols.">Selective loss of microvesicles is a major issue of the differential centrifugation isolation protocols.</a> Scientific Reports. 11, 3589 (2021).</li><li>Szatanek, R., 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=The+methods+of+choice+for+extracellular+vesicles+(EVs)+characterization.">The methods of choice for extracellular vesicles (EVs) characterization.</a> International Journal of Molecular Sciences. 18 (6), 1153 (2017).</li><li>Iliev, 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=Stimulation+of+exosome+release+by+extracellular+DNA+is+conserved+across+multiple+cell+types.">Stimulation of exosome release by extracellular DNA is conserved across multiple cell types.</a> The FEBS Journal. 285 (16), 3114-3133 (2018).</li><li>Zupan&#269;i&#269;, D., Zakraj&#353;ek, M., Zhou, G., Romih, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+and+localization+of+four+uroplakins+in+urothelial+preneoplastic+lesions.">Expression and localization of four uroplakins in urothelial preneoplastic lesions.</a> Histochemistry and Cell Biology. 136 (4), 491-500 (2011).</li><li>Wu, X. R., Manabe, M., Yu, J., Sun, T. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+scale+purification+and+immunolocalization+of+bovine+uroplakins+I,+II,+and+III.+Molecular+markers+of+urothelial+differentiation.">Large scale purification and immunolocalization of bovine uroplakins I, II, and III. Molecular markers of urothelial differentiation.</a> The Journal of Biological Chemistry. 265 (31), 19170-19179 (1990).</li><li>Kreft, M. E., Hudoklin, S., Jezernik, K., Romih, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+and+maintenance+of+blood-urine+barrier+in+urothelium.">Formation and maintenance of blood-urine barrier in urothelium.</a> Protoplasma. 246 (1-4), 3-14 (2010).</li><li>Kreft, M. E., Robenek, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Freeze-fracture+replica+immunolabelling+reveals+urothelial+plaques+in+cultured+urothelial+cells.">Freeze-fracture replica immunolabelling reveals urothelial plaques in cultured urothelial cells.</a> PLoS One. 7 (6), 38509 (2012).</li><li>Robenek, H., 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=Topography+of+lipid+droplet-associated+proteins:+Insights+from+freeze-fracture+replica+immunogold+labeling.">Topography of lipid droplet-associated proteins: Insights from freeze-fracture replica immunogold labeling.</a> Journal of Lipids. 2011, 409371 (2011).</li></ol>

The authors declare no conflicts of interest.

The characterization of MVs, or any other population of isolated EVs, is of prime importance to begin with before starting downstream analyses like &#34;omics&#34; studies or functional studies11,18. Herein, EVs from human invasive bladder cancer urothelial T24 cells were isolated by centrifugation, and following the provided protocol for analysis by freeze-fracture electron microscopy, we demonstrated that the isolated fraction was enriched in MVs11,13. The isolate of MVs was devoid of cell debris or organelles, confirming a successful isolation and purification protocol.
The combination of chemical fixation and freezing, which are critical steps of the protocol, retained the spherical shape of the EVs12. However, precautions are needed when assessing EV diameter by freeze-fracturing17. Since the fracture passes through the sample randomly, MV membranes are split into their equatorial and non-equatorial planes. To provide a rigorous method for analyzing the images of freeze-fractured and shadowed specimens, Hallett et al. showed that the mean vesicle diameter is 4/&#960; times the actual size of the vesicular diameter on the image17. Accounting for that, EVs from T24 cells were calculated to have a diameter of 304 nm, which fits in the MV&#39;s theoretical size distribution range of 100-1000 nm19.
Freeze-fracturing can supplement negative staining, the most extensively used TEM technique to visualize EVs. By negative staining, the sample is commonly chemically fixed, dried and attached to a TEM grid, and contrasted with uranyl solution. Without supporting media, EVs tend to collapse, which gives them a cup-shaped appearance. By freeze-fracturing, we show that MVs are spheres, which reflects their shape in extracellular spaces and body fluids12. By that, our results are also in agreement with observations of EVs in cryo-ultrathin sections20.
A crucial advantage of the freeze-fracture technique is its power to resolve the internal organization of the limiting membrane, which is a key factor in understanding how EVs are targeted to and interact with recipient membranes. Here, we analyzed the membranes of MVs, yet freeze-fracturing could reveal the membrane organization of any other population of EVs. MVs are formed by plasma membrane budding; therefore, plasma membrane proteins and protein clusters are expected to be found in the MV membrane. Our results supported that the MVs from T24 cells contained intramembrane particles, presenting integral membrane proteins. Based on particle distribution between the E-face and P-face, it is reasonable to expect that the particles observed in MVs are transmembrane proteins uroplakins, which are urothelial cell specific21,24. The observed particles were sparse, which is in accordance with previous studies reporting a reduction in uroplakins during urothelial carcinogenesis21,22,23. However, to further investigate the protein composition of MV membranes, the use of the freeze-fracture replica immune-labeling (FRIL) technique is recommended. FRIL is an upgrade of the presented freeze-fracture technique and is dedicated to revealing the identity of proteins in replicas by antibody recognition24,25. To sum up, the freeze-fracture technique is an electron microscopy technique suitable for the characterization of the EV limiting membrane, as well as the shape, size, and purity of the isolated EV fractions. The presented protocol can be used also for the assessment of other populations of isolated EVs; therefore, the freeze-fracturing technique merits inclusion in the International Society for Extracellular Vesicles guidelines for studies exploring the organization of EV limiting membranes.

