This research was supported by the Bio &#38; Medical Technology Development Program of the National Research Foundation (NRF) &#38; funded by the Korean government (MSIP&#38;MOHW) (No. 2016M3A9B6904244). We also thank members of the Medicinal Bioconvergence Research Center for purification of exosome.

1. Block Preparation, Sectioning, Staining and Imaging of the Exosome
<ol>
	<li>Pellet the exosomes from culture supernatant of HCT116 cells by centrifugation at 100,000 x g for 1.5 h23. Remove the culture supernatant and carefully fix the purified exosome pellet with 1 mL of 2.5% glutaraldehyde in 0.1 M sodium cacodylate solution (pH 7.0) for 1 h at 4 &#176;C. For 0.1 M sodium cacodylate, dissolve 4.28 g cacodylic acid in 160 mL of distilled water (DW). Adjust pH to 7.4 with 0.1 M HCl then make up to 200 mL with distilled water.</li>
	<li>Remove the fixative and rinse the pellets with 1 mL of 0.1 M sodium cacodylate buffer at room temperature. Repeat three times with each change lasting 10 min.</li>
	<li>Post-fix the samples with 1 mL of 2% Osmium tetroxide for 1 h at 4 &#176;C.</li>
	<li>Remove the fixative and rinse three times with 0.1 M sodium cacodylate buffer every 10 min.</li>
	<li>Incubate for 10 min with a graded acetone series (50%, 60%, 70%, 80%, 90%, 95%, 100%, respectively) on the shaker.</li>
	<li>Remove the acetone and incubate the solution of 3:1 acetone:low viscosity embedding mixture for 30 min. The exosome pellet is in the tube.</li>
	<li>Remove the medium and add 1:1 acetone:low viscosity embedding mixture medium, then incubate for 30 min.</li>
	<li>Remove the medium and add 1:3 acetone:low viscosity embedding mixture medium, then incubate for 30 min.</li>
	<li>Remove the medium and add 100% low viscosity embedding mixture and incubate overnight at room temperature.</li>
	<li>Embed the sample in pure low viscosity embedding mixture using the embedding mold and bake for 24 h at 65 &#176;C.</li>
	<li>Prepare sections with 60 nm thickness through an ultra-microtome.</li>
	<li>Double-stain with 2% uranyl acetate for 20 min and lead citrate for 10 min. 
	For Reynolds lead solution, add 1.33 g of lead nitrate and 1.76 g of sodium citrate to a total of 50 mL distilled water.</li>
	<li>Observe the grid under transmission electron microscopy at 80 kV.</li>
	<li>Click &#34;Acquire&#34; and then click &#34;File&#34; and &#34;Save as&#34; in CCD camera system under the electron microscope at 80 kV. Follow automatic settings for the exposure time.</li>
</ol>
2. Immuno-staining of Sections and Imaging (Figure 1)
<ol>
	<li>Cut 60 nm ultrathin sections using an ultra-microtome, and collect on the nickel grid.</li>
	<li>Incubate grids in 50 &#181;L drops of 0.02 M glycine for 10 min to quench free aldehyde groups.</li>
	<li>Rinse in 100 &#956;L of DW three times each for 10 min. Incubate at room temperature for 1 h in PBS containing 1% BSA.</li>
	<li>Incubate grids in 50 - 100 &#181;L drops of anti KRS antibody24 (1:100 in PBS containing 0.1% BSA) for 1 h (If necessary, this step should be carried out at 4 &#176;C overnight.)</li>
	<li>Wash grids with five separate drops (50 &#181;L) of PBS containing 0.1% BSA for 10 min each.</li>
	<li>Transfer grids to drops of 2nd antibody for 1 h (anti-rabbit IgG conjugated to 10 nm gold particle (1:100) in PBS containing 0.1% BSA).</li>
	<li>Wash grids with five separate drops (50 &#181;L) of PBS containing 0.1% BSA each 10 min.</li>
	<li>Double-stain with 2% uranyl acetate for 20 min under dark conditions and with Reynold&#39;s lead citrate for 10 min, respectively.</li>
