The authors would like to thank Dr. Daniel Baldassarre, SUNY Oswego for helpful discussions and comments on the manuscript. This study was supported by Rice Creek Associate Grants, Oswego; Challenge Grants SUNY Oswego and National Science Foundation (NSF) Small Grants to PGL and JG.

NOTE: Painted turtle (Chrysemys picta) eggs used in this study were collected during the nesting season of May through June 2015-16 from Rice Creek Field Station, Oswego New York with permission obtained from the New York State Department of Environmental Conservation (DEC).
1. Chemical drying method to process embryos for SEM
<ol>
	<li>Collect turtle eggs from field sites during the nesting season. Prepare the incubation chambers in advance, made of plastic boxes with lids (L x W x H) 6.7 cm x 25.4 cm x 10.2 cm filled with bedding medium prepared with a moist mixture of vermiculite and peat moss (1:1 ratio). Make 4-6 holes of approximately 0.25 cm along the sides of the boxes and on the lid to allow aeration.</li>
	<li>Gently remove the soil from the nest to uncover the eggs. Wipe the surface of turtle eggs with diluted iodine tincture (1:25,000) to control microbial contamination during incubation. Place the clutches separate from each other, clutch size of painted turtle may vary from 5-9 eggs and place a maximum of 8-9 eggs per box. 
	NOTE: Carefully handle the eggs during collection, wiping, labeling and placing inside the boxes. Position and alignment of eggs need to be the same as they were laid, any movement will inhibit embryo development.</li>
	<li>Manually bury the eggs half in the bedding, cover and place the box inside the incubator set at 30 &#176;C. Incubate the eggs for 10-17 days to obtain the embryonic stages 12, 13, and 18 respectively used in this study. Add distilled water to partially wet the bedding medium every other day to avoid dehydration and to maintain the moisture level for normal development of the embryos. 
	NOTE: Incubation and staging embryos are according to a complete developmental table published earlier39.</li>
	<li>Fix the embryos by making the first cut on one side of the dorsal eggshell and yolk membrane together, vertical to the long axis using pointed scissors. Insert the scissors into the yolk carefully to avoid cutting the embryonic disc. Now cut the lateral side of the egg along the long axis and then cut the other side of the egg along the short axis.</li>
	<li>Peel open the excised piece with the embryo side up using forceps. Cut the other lateral side of the eggshell and place the excised piece into phosphate buffered saline (PBS&#160;at a pH of 7.4). 
	NOTE: The turtle egg is filled with highly viscous egg yolk and the dorsal side of the embryo adheres to the eggshell membrane. The eggshell membrane near the embryo changes with the incubation from translucent white to an opaque, chalky white. This allows one to locate the embryo in the center of the long axis and be seen as a dark calcified spot from the exterior40,41.</li>
	<li>Use a stereomicroscope to isolate the embryos along with the yolk membrane by peeling them from the eggshell using forceps. Remove extra-embryonic membranes using forceps and micro-scissors. Transfer the embryo using an embryo spoon to fresh PBS in a Petri dish to wash any blood or yolk.</li>
	<li>Use clear 12-well plates to fix the embryos overnight in 4% paraformaldehyde in PBS at 4 &#176;C or in 2-3% glutaraldehyde in PBS. Place one to three embryos in each well depending on the size of the embryos. Ensure complete infiltration (samples will appear white) for older embryos by extending fixation time for 2-3 days. Rinse embryos 3x with fresh PBS for 5 min each rinse. 
	NOTE: Avoid damaging the surface of the embryo by using polystyrene inserts with polyester mesh bottoms for 12-well plates to transfer specimens from one solvent to another.</li>
	<li>Dehydrate samples using a series of ethanol concentration in distilled water: 30%, 50%, 70%, 80%, 95%, and 100% and treat samples for 1 h in each dehydration solution. Repeat the step with 100% ethanol twice to ensure complete dehydration. If not used immediately, store samples in 70% ethanol at -20 &#176;C for a longer period.</li>
	<li>Dry embryos using a series of hexamethyldisilazana (HMDS) to 100% ethanol concentration: 1:2, 2:1 and 100%. Leave the samples in each solution for 20 min and keep the Petri dish partially covered during the process. 
	NOTE: Carry out all steps involving HMDS in fume hood with necessary personal protection gear as HMDS is highly toxic.</li>
	<li>Leave the embryos in the final 100% HMDS solution covered completely or partially in a fume hood overnight aiding in evaporation of HMDS, leaving samples ready for mounting and sputter coating. Cover the dish to eliminate dust settling over the samples. 
	NOTE: The tissue will appear white after complete drying and partially dried samples will look yellow in color, leave these tissues in the fume hood for a longer time.</li>