Extracellular vesicles (EVs) are membrane-limited vesicles released from cells into the extracellular space. The three main populations of EVs are exosomes, microvesicles (MVs), and apoptotic bodies, which differ in their origin, size, and molecular composition1,2,3. The composition of EVs reflects the molecular profile of the donor cell and its physiological status (i.e., healthy or diseased)4,5. This gives EVs immense potential in the diagnosis, prognosis, and therapy of human diseases, and they have promising medical applications for the use in personalized medicine6,7,8.
EVs are mediators of intercellular communication. They contain biologically active proteins, lipids, and RNAs, which interfere with biological processes in the recipient cell and can change its behavior9,10. However, the composition of the EV limiting membrane is crucial for the interaction with the recipient cell membrane.
The sources of EVs are body fluids and conditioned culture media. To isolate an EV population, a suitable isolation technique must be used. For example, centrifugation at 10,000&#160;&#215;&#160;g&#160;yields a fraction enriched in MVs, whereas centrifugal forces of &#8805;100,000&#160;&#215;&#160;g yield a fraction enriched in exosomes11,12. The isolated fraction of EVs must be validated in terms of purity, size, and shape. For that purpose, the International Society for Extracellular Vesicles 2018 recommended three classes of high-resolution imaging techniques: electron microscopy, atomic force microscopy, and light microscopy-based super-resolution microscopy13. None of these techniques can provide information on the EV membrane interior.
Freeze-fracture electron microscopy is a technique of breaking frozen specimens to reveal their internal structures, particularly giving a view of the membrane interior. The steps of sample preparation are (1) rapid freezing, (2) fracturing, (3) making the replica, and (4) cleaning the replica14. In step 1, the sample is (optionally) chemically fixed, cryoprotected with glycerol, and frozen in liquid freon. In step 2, the frozen specimen is fractured in a freeze-fracture unit, which exposes the interior of the membrane bilayer. In step 3, the exposed fractured faces are shadowed with platinum (Pt) and carbon (C) to produce replicas. In step 4, the organic material is removed. The replica is analyzed in the transmission electron microscope (TEM). For accurate interpretation of the micrographs, one must follow guidelines for their proper orientation14,15. Briefly, the direction of shadows in the micrograph is a reference to orientate the micrograph (i.e., to determine the direction of Pt shadowing) and, consequently, to determine convex and concave shapes (Figure 1). Two interior views termed fractured faces of the membrane bilayer can be seen as a result of splitting the membrane by freeze-fracturing: the protoplasmic face (P-face) and the exoplasmic face (E-face). The P-face represents the membrane leaflet adjacent to the cell protoplasm, while the E-face represents the membrane leaflet adjacent to the extracellular space. Integral membrane proteins and their associations are seen as protruding intramembrane particles14,15.
Here, the goal is to apply the freeze-fracture technique to characterize MVs in terms of size, shape, and the structure of their limiting membrane. Here, we present a protocol for the isolation and freeze-fracturing of MVs originating from human invasive bladder cancerous urothelial cells.