	<li>Click &#34;Acquire&#34; and then click &#34;File&#34; and &#34;Save as&#34; in CCD camera system under a TEM at 80 kV. Exposure time followed automatic settings.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56482/56482fig1.jpg" /> 
Figure 1: Process of sample preparation and immunostaining. (A) Double fixed exosome is dehydrated and infiltrated with low viscosity embedding mixture resin. (B) Resin embedding. (C) Ultra-thin section with diamond knife. (D) Setup for Immuno-gold labelling. The grids with the ultrathin sections are put on the liquid droplets on a parafilm, which is placed with the wet paper towel. Each section is incubated with droplets of 50 - 100 &#181;L antibody, then washed with buffer. (E) Double staining with uranyl acetate and lead citrate. The lid is covered with aluminum foil for heavy metal staining. (F) Staining solution is removed by filter paper. <a href="//cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56482/56482fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Negative Staining
<ol>
	<li>Glow-discharge the thin formvar/carbon film coated 200 mesh copper EM grids for 1 min by glow discharger.</li>
	<li>Fix purified exosomes with 1 mL of 2% Paraformaldehyde (PFA) for 5 min. 
	Caution: Paraformaldehyde fumes are toxic. All work should be done in a ventilated fume hood.</li>
	<li>Load 5 - 7 &#181;L exosome suspension solution on the grid and incubate for 1 min. If the concentration of exosome is too high, dilute the concentration to 1/2 - 1/5.</li>
	<li>Immediately stain with the ~20 drops of filtered 1% uranyl acetate (UA) solution on the surface of the EM grid by syringe.</li>
	<li>Remove the excess UA solution on the grid by contacting the grid edge with filter paper.</li>
	<li>Quickly rinse the grid with a drop of water. This step will remove the excess staining solution.</li>
	<li>Place the grid on the table by holding with tweezers, and cover the grid partially with a culture dish to dry for 10 min under room temperature.</li>
	<li>Store the grid in an EM grid box for future observation by a TEM at 80 kV.</li>
</ol>
4. Whole Mount for Immunostaining
<ol>
	<li>Glow-discharge the thin formvar/carbon film coated 200 mesh copper EM grids for 30 s.</li>
	<li>Fix purified exosomes with 1 mL of 2% Paraformaldehyde (PFA) for 5 min. 
	Caution: Paraformaldehyde fumes are toxic. All work should be done in a ventilated fume hood.</li>
	<li>Load 5 - 7 &#181;L fixed exosome solution on the grid and incubate for 5 min. Rinse with 100 &#181;L of PBS three times each for 10 min.</li>
	<li>Treat grids with 50 &#181;L of 0.05 M glycine for 10 min to quench free aldehyde groups.</li>
	<li>Transfer grids to a drop of blocking buffer (PBS containing 1% BSA) for 30 min.</li>
	<li>Incubate grids with 50 - 100 &#181;L anti PD-L1 antibody (1:100 in PBS containing 0.1% BSA) for 1 h (if necessary, this step should be carried out at 4 &#176;C overnight).</li>
	<li>Wash grid with five separate drops (50 &#181;L) of PBS containing 0.1% BSA for 10 min each.</li>
	<li>Transfer grid to a drop of 2nd antibody for 1 h. Anti-mouse IgG conjugated to 9 - 11 nm gold particle diluted at 1:100 in PBS containing 0.1% BSA.</li>
	<li>Wash grid with five separate drops (50 &#181;L) of PBS containing 0.1% BSA for 10 min each. Wash grid with two separate drops (50 &#181;L) of DW. Perform negative staining with 2% uranyl acetate as described from 3.4 to 3.8.</li>
	<li>Store the grid in an EM grid box for future observation by a TEM at 80 kV.</li>
</ol>

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The authors have nothing to disclose.

