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 times depending on the fixation duration. Due to active interaction between the fixative and the tissue, the osmotic properties of cells would change considerably. Tissue fluids would dilute the fixative and thus an aldehyde effect contributing to the overall osmolality can be neglected as far as effects on cell volume are concerned. Also, formaldehyde would penetrate faster into tissues and is more effective as the cross-linker compared to glutaraldehyde. A buffered formaldehyde-glutaraldehyde mixture was also reported for fixing a wide variety of tissues, although for monolayers and cell suspensions diluted solutions were more suitable for fixing15,16. However, the diluted solutions are also hypertonic, thus the aldehydes are not themselves osmotically active. For delicate samples used in this study, PBS at pH 7.4 was used as a vehicle for the fixing agents, as phosphate buffers are thought to be more similar to the cytoplasmic environments of most biological samples. Further, the osmolarity is identified to be within the physiological range required for the sample while PBS is acting as a vehicle for the fixing agent. Post-fixation with osmium tetroxide is often recommended for various biological samples16,22,42&#160;but has been eliminated for all the biological samples used in this study. Samples without osmium tetroxide yielded crisp images and specimens fixed with osmium tetroxide yielded indistinguishable results (Figure 4A). Osmium tetroxide causes distortion of leaf tissue structures and yields poorer specimen preservation compared to glutaraldehyde and formaldehyde mixture43, and it has been suggested that a build-up of osmium molecules could inhibit infiltration of dehydrating agents and transitional agents used in CPD, and the solvents like HMDS used for chemical drying. The effect is tissue-specific and excluding osmium tetroxide will not have any effect on the image quality for the type of delicate specimens analyzed in this study. For other types of tissues use of osmium tetroxide may be eliminated with extended primary fixation.
HMDS dried turtle embryo specimens showed well-preserved surfaces, and less distortion or shrinkage compared to CPD (Figure 4). CPD minimizes the chance for distortion in cell morphology and artifacts production due to the zero or minimal surface tension created during the process. However, CPD can cause a potential physical hazard for delicate fragile samples. In line with our observation, HMDS yielded similar or higher quality imaging by minimizing the surface tension caused due to drying based on previous studies published44,45,46,47. Compared to CPD treatment, HMDS preserved structural details of the organic meshwork excellently in etched bivalves and barnacle shells48 and provided high resolution of the structural features of the complex internal organization of mermithid nematodes49 and insect internal tissues28. The drying plate artifacts were seen in high quantities on the specimens prepared by the CPD technique as compared to the HMDS technique. The membrane blebs and pellet artifacts were increased in cervical cells prepared by using CPD technique50. HMDS reacts with water to produce hexamethyldisiloxane and ammonia, both of which evaporate from the object. HMDS is commonly used in gas chromatography to create silyl ethers of compounds such as sugars, amino acids, alcohols, and numerous other compounds. It is not known if HMDS reacts with some of these compounds in tissues. HMDS might crosslink proteins and stiffen the tissue during the drying process28, although the exact reason that HMDS provides better preservation of embryos could not be explained except for tissue specificity. Based on the results of our investigation, CPD caused extensive shrinkage compared to HMDS, and is much more suitable for processing embryonic tissues, especially to analyze surface structure for early staged embryos. The mechanism by which HMDS works on tissue drying has not been elucidated, although slow drying with gradual evaporation in absolute anhydrous surroundings could be a reason for obtaining an excellent surface structure of the animals.
Small pieces of agar blocks with fungal colonies from previously established slide cultures were used in this study to avoid issues with maintaining the original fluffiness of the mycelial mat. CPD drying of agar pieces with fungal colonies resulted in a washing effect, while changes in the fixatives, buffers, and ethanol was found to be a better choice overcoming the difficulties with freeze drying. HMDS that works well for embryos of all stages was unable to resolve fungal cultures, while air drying with HMDS posed a significant problem: curling up and losing the structural rigidity. While CPD works well for fungal cultures, it induced instant damage to embryos especially at early stages in development, causing shrinkage on the surface. No charging or drying artifacts were observed when HMDS and CPD were used for embryos and fungal cultures, respectively. For rigid tissue like eggshell of turtles, no special processing is required except for washing and air drying at room temperature prior to mounting for imaging.
Regardless of the preparation method, gradual ethanol gradient steps should be used to reduce the potential for drying artifacts. Initially, both HMDS and CPD samples were found to have surface artifacts with shrinkage following the dehydration and drying protocols available in the literature. After a series of standardizations, we determined that the artifacts were the result of the alcohol dehydration. It is critical to use a slow dehydration process using more gradual ethanol increases during dehydration. Also, the duration of each iteration should be longer for embryos compared to fungal cultures. Further, an additional step at 100% ethanol ensured complete uniform saturation. HMDS can be removed completely or partially to allow air drying of samples, but extended exposure to HMDS and gradual air drying by evaporation were found to be effective for embryos to avoid disruption (Figure 4A,C) caused by large surface tension as observed for microbial cell attachment51. Immersing specimens in HMDS and slow drying overnight revealed an excellent surface structure in Daphnia species52 suggesting that soft delicate tissues could benefit from the slow drying process. Acceptable image quality for embryos can be obtained with slow chemical drying by gradually increasing the concentration of HMDS to ethanol following dehydration.
All samples were mounted onto aluminum stubs using a double-sided carbon tape to provide a conductive surface, a thin coat of colorless nail polish could also be used if the same sample needs to be imaged in a different orientation. Samples can be easily removed from the stubs when nail polish is used on the stub surface and positioning the samples in alternative planes is possible compared to using double-sided carbon tape. As the samples are nonconductive, it is necessary to sputter a thin layer of metal on the samples to increase conductance. A gold target was used in this study during sputter coating, and gold-palladium also yields similar results avoiding charging effects. The correct beam voltage is sample dependent, for lightly coated specimens, imaging with a field emission SEM at 2-5 kV will provide a good definition of surface structures. For tissues with adequate coating, 5 kV generally provides better signal without more beam penetration. For specimens that lack depth and are coated with gold alone, imaging at a higher voltage (10 kV) allowed better visualization of surface topography.
Preparation of delicate tissues for SEM poses distinctive challenges, and a range of sample preparation techniques can be used to overcome and minimize imaging artifacts. We provide comprehensive methods to process three different delicate biological tissue samples, for SEM imaging: embryos, rigid eggshells, and fungal cultures. In our investigation, subtle alterations to popular methods available from previously published studies were made, and we identified the most appropriate dehydration and drying methods specific to each delicate tissue type. These protocols could be adapted to obtain acceptable SEM image quality from similar biological samples.&#160;

