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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<li>Shulman, J. M., Feany, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genetic+modifiers+of+tauopathy+in+Drosophila%5BTitle%5D)+AND+%22Genetics%22%5BJournal%5D)">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/pubmed?term=((Implicating+calpain+in+tau-mediated+toxicity+in+vivo%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">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.

<table><tbody><tr><td>gold&amp;ndash;palladium</td><td>Ted Pella, Inc.</td><td>91212</td><td></td></tr><tr><td>Silver conductive adhesive 503</td><td>Electron Microscopy Sciences</td><td>12686-15</td><td></td></tr><tr><td>Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 8 &amp;mu;m, polycarbonate</td><td>Sigma</td><td>WHA110614</td><td></td></tr><tr><td>Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.8 &amp;mu;m, polycarbonate, black</td><td>Sigma</td><td>WHA110659</td><td></td></tr><tr><td>Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.2 &amp;mu;m, polycarbonate</td><td>Sigma</td><td>WHA110606</td><td></td></tr><tr><td>aluminum weighing dish&amp;nbsp;</td><td>Fisher Scientific</td><td>08-732-100</td><td></td></tr><tr><td>aluminum mounting stubs (12 mm)</td><td>Electron Microscopy Sciences</td><td>75210</td><td></td></tr><tr><td>aluminum mounting stubs (25 mm)</td><td>Electron Microscopy Sciences</td><td>75186</td><td></td></tr><tr><td>Adhesive tabs&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>76760</td><td></td></tr><tr><td>conductive carbon adhesive tabs</td><td>Electron Microscopy Sciences</td><td>77825</td><td></td></tr><tr><td>F/2 media</td><td>Culture Collection of Algae at the University of Texas at Austin</td><td>https://utex.org/products/f_2-medium</td><td>No Catalog number given - see link&amp;nbsp;</td></tr><tr><td>AF6 media</td><td>Bigelow - National Center for Marin Algae and Microbiota</td><td>https://ncma.bigelow.org/media/wysiwyg/Algal_recipes/NCMA_algal_medium_AF6_1.pdf</td><td>No Catalog number given - see link&amp;nbsp;</td></tr><tr><td>Soil water medium</td><td>Carolina Biological Supply Company</td><td>153785</td><td></td></tr><tr><td>Polyethylene glycol tert-octylphenyl ether (Triton X-100)</td><td>VWR</td><td>97062-208</td><td></td></tr><tr><td>Hummer 6.2 Sputter Coater</td><td>Anatech USA</td><td>http://www.anatechusa.com/hummer-sputter-systems/4</td><td>No Catalog number given - see link&amp;nbsp;</td></tr><tr><td>Hitachi 3400N-II SEM&amp;nbsp;</td><td>Hitachi</td><td>https://www.hitachi-hightech.com/us/product_list/?ld=sms2&amp;amp;md=sms2-1&amp;amp;sd=sms2-1&lt;br /&gt;-2&amp;amp;gclid=EAIaIQobChMIpq_jtJfj3A&lt;br /&gt;IVS7jACh2mdgkPEAAYASAAEgKA&lt;br /&gt;nfD_BwE</td><td>The company doesn&#039;t appear to sell this model any longer&amp;nbsp;</td></tr></tbody></table>

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

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Biology

19.1K 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

Sample Preparation Method of Scanning and Transmission Electron Microscope for the Appendages of Woodboring Beetle

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of the woodboring beetle, Chlorophorus caragana Xie &#38; Wang (2012), for both types of electron microscopy. The scanning electron microscopy (SEM) sample preparation protocol was based on sample chemical fixation, dehydration in a series of ethanol baths, drying, and sputter-coating. By adding Tween 20 (Polyoxyethylene sorbitan laurate) to the fixative and the wash solution, the insect body surface of woodboring beetle was washed more cleanly in SEM. This study&#39;s transmission electron microscopy (TEM) sample preparation involved a series of steps including fixation, ethanol dehydration, embedding in resin, positioning using fluorescence microscopy, sectioning, and staining. Fixative with Tween 20 enabled penetrate the insect body wall of woodboring beetle more easily than it would had been without Tween 20, and subsequently better fixed tissues and organs in the body, thus yielded clear transmission electron microscope observations of insect sensilla ultrastructures. The next step of this preparation was determining the positions of insect sensilla in the sample embedded in the resin block by using fluorescence microscopy to increase the precision of target sensilla positioning. This improved slicing accuracy.

In order to observe ultrastructure of insect sensilla, scanning and transmission electron microscopy (SEM and TEM, respectively) sample preparation protocol were presented in the study. Tween 20 was added into the fixative to avoid sample deformation in SEM. Fluorescence microscopy was helpful for improving slicing accuracy in TEM.

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of ...

Environment

Processing Embryo, Eggshell, and Fungal Culture for Scanning Electron Microscopy

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, methods involved in processing different biological samples involve unique practices. All conventional practices described in the literature for processing samples still find useful applications, but subtle changes in the sample preparation can alter image quality, as well as, introduce artifacts. Hence, using a unique sample preparation technique specific to the type of tissue analyzed is required to obtain a good quality image with ultrastructural resolution. The focus of this study is to provide the optimal sample preparation protocols for imaging embryos, rigid eggshells, and fungal cultures using SEM. The following optimizations were recommended to yield good results for the three different delicate biological samples studied. Use of milder fixatives like 4% paraformaldehyde or 3% glutaraldehyde followed by dehydration with ethanol series is mandatory. Fungal mycelium on agar blocks obtained by slide cultures yields a better ultrastructural integrity compared to cultures taken directly from agar plates. Chemical drying of embryos with HMDS provides drying without introducing surface tension artifacts compared to critical point drying. HMDS prevents cracking caused by shrinkage as samples are less brittle during drying. However, for fungal culture, critical point drying provides acceptable image quality compared to chemical drying. Eggshells can be imaged with no special preparation steps except for thorough washing and air drying prior to mounting. Preparation methodologies were standardized based on acceptable image quality obtained with each trial.&#160;

Here, we present detailed processing protocols for imaging delicate tissue samples using scanning electron microscopy (SEM). Three different processing methods, namely, hexamethyl disilazana (HMDS) chemical drying, simple air drying, and critical point drying are described for preparing rigid eggshells, embryos at early developmental stages, and fungal cultures respectively.