This article presents a protocol for observing the detailed exosome&#39;s structure and labeling of its specific proteins. Negative staining has been considered as the best method for exosome imaging17. This conventional technique has shown exosomes&#39; cup-shaped structure. However, this cup shape is a form of artefact that can occur due to the drying process. Cryo-TEM results have shown that exosomes have a perfectly spherical structure in aqueous solution25,26. The cryo-TEM technique is a very powerful method for examining the natural structure, but it is difficult to apply the immuno-gold method. Raso and co-workers pointed out drying in the conventional method (whole mount-negative staining method) resulted in a cup-shaped morphology. In this study, the standard method for cell preparation (routine EM; fixation, dehydration, embedding, and sectioning) was applied to reduce the exosome drying effect. The routine EM using plastic embedding can avoid or reduce artefacts (changes in volume and shape) caused by denaturation. For chemical fixation, glutaraldehyde (GA) was used for the cross-linking (covalent interactions between amino groups). Usually, osmium tetroxide (OsO4) is used for fixation of lipids as well as improved contrast.
In this study, negative staining helped to confirm the typical morphology of exosomes that has been reported in previous papers21,27,28,29. In addition to negative staining, block sectioning is also a good method for observation of exosomes (Figure 2). While sectioned images showed the inner structure of exosome, in contrast negative stained TEM showed mainly the surface of the exosome. In sectioned images (Figure 3), the vesicles showed the lumen structure that is known as a structural feature of exosomes30. In this protocol, we used block sectioning, immuno-staining, and TEM imaging. Specially, the block sectioning is useful for analysis at a later time when images may be required at different magnifications, for different microscopy technique, and for other analyses such as immuno-gold labelling. After sectioning, sections on the grid can be stored in the grid box, which can be used for immuno-staining for classification of exosomes. For example, we detected the known exosome markers14 in the isolated exosomes to study the functions of exosomes.
Immunogold labeling methods have some technical difficulties, including no labeling or high background labeling. When experiencing no labeling, we recommend checking the primary antibody concentration and primary incubation times. In the case of low concentration of primary antibodies, antibody titration may be helpful to find the optimized immunoreactivity. It may also be helpful to increase the incubation times to 4 &#176;C overnight. In contrast, if the concentration of the primary antibody is too high or the incubation time is too long at the high temperature, the background signal will be increased.

Extracellular vesicles (EVs) are lipid bilayer vesicles secreted by cells, and their size ranges between 30 and 300 nm. EVs were first reported in 1978, with evidence from vesicles of patients with Hodgkin&#39;s disease1. These vesicles were reported to be 40 to 120 nanometers in size. In 1980, it was reported that EVs are involved in the coagulation system and thrombosis, formation of a blood clot inside a blood vessel in cancer patients2. After 30 years, EVs have been reported to be an important factor to promote tumor invasion, immune escape, and angiogenesis3. In addition, the functions of EVs have been studied as regulators in cellular interactions in the areas of inflammation, immune disorder, neurological disease, and cancer4. Since EVs contain specific biomolecules such as protein, mRNA, and microRNA5, their potential application in diagnostics and therapeutics has been analyzed6,7. EVs are a category composed of subgroups including exosomes, prostasomes, oncosomes, dexosomes, microparticles, promininosomes, argosomes, and exosome-like vesicles, depending on their cellular origin and biological function3. In addition, based on their biogenesis, these EVs can be divided into microvesicles (shed microvesicles, 100 - 1000 nm) and exosomes (30 - 300 nm)8,9. Among these, the exosome has been reported as a cell communicator for immune response10, cancer11,12, and infectious disease13.
Interest in the exosomes as a biomarker for early diagnosis is increasing rapidly, and the purification and characterization of exosomes must be accompanied with molecular imaging techniques. The varying sizes and morphologies of exosomes, depending on their origin and function14, can be distinguished by microscopy techniques with high resolution, such as electron microscopy. Most exosomes were visualized by negative stained Transmission Electron Microscopy (TEM)15,16,17, and these results were confirmed by immunolabeling of a specific protein in whole mount vesicle18. Several research groups have reported the structure through Scanning Electron Microscopy, Atomic Force Microscopy19,20, and Cryo-TEM21,22. However, while these techniques are useful for studying the exosome structure, they are insufficient for observing the position of specific proteins located inside the exosome. Therefore, we introduced a protocol for imaging exosomes with a specific protein&#39;s labelling. We applied block preparation, ultrathin sectioning, and immunostaining for ascertaining the protein&#39;s detailed location in the exosome. This was compared with negative staining and whole mount immunostaining, which is traditionally used for characterization of the exosome.

