This project has received funding from the European Union&#39;s Horizon 2020 research and innovation programme under grant agreement No. 634429. This publication reflects the views only of the author, and the European Commission cannot be held responsible for any use which may be made of the information contained therein. We also acknowledge the financial contribution made by the Real Jard&#237;n Bot&#225;nico, CSIC. SR is grateful to the European Union [ITN-SAPRO-238550] for the support of her research in Saprolegnia. We also want to thank Francisco Calonge for kindly provide the Phellorinia herculanea images and B. Pueyo for processing samples (Figure 5). All images were taken by the SEM service at the Real Jard&#237;n Bot&#225;nico-CSIC in Madrid.

NOTE: This protocol includes six main sections, three devoted to specific organisms (sections 1-3), and three describing the procedures common to all (4-6). Asterisks (*) indicate steps modified by the experimenters.
1. Studies of Developing and Fully Formed Plant Structures
<ol>
	<li>Collection and fixation
	<ol>
		<li>If the plant material is collected in a place with no access to a fume cupboard, introduce and immerse the material in 70% ethanol in centrifuge tubes. Ideally, immerse the material after 48 h in FAA (steps 1.1.1-1.1.3) to avoid excessive dehydration in the ethanol. If a fume cupboard is accessible to the plant material, ignore this step and continue with 1.1.1.</li>
		<li>Prepare the formalin-acetic-alcohol (FAA) fixative in a fume cupboard fitted with an aldehyde filter. Add 85 parts of 70% denatured ethanol, 10 parts of 60% formaldehyde solution, and 5 parts of glacial acetic acid. Prepare the FAA just before fixing the material, as its long term storage is not recommended40.</li>
		<li>Under the fume cupboard, pour the stock of FAA into individual wide-mouth and leak-proof plastic bottles. Use as many bottles as there are samples available, and create labels for sample identification.</li>
		<li>Select the floral or vegetative meristems to fix, ensuring that they are not damaged by insects, fungi, or extreme weather conditions. Cut the branches, removing unwanted material, and deposit the sample immediately in the FAA solution.</li>
		<li>After 72-96 h, pour the FAA into a plastic container for chemical disposal. Immediately, wash the samples three times with fresh 70% ethanol to remove any residual FAA. Fixed material can be stored indefinitely in 70% ethanol.</li>
	</ol>
	</li>
	<li>Dissection and dehydration
	<ol>
		<li>Dissect the fixed material in 70% ethanol under the stereomicroscope using ultra fine tweezers, needles, forceps, brushes, and micro-scalpels (the maximum size of the tissue should be around 1 cm3, or 2 cm for flat material). Dissect the samples into a Petri dish covered with ethanol to prevent the tissues from drying. Use a Petri dish with the base covered with dry black silicon to better see the contrasting white tissues.</li>
		<li>Put the dissected material in specimen containers for the critical point dryer (CPD, Figure 2a). At this point, immerse the containers into the Petri dish with 70% ethanol, and include the sample identification labels (made with paper and pencil). For more effective drying for further manipulation, avoid mixing the young and mature samples in the same container.*</li>
		<li>Put the lids on the containers and deposit them in plastic centrifuge tubes with plenty of 70% ethanol. Store the tubes overnight if the material is not processed immediately.</li>
		<li>Transfer the dissected material through the following ethanol series in hermetic jars or centrifuge tubes: 70%, 90%, 100%, and 100%. Leave the samples in each solution for 1 h at least. Keep the samples overnight in a 100% ethanol solution.</li>
		<li>Transfer the containers with the material to the CPD (section 4).</li>
	</ol>
	</li>
	<li>Mounting and preparing plant tissues for SEM observation
	<ol>
		<li>Write the sample identification number underneath the SEM sample holders (i.e., aluminum stubs). Cover the top of the stubs with double-sided tape. Place the stubs into a specimen holder (Figure 2b).</li>
		<li>Under a stereomicroscope, carefully open the containers carrying the young and delicate samples already dried in the CPD. Bear in mind that after the CPD treatment, the samples become lighter and sensitive to electrostatics. Close the containers once the samples have been taken out to avoid dust or impurities.</li>
		<li>Put the samples on the sticky surface of the stubs, planning ahead the desired position (once the samples touch the surface, it is very difficult to remove them). Do not try to carry a major dissection at this point; just remove unwanted tissue that is easy to pick up. For palynological studies, dissect the anthers and open them to expose the pollen on the stubs.</li>
		<li>Put long samples (e.g., 2 cm long) such as inflorescences in the horizontal position. When possible, orient samples of the same structure for polar, side, and bottom views. Leave enough space between samples on the stub.</li>
		<li>If the samples cannot be processed immediately, keep them protected overnight in a hermetic container with silica gel to avoid rehydration (Figure 2c)*. Coat the samples using the sputter coater and transfer them to the SEM (sections 5 and 6).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/55031/55031fig2.jpg" /> 
Figure 2: Tools for sample manipulation and processing before SEM observation. (a) Steel-made specimen container with holed walls for the ethanol/CO2 interchange in the CPD chamber. (b) Steel stubs within a plastic specimen holder. (c) Glass container used to keep the samples protected from humidity and dust. At the base, there is a compartment for silica gel. (d) Critical Point Dryer. In the front, there are (from left to right) the manometer, the power switch, the temperature control system, and the temperature display. Usual working pressure for CO2-ethanol interchange is 60 bars (800 psi). In the top, there are four valves (inlet, drain, ventilation, and exhaust controls) flanking the central sample chamber. Photos were taken by Y. Ruiz-Le&#243;n and M.A. Bello. <a href="http://cloudfront.jove.com/files/ftp_upload/55031/55031fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Study of Cyst Behavior of Saprolegnia (Oomycetes) on Different Surfaces
<ol>
	<li>Growing and fixing the cysts
	<ol>
		<li>Prepare peptone and glucose (PG-l) media41 using D-(+)-glucose (6 g) and mycological peptone (3 g)*. Add up to 900 mL of tap water and autoclave 40 min at 121 &#176;C. Pour 50 mL of the previously autoclaved solution A (NaH2PO4, 0.13 M) and 50 mL of solution B (Na2HPO4, 0.13 M).</li>
		<li>From stock cultures of strains of Saprolegnia parasitica maintained on peptone, glucose, agar media (PGA, which is prepared as PG-l but adding 10 g of European bacteriological agar to the glucose and the peptone before autoclave), grow mycelia colonies in 0.5 mL of PG-l droplets for 24-48 h at 20 &#176;C in Petri dishes. Induce sporulation by washing the mycelia with autoclaved tap water three times and incubating them for 15 h at 20 &#176;C42,43.</li>
