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>

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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;</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&#160;</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&#160;</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

Nanyang Technological University

Research

JoVE Journal

Cancer Research

3.6K Views.  University of Ljubljana. 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.

Video: Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

A Preclinical Mouse Model of Osteosarcoma to Define the Extracellular Vesicle-mediated Communication Between Tumor and Mesenchymal Stem Cells

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under the influence of specific environmental signals, these adult stem cells can release paracrine mediators leading to accelerated tumor growth and metastasis. Defining the crosstalk between tumor and MSCs is of primary importance to understand the mechanisms underlying cancer progression and identify novel targets for therapeutic intervention. 
Cancer cells produce high amounts of extracellular vesicles (EVs), which can profoundly affect the behavior of target cells in the tumor microenvironment or at distant sites. Tumor EVs enclose functional biomolecules, including inflammatory RNAs and (onco)proteins, that can educate stromal cells to enhance the metastatic behavior of cancer cells or to participate in the pre-metastatic niche formation. In this article, we describe the development of a preclinical cancer mouse model that enables specific evaluation of the EV-mediated crosstalk between tumor and mesenchymal stem cells. First, we describe the purification and characterization of tumor-secreted EVs and the assessment of the EV internalization by MSCs. We then make use of a multiplex bead-based immunoassay to evaluate the alteration of the MSC cytokine expression profile induced by cancer EVs. Finally, we illustrate the generation of a bioluminescent orthotopic xenograft mouse model of osteosarcoma that recapitulates the tumor-MSC interaction, and show the contribution of EV-educated MSCs to tumor growth and metastasis formation. 
Our model provides the opportunity to define how cancer EVs shape a tumor-supporting environment, and to evaluate whether blockade of the EV-mediated communication between tumor and MSCs prevents cancer progression.

Direct injection of cancer-derived extracellular vesicles (EVs) leads to reprogramming of bone marrow supporting tumor progression; however, which cells mediate this effect is unclear. Herein, we describe a step-by-step protocol to investigate EV-mediated tumor-mesenchymal stem cell (MSC) interactions in vivo, revealing a crucial role for EV-educated MSCs in metastasis.

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under ...

Simple and Rapid Method to Obtain High-quality Tumor DNA from Clinical-pathological Specimens Using Touch Imprint Cytology

It is critical to determine the mutational status in cancer before administration and treatment of specific molecular targeted drugs for cancer patients. In the clinical setting, formalin-fixed paraffin-embedded (FFPE) tissues are widely used for genetic testing. However, FFPE DNA is generally damaged and fragmented during the fixation process with formalin. Therefore, FFPE DNA is sometimes not adequate for genetic testing because of low quality and quantity of DNA. Here we present a method of touch imprint cytology (TIC) to obtain genomic DNA from cancer cells, which can be observed under a microscope. Cell morphology and cancer cell numbers can be evaluated using TIC specimens. Furthermore, the extraction of genomic DNA from TIC samples can be completed within two days. The total amount and quality of TIC DNA obtained using this method was higher than that of FFPE DNA. This rapid and simple method allows researchers to obtain high-quality DNA for genetic testing (e.g., next generation sequencing analysis, digital PCR, and quantitative real time PCR) and to shorten the turnaround time for reporting results.

Obtaining high-quality genomic DNA from tumor tissues is an essential first step for analyzing genetic alterations using next generation sequencing. In this article, we present a simple and rapid method to enrich tumor cells and obtain intact DNA from touch imprint cytology specimens.

It is critical to determine the mutational status in cancer before administration and treatment of specific molecular targeted drugs for cancer patients. ...

Phospho Flow Cytometry with Fluorescent Cell Barcoding for Single Cell Signaling Analysis and Biomarker Discovery

Aberrant cell signaling plays a central role in cancer development and progression. Most novel targeted therapies are indeed directed at proteins and protein functions, and cell signaling aberrations may therefore serve as biomarkers to indicate personalized treatment options. As opposed to DNA and RNA analyses, changes in protein activity can more efficiently evaluate the mechanisms underlying drug sensitivity and resistance. Phospho flow cytometry is a powerful technique that measures protein phosphorylation events at the cellular level, an important feature that distinguishes this method from other antibody-based approaches. The method allows for simultaneous analysis of multiple signaling proteins. In combination with fluorescent cell barcoding, larger medium- to high-throughput data-sets can be acquired by standard cytometer hardware in short time. Phospho flow cytometry has applications both in studies of basic biology and in clinical research, including signaling analysis, biomarker discovery and assessment of pharmacodynamics. Here, a detailed experimental protocol is provided for phospho flow analysis of purified peripheral blood mononuclear cells, using chronic lymphocytic leukemia cells as an example.