	<li>Choose the size of aluminum stubs and carbon adhesive tape per size of the sample analyzed. Mount the dried samples carefully on a standard aluminum pin stub (12.7 mm x 8 mm) using double stick carbon conductive tape (12 mm).</li>
	<li>Introduce mounted samples into the chamber of the sputter coater to coat the specimen with a very thin film of gold to eliminate the charge effect. Gold plate the specimens for 60-120 s at a 35 mA sputter.</li>
	<li>Mount the stubs to corresponding pore-plates by securely fastening the setscrews. Transfer the sample holder into or out of the sample chamber of SEM using the sample exchange tool. Image the samples at high vacuum mode with an accelerating beam voltage of 10 kV and emission current 10 &#181;A. 
	NOTE: Always wear gloves while handling samples, sample holders, mounting stubs, and transfer tools to avoid grease contamination from hands to the SEM system.</li>
	<li>Test alternative techniques listed below to compare the resolution of specimens obtained from above procedure.
	<ol>
		<li>Include a post-fixation with 1% osmium tetroxide for 1 h at room temperature after step 1.7.</li>
		<li>Partially or completely remove the HMDS at step 1.10 for fast rapid evaporation of HMDS.</li>
		<li>Process embryos after step 1.8 using critical point drying (CPD) by following steps 3.3-3.4.</li>
	</ol>
	</li>
</ol>
2. Preparing the eggshell for SEM using an air-drying method
<ol>
	<li>Collect and incubate the painted turtle (Chrysemys picta) eggs to fix the embryos as specified in steps 1.1-1.7.</li>
	<li>Save eggshells in distilled water following embryo fixation in step 2.1. Clean the eggshells thoroughly by soaking in distilled water for at least 1 h to eliminate yolk and albumin contamination.</li>
	<li>Air-dry eggshells after washing on delicate antistatic wipes in the fume hood overnight. Store dried eggshells in clean specimen bottles labeled by number and stage.</li>
	<li>Mount, sputter coat and image the specimens by following steps specified in steps 1.11-1.13.</li>
</ol>
3. Critical point drying method for preparing fungal cultures for SEM
<ol>
	<li>Establish slide cultures
	<ol>
		<li>Prepare potato dextrose agar (PDA) media for fungal cultures: Add 39 g of PDA power in 1 L of distilled water in an Erlenmeyer flask. Mix well by swirling the flask and autoclave the media at 121 &#176;C for 30 min. Allow the media solution to cool and then add the antibiotic chloramphenicol (25 &#181;g/mL) using a sterile micropipette. 
		NOTE: After sterilization, the agar solution should be hot and it will not solidify soon. Cool it enough so that it will not inactivate the antibiotics.</li>
		<li>Mix it by swirling and pour plates (approximately 10-12 mL of media for each 10 cm Petri dish), carefully stack up and let solidify.</li>
		<li>Use a sterile scalpel blade to cut out small blocks of agar about &#189; to &#190; of an inch. Remove and place an agar block onto a clean glass microscope slide.</li>
		<li>Place the slide in a clean Petri dish to prevent contamination and preserve moisture during incubation.</li>
		<li>Raise the slide off the bottom of the Petri dish using a sterile toothpick to create surface tension between the plate and the slide to remove the glass slide without disrupting the delicate growth following incubation.</li>
		<li>Use a sterile loop or a needle to transfer some of the fungi from the specimen inoculum to each of the four sides of the agar block on the slide.</li>
		<li>Place a clean coverslip on the surface of the agar block following inoculation. Add a few drops of sterile distilled water to the Petri dish around the slide to ensure moisture for the growing fungi.</li>
		<li>Seal the plate partially using paraffin film and incubate the plate at 30 &#730;C for an appropriate length of time (for Fusarium species incubate for 36 to 48 h).</li>
		<li>Remove the slide from the Petri dish and separate the tightly adhered coverslip from the agar block using sterile forceps. Fix the agar blocks in 3% glutaraldehyde in PBS overnight at 4 &#730;C.</li>
	</ol>
	</li>
	<li>Dehydrate the samples by passing through an ethanol series: 10%, 25%, 50%, 75% and 90% with 15 min per change. Process the samples for final dehydration with two changes in 100% ethanol lasting 30 minutes each to ensure complete saturation.</li>
	<li>Critical point drying: Place dehydrated samples in the chamber of the CPD apparatus. Seal and cool the chamber by opening the valves to allow liquid CO2 in and vent ethanol out, until liquid CO2 completely fills the chamber.
	<ol>
		<li>Seal and heat the chamber slowly to achieve a critical point when the chamber pressure exceeds 1000 psi and the temperature exceeds 31 &#176;C, the liquid and gas phase of CO2 is in equilibrium. Slowly drain the CO2 from the chamber and the sample as gas to avoid effects of surface tension.</li>
	</ol>
	</li>
	<li>Perform mounting, gold plating and imaging the specimens by following steps specified in steps 1.11-1.13.</li>
	<li>Perform comparative analysis by chemically drying the specimens from step 3.2 using HMDS by following steps 1.9-1.13.</li>
</ol>

The authors have nothing to disclose

In our study, different fixation agents, dehydration and drying methods were tested to prepare three different delicate biological samples for SEM imaging: embryos, eggshells, and fungal cultures. SEM is commonly used for surface analysis, so fixative penetration is less concerning, but it must be understood that poorly fixed internal structures will cause inward shrinking or/and collapsed surface structures. Extended fixation time should also be considered for larger tissue samples, replacing the fixative solution a few...

Scanning electron microscope (SEM) ultrastructural analysis and intracellular imaging supplement light microscopy for three-dimensional profiling of prokaryotes, plants, and animals. The high spatial resolution of an SEM makes it one of the most versatile and powerful techniques available for the examination of microstructural characteristics of specimens at the nanometer to micrometer scale. Desiccated specimens are resolved to compositional and topographical structures with intense detail, which provides the foundation for developing valid conclusions about functional relationships1,2,3,4,5,6,7,8,9. When interpreting SEM images of biological specimens, it is a great challenge to distinguish between native structures and the artifacts that are created during processing. SEM is generally operated at very high vacuums to avoid any interference from gas molecules affecting the primary, secondary or backscattered electron beams emitted from the sample10,11. Also, biological materials are susceptible to radiation damage due to their poor or non-conducting properties. It is essential for the specimens loaded into the SEM to be completely dry and free of any organic contaminants to eliminate any possible outgassing in a high vacuum environment10,11. As biological specimens are mostly composed of water, additional preparative techniques are required to ensure that the native structures are retained.
The resolution obtained is based on optimizing preparation methods specific to specimen types and instrumental parameters utilized. Thus, it is necessary to avoid using generalized processing steps for all tissue types. Some biological specimens will require less stringent processing to preserve their structure while more time and care might be needed for delicate types of samples to avoid the introduction of drying artifacts, such as shrinkage and collapse. Sample preparation is a critical step in SEM imaging; the findings of morphometric studies are remarkably influenced by specimen preparation procedures12,13. Common preparation steps for many biological samples are fixation, dehydration and coating with a metal such as gold, platinum or palladium to convert their surfaces to be conductive for SEM analysis. The nature and combination of steps used will vary depending on the type of the tissue, and the specific goals of the study. Charge build-up, sensitivity to vacuum and electron beam damage pose problems when processing soft delicate biological samples, necessitating additional processing steps to retain the native structure of the object. Using conventional methods such as osmium tetroxide fixing, and dehydration cause shrinkage and the collapse of delicate tissues14,15,16,17. The aim of the study is to establish elegant methodologies derived by combining ideas from earlier studies with modifications to prepare and image soft delicate tissues (e.g., reptile embryos, eggshell of painted turtles, and fungal cultures).
Selection of a suitable fixing method is the first most important step for microscopic analysis of biological specimens. Fixing the tissues immediately after isolating from an organism is essential to prevent alteration in their morphology due to decomposition. An effective fixative should terminate cellular processes by permeating the cells quickly and maintaining the effect irreversibly to stabilize the structure of the sample to withstand both subsequent processing steps and examination under the SEM17,18. Although several chemical and physical fixation methods are known, chemical fixation is more commonly used for biological specimens to avoid any cellular changes due to autolysis, putrefaction, and drying effects. There are numerous fixative chemical formulations discussed in literature17,19,20,21,22,23, fixatives that work by denaturing and coagulating biological macromolecules, and those that fix by covalently cross-linking macromolecules. Alcohols are used as denaturing fixatives that preserve ultrastructure very poorly and are used mostly for light microscopy and not recommended for electron microscopic analysis. Cross-linking fixatives like formaldehyde, glutaraldehyde, and osmium tetroxide create intermolecular and intramolecular crosslinking between macromolecules within the tissues, providing excellent preservation of ultra-structures11,24,25,26. Biological samples are sensitive to temperature. The temperature at the beginning of fixation is recommended to be 4 &#176;C to reduce the lateral mobility of membrane proteins, to slow the diffusion of intercellular molecules, and to slow the rate of fixation11. The time required for fixing tissues largely depends on the size of the sample and the speed at which the fixative diffuses and reacts with the components of the specimen. An overnight fixation in 4% paraformaldehyde or 3% glutaraldehyde in PBS at 4 &#176;C is the preferred method for SEM analysis of specimens used in this study for their sequential penetrative properties, which allow smaller delicate samples to be processed17,18,19,20,27. A post-fixation step with osmium tetroxide is eliminated not only due to its toxic nature but also found to implement no added advantage to improve image quality for the samples analyzed in this study.