		<li>Collect the released secondary zoospores by gently pipetting the upper part of the suspension and pool them in 1 mL portions. Agitate vigorously the zoospores for 30 s by vortexing to produce secondary cysts44.</li>
		<li>To test the differential growth of the spines of the cysts, on separate Petri dishes (p60), put 0.5 mL of the secondary cyst suspension onto different surfaces (i.e., carbon, gold, and copper TEM grids; salmon and hake fish scales (previously bleached); and glass cover slips)*. Incubate the cysts at 20 &#176;C for 70 min, which favors the attachment of the cysts to the surface.</li>
		<li>Remove the liquid and add 0.5 mL of 2% glutaraldehyde to each surface for the fixation of the cysts. Keep the samples at room temperature under a fume cupboard for 1 h.</li>
		<li>Remove the glutaraldehyde and dehydrate the sample through an ethanol series (30%, 40%, 50%, 60%, 70%, 80%, 90%, 100%, and 100%), adding 5 mL of each ethanol solution for 15 min. Once in the last 100% ethanol solution, the sample can be stored for up to a month in a sealed Petri dish. At this stage, the samples are ready to be dried in the CPD.</li>
		<li>Carefully transfer the grids and the scales from the Petri dish to a holder suitable for the CPD (Section 4). For this step, use a CPD grid holder or a stacking specimen holder, which keep samples separated from each other. Take the grids and scales with tweezers, keeping in mind that the cysts should be facing up on the grids all the time.*</li>
	</ol>
	</li>
	<li>Mounting and preparing cyst samples for SEM observation
	<ol>
		<li>Mount the grids and the scales on the aluminum stubs that were previously covered with double-sided carbon tape and labelled underneath.</li>
		<li>Transfer the samples to the sputter coater (section 5).</li>
		<li>Observe the samples under the SEM (section 6).</li>
	</ol>
	</li>
</ol>
3. Study of Herbarium Fungal Spores of Phellorinia herculanea under SEM
<ol>
	<li>Rehydration and dehydration of spores
	<ol>
		<li>Wrap each sample carefully with filter paper, forming pencil-labeled envelopes ~0.5-1 cm2, taking care not to crush them. Seal the filter paper with paper clips. Transfer the packed samples to a Petri dish and immerse them in 10 mL of water to rehydrate tissues around the spores.</li>
		<li>Immediately put the samples in a microwave (600 W for approximately 20 s). Remove the material once the water starts to evaporate, and allow it to cool down at room temperature.</li>
		<li>Pass the samples through the following ethanol series: 30%, 50%, 70%, 80%, 90%, 95%, 100%, and 100%. Depending on the amount of samples, use a beaker or centrifuge tubes for this step. Leave the samples for 15 min in each solution.</li>
		<li>Place the samples to the CPD (section 4).</li>
	</ol>
	</li>
	<li>Mounting and preparing spores for SEM observation
	<ol>
		<li>Open the envelopes. Pour the spores on a previously-prepared stub with double-sided tape. Alternatively, collect the spores with the sticky surface of the stubs, taking care not to crush them*.</li>
		<li>If the samples contain few spores, in addition to the previous step, cut a small piece of the envelope (~1 mm2) and place it on a new stub*.</li>
		<li>Place the tissues into the sputter coater (section 5).</li>
		<li>Observe under the SEM (section 6).</li>
	</ol>
	</li>
</ol>
4. Drying of Material Using a Critical Point Dryer (CPD, Figure 2d)
<ol>
	<li>Use the CPD in a ventilated area and verify that all the valves of the machine are closed. Check if the sample chamber is empty and clean.</li>
	<li>Switch on the machine and verify that the temperature control system test takes place automatically. If the CPD has an external refrigeration bath system, check the water levels before switching it on.</li>
	<li>Follow the manufacturer&#39;s instructions of the specific CPD used for the ethanol and CO2 interchange. For safety, carry on this step under the supervision of someone trained for the use of the machine. Remember that it is exposed to rapid pressure changes, it could blow out violently.</li>
	<li>Take out the samples and continue with the step 1.3 if working with plant tissues, step 2.2 if working with oomycetes cysts, and steps 3.2 if working with fungi spores.</li>
</ol>
5. Coating the Samples with Gold Using the Sputter Coater (Figure 3a)
<ol>
	<li>Check the sputter coater. Verify that the gold cathode target is in good condition. Use a lint-free cloth drenched with 90% ethanol to clean the walls of the vacuum chamber and the chamber lid if necessary.</li>
	<li>Mark the sputter holder with numbers beside each stub hole for further identification of the samples under the microscope. Carefully, place the stubs loaded with the samples and secure them. Use a rotary planetary specimen stage to ensure a uniform coating on specimens with irregular surfaces.</li>
	<li>Follow the manufacturer&#39;s instructions to adjust settings such as the working distance (e.g., 30 mm), operation gas pressure (e.g., 5 x 10-1 - 7 x 10-1 mbar), the sputtering time (e.g., 50 s), thickness of the gold layer (e.g., 12 nm) the current (e.g., 15 mA) and the voltage supply (e.g., 600 V)45.</li>
	<li>Remove the stubs and take them to the SEM (section 6). Alternatively, place the stubs into a sealed container with silica gel (Figure 2c).</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/55031/55031fig3.jpg" /> 
Figure 3: Sputter coater (a) and scanning electron microscope (b). (a) Front view of the vacuum chamber (left), gas valve, timer, vacuum, and current controls. (b) Side view of the SEM main components (from left to right): the vacuum column with the sample chamber, the computer screen with the controls, and the chamber&#39;s monitor. Photos were taken by Y. Ruiz-Le&#243;n. <a href="http://cloudfront.jove.com/files/ftp_upload/55031/55031fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Observation under the Scanning Electron Microscope (SEM, Figure 3b)
<ol>
	<li>SEM start up
	<ol>
		<li>Follow the manufacturer&#39;s instructions to start and set the SEM, adjusting the sample height the objective aperture diameter (e.g., for plants 2 &#181;m and for fungi and oomycetes 4 &#181;m), the operating voltage (e.g., 15 kV).</li>
		<li>Check the correct alignment of the electron beam system and set the axial alignment and the stigmators according to the manufacturers indications. Adjust the working distance in order to obtain an adequate depth of field.</li>
	</ol>
	</li>
	<li>Image capture
	<ol>
		<li>Get a focused image of the sample and use it as a starting point. Increase the magnification close to the maximum level and focus the image again. Choose areas with surface irregularities such as holes. Correct astigmatism and adjust the optimum contrast and brightness.</li>
		<li>Capture the SEM Image with the high-resolution. Use the BSE detector if the image shows that the samples are charged. Otherwise, set the SE detector. Change the detectors following the manufacturer&#39;s instructions.</li>
	</ol>
	</li>
</ol>

