This work was funded by two ANR grants (ANR-11-BSV8-016 and ANR-10-IDEX-0001-02). We also acknowledge the PICT-IBiSA for providing access to chemical imaging equipment.

Caution: Consult the material safety data sheets (MSDSs) of the various reagents before using them. Several of the chemicals used during sample preparation are toxic, carcinogenic, mutagenic, and/or reprotoxic. Use personal protective equipment (gloves, lab coat, full-length pants, and closed-toe shoes) and work under a fume hood while handling the sample. Sectioning the sample with an ultramicrotome involves the use of sharp instruments, and careful use of these tools is mandatory. In the steps described below, the authors assume that the sample is a cell culture sample. The protocol may need to be modified according to the sample type, especially the centrifugation steps.
1. Preparation of Buffers and Mixtures
<ol>
	<li>Prepare phosphate-buffered saline (PBS) solution (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.76 mM KH2PO4).</li>
	<li>Prepare solutions containing 30%, 50%, and 70% ethanol in PBS for sample dehydration. Perform sample dehydration by gradually incubating the sample in increasing ethanol concentrations.</li>
	<li>Prepare epoxy resin by mixing bisphenol-A-(epichlorohydrin) (20 mL), dodecenyl succinic anhydride (9 mL), methyl-5-norbornene-2,3-dicarboxylic (11 mL), and 2,4,6-tris(dimethylaminomethyl)phenol (0.7 mL); other mixing ratios can be used depending on the desired hardness of the resin.</li>
	<li>Prepare solutions containing 30%, 50%, and 70% epoxy resin in ethanol for sample embedding. Perform resin embedding by gradually incubating the sample with increasing resin concentrations. 
	NOTE: The volumes of the prepared solutions depend on the type of sample that is studied. More solutions are needed for tissues than for cell cultures.</li>
</ol>
2. Fixation and Coloration of the Sample
<ol>
	<li>Centrifuge the cell culture for 5 min at 5,000 x g. Perform all of the following centrifugations using the same conditions.</li>
	<li>Discard the supernatant and fix the cell pellet by resuspending it in PBS (1 mL) with 4% paraformaldehyde and 2% glutaraldehyde. Incubate it at room temperature for 30 min (Figure 1A). Alternatively, fix the cells at 4 &#176;C overnight or over the weekend.</li>
	<li>After centrifugation, remove the supernatant and wash the pellet 3 times with PBS (1 mL) to completely remove the fixative agents.</li>
	<li>Post-fix the sample with 1% OsO4 in PBS (1 mL) for 30 min at room temperature.</li>
	<li>After centrifugation, remove the supernatant and wash the pellet 3 times with PBS (1 mL) to completely remove the OsO4. Deactivate the OsO4 in oil and discard.</li>
	<li>Add 2% uranyl acetate in PBS (1 mL) as a contrast agent and incubate the sample for 30 min at room temperature (Figure 1B).</li>
	<li>After centrifugation, remove the supernatant and wash the pellet 3 times with PBS (1 mL) to remove the unbound uranyl acetate; uranyl acetate contains radioactive elements and should be disposed of in a bin dedicated to radioactive waste. 
	NOTE: The incubation times can be modified according to the size of the sample. For example, fixation and staining of thicker samples, such as tissues samples, require longer incubation times. Longer incubation times may also be better for the dehydration and the embedding steps. Uranyl acetate can be replaced by a &#34;uranyl-less&#34; solution that contains Gadolinium salts.</li>
</ol>
3. Sample Dehydration
<ol>
	<li>After centrifugation, discard the supernatant and incubate the sample for 30 min at 4 &#176;C in PBS (1 mL) with 30% ethanol (Figure 1C).</li>
	<li>Repeat step 3.1 using solutions with gradually increasing ethanol concentrations (50%, 70%, and up to 100%). During these dehydration steps, maintain the sample at 4 &#176;C for sample preservation purposes.</li>
	<li>After centrifugation, discard the supernatant and resuspend the pellet in pure ethanol (1 mL); incubate overnight at 4 &#176;C.</li>
</ol>
4. Resin Embedding
<ol>
	<li>After centrifugation, discard the supernatant and resuspend the pellet in ethanol with 30% epoxy resin (1 mL) for 30 min at RT (Figure 1D).</li>
	<li>Repeat step 4.1 using solutions with gradually increasing epoxy resin concentrations (50%, 70%, and up to 100%).</li>