Biological samples contain fluids that interfere with the SEM operation; hence, the samples need to be dried before inserting it in the SEM sample chamber. Once dehydration is ensured, the solvent must be removed from the tissue without creating artifacts into the specimens due to the surface tension/drying. Three different drying methods were commonly used during processing tissues for SEM imaging: air drying, critical point drying, and freeze drying samples28,29,30,31. Few studies report all three drying methods producing identical results with animal tissue samples28,29,30,31. A general practice used for smaller specimens are chemical dehydration by ascending concentration series of alcohol and hexamethyldisilazane (HMDS), but larger specimens are dried using a critical point drying (CPD) instrument32. During the drying process, considerable forces formed in small cavities that are passed through the specimen by a liquid/gas interface; this can even lead to a complete collapse of the hollow structures33. Any deformation occurring due to the treatment could then be mistaken as a native structural feature of the specimen. Thus, the generalized phenomenon for processing should be eliminated and a unique drying process should be standardized for each type of tissue especially when delicate tissue specimens are analyzed.
In several trials conducted using various combination of all the above-mentioned processes, we standardized the methods that can be used for SEM analysis of three delicate tissues: reptile embryos, eggshells of painted turtles, and fungal cultures. Developmental biologists and morphologists describe normal and abnormal morphogenesis during embryo development in representative vertebrate animals. Investigations on gene signaling pathways depend on the morphological description of novel structures. To avoid any abrupt change in the vertebrate embryo structure during SEM analysis, we recommend chemical drying following dehydration. Chemical drying using HMDS is the relatively newest drying method and the advantages include relative quickness, ease of use, lost cost, and the limited expertise and equipment needed9. CPD is a commonly used drying technique using passaging CO2 across the specimens at a specific temperature and pressure. We identified that HMDS is suitable for drying soft delicate tissues and allows larger samples to be processed compared to critical point drying, which caused extensive deformation to embryonic tissues. Several methods have been used to prepare samples for SEM imaging to study the morphological characteristics of fungi34. Fungal specimens are commonly fixed in osmium tetroxide followed by ethanol dehydration and critical point drying, which may provide satisfactory results, although the toxic effects of osmium tetroxide6,7,35 and losing fungal materials while changing solutions during processing are pronounced disadvantages. The sample preparation technique using air-drying without fixation has also been practiced36 but results in shrunken and collapsed structures, and observation of such specimens can easily be misinterpreted while characterizing the species. Fungal hypha loses its integrity in contact with liquids and an even drying may not be achieved to restore the structure. Due to this effect, freeze-drying is commonly used for drying soft tissues like fungal mycelium. Freeze drying works well for clean materials but the presence of any salts or secretion will obscure surface detail that will be identified only at the SEM viewing stage. We coupled the slide culture method with glutaraldehyde fixing and critical point drying to yield structural details of intact fungal hyphae and spores. Although CPD drying caused shrinkage in embryos, it resulted in well preserved mycelial structures when coupled with glutaraldehyde fixation. The eggshell is of primary importance to the embryo of oviparous animals by not only acting as a protective covering but also providing mechanical stability, permeability to gas and water, and a calcium reserve for the developing embryo. Freshwater turtle eggshells are classified as &#34;rigid&#34;&#160;based on their structure, and due to their availability have received significant attention from biologists1,2,3,4,5,6,7,37,38.
We detail simple methods for easy examination of eggshell and shell membranes of painted turtle that can be applied to any rigid eggshell species. Preparation methodologies were evaluated based on resulting image quality and reduced potential artifacts.&#160;