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<li>Calonge, F. D., Martínez, A. J., Falcó, I., Samper, L. E. <a target="_blank" href="http://bddoc.csic.es:8080/detalles.html?id=209351&bd=ICYT&tabla=docu">Phellorinia herculanea f. stellata f. nova encontrada en España.</a> Bol. Soc. Micol.Madrid. 35 (1), 65-70 (2011).</li>
<li>Liu, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Deciphering+microbial+landscapes+of+fish+eggs+to+mitigate+emerging+diseases%5BTitle%5D)+AND+%22ISME+J%22%5BJournal%5D)">Deciphering microbial landscapes of fish eggs to mitigate emerging diseases.</a> ISME J. 8 (10), 2002-2014 (2014).</li>
<li>Sandoval-Sierra, J. V., Diéguez-Uribeondo, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+comprehensive+protocol+for+improving+the+description+of+Saprolegniales+%28Oomycota%29%3A+two+practical+examples+%28Saprolegnia+aenigmatica+sp.+nov.+and+Saprolegnia+racemosa+sp.+nov.%29%5BTitle%5D)+AND+%22PLOS+one%22%5BJournal%5D)">A comprehensive protocol for improving the description of Saprolegniales (Oomycota): two practical examples (Saprolegnia aenigmatica sp. nov. and Saprolegnia racemosa sp. nov.).</a> PLOS one. , (2015).</li>
<li>Endress, P. K. <a target="_blank" href="http://agris.fao.org/agris-search/search.do?recordID=US201303213009">Zur vergleichenden Entwicklungsmorphologie, Embryologie und Systematik bei Laurales.</a> Bot. Jahrb. Syst. 92 (2), 331-428 (1972).</li>
<li>Tucker, S. <a target="_blank" href="https://www.jstor.org/stable/2441644">Floral development in Saururus cernuus (Saururaceae):1. Floral initiation and stamen development.</a> Am. J. Bot. 62 (3), 993-1005 (1975).</li>
<li>Endress, P. K., Matthews, M. L. <a target="_blank" href="http://link.springer.com/article/10.1007/s00606-011-0576-2">Progress and problems in the assessment of flower morphology in higher-level systematics.</a> Plant Syst. Evol. 298 (2), 257-276 (2012).</li>
<li>Beakes, G. W., Glockling, S. L., Sekimoto, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+evolutionary+phylogeny+of+the+oomycete+%26%2334%3Bfungi%26%2334%5BTitle%5D)+AND+%22Protoplasma%22%5BJournal%5D)">The evolutionary phylogeny of the oomycete &#34;fungi&#34.</a> Protoplasma. 249 (1), 3-19 (2012).</li>
<li>Romansic, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effects+of+the+pathogenic+water+mold+Saprolegnia+ferax+on+survival+of+amphibian+larvae%5BTitle%5D)+AND+%22Dis.+Aquat.+Organ%22%5BJournal%5D)">Effects of the pathogenic water mold Saprolegnia ferax on survival of amphibian larvae.</a> Dis. Aquat. Organ. 83 (3), 187-193 (2009).</li>
<li>van West, P. <a target="_blank" href="http://www.sciencedirect.com/science/article/pii/S0269915X0600053X">Saprolegnia parasitica, an oomycete pathogen with a fishy appetite: new challengues for an old problem.</a> Mycologist. 20 (3), 99-104 (2006).</li>
<li>Johansen, D. A. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aJohansen%2C+DA+%22Plant+microtechnique%22">Plant microtechnique</a>. , McGrow-Hill. New York. (1940).</li>
<li>Unestam, T. <a target="_blank" href="http://onlinelibrary.wiley.com/doi/10.1111/j.1399-3054.1965.tb06911.x/full">Studies on the crayfish plague fungus Aphanomyces astaci. Some factors affecting growth in vitro.</a> Physiol. Plantarum. 18 (2), 483-505 (1965).</li>
<li>Cerenius, L., Söderhäll, K. <a target="_blank" href="https://www.researchgate.net/publication/222195448_Repeated_zoospore_emergence_from_isolated_spore_cysts_ofAphanomyces_astaci">Repeated zoospore emergence from isolated spore cysts of Aphanomyces astaci.</a> Exp. Mycol. 8 (4), 370-377 (1984).</li>
<li>Diéguez-Uribeondo, J., Cerenius, L., Söderhäll, K. <a target="_blank" href="http://www.sciencedirect.com/science/article/pii/S0953756209810605">Repeated zoospore emergence in Saprolegnia parasitica.</a> Mycol. Res. 98 (7), 810-815 (1994).</li>
<li>Söderhäll, K., Svensson, E., Unestam, T. <a target="_blank" href="http://link.springer.com/article/10.1007/BF00443081">Chitinase and protease activities in germinating zoospore cysts of a parasitic fungus, Aphanomyces astaci, Oomycetes.</a> Mycopathologia. 64 (1), 9-11 (1978).</li>
<li>Echlin, P. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aEchlin%2C+P+%22Handbook+of+sample+preparation+for+scanning+electron+microscopy+and+X-Ray+Microanalysis%22">Handbook of sample preparation for scanning electron microscopy and X-Ray Microanalysis</a>. , Springer Science + Business Media, LLC. NY. (2009).</li>
<li>Osumi, M., et al. <a target="_blank" href="https://academic.oup.com/jmicro/article-abstract/32/4/321/837464/Preparation-for-Observation-of-Fine-Structure-of">Preparation for observation of fine structure of biological specimens by high-resolution SEM.</a> Microscopy. 32 (4), 321-330 (1983).</li>
<li>Rezinciuc, S. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=%22The+Saprolegniales+morpho-molecular+puzzle%3A+an+insight+into+markers+identifying+specific+and+subspecific+levels+in+main+parasites%22">The Saprolegniales morpho-molecular puzzle: an insight into markers identifying specific and subspecific levels in main parasites</a>. , Universidad Internacional Menéndez Pelayo. Doctoral Thesis (2013).</li>
</ol>

The authors have nothing to disclose.

With respect to standard SEM protocols, the procedures presented here include relatively rapid, easy to follow, and low-cost methodologies. Depending on the amount of samples and on the ease of processing, it takes four to five days to acquire good quality images. Including adequate safety precautions for the CPD and SEM operation, the procedures are easy to handle. Particular caution should be taken with formalin and the glutaraldehyde (see steps 1.1.1 to 1.1.3 and 2.1.5 of the protocol). There are certain steps where, if necessary, the process can be stopped for a long time without damaging the samples or ruining previous steps (e.g., steps 1.1.5, 1.2.3, and 1.3.5). In terms of cost, much of the equipment can be reused for several preparations (e.g., CPD aluminum containers and stub holders), and the reagents are non-expensive chemicals available from all commercial suppliers.
Disadvantages of these procedures are the need for proper chemical waste disposal for the aldehydes and the ethanol and the lack of key supplies and the limitations of the technique. Materials such as the gold disk for the sputter coater and the CO2 cylinder for the CPD should be checked well in advance. If they are in continuous use, the laboratory will require multiple stocks of them, which eventually elevates the costs. Because individual researchers often cannot justify the expenses derived from the maintenance of these types of supplies, or they simply do not need to use the SEM continuously, these procedures nowadays tend to be performed in laboratories with external electron microscopy services.
The observation technique of the SEM is limited to a high magnification study of surfaces. If internal tissues need to be observed, the samples should be cut in the proper way to explore the external appearance of the internal tissues. To explore internal aspects of the cells at high magnification, a transmission electron microscope (TEM) is required. Critical steps within the protocol are the proper fixation and dehydration of the samples before CPD treatment, where it is crucial to keep the tissues safe from shocking changes and direct contact with the air. Also, careful management of the pressure change in the CPD sample chamber and the amount of coating deposited on different types of samples is important.
Despite these limitations and special manipulations, the study of biological samples with SEM following the protocols described here allows for the resolution of some common problems, such as the cell wall and organ distortion (Figures 1, 4, 5), the restrictions on the observation and growth of structures coming from wet and liquid environments (Figures 5, 6), and the destruction of sensitive cells (Figures 1c, 1d).
The protocols presented here have allowed the reinterpretation and questioning of traditional taxonomic characteristics, such as the spine structure in cysts of Saprolegnia47. The differential growth pattern of the Saprolegnia spines on different surfaces demonstrates the lability of this feature and its limitations for species diagnosis. In addition to the capture of relevant characteristics for taxonomic studies, stages of organ development, and infectious diseases, functions of tissues never observed before have been explored thanks to these techniques. Now, this protocol can be extended to thousands of case studies waiting to be examined33. According to the peer-reviewed literature database Scopus, in the last the last five years there have been 7,425 publications of papers dealing with SEM imaging of plants (4,914), oomycetes (21) and fungi (2,490). This fact suggests that research in oomycetes using the SEM imaging is still very scarce in comparison with plants and fungi. With ESEM, the numbers are even lower. There are 588 manuscripts in the same time span: 337 in plants, 1 in oomycetes and 250 in fungi.

In biology, the use of scanning electron microscopy (SEM) has been extended to studies of structural evolution, comparative morphology, organ development, and characterization of populations or species1. With its two-dimensional view of microscopic structures, areas such as micromorphology and systematics profited from SEM technique advances since the second half of the 20th century. For example, the introduction of the sputter coating methodology in the 1970s made possible observations of delicate materials such as shoot apices and flowers enhancing the imaging of non-conductive tissues2,3. SEM uses electrons ejected from the surface of the specimen to reproduce the topography in a high-vacuum environment4.
Studies involving SEM are focused in both the inference of structural characters and the reconstruction of growth processes. New structural characters relevant to the taxonomy and systematics of a wide range of organisms have been discovered from SEM observations. For example, plant traits used for species diagnosis or supraspecific classifications, such as the vestured pits of wood5, stigma diversity6, nectary and floral morphology7,8, trichome details9, and pollen grains10,11, cannot be properly visualized without SEM. Successful observations with conventional SEM have been also achieved for long-time formalin-fixed organisms12 and plant herbarium specimens13.
On the other hand, studies of reconstruction of growth processes using SEM involve a wide range of topics, such as organ development14, infections induced by bacteria15, plant root physiology16, parasite-host attachment mechanisms17,18, drug effects on parasites19, mycoparasitism and antibiosis20,21, growth malformation22, comparative development of wild and mutant individuals23, and entire life cycles24. Although environmental scanning electron microscopes (ESEM)25 may have important advantages for the observation of wet biological samples in growth processes, delicate material may still be compromised even in the low vacuum condition of the ESEM), and need to be processed adequately to avoid loss of valuable morphological observation.
In this paper, a review of specific protocols for SEM observation of three different types of samples is presented: floral meristems, oomycetes (Saprolegnia), and fungal material. These protocols compile the experience of our previous SEM-based studies26,27,28,29,30,31,32,33, where specific difficulties and alternative solutions have been found. In the case of plant comparative developmental and structural studies, the use of SEM started in the 1970s34,35, and since then, researchers discovered that certain floral features are more labile than previously thought36. Reconstruction of floral development involves the capture of all stages between young floral meristems and anthesis. To reach this aim, it is essential that the sample topography and the cell wall integrity are not compromised after the fixation and subsequent dehydration. Young floral meristems are particularly vulnerable to cell wall collapse (Figures 1a, 1b). Similarly, delicate structures such as nectaries, petals, stigmas and sporangia require effective and undamaging protocols. This review summarizes an optimal protocol to keep young and delicate tissues intact for SEM imaging.
In the case of the oomycetes (Stramenopiles)-one of the most diverse and widespread groups of parasites, with hosts ranging from microbes and plants to invertebrates and vertebrates37- there are spores that grow and develop in a wet environment. This condition represents a challenge for SEM observation because the spores need an adequate substrate not suitable for standard SEM protocols. Among the oomycetes, species of Saprolegnia are of particular interest because they can cause severe reductions in aquacultures, fisheries, and amphibian populations38. Micromorphological characteristics, such as the hooked spines of cysts, have been found to be useful to identify species of Saprolegnia, which is fundamental to establish infection controls and potential treatments39. Here, there is an experimental protocol to compare the patterns of the spine growth of cysts on different substrates and to manipulate the sample for critical point dryer (CPD) preparation and subsequent SEM observation.
In a third case, there are interesting findings that came up after an inspection of the spores of the fungi Phellorinia herculanea f. stellata f. nova (Agaricales)31. Together with the spores, a group of unexpected nursery cells was identified under the SEM. With previous traditional protocols and untreated material, the nurse cells came out completely collapsed (Figure 1c). Further inferences about particular tissues associated to the spores can be made with the simple but crucial modifications to the standard approaches described here (Figure 1d).
In this review, there are detailed SEM protocols that can be used to deal with different problems associated with SEM observation in angiosperms, oomycetes, and Agaricales, such as cell collapse and meristematic tissue shrinking, non-optimal growth of cyst spines, and destruction of ephemeral tissues, respectively.
<img alt="Figure 1" src="/files/ftp_upload/55031/55031fig1.jpg" /> 
Figure 1: Comparison of samples treated without (a, c) and with (b, d) the protocol FAA-ethanol-CPD. (a-b) Floral buds of Anacyclus clavatus, mid-development. Bud treated with osmium tetroxide46 (a) and bud treated with the FAA-CPD protocol (b). (c-d) Nurse cells with spores of Phellorinia herculanea f. stellata. Dried samples without any treatment (c) and with the protocol here described for Agaricales (d). Spores in orange. Scales: (a-b) 100 &#181;m, (c-d) 50 &#181;m. Photos were taken by Y. Ruiz-Le&#243;n. <a href="http://cloudflare.jove.com/files/ftp_upload/55031/55031fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>Acetic acid</td><td>No specific supplier</td><td></td><td>Skin irritation, eye irritation</td></tr><tr><td>aluminium stubs</td><td>Ted Pella, Inc.</td><td>16221</td><td>www.tedpella.com</td></tr><tr><td>Centrifuge tubes</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Critical Point Dryer</td><td>Polaron Quatum Technologies</td><td>CPD7501</td><td></td></tr><tr><td>D-(+)-Glucose</td><td>Merck</td><td>1,083,421,000</td><td></td></tr><tr><td>Double sided sellotape</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Ethanol absolute</td><td>No specific supplier</td><td></td><td>Flammable</td></tr><tr><td>European bacteriological agar</td><td>Conda</td><td>1800.00</td><td>www.condalab.com</td></tr><tr><td>Filter paper</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Forceps</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Formalin 4%</td><td>No specific supplier</td><td></td><td>Harmful, acute toxicity, skin sensitisation, carcinogenicity. Flammable</td></tr><tr><td>Glass cover slips</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Glass hermetic container&amp;nbsp;</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Glutaraldehyde 25% DC 253857.1611&amp;nbsp; (L)</td><td>Dismadel S.L.</td><td>3336</td><td>www.dismadel.com</td></tr><tr><td>Mycological peptone</td><td>Conda</td><td>1922.00</td><td>www.condalab.com</td></tr><tr><td>needles</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Petri dishes</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Plastic containers</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Sample holder with lid&amp;nbsp; for the critical point dryer&amp;nbsp;</td><td>Ted Pella, Inc.</td><td>4591</td><td>www.tedpella.com</td></tr><tr><td>scalpels</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Scanning Electron Microscope</td><td>Hitachi</td><td>S3000N</td><td></td></tr><tr><td>Software for SEM</td><td></td><td></td><td></td></tr><tr><td>Solution A: NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td></td><td></td><td></td></tr><tr><td>Solution B: Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td></td><td></td><td></td></tr><tr><td>Specimen holders</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Sputter coater</td><td>Balzers</td><td>SCD 004</td><td></td></tr><tr><td>Stereomicroscope</td><td>No specific supplier</td><td></td><td></td></tr><tr><td>Transmission Electron Microscope (TEM) grids</td><td>Electron Microscopy Sciences</td><td>G200 (Square Mesh)</td><td>www.emsdiassum.com</td></tr><tr><td>Tweezers</td><td>No specific supplier</td><td></td><td></td></tr></tbody></table>

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.

Floral Development and Fixation of Developing and Fully Formed Plant Structures
Using the FAA-CPD protocol described here, young and mature plant tissues are optimally fixed and dehydrated for SEM imaging. Processes such as floral development can be reconstructed because the topography and shape of the buds is not distorted by cell shrinking (Figures 1b, 1d, 4a-f). Structures with complex shapes can be successfully covered with a uniform layer of conductive material (the metal from the sputter coater), allowing the recovery of otherwise unreachable details (Figures 4g-I, 5e). Good quality images of samples coming from wet microenvironments, which used to be overcharged with the electron current if badly dehydrated and coated, can be achieved (Figures 4f, 5a, 5b). Important details such as the pollen wall surface (Figure 5b, 5c) and different types of indumenta (Figures 5d-g) can be studied in depth without artificial or undesirable particles or distorted shapes.
<img alt="Figure 4" src="/files/ftp_upload/55031/55031fig4.jpg" /> 
Figure 4: Early (a, b), mid (c-e), and late (f-i) floral development captured under the SEM. (a) Raceme of Caesalpinia spinosa (Molina) Kuntze with several floral meristems. (b) Top view of a young raceme Polygala violacea Aubl. (c) Floral bud of Krameria ixine Loefl. during gynoecium differentiation. (d-e) Buds of Erythrina sp. (f) Side view of a flower of Krameria ixine. The anthers and style protrude. (g-i) Hoya carnosa (L.) R. Brown. (g) Flower at anthesis (digital image taken with a Olympus SP-590UZ 26X). (h) Top view of the stamens and split carpels. (i) Side view of a pair of stamens flanking two carpels. Asterisks indicate stamens. g = gynoecium. Scales: (a,i) 1 mm, (b, d) 400 &#181;m, (c) 200 &#181;m, (e) 500 &#181;m, (f, h) 2 mm, (g) 0.25 cm. Photos were taken by M.A. Bello. <a href="http://cloudflare.jove.com/files/ftp_upload/55031/55031fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55031/55031fig5.jpg" /> 
Figure 5: SEM micrographs of sporangia and pollen (a, b), detailed surfaces (c-e), and indumenta (f-g). (a) Sporangia of Dryopteris sp. (b) Pollen of Prunus dulcis D. A. Webb landing on stigma. (c) Pollen of Nepenthes alata Blanco. (d) Side view of an inflorescence of Erigeron karvinskianus DC. Image taken with BSE detector option. (e) Side view of the bryophyte Peristoma sp. (f) Glandular and no-glandular trichomes on the abaxial surface of a leaf of Rosmarinus officinalis L. (g) Top view of the leaf flat scales of Olea europaea L. Scales: (a) 400 &#181;m, (b) 60 &#181;m, (c) 5 &#181;m, (d) 4 mm, (e) 600 &#181;m, (f) 200 &#181;m, (g) 600 &#181;m. Photos were taken by Y. Ruiz-Le&#243;n. <a href="http://cloudflare.jove.com/files/ftp_upload/55031/55031fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Effect of Different Substrates on the Spine Growth of Secondary Cysts Saprolegnia parasitica 
The spine growth pattern was different on different substrates. On glass, the spines were elongated and showed loops at their tips (Table 1, Figure 6a). On the carbon and gold grids the spines are shorter, curled and grow more abundantly around the cysts without forming loops (Figures 6b, 6d). Although on copper grids, the spines tended to curl without looping, some of them elongated (Figure 6c). On fish scales, the spines were curly, abundant around the surface and sometimes formed looped ends (Figure 6e-f).
<img alt="Figure 6" src="/files/ftp_upload/55031/55031fig6.jpg" /> 
Figure 6: Differential growth patterns of the spines of Saprolegnia parasitica in cysts immersed in liquid media. (a) Glass. (b) Carbon. (c) Copper. (d) Gold. (e) Hake scale. (f) Salmon scale. Straight spines were developed on glass and copper, whereas curled spines were formed on carbon, gold, and fish scales. The hooked tips (white arrows) of the spines are observable in all spines growing on glass and in few spines on fish scales. The cyst walls are in yellow. Scale: 20 &#181;m. Photos were taken by Y. Ruiz-Le&#243;n and S. Rezinciuc. <a href="http://cloudflare.jove.com/files/ftp_upload/55031/55031fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Surface</td>
			<td>Spine morphology</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>Elongation</td>
			<td>Loop at the tip</td>
		</tr>
		<tr>
			<td>Glass cover</td>
			<td>Good elongation, spines easy to count</td>
			<td>Present</td>
		</tr>
		<tr>
			<td>Fish scales</td>
			<td>Spines with curly shape not easy to count</td>
			<td>Present</td>
		</tr>
		<tr>
			<td>TEM grids</td>
			<td>Short and curly spines not easy to count</td>
			<td>Absent</td>
		</tr>
	</tbody>
</table>
Table 1: Spine morphology using different types of substrate.
Unexpected Observations in Phellorinia herculanea f. stellata
Apart from the unexpected finding of laticiferous hyphae on the exoperidium of P. herculanea, Calonge et al.31 found globose and smooth nurse cells (8-13 &#181;m) mixed with spores in the material dehydrated with CPD thanks to the rehydration step performed in the microwave. (Figure 1d). The structure and details of the walls of the nurse cells and the spores are well preserved despite their differential composition (Figures 1c-d).