<table><tbody><tr><td>A-DMEM</td><td>ThermoFisher Scientific, USA</td><td>12491015</td><td></td></tr><tr><td>Balzers freeze-fracture device</td><td>Balzers AG, Liechtenstein</td><td>Balzers BAF 200</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf, Germany</td><td>model 5810R</td><td></td></tr><tr><td>CO2 incubator HeraCell&amp;nbsp;</td><td>Heraues, Germany</td><td></td><td></td></tr><tr><td>Copper carriers</td><td>Balzers</td><td>BB 113 142-2</td><td>100+ mesh</td></tr><tr><td>Copper grids</td><td>SPI, United States</td><td>2010C</td><td></td></tr><tr><td>Culture flask T75</td><td>TPP, Switzerland</td><td></td><td>growing surface 75 cm2</td></tr><tr><td>F12 (HAM)</td><td>Sigma-Aldrich, Germany</td><td>21765029</td><td></td></tr><tr><td>FBS</td><td>Gibco, Life Technologies, USA</td><td>10500064</td><td></td></tr><tr><td>Glazed Porcelain Plate 6 Well</td><td>Fischer Sientific, United States</td><td>50-949-072</td><td></td></tr><tr><td>Glycerol</td><td>Kemika, Croatia</td><td>711901</td><td>final concentration 30 % (v/v)</td></tr><tr><td>Glutaradehyde&amp;nbsp;</td><td>Serva, Germany</td><td>23114.01</td><td>final concentration 2.5 % (v/v)</td></tr><tr><td>GlutaMAX</td><td>Gibco, Life Technologies</td><td>35050-079</td><td>final concentration 4 mM</td></tr><tr><td>Penicillin</td><td>Gibco, Life Technologies</td><td>15140163</td><td>final concentration 100 U/ml</td></tr><tr><td>Phase-contast inverted microscope</td><td>Nikon, Japan</td><td>model Eclipse</td><td></td></tr><tr><td>Rotor</td><td>Eppendorf, Germany</td><td>A 4-44</td><td></td></tr><tr><td>Rotor&amp;nbsp;</td><td>Eppendorf, Germany</td><td>F-34-6-38</td><td></td></tr><tr><td>Serological pipetts</td><td>TPP, Switzerland</td><td></td><td>50mL volume</td></tr><tr><td>Sodium hypochloride solution in water</td><td>Carlo Erba, Italy</td><td>481181</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica, Germany</td><td></td><td></td></tr><tr><td>Streptomycin</td><td>Gibco, Life Technologies</td><td>35050038</td><td>final concentration 100 mg/mL</td></tr><tr><td>Transmission electron microscope</td><td>Philips, The Netherlands</td><td>model CM100</td><td>working at 80 kV</td></tr><tr><td>Tweezers</td><td>SPI, United States</td><td></td><td></td></tr></tbody></table>

freeze-fracture electron microscopy for extracellular vesicle analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

MVs were isolated from the conditioned medium of cancer urothelial T24 cells after differential centrifugations. Following the protocol, the EV fraction was first detected after centrifugation at 10,000 &#215; g when it was seen as a white pellet (Figure 2G).
Next, the EVs were processed according to the above protocol and examined under TEM. Pt/C shadowing produced (Figure 4L) relatively large replicas that frequently broke up into smaller fragments during the cleaning step. Large replicas and their smaller fragments were picked on the TEM grid and compared under the microscope. The results showed no differences in the quality of the replica surfaces regardless of their size, and they can, therefore, all be used. Low magnification examinations showed regions of the replica with i) a homogeneous surface (background), ii) concave and convex spherical profiles (vesicles), and iii) regions of damaged surface (artifacts; Figure 5A). Typical artifacts were seen as ruptures, folds, and irregular dimmer shadows (i.e., replicas of ice-crystal deposits on the specimen). It is also important to note that no cell debris or cellular organelles were seen, confirming the purity of the sample (Figure 5B).
The isolated vesicles were commonly gathered either in clusters of three or more or were individually distributed (Figure 5B). The vesicles were spherical, which points to good preservation of the ultrastructure during isolation, fixation, and freezing (Figure 5C). &#34;Flat-ball&#34; and elongated vesicles (Figure 5C), which were seen occasionally, were presumably artifacts of preparation and were not included in the subsequent measurements of vesicle diameter. The diameter of the visible profiles was 238.5 nm (&#177;8.0 nm, n = 190), which, taking into account the correction factor proposed by Hallett et al.17, corresponds to the mean vesicle diameter of 304 nm (&#177;10 nm; Figure 5D). The size correlates to a diameter range of MVs between 100 nm and 1000 nm and proves the effectiveness of the used isolation protocol. The images of the vesicles together with diameter determination unequivocally confirmed that the EVs in the isolate were enriched with MVs.
The analysis of the replicas revealed the organization of P-face and E-face of the MV limiting membranes. MVs were observed as concave and convex round shapes (Figure 5C,E,F), reflecting the fracture plane. The convex shapes represent E-faces (i.e., fractured exoplasmic leaflet of the membranes; Figure 5E). The exoplasmic faces of the EVs had a smooth, uniform appearance. The concave shapes correspond to P-faces (Figure 5F). In the P-faces, a few protruding intramembrane particles were seen within smooth membranes (Figure 5C,F). This implies that MVs isolated from cancer urothelial T24 cells contain only a low amount of membrane proteins.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63550/63550fig01.jpg" /> 
Figure 1: Schematic presentation of membrane determination after freeze-fracturing. (A) Microvesicles bud from the plasma membrane into the extracellular space. (B) Microvesicle is limited with P- and E- membrane leaflet until freeze-fractioning, which splits the leaflets and exposes the interior views of leaflets, termed fractured faces. (C) After Pt/C shadowing, two fractured faces are discernible: the protoplasmic face (P-face) with a convex shape facing the cytoplasm (protoplasm) and an exoplasmic face (E-face) with a concave shape facing the extracellular space. Figure 1 was created using Biorender.com. <a href="https://www.jove.com/files/ftp_upload/63550/63550fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63550/63550fig02.jpg" /> 
Figure 2: Isolation of EVs. (A) T24 cells grown in a CO2 incubator are examined with (B) a light microscope to confirm their viability and confluence before MV isolation. (C) The cell culture medium is collected and (D) consecutively centrifuged at 300 &#215; g and at 2,000 &#215; g (E) each time the supernatant is collected. (F,G) After the centrifugation at 10,000 &#215; g, a white patch indicating a pellet is visible and marked. The supernatant is removed and (H) fixative is carefully added to the pellet without resuspending it. <a href="https://www.jove.com/files/ftp_upload/63550/63550fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63550/63550fig03.jpg" /> 
Figure 3: Freezing of EVs. (A) Air-dried clean copper carriers with a central pit are (B) marked before processing. Microvesicles are resuspended in glycerol to get a homogenous sample and (C) added to a central pit of a cooper carrier under a stereomicroscope. (D) One has to add a volume of the sample such that it forms a convex drop in the central pit. (E) Immediately before freezing, mix LN2-cooled freon with a metal rod to liquefy the freon. (F) Freeze the sample by submerging it in freon, and then (G) transfer it into LN2. (H) Carriers with the frozen sample can be collected into cryovials and stored in an LN2 Dewar container. <a href="https://www.jove.com/files/ftp_upload/63550/63550fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63550/63550fig04.jpg" /> 
Figure 4: Freeze-fracturing and making a replica. (A) Freeze-fracturing unit. Inside the unit chamber (B) is a platinum (Pt) and carbon (C) electron gun, a knife, and the sample table. (C) To start fracturing, carriers with the sample are transferred to the sample table, (D) the freeze-fracturing unit is cooled, and a vacuum is established. (E) Sectioning is done by motorized (F) movement of the knife, but fracturing is preferably done manually. (G) Sample before sectioning and (H) after fracturing. (I) Immediately after fracturing, the replica is made. (J) During this process, Pt is shadowed on the sample. Platinum shadowing is seen as a bright light (sparks) in the chamber. (K) Sample carriers are collected into a porcelain well filled with water. (L) Replica (arrow) floating in the sodium hypochlorite solution during the cleaning step. <a href="https://www.jove.com/files/ftp_upload/63550/63550fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63550/63550fig05.jpg" /> 
Figure 5: Electron micrographs of freeze-fractured and Pt/C shadowed MVs. (A) An overview of the freeze-fractured and shadowed EV pellet (i) at lower magnification. The homogeneous surface is background (ii), bright areas are ruptures in the replicas (iii), darker areas are folds of replicas (asterisk), and irregular dimmer shadows are due to ice crystals (two asterisks). (B) Cluster of round-shaped EVs with concave and convex surfaces. (C) High magnification of EVs. In convex fractures, which exhibit P-face of the EV membranes, intramembrane particles (arrows) and patches of a smooth surface (arrowheads) are seen. Extracellular vesicles with elongated (star) and flat-ball shapes (two stars) are found. (D) The mean diameter of isolated urothelial MVs is 304 nm &#177; 10 nm according to the size measurements as proposed by Hallett et al.17. Data are presented as mean &#177; SEM. (E,F) E-face and P-face of MVs. Legend: arrows = intramembrane particles in P-face, encircled arrows = the direction of Pt/C shadowing. Scale bars: A = 10 &#181;m, B-F = 400 nm. <a href="https://www.jove.com/files/ftp_upload/63550/63550fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis at JoVE.com