Here, a protocol for medium- to high-throughput analysis of protein phosphorylation events at the cellular level is presented. Phospho flow cytometry is a powerful approach to characterize signaling aberrations, identify and validate biomarkers, and assess pharmacodynamics.

Aberrant cell signaling plays a central role in cancer development and progression. Most novel targeted therapies are indeed directed at proteins and ...

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 ...

Three-Dimensional Bone Extracellular Matrix Model for Osteosarcoma

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with diagnostic or prognostic difficulties. OS comprises genotypically and phenotypically heterogeneous cancer cells. Bone microenvironment elements are proved to account for tumor heterogeneity and disease progression. Bone extracellular matrix (BEM) retains the microstructural matrices and biochemical components of native extracellular matrix. This tissue-specific niche provides a favorable and long-term scaffold for OS cell seeding and proliferation. This article provides a protocol for the preparation of BEM model and its further experimental application. OS cells can grow and differentiate into multiple phenotypes consistent with the histopathological complexity of OS clinical specimens. The model also allows visualization of diverse morphologies and their association with genetic alterations and underlying regulatory mechanisms. As homologous to human OS, this BEM-OS model can be developed and applied to the pathology and clinical research of OS.

The bone extracellular matrix (BEM) model for osteosarcoma (OS) is well established and shown here. It can be used as a suitable scaffold for mimicking primary tumor growth in vitro and providing an ideal model for studying the histologic and cytogenic heterogeneity of OS.

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with ...

Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue and its role in local recurrence at the primary tumor site are unknown. Here we present a method for the decellularization, lyophilization, and fabrication of ECM hydrogels derived from murine mammary fat pads. Results are presented on the effectiveness of the decellularization process, and rheological parameters were assessed. GFP- and luciferase-labeled breast cancer cells encapsulated in the hydrogels demonstrated an increase in proliferation in irradiated hydrogels. Finally, phalloidin conjugate staining was employed to visualize cytoskeleton organization of encapsulated tumor cells. Our goal is to present a method for fabricating hydrogels for in vitro study that mimic the in vivo breast tissue environment and its response to radiation in order to study tumor cell behavior.

This protocol presents a method for decellularization and subsequent hydrogel formation of murine mammary fat pads following ex vivo irradiation.

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue ...

Evaluating Cell Death Using Cell-Free Supernatant of Probiotics in Three-Dimensional Spheroid Cultures of Colorectal Cancer Cells

This manuscript describes a protocol to evaluate cancer cell deaths in three dimensional (3D) spheroids of multicellular types of cancer cells using supernatants from Lactobacillus fermentum cell culture, considered as probiotics cultures. The use of 3D cultures to test Lactobacillus cell-free supernatant (LCFS) are a better option than testing in 2D monolayers, especially as L. fermentum can produce anti-cancer effects within the gut. L. fermentum supernatant was identified to possess increased anti-proliferative effects against several colorectal cancer (CRC) cells in 3D culture conditions. Interestingly, these effects were strongly related to the culture model, demonstrating the notable ability of L. fermentum to induce cancer cell death. Stable spheroids were generated from diverse CRCs (colorectal cancer cells) using the protocol presented below. This protocol of generating 3D spheroid is time saving and cost effective. This system was developed to easily investigate the anti-cancer effects of LCFS in multiple types of CRC spheroids. As expected, CRC spheroids treated with LCFS strongly induced cell death during the experiment and expressed specific apoptosis molecular markers as analyzed by qRT-PCR, western blotting, and FACS analysis. Therefore, this method is valuable for exploring cell viability and evaluating the efficacy of anti-cancer drugs.

Here methods are presented to understand anti-cancer effects of Lactobacillus cell-free supernatant (LCFS). Colorectal cancer cell lines show cell deaths when treated with LCFS in 3D cultures. The process of generating spheroids can be optimized depending on the scaffold and the analysis methods presented are useful for evaluating the involved signaling pathways.

This manuscript describes a protocol to evaluate cancer cell deaths in three dimensional (3D) spheroids of multicellular types of cancer cells using ...

Generation of Zebrafish Larval Xenografts and Tumor Behavior Analysis

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly visualizing the response to anti-cancer therapies (chemo, radiotherapy, and biologicals), angiogenesis and metastasis with single cell resolution, places the zebrafish xenograft model as a top choice to develop preclinical studies.

The zebrafish larval xenograft assay presents several experimental advantages compared to other models, but probably the most striking is the reduction of size scale and consequently time. This reduction of scale allows single cell imaging, the use of a relatively low number of human cells (compatible with biopsies), medium-high-throughput drug screenings, but most importantly enables a significant reduction of the time of the assay. All these advantages make the zebrafish xenograft assay extremely attractive for future personalized medicine applications.