Watch this Scientific Journal Video about Scanning Electron Microscopy SEM Protocols for Problematic Plant, Oomycete, and Fungal Samples at JoVE.com

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.

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 ...

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Biology

28.9K Views.  Real Jardín Botánico, CSIC. 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. 

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

Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample preparation for transmission electron microscopy (TEM) such investigations can take several days1,2 and are therefore not suited for a rapid diagnosis of plant virus diseases. Microwave fixation can be used to drastically reduce sample preparation time for TEM investigations with similar ultrastructural results as observed after conventionally sample preparation3-5. Many different custom made microwave devices are currently available which can be used for the successful fixation and embedding of biological samples for TEM investigations5-8. In this study we demonstrate on Tobacco Mosaic Virus (TMV) infected Nicotiana tabacum plants that it is possible to diagnose ultrastructural alterations in leaves in about half a day by using microwave assisted sample preparation for TEM. We have chosen to perform this study with a commercially available microwave device as it performs sample preparation almost fully automatically5 in contrast to the other available devices where many steps still have to be performed manually6-8 and are therefore more time and labor consuming. As sample preparation is performed fully automatically negative staining of viral particles in the sap of the remaining TMV-infected leaves and the following examination of ultrastructure and size can be performed during fixation and embedding.

This study describes a method that allows the rapid and clear diagnosis of plant virus diseases in about half a day by using a combination of microwave assisted plant sample preparation for transmission electron microscopy and negative staining methods.

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample ...

Immunology and Infection

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

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

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

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

Environment

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

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

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

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

Leveraging Micro-CT Scanning to Analyze Parasitic Plant-Host Interactions

Micro-CT scanning has become an established tool in investigating plant structure and function. Its non-destructive nature, combined with the possibility of three-dimensional visualization and virtual sectioning, has allowed novel and increasingly detailed analysis of complex plant organs. Interactions among plants, including between parasitic plants and their hosts, can also be explored. However, sample preparation before scanning becomes crucial due to the interaction between these plants, which often differ in tissue organization and composition. Furthermore, the broad diversity of parasitic flowering plants, ranging from highly reduced vegetative bodies to trees, herbs, and shrubs, must be considered during the sampling, treatment, and preparation of parasite-host material. Here two different approaches are described for introducing contrast solutions into the parasite and/or host plants, focusing on analyzing the haustorium. This organ promotes connection and communication between the two plants. Following a simple approach, details of haustorium tissue organization can be explored three-dimensionally, as shown here for&#160;euphytoid, vine, and mistletoe parasitic species. Selecting specific contrasting agents and application approaches also allow detailed observation of endoparasite spread within the host body and detection of direct vessel-to-vessel connection between parasite and host, as shown here for an obligate root parasite. Thus, the protocol discussed here can be applied to the broad diversity of parasitic flowering plants to advance the understanding of their development, structure, and functioning.

Micro-CT is a non-destructive tool that can analyze plant structures in three dimensions. The present protocol describes the sample preparation to leverage micro-CT to analyze parasitic plant structure and function. Different species are used to highlight the advantages of this method when coupled with specific preparations.

Micro-CT scanning has become an established tool in investigating plant structure and function. Its non-destructive nature, combined with the possibility ...

A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic cooperation, competition, or predation. The study of biofilms is important in many biological fields, including environmental science, food production, and medicine. However, few studies have focused on such bacterial biofilms, partially due to the difficulty of investigating them. Most of the methods for qualitative and quantitative biofilm analyses described in the literature are only suitable for biofilms forming on abiotic surfaces or on homogeneous and thin biotic surfaces, such as a monolayer of epithelial cells.
While laser scanning confocal microscopy (LSCM) is often used to analyze in situ and in vivo biofilms, this technology becomes very challenging when applied to bacterial biofilms on fungal hyphae, due to the thickness and the three dimensions of the hyphal networks. To overcome this shortcoming, we developed a protocol combining microscopy with a method to limit the accumulation of hyphal layers in fungal colonies. Using this method, we were able to investigate the development of bacterial biofilms on fungal hyphae at multiple scales using both LSCM and scanning electron microscopy (SEM). This report describes the protocol, including microorganism cultures, bacterial biofilm formation conditions, biofilm staining, and LSCM and SEM visualizations.

This protocol describes a new method to grow and qualitatively analyze bacterial biofilms on fungal hyphae by confocal and electron microscopy.

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic ...

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 ...

Targeted Studies Using Serial Block Face and Focused Ion Beam Scan Electron Microscopy

This protocol allows for the efficient and effective imaging of cell or tissue samples in three dimensions at the resolution level of electron microscopy. For many years electron microscopy (EM) has remained an inherently two-dimensional technique. With the advent of serial scanning electron microscope imaging techniques (volume EM), using either an integrated microtome or focused ion beam to slice then view embedded tissues, the third dimension becomes easily accessible. Serial block face scanning electron microscopy (SBF-SEM) uses an ultramicrotome enclosed in the SEM chamber. It has the capability to handle large specimens (1,000 &#181;m x 1,000 &#181;m) and image large fields of view at small X,Y pixel size, but is limited in the Z dimension by the diamond knife. Focused ion beam SEM (FIB-SEM) is not limited in 3D resolution, (isotropic voxels of &#8804;5 nm are achievable), but the field of view is much more limited. This protocol demonstrates a workflow for combining the two techniques to allow for finding individual regions of interest (ROIs) in a large field and then imaging the subsequent targeted volume at high isotropic voxel resolution. Preparing fixed cells or tissues is more demanding for volume EM techniques due to the extra contrasting needed for efficient signal generation in SEM imaging. Such protocols are time consuming and labor intensive. This protocol also incorporates microwave assisted tissue processing facilitating the penetration of reagents, which reduces the time needed for the processing protocol from days to hours.

Here, we present a protocol for efficiently combining serial block face and focused ion beam scanning electron microscopy to target an area of interest. This allows for efficient searching, in three dimensions, and locating rare events in a large field of view.

This protocol allows for the efficient and effective imaging of cell or tissue samples in three dimensions at the resolution level of electron microscopy. ...

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

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

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

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

 Microscale SEM Analysis of Microbial Communities in Fungus-Growing Ant Cultures

Microbiota of Attine Ants' Gardens: Visualizing a Microbial Landscape by Scanning Electron Microscopy

In macroscale ecosystems, such as rainforests or coral reefs, the spatial localization of organisms is the basis of our understanding of community ecology. In the microbial world, likewise, microscale ecosystems are far from a random and homogeneous mixture of organisms and habitats. Accessing the spatial distribution of microbes is fundamental for understanding the functioning and ecology of the microbiota, as cohabiting species are more likely to interact and influence each other&#39;s physiology.
An interkingdom microbial ecosystem is at the core of fungus-growing ant colonies, which cultivate basidiomycete fungi as a nutritional resource. Attine ants forage for diverse substrates (mostly plant-based), metabolized by the cultivated fungus while forming a spongy structure, a &#34;microbial garden&#34; that acts as an external gut. The garden is an intertwined mesh of fungal hyphae growing by metabolizing the substrate, opening niches for a characteristic and adapted microbiota to establish. The microbiota is thought to be a contributor to substrate degradation and fungal growth, though its spatial organization is yet to be determined.
Here, we describe how we employ Scanning Electron Microscopy (SEM) to investigate, with unprecedented detail, the microbiota and biofilm spatial organization across different fungiculture systems of fungus-growing ants. SEM imaging has provided a description of the microbiota spatial structure and organization. SEM revealed that microbiota commonly assemble in biofilms, a widespread structure of the microbial landscapes in fungiculture. We present the protocols employed to fix, dehydrate, dry, sputter coating, and image such a complex community. These protocols were optimized to deal with delicate and heterogeneous samples, comprising plant and fungal biomass, as well as the microbiota and the biofilm.

We propose an optimized Scanning Electron Microscopy protocol for visualizing highly heterogeneous and delicate samples containing plant and fungal biomass, together with microbiota and biofilm. This protocol allows describing the spatial dimensions of the microbiota organization.

In macroscale ecosystems, such as rainforests or coral reefs, the spatial localization of organisms is the basis of our understanding of community ...

SEM Imaging of Biological Samples

Source: <a href="https://www.jove.com/author/Peiman_Shahbeigi-Roodposhti">Peiman Shahbeigi-Roodposhti</a> and Sina Shahbazmohamadi, Biomedical Engineering Department, University of Connecticut, Storrs, Connecticut
A scanning electron microscope (SEM) is an instrument that uses an electron beam to nondestructively image and characterize conductive materials in a vacuum. As an analogy, an electron beam is to the SEM as light is to the optical microscope. The difference is that the electron microscope yields images of much higher resolution and magnification. The best optical microscopes typically have a resolution down to 200 nm, whereas SEMs usually claim a resolution of 0.5 nm. This is due to the fact that optical microscopes are limited by the diffraction of waves, a function of the wavelength, which is around 500 nm for visible light. Conversely, the SEM uses an energized electron beam, which as a wavelength of 1 nm. This characteristic makes them very dependable tools for the study of nano and microstructures. Electron microscopes also enable the study of biological samples with feature sizes too small for optical microscopy.
This demonstration provides an introduction to sample preparation and initial image acquisition of biological samples using a scanning electron microscope. In this case, a collagen-hydroxyapatite (HA) cellular scaffold will be studied. The vacuum environment of the SEM and the induced charging by the electron beam on non-conductive samples (such as organic matter) creates challenges that will be addressed in the preparation. The advantages and disadvantages of different imaging methods as they relate to resolution, depth of focus and sample type will also be discussed. The purpose of this demonstration is to give the participant more information on SEM to determine if this microscopy module is the best fit for a type of biological sample.

Scanning Electron Microscopy: Operations and Sample Preparation

Source: Peiman Shahbeigi-Roodposhti and Sina Shahbazmohamadi, Biomedical Engineering Department, University of Connecticut, Storrs, Connecticut
A scanning ...

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

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