<table border="0" cellpadding="0" cellspacing="0" width="583">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Glutaraldehyde</td>
			<td style="width:81px;">EMS</td>
			<td style="width:119px;">16200</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Sodium cacodylate&#160;</td>
			<td>EMS</td>
			<td style="width:119px;">12300</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Osmium tetroxide 1 g crystal</td>
			<td>EMS</td>
			<td style="width:119px;">19100</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">acetone</td>
			<td>JUNSEI</td>
			<td>1B2031</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Spurr medium</td>
			<td>TED PELLA</td>
			<td style="width:119px;">18108</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Ultra-microtome</td>
			<td style="width:81px;">Leica</td>
			<td style="width:119px;">UCT</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Uranyl acetate</td>
			<td>EMS</td>
			<td style="width:119px;">22400</td>
			<td>Hazardous chemical</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Lead citrate</td>
			<td>EMS</td>
			<td style="width:119px;">17900</td>
			<td></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Transmission Electron Microscopy</td>
			<td style="width:81px;">Hitachi</td>
			<td style="width:119px;">H7600</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">nickel grid</td>
			<td style="width:81px;">EMS</td>
			<td style="width:119px;">G200-Ni</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Copper grid</td>
			<td>EMS</td>
			<td style="width:119px;">G200-Cu</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">glycine</td>
			<td>SIGMA</td>
			<td>022K5404</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Phosphate buffer saline</td>
			<td style="width:81px;">SIGMA</td>
			<td style="width:119px;">P4417</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Bovine serum albumin</td>
			<td style="width:81px;">Aurion</td>
			<td style="width:119px;">25557</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">1st Antibody</td>
			<td style="width:81px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">purification in manually&#160;</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Purified anti-human CD274 (B7-H1, PD-L1) Antibody</td>
			<td style="width:81px;">BioLegend</td>
			<td style="width:119px;">329710</td>
			<td style="width:167px;">Whole mount Immumogold</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Purified Mouse IgG2b, &#954; Isotype Ctrl</td>
			<td style="width:81px;">BioLegend</td>
			<td style="width:119px;">401202</td>
			<td style="width:167px;">Whole mount Immumogold (Control)</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">2nd Anti-mouse igG conjugated 9-11 nm gold particle</td>
			<td style="width:81px;">sigma</td>
			<td style="width:119px;">G7652</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">silver paint</td>
			<td style="width:81px;">CANS</td>
			<td style="width:119px;">CTP100</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">aluminum stub</td>
			<td style="width:81px;">EMS</td>
			<td style="width:119px;">75622</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">FIB-SEM</td>
			<td style="width:81px;">Zeiss</td>
			<td style="width:119px;">AURIGA</td>
			<td></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Transmission Electron Microscopy</td>
			<td style="width:81px;">JEOL</td>
			<td style="width:119px;">JEM2200FS</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Glow discharger</td>
			<td style="width:81px;">JEOL</td>
			<td style="width:119px;">JFC1100E</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Carbon grid</td>
			<td>EMS</td>
			<td style="width:119px;">121119</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Paraformaldehyde</td>
			<td>EMS</td>
			<td style="width:119px;">19210</td>
			<td></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Formvar carbon coated Copper Grid (200 meh)</td>
			<td style="width:81px;">EMS</td>
			<td style="width:119px;">FCF200-CU</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Formvar carbon coated&#160; Nickel Grid (200 mesh)</td>
			<td style="width:81px;">EMS</td>
			<td style="width:119px;">FCF200-NI</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

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.