	<li>After centrifugation, discard the supernatant and resuspend the pellet in pure epoxy resin (1 mL); incubate overnight at RT. The presence of hardening DMP-30 might trigger the polymerization of the resin; if this happens, prepare two batches of epoxy resin, one with and one without DMP-30.</li>
	<li>After centrifugation, resuspend the pellet in 200 &#181;L of pure epoxy resin (containing the hardener) in a plastic capsule; the sample should be at the bottom of the plastic capsule.</li>
</ol>
5. Resin Polymerization
<ol>
	<li>Incubate the plastic capsule containing the sample in an incubator set at 60 &#176;C for 48 h; other kinds of resin might polymerize with different settings.</li>
	<li>Remove the capsule from the incubator and verify that the resin has polymerized; the resin should be hard if pressed against a hard surface.</li>
	<li>Add pure epoxy resin (containing the hardener) on top of the polymerized resin containing the sample; this extra layer of resin will make it easier to cut the sample. Incubate the capsule at 60 &#176;C for 48 h.</li>
</ol>
6. Sample Sectioning
<ol>
	<li>Mount the sample upside-down in a specimen holder so that the sample is towards the top and fix it on the trimming block of the ultramicrotome. Tightly lock the lever to secure the trimming block to the ultramicrotome.</li>
	<li>Using a razor blade or a similar sharp cutting instrument, cut out the plastic capsule to separate it from the polymerized resin. Cut the resin so that the sample is contained in resin in roughly the shape of a pyramid. There is no need to remove the entire plastic capsule; only remove the part covering the polymerized sample.</li>
	<li>Take the capsule and the specimen holder off of the trimming block and install them on the moving arm of the ultramicrotome.</li>
	<li>Install a trimming knife on the knife block and accurately trim the faces of the pyramid. The use of binoculars is advised to give a perfect shape to the pyramid; the better the pyramid, the better the sections will be.</li>
	<li>Take off the trimming knife and replace it with a histology knife that can be used to cut sections that are thicker than 0.5 &#181;m.</li>
	<li>After filling the cuvette of the histology knife with the correct amount of water, bring the cutting edge of the knife to the surface of the sample and use the binoculars to adjust the capsule pitch angle so that the knife will slice perpendicularly to the pyramid.</li>
	<li>Adjust the cutting speed according to the desired section thickness to recover homogeneous sections.</li>
	<li>Let the section float over the water and fish it with a loop.</li>
	<li>Move the loop towards an electron microscopy copper grid that has been placed on top of filter paper. 
	NOTE: The grid should have been treated previously to increase its hydrophilicity. This can be performed with a grid-coating pen.</li>
	<li>Due to water absorption by the filter paper, let the section stick to the grid. Let it dry for few minutes before inserting it into the electron microscope.</li>
</ol>
7. Design of the Through-focal Tilt-series
<ol>
	<li>Collect through-focal images at constant focus intervals. 
	NOTE: The through-focal tilt-series consist of a set of through-focal images that are collected at each tilt angle of the tilt-series. The whole focus amplitude should be sufficient to cover the whole sample thickness.</li>
	<li>Determine the value of the focus interval. If the interval is equal to the depth-of-field, collect the entire sample in focus; this setting represents the highest sampling possible. If the interval is greater than the depth-of-field, some parts of the sample will not be acquired in focus, meaning that some parts of the sample will not be collected in focus.</li>
	<li>Calculate the depth-of-field of an electron beam from the electron&#39;s wavelength and the convergence semi-angle of the electron beam. Whereas the wavelength of electrons is directly linked to the acceleration voltage, use the convergence semi-angle provided by the electron microscope manufacturer or determine it experimentally from the diffraction pattern of a known specimen16. Because electrons accelerated at a wavelength &#955; hit the sample with a convergence semi-angle of &#945; in an electron microscope, use the following equation to determine the depth-of-field (DOF): 
	 
	<img alt="Equation" src="/files/ftp_upload/55215/55215eq1.jpg" /></li>
	<li>Design the focal series so that the whole focus amplitude will cover the sample at its maximal apparent thickness (i.e., at its highest tilt). 
	NOTE: A 0.75 &#181;m-thick sample will have an apparent thickness of 2.19 &#181;m at a 70&#176; tilt, so the focus amplitude should be greater than 2.19 &#181;m. For this sample, if the focus interval determined above is equal to 50 nm, it will require 44 images (2.19 &#181;m/50 nm) to cover the sample at its maximum apparent thickness. The whole focus amplitude is equal to the value of the focus interval multiplied by the number of focal steps. At the highest tilt (&#952;), the sample apparent thickness is defined as the following: 
	 
	<img alt="Equation" src="/files/ftp_upload/55215/55215eq2.jpg" /></li>
</ol>
8. Sample Imaging
NOTE: It is beyond the scope of this protocol to describe how to set up an electron microscope in STEM mode; most of this information can be found in the reference manual provided by the electron microscope manufacturer. However, note the following: i) the spot size should be chosen according to the desired magnification; ii) the Ronchigram must be well-aligned; and iii) in BF mode, the camera length must be adjusted to select unscattered electrons while maintaining good illumination of the detector. Resin sections deposited on electron microscopy copper grids should be illuminated before image acquisition to prevent shrinkage. Resin shrinkage and charging effects might be confounded during image acquisition. The use of an objective lens aperture might improve sample stability when charging effects occur. However, in the specific case of STEM setups, the objective aperture is not compatible with HAADF imaging mode, and very small objective apertures are not compatible with BF imaging mode.
<ol>
	<li>Using low magnification (around 20,000X in STEM), browse through the sample and locate the region of interest.</li>
	<li>Adjust the eucentric height (Z-axis) so that the region of interest does not drift away while rotating the sample.</li>
	<li>Verify that the object of interest will remain visible throughout the entire tilt range.</li>
	<li>Because of the substantial number of images acquired during a through-focal tilt-series, use an automatic collection software. Either use a commercial solution or design a custom data collection software if desired. Ensure that the software program is able to perform the three following steps:
	<ol>
		<li>Track and focus as in a standard tilt-series collection scheme.</li>
		<li>Ensure that the through-focal images overlap with the Z-position of the sample.</li>
		<li>Automatically acquire a series of through-focal images at each tilt angle.</li>
	</ol>
	</li>
	<li>Give the software program the main parameters of the through-focal tilt-series (i.e., focal intervals, focal step, minimum tilt angle, maximum tilt angle, tilt step, and scanning speed).</li>
	<li>Start the image collection process and wait for the software program to finish.</li>
</ol>
9. Image Processing
NOTE: In this protocol, the through-focal tilt-series processing is performed using ImageJ plugins17. Supplementary Data 1 shows an ImageJ macro for the alignment step (Turboreg) and for merging in-focus information (Extended Depth of Field) for a through-focal tilt-series designed with three focus values.
<ol>
	<li>Align the images collected at the same tilt angle (i.e., the through-focal images). Since images do not have the same in-focus information, they do not harbor the same features for alignment; perform alignment by aligning neighboring images two by two (i.e., following a pyramidal hierarchy, with the closest focus values first) using Turboreg18, an ImageJ plugin.</li>
	<li>Consistently verify the alignment of the through-focal images at each tilt angle. Stack the aligned through-focal images to improve visual inspection. 
	NOTE: Only the zone that is in focus should move while browsing through the stack. Accurate alignment is the most important thing, since the following step consists of the recovery of the information that is in focus from different images.</li>
	<li>Merge the information that is in focus using Extended Depth of Field19, an ImageJ plugin; this will eventually generate a single image from the stack of images. Several settings can be used during the depth-of-field recovery, but use the highest settings to preserve the image information. 
	NOTE: The tilt-series is now composed of a single image per tilt angle, each image containing the in-focus information recovered from the through-focal images.</li>
	<li>Process the data. 
	NOTE: Standard tools for alignment and reconstruction can be used since the data has now been reduced to a standard tilt-series. In this protocol, TomoJ is used for data alignment and data reconstruction20,21. More details about how to use TomoJ can be found in the original publication22.</li>
	<li>Perform fine alignment of the tilt-series. 
	NOTE: A good strategy is to use gold beads as fiducial markers to accurately track shifts between images. Local minima can also be used if gold beads cannot be added to the sample.</li>
	<li>Perform a 3D reconstruction of the aligned data; several algorithms are available for performing the reconstructions in real space or Fourier space, each with advantages and limitations.</li>
	<li>If the final alignment of the tilt-series is poor, make some improvements by optimizing the initial steps. Reiterate the alignment steps if the final 3D reconstruction appears blurred or deformed; in practice, the greater the number of collected focal planes, the better the contrast and details of the 3D reconstruction. 
	NOTE: Several software programs can be used to process the through-focal tilt-series. However, the tools used in this protocol were chosen because, in our experience, they performed the best. Turboreg18 performed better in terms of aligning the images when a gradient of focus was present. More information can be found on the dedicated website23. The Extended Depth of Field plugin19 performed very well in terms of recovering the in-focus information from different images, whereas other tools resulted in visible pixels modifications. More information, especially regarding script writing, can be found on this dedicated website24.</li>
</ol>

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We have no conflict of interest to disclose.

In this article, we present a conventional sample preparation protocol along with a step-by-step guide for performing 3D analysis on thick biological samples using STEM through-focal tomography. Resin embedding of biological specimens has been used for decades26, and alternative protocols that are adapted to different kinds of samples can be found throughout the literature27. In contrast, imaging thick specimens in STEM using through-focus methods is a new undertaking, and ...

Since the early 1970s, tomography approaches in transmission electron microscopy (TEM) have been widely used for the structural characterization of biological specimens1,2,3. The appeal of transmission electron tomography was that it could be used to study a wide range of biological structures at the nanometer scale, from the architecture of cells and the ultrastructure of organelles to the structure of macromolecular complexes and proteins. Nevertheless, transmission electron tomography could not be used to study very thick samples (greater than 0.5 &#181;m). Indeed, thick specimens produce too many scattered electrons, generating low signal-to-noise ratio (SNR) images. In addition, tomography involves the collection of images of tilted specimens, with the apparent thickness of the sample increasing with the tilt angle. Even though inelastic scattering can be filtered out using energy filters, the critical amount of electrons needed for high SNR images is barely reached in TEM. Hence, thick biological specimens have only been studied using sectioning4.
Some samples cannot be sliced: some might degrade when they are cut, and others need to be studied in their entirety in order to understand their complexity. An alternative approach is to use TEM in scanning mode5,6,7,8. In scanning transmission electron microscopy (STEM), the optical path of the electrons is different from that in conventional TEM. Electrons passing through the sample without scattering can be collected on the optical axis with a bright-field (BF) detector9, whereas those scattered elastically can be collected at a certain angle from the optical axis with a dark-field (DF) detector. The other advantage of STEM is that the focused electron beam is scanned at the surface of the sample, enabling the pixel-by-pixel collection of the images. Even though the electron beam broadens while passing through the sample10, this particular collection scheme is less sensitive to inelastically scattered electrons than conventional TEM. Furthermore, there is no lens post-specimen in STEM, thereby avoiding the chromatic aberrations that can occur in TEM. The camera length can be adjusted so that the BF detector mainly detects unscattered electrons. Using the DF detector to study thick samples is not recommended because of multiple scattering, which produces inaccurate images. Instead, the BF detector can be used11. While STEM can produce high SNR images, it has a relatively low depth-of-field because of the relatively high beam convergence compared to TEM, decreasing the amount of in-depth information that can be recovered from thick specimens. In the case of aberration-corrected STEM microscopes, where the convergence angle can be as high as 30 mrad, the depth-of-field can be low enough so that the information that is in focus originates from a focal plane of only a few nanometers. Setting up the electron beam in parallel mode enhances the depth-of-field of the electron beam to the detriment of the resolution12. However, this setup is not always possible.