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 border="0" cellpadding="0" cellspacing="0" style="width:712px;" width="711">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Agar</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">S25127A</td>
			<td style="width:229px;">for slide cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Aluminum pin stub</td>
			<td style="width:123px;">Tedpella</td>
			<td style="width:145px;">16111</td>
			<td style="width:229px;">12.7 mm x 8 mm</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:215px;">BD Difco Dehydrated Culture Media: Potato Dextrose Agar</td>
			<td style="width:123px;">BD 213400</td>
			<td style="width:145px;">DF0013-17-6</td>
			<td style="width:229px;">Media for isolation and cultivation of Fungi, yeast and molds</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Chloramphenicol</td>
			<td style="width:123px;">Fischer BioReagents</td>
			<td style="width:145px;">BP904-100</td>
			<td style="width:229px;">Antibiotic for media</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Coarse Vermiculite</td>
			<td style="width:123px;">Greenhouse Megastore</td>
			<td style="width:145px;">SO-VER-12</td>
			<td style="width:229px;">bedding medium</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Clear 12- well plate</td>
			<td style="width:123px;">Corning</td>
			<td style="width:145px;">07-201-589</td>
			<td style="width:229px;">for fixing embryo</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Coverslips</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">S17525B</td>
			<td style="width:229px;">for slide culture</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Critical Point Dryer</td>
			<td style="width:123px;">Quorum CPD</td>
			<td style="width:145px;">EMS850</td>
			<td style="width:229px;">critical point drying</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Culture dishes</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">08 747B</td>
			<td style="width:229px;">DISH PETRI 100X10MM 12/PK</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Ethanol</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">A406P 4</td>
			<td style="width:229px;">dehydration agent</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Forceps- Aquarius Tweezers</td>
			<td style="width:123px;">Tedpella</td>
			<td style="width:145px;">5804</td>
			<td style="width:229px;">style 4, length 108mm, widh x thickness 0.017 x 0.17 mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Glutaraldehyde</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">G151-1</td>
			<td style="width:229px;">fixative</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Gold target for sputter coater</td>
			<td style="width:123px;">DENTON VACUUM</td>
			<td style="width:145px;">TAR001-0158</td>
			<td style="width:229px;">Gold Target, 2.375&#8243; D X .002&#8243;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Hexamethyldisilazana</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">C19479-5000</td>
			<td style="width:229px;">chemical drying agent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Kim wipes</td>
			<td style="width:123px;">Kimtech</td>
			<td style="width:145px;">S-8115</td>
			<td style="width:229px;">cleaning</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Microscope slides</td>
			<td style="width:123px;">Thermo Scientific</td>
			<td style="width:145px;">67-762-16</td>
			<td style="width:229px;">for slide culture</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Microscopy Scissors</td>
			<td style="width:123px;">Tedpella</td>
			<td style="width:145px;">1327</td>
			<td style="width:229px;">Double pointed, stainless steel, 100 mm L (3-5/8&#34;).</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Micro-scissors</td>
			<td style="width:123px;">Tedpella</td>
			<td style="width:145px;">1346</td>
			<td style="width:229px;">Vannas-type, straight, 80mm L</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Moria Perforated Embryo Spoon</td>
			<td style="width:123px;">Fine Science Tools</td>
			<td style="width:145px;">10370-17</td>
			<td style="width:229px;">Length 14.5 cm, tip diameter 20 mm, spoon depth 5 mm</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:215px;">Netwell Inserts</td>
			<td style="width:123px;">Corning</td>
			<td style="width:145px;">0330B09</td>
			<td style="width:229px;">15 mm Inserts with 74 &#181;m Mesh Size Polyester Membrane act as handy carriers during specimen processing into different solvents</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Paraformaldehyde</td>
			<td style="width:123px;">Fischer Scientific</td>
			<td style="width:145px;">T353 500</td>
			<td style="width:229px;">fixative</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Peat moss</td>
			<td style="width:123px;">Walmart- Miracle Gro</td>