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, ...

Cell Culture on Silicon Nitride Membranes and Cryopreparation for Synchrotron X-ray Fluorescence Nano-analysis

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions and their disturbed homeostasis is involved in various diseases. State-of-the-art synchrotron X-ray fluorescence nanoprobes provide the required sensitivity and spatial resolution to elucidate the two-dimensional (2D) and three-dimensional (3D) distribution and concentration of metals inside entire cells at the organelle level. This opens new exciting scientific fields of investigation on the role of metals in the physiopathology of the cell. The cellular preparation is a key and often complex procedure, particularly for basic analysis. Although X-ray fluorescence techniques are now widespread and various preparation methods have been used, very few studies have investigated the preservation of the elemental content of cells at best, and no stepwise detailed protocol for the cryopreparation of adherent cells for X-ray fluorescence nanoprobes has been released so far. This is a description of a protocol that provides the stepwise cellular preparation for fast cryofixation to enable synchrotron X-ray fluorescence nano-analysis of cells in a frozen hydrated state when a cryogenic environment and transfer is available. In case nano-analysis has to be performed at room temperature, an additional procedure for freeze-drying the cryofixed adherent cellular preparation is provided. The proposed protocols have been successfully used in previous works, most recently in studying the 2D and 3D intracellular distribution of an organometallic compound in breast cancer cells.

Presented here is a protocol for cell culture on silicon nitride membranes and plunge-freezing prior to X-ray fluorescence imaging with a synchrotron cryogenic X-ray nanoprobe. When only room temperature nano-analysis is provided, the frozen samples can be further freeze-dried. These are critical steps to obtain information on the intracellular elemental composition.

Very little is known about the distribution of metal ions at the subcellular level. However, those chemical elements have essential regulatory functions ...

Biochemistry

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in situ. Owing to the strong interaction of an electron with the matter, high-resolution cryo-ET studies are limited to specimens with a thickness of less than 200 nm, which restricts the applicability of cryo-ET only to the peripheral regions of a cell. A complex workflow that comprises the preparation of thin cellular cross-sections by cryo-focused ion beam micromachining (cryo-FIBM) was introduced during the last decade to enable the acquisition of cryo-ET data from the interior of larger cells. We present a protocol for the preparation of cellular lamellae from a sample vitrified by plunge freezing utilizing Saccharomyces cerevisiae as a prototypical example of a eukaryotic cell with wide utilization in cellular and molecular biology research. We describe protocols for vitrification of S. cerevisiae into isolated patches of a few cells or a continuous monolayer of the cells on a TEM grid and provide a protocol for lamella preparation by cryo-FIB for these two samples.

Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

We present a protocol for lamella preparation of plunge frozen biological specimens by cryo-focused ion beam micromachining for high-resolution structural studies of macromolecules in situ with cryo-electron tomography. The presented protocol provides guidelines for the preparation of high-quality lamellae with high reproducibility for structural characterization of macromolecules inside the&#160;Saccharomyces cerevisiae.

Today, cryo-electron tomography (cryo-ET) is the only technique that can provide near-atomic resolution structural data on macromolecular complexes in ...

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

Neuroscience

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 ...

Optimized Protocol for Retinal Organoid Sample Preparation for TEM

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem cells (iPSCs) under specific conditions. Synapse development and maturation in ROs have been studied immunocytochemically and functionally. However, the direct evidence of the synaptic contact ultrastructure is limited, containing both special ribbon synapses and conventional chemical synapses. Transmission electron microscopy (TEM) is characterized by high resolution and a respectable history elucidating retinal development and synapse maturation in humans and various species. It is a powerful tool to explore synaptic structure in ROs and is widely used in the research field of ROs. Therefore, to better explore the structure of RO synaptic contacts at the nanoscale and obtain high-quality microscopic evidence, we developed a simple and repeatable method of RO TEM sample preparation. This paper describes the protocol, reagents used, and detailed steps, including RO fixation preparation, post fixation, embedding, and visualization.

Author Spotlight: Analyzing the Synaptic Ultrastructure in Mature Retinal Organoids Using TEM

This protocol provides an optimized and elaborate preparation procedure for retinal organoid samples for transmission electron microscopy. It is suitable for applications that involve the analysis of synapses in mature retinal organoids.

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem ...

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

Chemical Drying: Biological Sample Preparation for Scanning Electron Microscopy (SEM)

Source: Koon, M. A., et al. <a href="https://www.jove.com/video/58761/" target="_blank">Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)</a>. J. Vis. Exp. (2019).
Scanning electron microscopy (SEM) is a powerful tool to image&#8212;primarily&#8212;the surface features of a biological sample. Proper sample preparation, however, is critical to obtaining quality images. This video focuses on the dehydration and drying steps of preparing intact adult Drosophila for SEM.

Source: Koon, M. A., et al. Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron ...

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

Summary

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Sample Preparation Method of Scanning and Transmission Electron Microscope for the Appendages of Woodboring Beetle

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Preparation and Cryo-FIB micromachining of Saccharomyces cerevisiae for Cryo-Electron Tomography

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

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

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

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

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