Many zebrafish xenograft protocols have been developed with a wide diversity of human tumors; however, a general and standardized protocol to efficiently generate zebrafish larval xenografts is still lacking. Here we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Here, we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly ...

In Vivo Immunogenicity Screening of Tumor-Derived Extracellular Vesicles by Flow Cytometry of Splenic T Cells

Immunogenic cell death of tumors, caused by chemotherapy or irradiation, can trigger tumor-specific T cell responses by releasing danger-associated molecular patterns and inducing the production of type I interferon. Immunotherapies, including checkpoint inhibition, primarily rely on preexisting tumor-specific T cells to unfold a therapeutic effect. Thus, synergistic therapeutic approaches that exploit immunogenic cell death as an intrinsic anti-cancer vaccine may improve their responsiveness. However, the spectrum of immunogenic factors released by cells under therapy-induced stress remains incompletely characterized, especially regarding extracellular vesicles (EVs). EVs, nano-scale membranous particles emitted from virtually all cells, are considered to facilitate intercellular communication and, in cancer, have been shown to mediate cross-priming against tumor antigens. To assess the immunogenic effect of EVs derived from tumors under various conditions, adaptable, scalable, and valid methods are sought-for. Therefore, herein a relatively easy and robust approach is presented to assess EVs' in vivo immunogenicity. The protocol is based on flow cytometry analysis of splenic T cells after in vivo immunization of mice with EVs, isolated by precipitation-based assays from tumor cell cultures under therapy or steady-state conditions. For example, this work shows that oxaliplatin exposure of B16-OVA murine melanoma cells resulted in the release of immunogenic EVs that can mediate the activation of tumor-reactive cytotoxic T cells. Hence, screening of EVs via in vivo immunization and flow cytometry identifies conditions under which immunogenic EVs can emerge. Identifying conditions of immunogenic EV release provides an essential prerequisite to testing EVs' therapeutic efficacy against cancer and exploring the underlying molecular mechanisms to ultimately unveil new insights into EVs' role in cancer immunology.

In Vivo Immunogenicity Screening of Tumor-Derived Extracellular Vesicles by Flow Cytometry of Splenic T Cells

This manuscript describes how to assess in vivo immunogenicity of tumor cell-derived extracellular vesicles (EVs) using flow cytometry. EVs derived from tumors undergoing treatment-induced immunogenic cell death seem particularly relevant in tumor immunosurveillance. This protocol exemplifies the assessment of oxaliplatin-induced immunostimulatory tumor EVs but can be adapted to various settings.

Immunogenic cell death of tumors, caused by chemotherapy or irradiation, can trigger tumor-specific T cell responses by releasing danger-associated ...

Exploring Mitochondrial Energy Metabolism of Single 3D Microtissue Spheroids Using Extracellular Flux Analysis

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing cells as two-dimensional, single-cell monolayers (2D culture), spheroid cell culture promotes, regulates, and supports physiological cellular architecture and characteristics that exist in vivo, including the expression of extracellular matrix proteins, cell signaling, gene expression, protein production, differentiation, and proliferation. The importance of 3D culture has been recognized in many research fields, including oncology, diabetes, stem cell biology, and tissue engineering. Over the last decade, improved methods have been developed to produce spheroids and assess their metabolic function and fate.
Extracellular flux (XF) analyzers have been used to explore mitochondrial function in 3D microtissues such as spheroids using either an XF24 islet capture plate or an XFe96 spheroid microplate. However, distinct protocols and the optimization of probing mitochondrial energy metabolism in spheroids using XF technology have not been described in detail. This paper provides detailed protocols for probing mitochondrial energy metabolism in single 3D spheroids using spheroid microplates with the XFe96 XF analyzer. Using different cancer cell lines, XF technology is demonstrated to be capable of distinguishing between cellular respiration in 3D spheroids of not only different sizes but also different volumes, cell numbers,&#160;DNA content and type.
The optimal mitochondrial effector compound concentrations of oligomycin, BAM15, rotenone, and antimycin A are used to probe specific parameters of mitochondrial energy metabolism in 3D spheroids. This paper also discusses methods to normalize data obtained from spheroids and addresses many considerations that should be considered when exploring spheroid metabolism using XF technology. This protocol will help drive research in advanced in vitro spheroid models.

These protocols will help users probe mitochondrial energy metabolism in 3D cancer cell-line-derived spheroids using Seahorse extracellular flux analysis.

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing ...