This work was funded by a grant to MLS and a Summer Scholar Award to MAK from the Office of Research and Graduate Studies at Central Michigan University. Cyanobacteria were supplied by the Zimba and plankton lab, part of the Center for Coastal Studies, Texas A&#38;M University at Corpus Christi. Euglenoids were supplied by the Triemer lab, Michigan State University.

1. Preparation and Fixation
<ol>
	<li>Prepare cyanobacteria.
	<ol>
		<li>Grow unialgal cultures in F/2 media24,25 at a temperature of 28 &#176;C on a 14:10 h light dark cycle. Transfer sufficient culture to a 1.5 mL microcentrifuge tube such that after allowing 15 min to settle, the bacterial pellet is approximately 0.05 mL in size. Remove the media and replace with 1.5 mL of fixative (1.25% glutaraldehyde, 0.1 M phosphate buffer pH 7.0), gently invert several times, and incubate overnight at 4 &#176;C.</li>
		<li>Transfer the fixed cells with a glass pipette into a 10 mL filtration rig (Figure 1) containing a polycarbonate 25 mm filter with 0.8 or 0.2 &#181;m pores, depending on the size of the cells. Seal the side arm and funnel with rubber stoppers to contain the culture in the funnel. Remove the fixative by gentle vacuum on the filtration flask, after removing both stoppers. 
		Note: If the cell density on the polycarbonate filter is not optimal, adjust the amount of starting culture used.</li>
	</ol>
	</li>
	<li>Prepare single cell algae (euglenoids).
	<ol>
		<li>Grow unialgal cultures in AF-6 media26 with 150 mL L-1 of commercially-available soil-water medium added to the media, at a temperature of 22 &#176;C on a 14:10 h light dark cycle. Transfer 2 mL of low density (OD600 &#60; 0.5) culture with a glass pipette into a 10 mL filtration rig (Figure 1) containing a polycarbonate 25 mm filter with 8 &#181;m pores. Seal the side arm and funnel with rubber stoppers to contain the culture in the funnel. 
		Note: If the cell density on the polycarbonate filter is not optimal, adjust the amount of starting culture used.</li>
		<li>Add three drops of 4% osmium tetroxide (OsO4) directly to the culture and allow to incubate for 30 min. Remove the fixative by gentle vacuum on the filtration flask, after removing both stoppers. 
		Note: OsO4 is an acute toxin (dermal, oral, and inhalation routes), corrosive to the skin, damaging to the eyes, and a respiratory sensitizer. OsO4 should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.</li>
	</ol>
	</li>
	<li>Prepare Drosophila melanogaster (fruit fly)23.
	<ol>
		<li>Anesthetize adult flies using 100% carbon dioxide. Place anesthetized adults (about 10 to 30 flies) in a small plastic screw cap vial or 1.5 mL centrifuge tube.</li>
		<li>Immerse anesthetized flies in 1 mL of fixative (1.25% glutaraldehyde, 0.1 M phosphate buffer pH 7.2) for 2 h (or overnight) at 4 &#176;C. If the flies float to the surface of the fixative, add a few drops of 2.5% polyethylene glycol tert-octylphenyl ether to weaken the surface tension of the fixative allowing for total submersion of the tissue. Remove the fixative using a glass pipet.</li>
	</ol>
	</li>
</ol>
2. Washing and Dehydration
<ol>
	<li>Wash and dehydrate cyanobacteria.
	<ol>
		<li>Wash the fixed sample three times with 5 mL of 0.1 M phosphate buffer pH 7.0 at room temperature for 10 min each. Keep the flask and funnel stoppered to hold the wash. Remove each wash by gentle vacuum on the filtration flask, after removing both stoppers.</li>
		<li>Dehydrate the sample while maintaining continuous immersion using a graded ethanol series, for 10 min in a volume of 5 mL in the funnel. The graded ethanol concentrations are: 25%, 50%, 75%, 95%, and 2 changes of 100% ethanol. Keep the flask and funnel stoppered to contain the ethanol in the funnel, preventing loss both via evaporation and passive flow through the filter.</li>
		<li>Remove ethanol by gentle vacuum on the filtration flask, after removing both stoppers. Transfer the filter/sample to a disposable aluminum weighing dish containing just enough 100% ethanol to cover the sample before drying.</li>
	</ol>
	</li>
	<li>Wash and dehydrate single cell algae (euglenoids).
	<ol>
		<li>Wash the fixed sample three times with 5 mL of deionized water at room temperature for 10 min each. Keep the flask and funnel stoppered to contain the wash in the funnel. Remove each wash by gentle vacuum on the filtration flask, after removing both stoppers.</li>
		<li>Dehydrate the sample while maintaining continuous immersion using a graded ethanol series for 10 min in a volume of 5 mL in the funnel. The graded ethanol concentrations are: 25%, 50%, 75%, 95%, and 2 changes of 100% ethanol. Keep the flask and funnel stoppered to contain the ethanol in the funnel, preventing loss both via evaporation and passive flow through the filter.</li>
		<li>Remove the ethanol by gentle vacuum on the filtration flask, after removing both stoppers. Transfer the filter/sample to a disposable aluminum weighing dish containing just enough 100% ethanol to cover the sample before drying.</li>
	</ol>
	</li>
	<li>Wash and dehydrate Drosophila melanogaster (fruit fly).
	<ol>
		<li>Wash the fixed sample three times with 1 mL of 0.1 M phosphate buffer pH 7.2 at room temperature for 10 min in a 1.5 mL microcentrifuge tube. Remove each wash with a glass pipette, being careful not to remove the flies.</li>
		<li>Dehydrate the sample using a graded ethanol series, for 10 min in a volume of 1 mL in a microfuge tube. The ethanol concentrations are: 25%, 50%, 75%, 80%, 95%, 100%. Remove the ethanol with a glass pipette, being careful not to remove the flies.</li>
		<li>Retain the sample in the 1.5 mL microcentrifuge tube with just enough 100% ethanol to cover the sample before drying.</li>
	</ol>
	</li>
</ol>
3. Drying
<ol>
	<li>Perform chemical drying using t-butyl alcohol (TBA)27.
	<ol>
		<li>Replace the 100% ethanol solution with a 1:1 solution of TBA and 100% ethanol for 20 min. Replace the 1:1 solution with 100% TBA for 20 min. Repeat twice. Keep the solution at 37 &#176;C so that TBA does not freeze. 
		Note: 100% TBA has a freezing point of 25.5 &#176;C; the 1:1 solution will not freeze at room temperature. TBA is flammable, causes serious eye irritation, is harmful if inhaled, and may cause respiratory irritation, drowsiness, or dizziness. TBA should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.</li>
		<li>Transfer the sample in TBA, if in a 1.5 mL microcentrifuge tube, into a disposable aluminum weighing dish. Once in the aluminum weighing dish, remove TBA and replace with just enough fresh 100% TBA to cover the sample. Freeze the TBA at 4 &#176;C for 10 min.</li>
		<li>Transfer to a vacuum desiccator (bell jar) with frozen gel packs to keep TBA frozen. Evacuate and maintain vacuum with a rotary pump to allow the sample to dry by vacuum sublimation of the frozen TBA for at least 3 h or overnight.</li>
	</ol>
	</li>
	<li>Perform chemical drying using hexamethyldisilazane (HMDS)20.
	<ol>
		<li>Replace the 100% ethanol solution with a 1:2 solution of HMDS and 100% ethanol for 20 min. Replace the 1:2 solution with a 2:1 solution of HMDS and 100% ethanol for 20 min. Replace the 2:1 solution with 100% HMDS for 20 min. Repeat once. 
		Note: HMDS is flammable and an acute toxin (dermal route). HMDS should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.</li>
		<li>Transfer the sample in HMDS, if in a 1.5 mL microcentrifuge tube, into a disposable aluminum weighing dish. Once in the aluminum weighing dish, replace the 100% HMDS with just enough fresh 100% HMDS to cover the sample.</li>
		<li>Transfer the sample to a plastic or glass non-vacuum desiccator with fresh desiccant (5-6 cm deep) and place into in a chemical fume hood. Alternatively, place the sample directly in a chemical fume hood to dry with a loose lid, such as a box, to prevent debris from falling on the sample. Allow the sample to dry for 12 to 24 h.</li>
	</ol>
	</li>
</ol>
4. Mounting
<ol>
	<li>Mount cyanobacteria and euglenoids.
	<ol>
		<li>Label the bottom of either a 12- or 25-mm aluminum mounting stub to indicate what is being placed on top. Cut the polycarbonate filter with the dried cells into quarters with a clean razor blade or scalpel if using a 12 mm stub, or place the entire filter if using a 25 mm stub.</li>
		<li>Place each filter on adhesive or a carbon adhesive tab secured to the top of a stub.</li>
	</ol>
	</li>
	<li>Mount Drosophila melanogaster (fruit fly).
	<ol>
		<li>Label the bottom of the aluminum mounting stub to indicate what is being placed on top. 
		Place the dried flies in the desired position on adhesive or carbon adhesive tab secured to the top of a stub under a dissecting microscope with precision tweezers.</li>
		<li>Apply silver conductive adhesive, i.e., silver paint, around the outer edges of the stubs. Connect the silver paint to the flies using a toothpick to ensure conductivity. Do not allow the paint to touch the desired imaging area.</li>
		<li>Place the stubs in a stub holder box and place the open stub holder box in a desiccator. Allow the silver paint to dry at least 3 h or overnight for best results.</li>
	</ol>
	</li>
</ol>
5. Sputter Coating
<ol>
	<li>Prepare the sputter coating apparatus by performing four to five flushes of argon gas. If the humidity is high (i.e., the room air feels moist, usually about 50% relative humidity), perform six to seven flushes. Sputter coat the stubs following the manufacturer&#8217;s instructions.</li>
	<li>Coat samples based on specimen used:
	<ol>
		<li>For filters with cyanobacteria, set the timer to sputter coat for 180 s straight on.</li>
		<li>For filters with eukaryotic algae, i.e., euglenoids, set the timer to sputter coat for 120 s straight on, then 20 s on three sides at a 45&#176; angle.</li>
		<li>For large samples, i.e., fly heads, set the timer to sputter coat for 60 s straight on, then 30 s on each side at a 45&#176; angle. 
		Note: After coating is complete, stubs should be light gray in color.</li>
	</ol>
	</li>
</ol>
6. Imaging
<ol>
	<li>Image samples using a scanning electron microscope that includes a secondary electron (SE) detector.
	<ol>
		<li>For cyanobacteria, apply the following settings: 3-5 kV accelerator voltage (AC), 20-30 probe current (PC), and 5 mm working distance (WD). For euglenoids, use 3 kV AC, 30 PC, and 5 mm WD. For flies, use 5 kV AC, 30 PC, and 5 to 10 mm WD.</li>
	</ol>
	</li>
	<li>Adjust magnification depending on the size of the detail or object to be visualized. Adjust the stigmators, apertures, and focus until a clear image is produced. Capture the image using high-resolution settings according to the manufacturer&#8217;s instructions of the SEM.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Trends+in+the+evolution+of+the+euglenid+pellicle.">Trends in the evolution of the euglenid pellicle.</a> Evolution. 55 (11), 2215-2235 (2001).</li><li>Shulman, J. M., Feany, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+modifiers+of+tauopathy+in+Drosophila.">Genetic modifiers of tauopathy in Drosophila.</a> Genetics. 165 (3), 1233-1242 (2003).</li><li>Reinecke, J. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Implicating+calpain+in+tau-mediated+toxicity+in+vivo.">Implicating calpain in tau-mediated toxicity in vivo.</a> PLoS One. 6 (8), e23865 (2011).</li></ol>

The authors have nothing to disclose.

Here we described a protocol using SEM to obtain detailed information about external morphological characteristics of three types of organisms that others can apply to examine features of many types of organisms or tissues. Within each step of the protocol, there are potential points of error that may arise and are discussed in detail below.
While the volumes for fixative and washes given here are specific, in general the fixative and washes should be 5-10x the volume of the specimen. All fixa...

A scanning electron microscope (SEM) uses a focused beam of high-energy electrons to generate an image from secondary electrons that shows the morphology and topography of a sample1. SEM can be used to directly determine the physical size of a sample, the surface structure, and the three-dimensional shape, and offers greater resolution and larger depth of field compared to light microscopy. Another form of electron microscopy (EM), transmission electron microscopy (TEM) uses focused electrons that pass through the sample, generating images with fine details of internal structure. While TEM has higher resolution than light or SEM and can be used to resolve structures as small as single atoms, it has three major disadvantages: extensive sample preparation, a small field of view, and a shallow depth of field2,3. Although other visualized protocols exist using SEM to examine specific cells, organelles, or tissues4,5,6,7,8,9,10 , this protocol is unique in that we describe methods that can be broadly applied to a large range of organisms for morphological assessment.
SEM has found broad applications for examining inorganic materials including nanoparticles11,12, polymers13, and numerous applications in geological, industrial, and material sciences14,15,16. In biology, SEM has long been used as a method for examining biological samples ranging from individual proteins to whole organisms17,18. SEM is of particular value because morphological surface details can be used to inform scientific discovery. SEM analysis is a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of a wide range of biological samples.
Because an SEM normally operates under high vacuum (10-6 Torr minimum) to support a coherent beam of high-speed electrons, no liquids (water, oils, alcohols) are permitted in the sample chamber, as liquids prevent a vacuum from forming. Thus, all samples examined using SEM must be dehydrated, typically using a graded ethanol series followed by a drying process to remove the ethanol. There are several methods of drying biological tissues for use in the SEM, including air drying, lyophilization, use of a critical point drying (CPD) device, or chemical drying using t-butyl alcohol (TBA) or hexamethyldisilazane (HMDS)19,20,21,22. Most often, selection of a drying method is empirical since each biological sample may react differently to each drying method. For any given sample, all of these methods may be appropriate, so comparing the advantages and disadvantages of each is useful in selecting the appropriate method.
While air drying a sample at room temperature or in a drying oven (60 &#176;C) is the simplest method, most biological samples show drying-induced damage such as shriveling and collapse, resulting in distortion of the specimen. The process of lyophilization also removes water (or ice) from a sample, but requires samples to be flash-frozen and placed under vacuum to remove the ice via the process of sublimation, potentially damaging the sample. In addition, the user must have access to a lyophilizer. The most commonly used method for dehydrating samples for SEM is critical point drying (CPD). In CPD, the ethanol in a sample is replaced with liquid carbon dioxide (CO2) and, under specific temperature and pressure conditions known as the critical point (31.1 &#176;C and 1,073 psi), CO2 vaporizes without creating surface tension, thereby effectively maintaining the morphological and structural features of the sample. While CPD is generally the standard method, it has several drawbacks. First, the process requires access to a critical point dryer, which is not only expensive, but also necessitates the use of liquid carbon dioxide. Second, the size of the sample that can be dried is limited to the chamber size of the critical point dryer. Third, the exchange of liquids during CPD can cause turbulence that can damage the sample.
Chemical drying offers many advantages over CPD and serves as a suitable alternative that is becoming widely used in SEM sample preparation. The use of chemical dehydrants such as TBA and HMDS offers a fast, inexpensive, and simple alternative to other methods, while still maintaining the structural integrity of the sample. We recently showed that there was no difference in the integrity of the tissue or the quality of the final image captured when using CPD or TBA as the drying method in adult Drosophila retinal tissue23. Unlike CPD, TBA and HMDS do not require a drying instrument or liquid CO2 and there is no limitation on the size of the sample to be dried. In addition to obtaining the chemicals, only a standard chemical fume hood and appropriate personal protective equipment (gloves, lab coat, and safety goggles) are required to complete the drying process. While both TBA and HMDS are flammable, TBA is less toxic and less expensive (approximately 1/3 the cost of HMDS) than HMDS.
In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). These organisms represent a wide range in size (0.5 &#181;m to 4 mm) and cellular diversity (single-celled to multicellular), yet all are easily amenable to SEM analysis with only small variations needed for specimen preparation. This protocol describes the methods for using chemical dehydration and SEM analysis to examine morphological details of three types of organisms and would be broadly applicable to examining many organismal and tissue types.

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			<td height="40" style="height:40px;width:351px;">Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.8 &#956;m, polycarbonate, black</td>
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			<td height="40" style="height:40px;width:351px;">Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.2 &#956;m, polycarbonate</td>
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			<td style="width:373px;">Bigelow - National Center for Marin Algae and Microbiota</td>
			<td style="width:395px;">https://ncma.bigelow.org/media/wysiwyg/Algal_recipes/NCMA_algal_medium_AF6_1.pdf</td>
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			<td>Carolina Biological Supply Company</td>
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			<td height="40" style="height:40px;">Hitachi 3400N-II SEM&#160;</td>
			<td>Hitachi</td>
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preparation of prokaryotic and eukaryotic organisms using chemical drying for morphological analysis in scanning electron microscopy (sem)

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to cells, tissues, organelles, and even whole organisms. This protocol focuses on two chemical drying methods, hexamethyldisilazane (HMDS) and t-butyl alcohol (TBA), and their application to imaging of both prokaryotic and eukaryotic organisms using SEM. In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). The purpose of this protocol is to describe a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of specimens that can be broadly applied to a large range of organisms for morphological assessment. Successful completion of this protocol will allow others to use SEM to visualize samples by applying these techniques to their system.

Cyanobacteria are a prokaryotic group of organisms that are critical to the global carbon, oxygen, and nitrogen cycles28,29. Of the estimated 6000 species of cyanobacteria30, most have a mucilaginous sheath that cover and connect the cells together and to other structures31, which along with the shape, can be resolved microscopically32. Cell size, shape, and p...

Watch this Scientific Journal Video about Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM at JoVE.com

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM

SEM analysis is an effective method to aid in species identification or phenotypic discrimination. This protocol describes the methods for examining specific morphological details of three representative types of organisms and would be broadly applicable to examining features of many organismal and tissue types.

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to ...

preparation-prokaryotic-eukaryotic-organisms-using-chemical-drying

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

18.9K Views.  Central Michigan University. SEM analysis is an effective method to aid in species identification or phenotypic discrimination. This protocol describes the methods for examining specific morphological details of three representative types of organisms and would be broadly applicable to examining features of many organismal and tissue types.

Video: Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM

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 ...

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

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 ...

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 ...

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 ...

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 ...

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)

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