Whenever it is necessary to use a convergent electron beam, one must use techniques that enhance the depth-of-field of the electron beam when studying thick samples. Recent studies have reported the acquisition of multiple images at various focal planes throughout the sample to recover the maximum amount of in-focus information13,14. The two studies describe different ways to handle the information from the different focal planes. Hovden et al. combined in Fourier space the images that were collected at various focal planes, and the final reconstruction was obtained directly from the 3D inverse Fourier transform13. In contrast, Dahmen et al. developed a convergent beam reconstruction engine to reconstruct in real space the 3D volume from the various focal planes14. Our laboratory also developed a method for imaging thick biological specimens. Our strategy was different from the two methods described above in that we merged the information that is in focus in the various focal planes and reconstructed the final 3D volume in real space using a parallel beam projector15. Our aim was to develop a method that can easily be performed in any electron microscopy laboratory. To this end, we aimed to collect the focal images in a limited amount of time, comparable to the time frame of conventional tomography experiments. In addition, our proposed method could be adapted for use with different types of alignment and reconstruction software.
In the context of our publication from 201515, we wanted to visualize and characterize the recovery of the depth-of-field, so we used a large convergence semi-angle of 25 mrad. Here, we present a step-by-step protocol for performing through-focal imaging in STEM according to the method developed in our laboratory in 201515, and we present how the data from 2015 was processed. This method recovers in-focus information from several focal planes throughout a thick (750 nm) biological sample and enables high-quality 3D reconstructions. Where relevant, the differences in this methodology versus the methods used by other groups are also presented.

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			<td>Phosphate Buffered Saline</td>
			<td>Sigma-Aldrich</td>
			<td>P4417</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma-Aldrich</td>
			<td>2860</td>
			<td></td>
		</tr>
		<tr>
			<td>Epoxy resin</td>
			<td>EMS</td>
			<td>14120</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>P6148</td>
			<td>Add paraformaldehyde powder to PBS heated at approximately 60 &#176;C. 
			Increase pH by adding 1 N NaOH until no PFA powder is visible.</td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>G5882&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium Tetroxyde</td>
			<td>EMS</td>
			<td>19150</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl Acetate</td>
			<td>Sigma-Aldrich</td>
			<td>73943</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin Capsule</td>
			<td>EMS</td>
			<td>70110</td>
			<td></td>
		</tr>
		<tr>
			<td>Triming and Histo knives</td>
			<td>LFG Distribution</td>
			<td>Diatome diamond knives</td>
			<td></td>
		</tr>
		<tr>
			<td>Electron Microscopy copper grid</td>
			<td>LFG Distribution</td>
			<td>G200-Cu</td>
			<td></td>
		</tr>
		<tr>
			<td>Grid Coating Pen</td>
			<td>LFG Distribution</td>
			<td>70624</td>
			<td></td>
		</tr>
		<tr>
			<td>Specimen Holder</td>
			<td>JEOL</td>
			<td>EM-21311 HTR</td>
			<td></td>
		</tr>
		<tr>
			<td>Electron Microscope</td>
			<td>JEOL</td>
			<td>JEM-2200FS</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

In our study, electrons were accelerated to 200 kV in the field emission gun transmission electron microscope. Images were collected in STEM BF mode using a 20-&#181;s dwell time. Regarding the design of the through-focal tilt-series, we found that focus intervals of 150 nm gave satisfactory results for the ultrastructural study of a 750 nm-thick biological sample-even though the depth-of-field of the electron beam was only 3 nm-since we used a very large convergence semi-angle of 25 mrad...