			<td style="width:145px;">551705263</td>
			<td style="width:229px;">bedding medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">PELCO tabs double stick carbon conductive tape</td>
			<td style="width:123px;">Tedpella</td>
			<td style="width:145px;">5000</td>
			<td style="width:229px;">12 mm OD</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Sputter coater</td>
			<td style="width:123px;">DENTON VACUUM</td>
			<td style="width:145px;">DESK V</td>
			<td style="width:229px;">thin metal coating</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:215px;">SEM</td>
			<td style="width:123px;">JEOL USA</td>
			<td style="width:145px;">JEOL JSM 6610LV scanning electron scope</td>
			<td style="width:229px;">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 arches were also observed (Figure 1F). Newly formed somites at the posterior tail region of the embryo are easily countable compared to the segmented body somites. Forelimb buds appear longer compared to hind limb buds and points more caudally than ventrally. A well-defined outgrowth of a carapace ridge is seen along the entire inter-limb flank region of the stage 15 embryo (Figure 1G. At stage 18, painted turtles possess a short, feebly projecting snout. The lower beak rests within the upper jaw and is slightly upturned with a terminal hook that fits into the central notch of the upper beak (Figure 1H). The upper jaw of the painted turtles shows a notched appearance forming a medial U-V shaped distal tip with a small cusp on each side, a tiny egg tooth begins to form at the tip of the upper jaw. Limb buds that appeared longer during earlier stages form a paddle-like structure during stages 13-14 with digital plate vaguely indicated. Limb mesenchyme demarcated by the apical ectodermal ridge is seen along the anterior-posterior margins (Figure 1I-J).
Figure 2 shows ultra-structural analysis of Chrysemys picta eggshell and shell membrane using SEM imaging. Painted turtle eggshells were obtained as described above and were subjected to air-drying following washing with double distilled water (Figure 2A,B). Eggshells mounted on aluminum stubs with double-sided carbon tape (Figure 2C) were imaged using SEM. A lateral view of the shell showed outer calcareous eggshell layer firmly attaching to the inner filamentous shell membrane (Figure 2D). The outer surface of the eggshell consists of well-distinguished mineralized shell units, made of globular/spherical nodules arranged in groups and in between adjacent shell units a concentration of small rounded depressions or pores of various sizes were observed (Figure 2E). Nodules from each group of shell units meet at a connection junction, the center point (Figure 2F). The outer surface was manually peeled using forceps to observe the surface of the shell membrane. Rows of central plaques were seen which provides the attachment point between the shell units and underlying multilayered fibrous membrane (Figure 2G).
Figure 3 shows the morphological characterization of ascomycete fungal isolate from slide cultures observed using SEM imaging. Scheme of slide culture setup established (Figure 3A) showing fungal growth on the agar block within two days of inoculation. Colonies growing rapidly with white to cream-colored aerial mycelium (Figure 3B). SEM imaging following critical point drying and sputter coating of agar slices revealed fungal hyphae, curved conidia, and rice like spores. Conidiophores were seen arising laterally from the septate aerial hyphae (Figure 3C). Macroconidia produced on shorter, branched conidiophores are moderately curved, with short, blunt apical and indistinctly pedicellate basal cells mostly septate (Figure 3D).
Figure 4 shows the comparative results for processing similar specimens using alternative techniques. Immersing heads of stage 15 embryos in HMDS and drying by gradual evaporation show well preserved craniofacial structures, almost no shrinkage or distortion is seen with extended times of slow evaporation from HMDS (Figure 4A). Embryos post-fixed with osmium tetroxide showed no distinguishable difference in image quality except for slight shrinkage of tissue compared to HMDS slow processing (Figure 4B). Drying embryos by removing HMDS partially or completely allowing fast evaporation results in structural distortion and shrinkage (Figure 4C), whereas slow evaporation resolved excellent surface structures of a similarly staged embryo (Figure 4A). The drying artifacts were seen in the embryos treated with the CPD technique causing extensive shrinkage and destruction of tissue (Figure 4D,E). Agar slices from slide cultures show incomplete drying of fungal isolate with HMDS treatment compared to CPD (Figure F). Complete drying is achieved with CPD treatment of agar slices with a well dried, white specimen showing intact high-resolution structural features of fungal hyphae and spores (Figure 4G-G&#39;). Agar blocks with fungal cultures appear shrunken and yellow in color making them not suitable for SEM imaging (Figure 4H-H&#39;).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60018/60018fig1.jpg" /> 
Figure 1: Scanning electron micrographs of Chrysemys picta embryos prepared by chemical drying. (A) Chrysemys picta, nesting during the breeding season at Rice Creek Field Station, Oswego, NY. (B) Well-constructed nest from outside. (C) Surface soil removed with care to exposure the eggs laid. (D) Eggs placed into the incubation chamber with bedding medium. (E) Mounting embryos onto aluminum stubs following chemical drying, E4 showing the sputter coated embryo. (F) Lateral view of stage 12 embryo showing facial structures, limb buds and somites. (G) Well defined carapace ridge and paddle-shaped limb buds are seen at stage 15. (H) Craniofacial structures of stage 18 embryo showing upper and lower jaw, note a small egg tooth formed at the tip of the upper beak. (I) Dorsal view of right forelimb at stage 13-14 and a close-up view of AER (J) at the dorsal-ventral boundary. Scale bars: F-H, 500 &#181;m; I, 100 &#181;m, and J, 10 &#181;m. Keys: mb, midbrain; np, nasal pit; mxp, maxillary prominence; pa, pharyngeal arches; h, heart; fl, forelimb; s, somite; hl, hind limb; t, tail tip; cr, carapacial ridge; et, egg tooth; oc, oral cavity; e, eye; uj, upper jaw; lj, lower jaw; m, mesenchyme, AER, apical ectodermal ridge. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60018/60018fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60018/60018fig2.jpg" /> 
Figure 2: Ultrastructural analyses of the air-dried eggshell and the shell membrane of painted turtle eggs. (A) After fixing the embryos, eggshells were washed thoroughly in distilled water to remove yolk and albumin. (B) Clean eggshells were air dried at least overnight. (C) Mounting of an eggshell onto aluminum stubs. (D) Radial view of shell fragment showing outer calcareous layer and inner shell membrane. (E) Outer calcareous layer shows globular shell units (su) arranged in groups and pores seen in between these units (asterisks). (F) Magnified view of a nodule showing the center point (c) between the shell units. (G) Removal of calcareous layer shows the outer surface of the shell membrane with depressions left broken from shell units. Scale bars: E, 100 &#181;m; F, 10 &#181;m, and G, 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60018/60018fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60018/60018fig3.jpg" /> 
Figure 3: Preparing fungal culture for SEM imaging. (A) Schematic diagram to show the setup of slide culture. (B) White colored fungal colonies seen on the agar block after two days of incubation at 30&#176;C. (C) SEM image showing a section of mycelium of Fusarium solani, black asterisks marking the macroconidia in sporodochia, an arrow pointing to phialide, and arrowheads directed towards microconidia, scale bar, 10 &#181;m. (D) Magnified image of septate chlamydospores, scale bar 5 &#181;m. Keys: ab, agar block; cs, coverslip; f, fungal culture. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60018/60018fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60018/60018fig4.jpg" /> 
Figure 4: Comparative analysis of alternative techniques to process similar specimens. (A) Frontal view of painted turtle head dried with slow evaporation of HMDS (A), after post-fixing with osmium tetroxide (B), rapid faster evaporation of HMDS (C), and CPD (D-E). Including a post-fixation step using osmium tetroxide do not show any visible difference in image quality compared to HMDS treatment without post-fixation. Deformations are seen after fast evaporation of HMDS, while gradual drying with slow evaporation provides a better surface structure. Different degrees of shrinkage and collapsed structures are seen in the regions of brain, eyes and facial prominences in embryos treated with CPD. (F) Mounting of slide cultures processed with HMDS (1), CPD (2) and sputter coated CPD treated specimen (3) on aluminum stubs. (G-G&#39;) Complete drying seen as intact white colored agar slice with fungal inoculate achieved by CPD preserving structures of fungal mycelium and spores of Fusarium solani. (H-H&#39;) Inadequate drying of slide culture, shrinkage and damaged structural integrity seen with HMDS preservation technique. Scale bars: A-E, 500 &#181;m; G-H, 2 mm, and G&#39;-H&#39;, 20 &#181;m. Keys: et, egg tooth; oc, oral cavity; e, eye; uj, upper jaw; lj, lower jaw; black asterisks, macroconidia in sporodochia; arrow, phialide; and arrowheads, microconidia. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60018/60018fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Processing Embryo, Eggshell, and Fungal Culture for Scanning Electron Microscopy at JoVE.com

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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9.2K 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

Surface Renewal: An Advanced Micrometeorological Method for Measuring and Processing Field-Scale Energy Flux Density Data

Advanced micrometeorological methods have become increasingly important in soil, crop, and environmental sciences. For many scientists without formal training in atmospheric science, these techniques are relatively inaccessible. Surface renewal and other flux measurement methods require an understanding of boundary layer meteorology and extensive training in instrumentation and multiple data management programs. To improve accessibility of these techniques, we describe the underlying theory of surface renewal measurements, demonstrate how to set up a field station for surface renewal with eddy covariance calibration, and utilize our open-source turnkey data logger program to perform flux data acquisition and processing. The new turnkey program returns to the user a simple data table with the corrected fluxes and quality control parameters, and eliminates the need for researchers to shuttle between multiple processing programs to obtain the final flux data. An example of data generated from these measurements demonstrates how crop water use is measured with this technique. The output information is useful to growers for making irrigation decisions in a variety of agricultural ecosystems. These stations are currently deployed in numerous field experiments by researchers in our group and the California Department of Water Resources in the following crops: rice, wine and raisin grape vineyards, alfalfa, almond, walnut, peach, lemon, avocado, and corn.

Surface renewal is a micrometeorological method that is being used increasingly to determine energy fluxes, but its technical complexity makes it inaccessible to a broad audience. We describe the steps needed to set up and calibrate a surface renewal field station, to acquire and process data, and to correctly interpret results.

Advanced micrometeorological methods have become  increasingly important in soil, crop, and environmental sciences.  For many scientists without formal ...

Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of natural proteins have a molecular mass between 40 - 200 kDa1,2, a robust and high-throughput method with a nanometer resolution capability is needed. Negative staining (NS) electron microscopy (EM) is an easy, rapid, and qualitative approach which has frequently been used in research laboratories to examine protein structure and protein-protein interactions. Unfortunately, conventional NS protocols often generate structural artifacts on proteins, especially with lipoproteins that usually form presenting rouleaux artifacts. By using images of lipoproteins from cryo-electron microscopy (cryo-EM) as a standard, the key parameters in NS specimen preparation conditions were recently screened and reported as the optimized NS protocol (OpNS), a modified conventional NS protocol 3 . Artifacts like rouleaux can be greatly limited by OpNS, additionally providing high contrast along with reasonably high&#8208;resolution (near 1 nm) images of small and asymmetric proteins. These high-resolution and high contrast images are even favorable for an individual protein (a single object, no average) 3D reconstruction, such as a 160 kDa antibody, through the method of electron tomography4,5. Moreover, OpNS can be a high&#8208;throughput tool to examine hundreds of samples of small proteins. For example, the previously published mechanism of 53 kDa cholesteryl ester transfer protein (CETP) involved the screening and imaging of hundreds of samples 6. Considering cryo-EM rarely successfully images proteins less than 200 kDa has yet to publish any study involving screening over one hundred sample conditions, it is fair to call OpNS a high-throughput method for studying small proteins. Hopefully the OpNS protocol presented here can be a useful tool to push the boundaries of EM and accelerate EM studies into small protein structure, dynamics and mechanisms.

More than half of proteins are small proteins (molecular mass &#60;200 kDa) that are challenging for both electron microscope imaging and three-dimensional reconstructions. Optimized negative staining is a robust and high-throughput protocol to obtain high contrast and relatively high resolution (~1 nm) images of small asymmetric proteins or complexes under different physiological conditions.

Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of ...

Multimodal Optical Microscopy Methods Reveal Polyp Tissue Morphology and Structure in Caribbean Reef Building Corals

An integrated suite of imaging techniques has been applied to determine the three-dimensional (3D) morphology and cellular structure of polyp tissues comprising the Caribbean reef building corals Montastraeaannularis and M. faveolata. These approaches include fluorescence microscopy (FM), serial block face imaging (SBFI), and two-photon confocal laser scanning microscopy (TPLSM). SBFI provides deep tissue imaging after physical sectioning; it details the tissue surface texture and 3D visualization to tissue depths of more than 2 mm. Complementary FM and TPLSM yield ultra-high resolution images of tissue cellular structure. Results have: (1) identified previously unreported lobate tissue morphologies on the outer wall of individual coral polyps&#160;and (2) created the first surface maps of the 3D distribution and tissue density of chromatophores and algae-like dinoflagellate zooxanthellae endosymbionts. Spectral absorption peaks of 500 nm and 675 nm, respectively, suggest that M. annularis and M. faveolata contain similar types of chlorophyll and chromatophores. However, M. annularis and M. faveolata exhibit significant differences in the tissue density and 3D distribution of these key cellular components. This study focusing on imaging methods indicates that SBFI is extremely useful for analysis of large mm-scale samples of decalcified coral tissues. Complimentary FM and TPLSM reveal subtle submillimeter&#160;scale changes in cellular distribution and density in nondecalcified coral tissue samples. The TPLSM technique affords: (1) minimally invasive sample preparation,&#160;(2) superior optical sectioning ability,&#160;and (3) minimal light absorption and scattering, while still permitting deep tissue imaging.

An integrated suite of imaging techniques has been applied to determine polyp morphology and tissue structure in the Caribbean corals Montastraeaannularis and M. faveolata. Fluorescence, serial block face,&#160;and two-photon confocal laser scanning microscopy have identified lobate structure, polyp walls, and estimated chromatophore and zooxanthellae densities and distributions.

An integrated suite of imaging techniques has been applied to determine the three-dimensional (3D) morphology and cellular structure of polyp tissues ...

Data Processing Methods for 3D Seismic Imaging of Subsurface Volcanoes: Applications to the Tarim Flood Basalt

The morphology and structure of plumbing systems can provide key information on the eruption rate and style of basalt lava fields. The most powerful way to study subsurface geo-bodies is to use industrial 3D&#160;reflection seismological imaging. However, strategies to image subsurface volcanoes are very different from that of oil and gas reservoirs. In this study, we process seismic data cubes from the Northern Tarim Basin, China, to illustrate how to visualize sills through opacity rendering techniques and how to image the conduits by time-slicing. In the first case, we isolated probes by the seismic horizons marking the contacts between sills and encasing strata, applying opacity rendering techniques to extract sills from the seismic cube. The resulting detailed sill morphology shows that the flow direction is from the dome center to the rim. In the second seismic cube, we use time-slices to image the conduits, which corresponds to marked discontinuities within the encasing rocks. A set of time-slices obtained at different depths show that the Tarim flood basalts erupted from central volcanoes, fed by separate pipe-like conduits.

Three-dimensional (3D) reflection seismology is a powerful method for imaging subsurface volcanoes. By using industrial 3D seismological data from the Tarim Basin, we illustrate how to extract the sills and the conduits of subsurface volcanoes from seismic data cubes.

The morphology and structure of plumbing systems can provide key information on the eruption rate and style of basalt lava fields. The most powerful way ...

Thermal Scanning Conductometry (TSC) as a General Method for Studying and Controlling the Phase Behavior of Conductive Physical Gels

The thermal scanning conductometry protocol is a new approach in studying ionic gels based on low molecular weight gelators. The method is designed to follow the dynamically changing state of the ionogels, and to deliver more information and details about the subtle change of conductive properties with an increase or decrease in the temperature. Moreover, the method allows the performance of long term (i.e. days, weeks) measurements at a constant temperature to investigate the stability and durability of the system and the aging effects. The main advantage of the TSC method over classical conductometry is the ability to perform measurements during the gelation process, which was impossible with the classical method due to temperature stabilization, which usually takes a long time before the individual measurement. It is a well-known fact that to obtain the physical gel phase, the cooling stage must be fast; moreover, depending on the cooling rate, different microstructures can be achieved. The TSC method can be performed with any cooling/heating rate that can be assured by the external temperature system. In our case, we can achieve linear temperature change rates between 0.1 and approximately 10 &#176;C/min. The thermal scanning conductometry is designed to work in cycles, continuously changing between heating and cooling stages. Such an approach allows study of the reproducibility of the thermally reversible gel-sol phase transition. Moreover, it allows the performance of different experimental protocols on the same sample, which can be refreshed to initial state (if necessary) without removal from the measuring cell. Therefore, the measurements can be performed faster, in a more efficient way, and with much higher reproducibility and accuracy. Additionally, the TSC method can be also used as a tool to manufacture the ionogels with targeted properties, like microstructure, with an instant characterization of conductive properties.

Thermal Scanning Conductometry TSC as a General Method for Studying and Controlling the Phase Behavior of Conductive Physical Gels

The kinetics of the cooling process defines the properties of ionic gels based on low molecular weight gelators. This manuscript describes the usage of thermal scanning conductometry (TSC), which obtains full control over the gelation process, along with in situ measurements of the samples' temperature and conductivity.

The thermal scanning conductometry protocol is a new approach in studying ionic gels based on low molecular weight gelators. The method is designed to ...

Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several limitations that have resulted in the exclusion of many species. Advances in the field over the last two decades have led occupational health researchers to turn to molecular-based approaches for identifying fungal hazards. These methods have resulted in the detection of many species within indoor and occupational environments that have not been detected using traditional methods. This protocol details an approach for determining fungal diversity within air samples through genomic DNA extraction, amplification, sequencing, and taxonomic identification of fungal internal transcribed spacer (ITS) regions. ITS sequencing results in the detection of many fungal species that are either not detected or difficult to identify to species level using culture or microscopy. While these methods do not provide quantitative measures of fungal burden, they offer a new approach to hazard identification and can be used to determine overall species richness and diversity within an occupational environment.

Determining the fungal diversity within an environment is a method utilized in occupational health studies to identify health hazards. This protocol describes DNA extraction from occupational air samples for amplification and sequencing of fungal ITS regions. This approach detects many fungal species that can be overlooked by traditional assessment methods.

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several ...

Quantification of Endogenous Auxin and Cytokinin During Internode Culture of Ipecac

Adventitious shoot formation is an important technique for the propagation of economically important crops and for the regeneration of transgenic plants. Phytohormone treatment is required for the induction of adventitious shoots in most species. Whether adventitious shoots can be induced is determined by the balance between auxin and cytokinin (CK) levels. Much effort goes into determining optimum concentrations and combinations of phytohormones in each tissue used as explants and in each plant species. In ipecac, however, adventitious shoots can be induced on internodal segments in culture medium without phytohormone treatment. This allows the inherent plasticity of ipecac for cell differentiation to be evaluated. To induce adventitious shoots in ipecac, we cultured internodal segments at 24 &#176;C under 15 &#181;mol m&#8722;2 s&#8722;1 of light in a 14-h light/10-h dark cycle on phytohormone-free B5 medium solidified with 0.2% gellan gum for 5 weeks. To investigate phytohormone dynamics during adventitious shoot formation, we measured endogenous indole-3-acetic acid and CKs in the segments by liquid chromatography-tandem mass spectrometry LC-MS/MS. This method allows analysis of endogenous indole-3-acetic acid and CKs levels in a simple manner. It can be applied to investigate the dynamics of endogenous auxin and CK during organogenesis in other plant species.

Adventitious shoots can be induced on internodal segments of ipecac without phytohormone treatment. To evaluate phytohormone dynamics during adventitious shoot formation, we measured endogenous auxin and cytokinin in internodal segments by LC-MS/MS.

Adventitious shoot formation is an important technique for the propagation of economically important crops and for the regeneration of transgenic plants. ...

High-throughput, Microscale Protocol for the Analysis of Processing Parameters and Nutritional Qualities in Maize (Zea mays L.)

Maize is an important grain crop in the United States and worldwide. However, maize grain must be processed prior to human consumption. Furthermore, whole grain composition and processing characteristics vary among maize hybrids and can impact the quality of the final processed product. Therefore, in order to produce healthier processed food products from maize, it is necessary to know how to optimize processing parameters for particular sets of germplasm to account for these differences in grain composition and processing characteristics. This includes a better understanding of how current processing techniques impact the nutritional quality of the final processed food product. Here, we describe a microscale protocol that both simulates the processing pipeline to produce cornflakes from large flaking grits and allows for the processing of multiple grain samples simultaneously. The flaking grits, the intermediate processed products, or final processed product, as well as the corn grain itself, can be analyzed for nutritional content as part of a high-throughput analytical pipeline. This procedure was developed specifically for incorporation into a maize breeding research program, and it can be modified for other grain crops. We provide an example of the analysis of insoluble-bound ferulic acid and p-coumaric acid content in maize. Samples were taken at five different processing stages. We demonstrate that sampling can take place at multiple stages during microscale processing, that the processing technique can be utilized in the context of a specialized maize breeding program, and that, in our example, most of the nutritional content was lost during food product processing.

High-throughput, Microscale Protocol for the Analysis of Processing Parameters and Nutritional Qualities in Maize Zea mays L.

Here, we present a microscale protocol for processing grain samples and for incorporating this microscale approach into a high-throughput analytical pipeline. This is a higher throughput adaptation of currently available protocols.

Maize is an important grain crop in the United States and worldwide. However, maize grain must be processed prior to human consumption. Furthermore, whole ...

A 3-dimensional (3D)-printed Template for High Throughput Zebrafish Embryo Arraying

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related research fields. Thanks to its unique features, including large fecundity, embryo translucency, rapid and simultaneous development, etc., zebrafish embryos are often used for large scale toxicity assessment of chemicals and drug/compound screening. A typical screening procedure involves adult zebrafish spawning, embryos selection, and arraying the embryos into multi-well plates. From there, embryos are subjected to exposure and the toxicity of chemical, or the effectiveness of the drugs/compounds can be evaluated relatively quickly based on phenotypic observations. Among these processes, embryos arraying is one of the most time-consuming and labor-intensive steps that limits the throughput level. In this protocol, we present an innovative approach that makes use of a 3D-printed arraying template coupled with vacuum manipulation to speed up this laborious step. The protocol herein describes the overall design of the arraying template, a detailed experimental setup and step-by-step procedure, followed by representative results. When implemented, this approach should prove beneficial in a variety of research applications using zebrafish embryos as testing subjects.

A 3-dimensional 3D-printed Template for High Throughput Zebrafish Embryo Arraying

Here, we present a protocol to design and fabricate a zebrafish embryo arraying template, followed by a detailed procedure on the use of such template for high throughput zebrafish embryo arraying into a 96-well plate.

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related ...

Xylem Water Distribution in Woody Plants Visualized with a Cryo-scanning Electron Microscope

A scanning electron microscope installed cryo-unit (cryo-SEM) allows specimen observation at subzero temperatures and has been used for exploring water distribution in plant tissues in combination with freeze fixation techniques using liquid nitrogen (LN2). For woody species, however, preparations for observing the xylem transverse-cut surface involve some difficulties due to the orientation of wood fibers. Additionally, higher tension in the water column in xylem conduits can occasionally cause artifactual changes in water distribution, especially during sample fixation and collection. In this study, we demonstrate an efficient procedure to observe the water distribution within the xylem of woody plants in situ&#160;by using a cryostat and cryo-SEM. At first, during sample collection, measuring the xylem water potential should determine whether high tension is present in the xylem conduits. When the xylem water potential is low (&#60; ca. &#8722;0.5 MPa), a tension relaxation procedure is needed to facilitate better preservation of the water status in xylem conduits during sample freeze fixation. Next, a watertight collar is attached around the tree stem and filled with LN2&#160;for freeze fixation of the water status of xylem. After harvesting, care should be taken to ensure that the sample is preserved frozen while completing the procedures of sample preparation for observation. A cryostat is employed to clearly expose the xylem transverse-cut surface. In cryo-SEM observations, time adjustment for freeze-etching is required to remove frost dust and accentuate the edge of the cell walls on the viewing surface. Our results demonstrate the applicability of cryo-SEM techniques for the observation of water distribution within xylem at cellular and subcellular levels. The combination of cryo-SEM with non-destructive in situ observation techniques will profoundly improve the exploration of woody plant water flow dynamics.

Observing the water distribution within the xylem provides significant information regarding water flow dynamics in woody plants. In this study, we demonstrate the practical approach to observe xylem water distribution in situ&#160;by using a cryostat and cryo-SEM, which eliminates artifactual changes in the water status during sample preparation.

A scanning electron microscope installed cryo-unit (cryo-SEM) allows specimen observation at subzero temperatures and has been used for exploring water ...