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

freeze-fracture-electron-microscopy-for-extracellular-vesicle-analysis

Research

JoVE Journal

Cancer Research

3.6K Views.  University of Ljubljana. In video presenteren we een protocol voor het in vitro breken van extracellulaire blaasjes afkomstig van blaaskankercellen. Extracellulaire blaasjes zijn membraanbeperkte blaasjes die afkomstig zijn van cellen die worden vrijgegeven in de extracellulaire ruimte en een rol spelen in de intercellulaire communicatie. Binnen extracellulaire blaasjes onderscheiden we drie populaties, exosomen, microvesicles en apoptotische lichamen, die verschillen door hun oorsprong, grootte en moleculaire samens...

Video: Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extraction of Extracellular Vesicles from Whole Tissue

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or prognostic biomarker assays, and likely serve as important players in the progression of a vast spectrum of diseases. Current research is focused on the characterization of vesicles secreted from multiple cell and tissue types in order to better understand the role of EVs in the pathogenesis of conditions including neurodegeneration, inflammation, and cancer. However, globally consistent and reproducible techniques to isolate and purify vesicles remain in progress. Moreover, methods for extraction of EVs from solid tissue ex vivo are scarcely described. Here, we provide a detailed protocol for extracting small EVs of interest from whole fresh or frozen tissues, including brain and tumor specimens, for further characterization. We demonstrate the adaptability of this method for multiple downstream analyses, including electron microscopy and immunophenotypic characterization of vesicles, as well as quantitative mass spectrometry of EV proteins.

Here, we provide a detailed protocol to isolate small extracellular vesicles (EVs) from whole tissues, including brain and tumor specimens. This method offers a reproducible technique to extract EVs from solid tissue for further downstream analyses.

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or ...

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;

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 ...

Biochemistry

Evaluation of the Storage Stability of Extracellular Vesicles

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical development, not only their pharmaceutical activity is important but also their production needs to be evaluated. In this context, research focuses on the isolation of EVs, their characterization, and their storage. The present manuscript aims at providing a facile procedure for the assessment of the effect of different storage conditions on EVs, without genetic manipulation or specific functional assays. This makes it possible to quickly get a first impression of the stability of EVs under a given storage condition, and EVs derived from different cell sources can be compared easily. The stability measurement is based on the physicochemical parameters of the EVs (size, particle concentration, and morphology) and the preservation of the activity of their cargo. The latter is assessed by the saponin-mediated encapsulation of the enzyme beta-glucuronidase into the EVs. Glucuronidase acts as a surrogate and allows for an easy quantification via the cleavage of a fluorescent reporter molecule. The present protocol could be a tool for researchers in the search for storage conditions that optimally retain EV properties to advance EV research toward clinical application.

Here we present a readily applicable protocol to assess the storage stability of extracellular vesicles, a group of naturally occurring nanoparticles produced by cells. The vesicles are loaded with glucuronidase as a model enzyme and stored under different conditions. After storage, their physicochemical parameters and the activity of the encapsulated enzyme are evaluated.

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical ...

Bioengineering

Labeling of Extracellular Vesicles for Monitoring Migration and Uptake in Cartilage Explants

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their cellular source, such as platelet lysate (PL). When used as treatment, EVs are expected to enter the target cells and effect a response from these. In this research, PL-derived EVs have been studied as a cell-free treatment for osteoarthritis (OA). Thus, a method was set up to label EVs and test their uptake on cartilage explants. PL-derived EVs are labeled with the lipophilic dye PKH26, washed twice through a column, and then tested in an in vitro inflammation-driven&#160;OA model for 5 h after particle quantification by nanoparticle tracking analysis (NTA). Hourly, cartilage explants are fixed, paraffined, cut into 6 &#181;m sections to mount on slides, and observed under a confocal microscope. This allows verification of whether EVs enter the target cells (chondrocytes) during this period and analyze their direct effect.

Here, we present a protocol to label platelet lysate-derived extracellular vesicles to monitor their migration and uptake in cartilage explants used as a model for osteoarthritis.

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their ...