Watch this Scientific Journal Video about Preparation and Observation of Thick Biological Samples by Scanning Transmission Electron Tomography at JoVE.com

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

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 ...

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9.2K Views.  Campus Universitaire d'Orsay. This report describes a sample preparation protocol and specific imaging conditions for performing scanning transmission electron tomography of thick biological specimens.

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

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Scanning Electron Microscopy (SEM) Protocols for Problematic Plant, Oomycete, and Fungal Samples

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of samples from wet microenvironments and cell destruction. Using young floral tissues, oomycete cysts, and fungi spores (Agaricales) as examples, specific protocols to process delicate samples are described here that overcome some of the main challenges in sample treatment for image capture under the SEM.
Floral meristems fixed with FAA (Formalin-Acetic-Alcohol) and processed with the Critical Point Dryer (CPD) did not display collapsed cellular walls or distorted organs. These results are crucial for the reconstruction of floral development. A similar CPD-based treatment of samples from wet microenvironments, such as the glutaraldehyde-fixed oomycete cysts, is optimal to test the differential growth of diagnostic characteristics (e.g., the cyst spines) on different types of substrates. Destruction of nurse cells attached to fungi spores was avoided after rehydration, dehydration, and the CPD treatment, an important step for further functional studies of these cells.
The protocols detailed here represent low-cost and rapid alternatives for the acquisition of good-quality images to reconstruct growth processes and to study diagnostic characteristics.

Scanning Electron Microscopy SEM Protocols for Problematic Plant, Oomycete, and Fungal Samples

Problems in the processing of biological samples for scanning electron microscopy observation include cell collapse, treatment of samples from wet microenvironments and cell destruction. Low-cost and relatively rapid protocols suited for preparing challenging samples such as floral meristems, oomycete cysts, and fungi (Agaricales) are compiled and detailed here.

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

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

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

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

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

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 ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

Serial Block-Face Scanning Electron Microscopy (SBF-SEM) of Biological Tissue Samples

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, offering an unprecedented three-dimensional view of tissue microanatomy. While SBF-SEM has seen an exponential increase in use in recent years, technical aspects such as proper tissue preparation and imaging parameters are paramount for the success of this imaging modality. This imaging system benefits from the automated nature of the device, allowing one to leave the microscope unattended during the imaging process, with the automated collection of hundreds of images possible in a single day. However, without appropriate tissue preparation cellular ultrastructure can be altered in such a way that incorrect or misleading conclusions might be drawn. Additionally, images are generated by scanning the block-face of a resin-embedded biological sample and this often presents challenges and considerations that must be addressed. The accumulation of electrons within the block during imaging, known as "tissue charging," can lead to a loss of contrast and an inability to appreciate cellular structure. Moreover, while increasing electron beam intensity/voltage or decreasing beam-scanning speed can increase image resolution, this can also have the unfortunate side effect of damaging the resin block and distorting subsequent images in the imaging series. Here we present a routine protocol for the preparation of biological tissue samples that preserves cellular ultrastructure and diminishes tissue charging. We also provide imaging considerations for the rapid acquisition of high-quality serial-images with minimal damage to the tissue block.

Serial Block-Face Scanning Electron Microscopy SBF-SEM of Biological Tissue Samples

This protocol outlines a routine method for using serial block-face scanning electron microscopy (SBF-SEM), a powerful 3D imaging technique. Successful application of SBF-SEM hinges on proper fixation and tissue staining techniques, as well as careful consideration of imaging settings. This protocol contains practical considerations for the entirety of this process.

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, ...