<table><tbody><tr><td>Agar</td><td>Fischer Scientific</td><td>S25127A</td><td>for slide cultures</td></tr><tr><td>Aluminum pin stub</td><td>Tedpella</td><td>16111</td><td>12.7 mm x 8 mm</td></tr><tr><td>BD Difco Dehydrated Culture Media: Potato Dextrose Agar</td><td>BD 213400</td><td>DF0013-17-6</td><td>Media for isolation and cultivation of Fungi, yeast and molds</td></tr><tr><td>Chloramphenicol</td><td>Fischer BioReagents</td><td>BP904-100</td><td>Antibiotic for media</td></tr><tr><td>Coarse Vermiculite</td><td>Greenhouse Megastore</td><td>SO-VER-12</td><td>bedding medium</td></tr><tr><td>Clear 12- well plate</td><td>Corning</td><td>07-201-589</td><td>for fixing embryo</td></tr><tr><td>Coverslips</td><td>Fischer Scientific</td><td>S17525B</td><td>for slide culture</td></tr><tr><td>Critical Point Dryer</td><td>Quorum CPD</td><td>EMS850</td><td>critical point drying</td></tr><tr><td>Culture dishes</td><td>Fischer Scientific</td><td>08 747B</td><td>DISH PETRI 100X10MM 12/PK</td></tr><tr><td>Ethanol</td><td>Fischer Scientific</td><td>A406P 4</td><td>dehydration agent</td></tr><tr><td>Forceps- Aquarius Tweezers</td><td>Tedpella</td><td>5804</td><td>style 4, length 108mm, widh x thickness 0.017 x 0.17 mm</td></tr><tr><td>Glutaraldehyde</td><td>Fischer Scientific</td><td>G151-1</td><td>fixative</td></tr><tr><td>Gold target for sputter coater</td><td>DENTON VACUUM</td><td>TAR001-0158</td><td>Gold Target, 2.375&amp;Prime; D X .002&amp;Prime;</td></tr><tr><td>Hexamethyldisilazana</td><td>Fischer Scientific</td><td>C19479-5000</td><td>chemical drying agent</td></tr><tr><td>Kim wipes</td><td>Kimtech</td><td>S-8115</td><td>cleaning</td></tr><tr><td>Microscope slides</td><td>Thermo Scientific</td><td>67-762-16</td><td>for slide culture</td></tr><tr><td>Microscopy Scissors</td><td>Tedpella</td><td>1327</td><td>Double pointed, stainless steel, 100 mm L (3-5/8&quot;).</td></tr><tr><td>Micro-scissors</td><td>Tedpella</td><td>1346</td><td>Vannas-type, straight, 80mm L</td></tr><tr><td>Moria Perforated Embryo Spoon</td><td>Fine Science Tools</td><td>10370-17</td><td>Length 14.5 cm, tip diameter 20 mm, spoon depth 5 mm</td></tr><tr><td>Netwell Inserts</td><td>Corning</td><td>0330B09</td><td>15 mm Inserts with 74 &amp;micro;m Mesh Size Polyester Membrane act as handy carriers during specimen processing into different solvents</td></tr><tr><td>Paraformaldehyde</td><td>Fischer Scientific</td><td>T353 500</td><td>fixative</td></tr><tr><td>Peat moss</td><td>Walmart- Miracle Gro</td><td>551705263</td><td>bedding medium</td></tr><tr><td>PELCO tabs double stick carbon conductive tape</td><td>Tedpella</td><td>5000</td><td>12 mm OD</td></tr><tr><td>Sputter coater</td><td>DENTON VACUUM</td><td>DESK V</td><td>thin metal coating</td></tr><tr><td>SEM</td><td>JEOL USA</td><td>JEOL JSM 6610LV scanning electron scope</td><td>electron microscopy</td></tr></tbody></table>