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated to generate a carbon/platinum &#8220;cast&#8221; intended for examination by transmission electron microscopy. Specimens are subjected to ultrarapid freezing rates, often in the presence of cryoprotective agents to limit ice crystal formation, with subsequent fracturing of the specimen at liquid nitrogen cooled temperatures under high vacuum. The resultant fractured surface is replicated and stabilized by evaporation of carbon and platinum from an angle that confers surface three-dimensional detail to the cast. This technique has proved particularly enlightening for the investigation of cell membranes and their specializations and has contributed considerably to the understanding of cellular form to related cell function. In this report, we survey the instrument requirements and technical protocol for performing freeze-fracture, the associated nomenclature and characteristics of fracture planes, variations on the conventional procedure, and criteria for interpretation of freeze-fracture images. This technique has been widely used for ultrastructural investigation in many areas of cell biology and holds promise as an emerging imaging technique for molecular, nanotechnology, and materials science studies.

Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated ...

Biology

Size Exclusion Chromatography for Separating Extracellular Vesicles from Conditioned Cell Culture Media

Extracellular vesicles (EVs) are nano-sized lipid-membrane bound structures that are released from all cells, are present in all biofluids, and contain proteins, nucleic acids, and lipids that are reflective of the parent cell from which they are derived. Proper separation of EVs from other components in a sample allows for characterization of their associated cargo and lends insight into their potential as intercellular communicators and non-invasive biomarkers for numerous diseases. In the current study, oligodendrocyte derived EVs were isolated from cell culture media using a combination of state-of-the-art techniques, including ultrafiltration and size exclusion chromatography (SEC) to separate EVs from other extracellular proteins and protein complexes. Using commercially available SEC columns, EVs were separated from extracellular proteins released from human oligodendroglioma cells under both control and endoplasmic reticulum (ER) stress conditions. The canonical EV markers CD9, CD63, and CD81 were observed in fractions 1-4, but not in fractions 5-8. GM130, a protein of the Golgi apparatus, and calnexin, an integral protein of the ER, were used as negative EV markers, and were not observed in any fraction. Further, when pooling and concentrating fractions 1-4 as the EV fraction, and fractions 5-8 as the protein fraction, expression of CD63, CD81, and CD9 in the EV fraction was observed. The expression of GM130 or calnexin was not observed in either of the fraction types. The pooled fractions from both control and ER stress conditions were visualized with transmission electron microscopy and vesicles were observed in the EV fractions, but not in the protein fractions. Particles in the EV and protein fractions from both conditions were also quantified with nanoparticle tracking analysis. Together, these data demonstrate that SEC is an effective method for separating EVs from conditioned cell culture media.

The protocol here demonstrates that extracellular vesicles can be adequately separated from conditioned cell culture media using size exclusion chromatography.

Extracellular vesicles (EVs) are nano-sized lipid-membrane bound structures that are released from all cells, are present in all biofluids, and contain ...

Neuroscience

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

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 ...

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.

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 ...

Single Extracellular Vesicle Transmembrane Protein Characterization by Nano-Flow Cytometry

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and first-generation particle analysis to comprehensive multi-parameter measurements such as nano-flow cytometry (nFCM). nFCM is a form of flow cytometry that utilizes instrumentation specifically designed for nano-particle analysis, allowing for thousands of EVs to be characterized per minute both with and without the use of staining techniques. High resolution side scatter (SS) detection allows for size and concentration to be determined for all biological particles larger than 45 nm, while simultaneous fluorescence (FL) detection identifies the presence of labeled markers and targets of interest. Labeled subpopulations can then be described in quantitative units of particles/mL or as a percentage of the total particles identified by side scatter.
Here, EVs derived from conditioned cell culture media (CCM) are labeled with both a lipid dye, to identify particles with a membrane, and antibodies specific for CD9, CD63, and CD81 as common EV markers. Measurements of comparison material, a concentration standard and a size standard of silica nanospheres, as well as labeled sample material are analyzed in a 1-minute analysis. The software is then used to measure the concentration and size distribution profile of all particles, independent of labeling, before determining the particles that are positive for each of the labels.
Simultaneous SS and FL detection can be utilized flexibly with many different EV sources and labeling targets, both external and internal, describing EV samples in a comprehensive and quantitative manner.

The latest generation of EV characterization tools are capable of single EV analysis across multiple parameters simultaneously. Nano-flow cytometry measures all biological particles larger than 45 nm without labeling and identifies specific characteristics of subpopulations by a variety of fluorescent labeling techniques.

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and ...

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

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