Currently, exosomes are classified into size and shape categories by transmission electron microscopy. Figure 2 shows negative stained exosome and immune-labelled exosome in whole mount status. Figure 3 shows sectioned exosome and immuno-labelled exosomes after thin sectioning. Immuno-gold staining using antibodies of specific proteins is used to positively identify an exosome and classify the types of proteins in the exosome. The protocol in this paper uses whole mount immunostaining and immunostaining with plastic sectioned exosome.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56482/56482fig2.jpg" /> 
Figure 2: Negative staining and whole mount immuno-staining. (A) Exosome morphology is observed by negative staining. Exosomes shows their cup-shaped morphology. (B, C) Whole mount immuno-gold staining shows the location of specific protein (anti human CD274; PD-L1) in exosome. White arrows indicate the location of the gold. Black arrows indicate the location of the background signal. (D) Negative control of immunostaining result. Isotype control was used by primary antibody in this immunostaining process. Scale bar = 100 nm.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56482/56482fig3.jpg" /> 
Figure 3: Electron micrograph of sectioned exosomes. (A) Exosomes&#39; round shape morphology. (B) Boxed exosome using the &#34;boxer&#34; tool in EMAN program. (C, D) Immuno-gold labelled exosome with heavy metal staining. Black arrows indicate gold particles (A-D) Scale bar = 100 nm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56482/56482fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy at JoVE.com

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

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 ...

sample-preparation-imaging-exosomes-transmission-electron

Nanyang Technological University

Research

JoVE Journal

Biology

38.6K Views.  Asan Medical Center. 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.

Video: Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

Do's and Don'ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state by transmission electron microscopy (TEM). This can be achieved with very low quantity of protein and in the buffer of choice, without the use of any stain, which is very useful to determine structure-function correlations of macromolecules. When combined with single-particle image processing, the technique has found widespread usefulness for 3D structural determination of purified macromolecules. 
The protocol presented here explains how to perform cryoEM and examines the causes of most commonly encountered problems for rational troubleshooting; following all these steps should lead to acquisition of high quality cryoEM images. The technique requires access to the electron microscope instrument and to a vitrification device. Knowledge of the 3D reconstruction concepts and software is also needed for computerized image processing. Importantly, high quality results depend on finding the right purification conditions leading to a uniform population of structurally intact macromolecules. 
The ability of cryoEM to visualize macromolecules combined with the versatility of single particle image processing has proven very successful for structural determination of large proteins and macromolecular machines in their near-native state, identification of their multiple components by 3D difference mapping, and creation of pseudo-atomic structures by docking of x-ray structures. The relentless development of cryoEM instrumentation and image processing techniques for the last 30 years has resulted in the possibility to generate de novo 3D reconstructions at atomic resolution level.

This paper describes the cryo-electron microscopy methodology, which is used to obtain high quality microscopic images of macromolecules in their near-native state. The method yields images suitable for further computerized processing using the single particle approach, devised to generate the 3D reconstruction of a macromolecule.

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state ...

Preparation and Observation of Thick Biological Samples by Scanning Transmission Electron Tomography

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It also describes an imaging method for studying the 3D structure of thick biological specimens by scanning transmission electron tomography. The sample preparation protocol is based on conventional methods in which the sample is fixed using chemical agents, treated with a heavy atom salt contrasting agent, dehydrated in a series of ethanol baths, and embedded in resin. The specific imaging conditions for observing thick samples by scanning transmission electron microscopy are then described. Sections of the sample are observed using a through-focus method involving the collection of several images at various focal planes. This enables the recovery of in-focus information at various heights throughout the sample. This particular collection pattern is performed at each tilt angle during tomography data collection. A single image is then generated, merging the in-focus information from all the different focal planes. A classic tilt-series dataset is then generated. The advantage of the method is that the tilt-series alignment and reconstruction can be performed using standard tools. The collection of through-focal images allows the reconstruction of a 3D volume that contains all of the structural details of the sample in focus.

This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

This report describes a protocol for preparing thick biological specimens for further observation using a scanning transmission electron microscope. It ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.