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;

Figure 1 show scanning electron micrographic analysis of painted turtle (Chrysemys picta) embryos. Painted turtle eggs collected and incubated on a bedding medium, mounted on aluminum stubs following chemical drying were used for SEM imaging (Figure 1A-E). A lateral view of a stage 12 embryo shows the craniofacial structures; maxillary prominence extends beyond the mandibular and limits a well-marked nasal pit medially; five pharyngeal ...

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

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, ...

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9.3K Views.  SUNY Oswego. 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.

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

Rapid Acquisition of 3D Images Using High-resolution Episcopic Microscopy

High-resolution episcopic microscopic (HREM) technology enables rapid acquisition of high-resolution digital volumetric and three-dimensional (3D) morphometric data. Here, we describe the detailed protocol to image the entire mouse embryo. The protocol consists of four major sections: sample preparation, embedding, image acquisition and finally, 3D visualization. The technology requires specimens to be stained with a fluorescent dye, which can be problematic for large or dense specimens. To overcome this limitation, we have improved the existing protocol to enhance tissue penetration of the dye by pretreating the specimen with a solution containing urea and sodium dodecyl sulfate. The protocol uses only routine laboratory equipment and reagents for easy adaptation in standard laboratory settings. We show that the resulting high-resolution 3D images faithfully recapitulate the detailed morphologic features of the internal organs of mouse embryos, thereby permitting morphometric analyses. Together, we present a detailed and improved protocol using standard laboratory equipment to acquire high-resolution 3D images of small and large sized specimens.

We describe a detailed protocol using high-resolution episcopic microscopy to acquire three-dimensional (3D) images of mouse embryos. This improved protocol utilizes a modified tissue preparation method to enhance penetration of the fluorescent dye, thereby permitting morphometric analysis of both small and large-sized specimens.

High-resolution episcopic microscopic (HREM) technology enables rapid acquisition of high-resolution digital volumetric and three-dimensional (3D) ...

Developmental Biology

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

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 ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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 ...

Biochemistry

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 ...

Biology

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

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 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.

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.

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

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 ...

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

Summary

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

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

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

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

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

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