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

Biology

Your institution must subscribe to JoVE's Biology section to access this content.

Fill out the form below to receive a free trial or learn more about access:

 

Summary

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.

Cite this Article

Copy Citation | Download Citations | Reprints and Permissions

Koon, M. A., Almohammed Ali, K., Speaker, R. M., McGrath, J. P., Linton, E. W., Steinhilb, M. L. Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM). J. Vis. Exp. (143), e58761, doi:10.3791/58761 (2019).

Abstract

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.

Introduction

A scanning electron microscope (SEM) uses a focused beam of high-energy electrons to generate an image from secondary electrons that shows the morphology and topography of a sample1. SEM can be used to directly determine the physical size of a sample, the surface structure, and the three-dimensional shape, and offers greater resolution and larger depth of field compared to light microscopy. Another form of electron microscopy (EM), transmission electron microscopy (TEM) uses focused electrons that pass through the sample, generating images with fine details of internal structure. While TEM has higher resolution than light or SEM and can be used to resolve structures as small as single atoms, it has three major disadvantages: extensive sample preparation, a small field of view, and a shallow depth of field2,3. Although other visualized protocols exist using SEM to examine specific cells, organelles, or tissues4,5,6,7,8,9,10 , this protocol is unique in that we describe methods that can be broadly applied to a large range of organisms for morphological assessment.

SEM has found broad applications for examining inorganic materials including nanoparticles11,12, polymers13, and numerous applications in geological, industrial, and material sciences14,15,16. In biology, SEM has long been used as a method for examining biological samples ranging from individual proteins to whole organisms17,18. SEM is of particular value because morphological surface details can be used to inform scientific discovery. SEM analysis is a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of a wide range of biological samples.

Because an SEM normally operates under high vacuum (10-6 Torr minimum) to support a coherent beam of high-speed electrons, no liquids (water, oils, alcohols) are permitted in the sample chamber, as liquids prevent a vacuum from forming. Thus, all samples examined using SEM must be dehydrated, typically using a graded ethanol series followed by a drying process to remove the ethanol. There are several methods of drying biological tissues for use in the SEM, including air drying, lyophilization, use of a critical point drying (CPD) device, or chemical drying using t-butyl alcohol (TBA) or hexamethyldisilazane (HMDS)19,20,21,22. Most often, selection of a drying method is empirical since each biological sample may react differently to each drying method. For any given sample, all of these methods may be appropriate, so comparing the advantages and disadvantages of each is useful in selecting the appropriate method.

While air drying a sample at room temperature or in a drying oven (60 °C) is the simplest method, most biological samples show drying-induced damage such as shriveling and collapse, resulting in distortion of the specimen. The process of lyophilization also removes water (or ice) from a sample, but requires samples to be flash-frozen and placed under vacuum to remove the ice via the process of sublimation, potentially damaging the sample. In addition, the user must have access to a lyophilizer. The most commonly used method for dehydrating samples for SEM is critical point drying (CPD). In CPD, the ethanol in a sample is replaced with liquid carbon dioxide (CO2) and, under specific temperature and pressure conditions known as the critical point (31.1 °C and 1,073 psi), CO2 vaporizes without creating surface tension, thereby effectively maintaining the morphological and structural features of the sample. While CPD is generally the standard method, it has several drawbacks. First, the process requires access to a critical point dryer, which is not only expensive, but also necessitates the use of liquid carbon dioxide. Second, the size of the sample that can be dried is limited to the chamber size of the critical point dryer. Third, the exchange of liquids during CPD can cause turbulence that can damage the sample.

Chemical drying offers many advantages over CPD and serves as a suitable alternative that is becoming widely used in SEM sample preparation. The use of chemical dehydrants such as TBA and HMDS offers a fast, inexpensive, and simple alternative to other methods, while still maintaining the structural integrity of the sample. We recently showed that there was no difference in the integrity of the tissue or the quality of the final image captured when using CPD or TBA as the drying method in adult Drosophila retinal tissue23. Unlike CPD, TBA and HMDS do not require a drying instrument or liquid CO2 and there is no limitation on the size of the sample to be dried. In addition to obtaining the chemicals, only a standard chemical fume hood and appropriate personal protective equipment (gloves, lab coat, and safety goggles) are required to complete the drying process. While both TBA and HMDS are flammable, TBA is less toxic and less expensive (approximately 1/3 the cost of HMDS) than HMDS.

In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). These organisms represent a wide range in size (0.5 µm to 4 mm) and cellular diversity (single-celled to multicellular), yet all are easily amenable to SEM analysis with only small variations needed for specimen preparation. This protocol describes the methods for using chemical dehydration and SEM analysis to examine morphological details of three types of organisms and would be broadly applicable to examining many organismal and tissue types.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

1. Preparation and Fixation

  1. Prepare cyanobacteria.
    1. Grow unialgal cultures in F/2 media24,25 at a temperature of 28 °C on a 14:10 h light dark cycle. Transfer sufficient culture to a 1.5 mL microcentrifuge tube such that after allowing 15 min to settle, the bacterial pellet is approximately 0.05 mL in size. Remove the media and replace with 1.5 mL of fixative (1.25% glutaraldehyde, 0.1 M phosphate buffer pH 7.0), gently invert several times, and incubate overnight at 4 °C.
    2. Transfer the fixed cells with a glass pipette into a 10 mL filtration rig (Figure 1) containing a polycarbonate 25 mm filter with 0.8 or 0.2 µm pores, depending on the size of the cells. Seal the side arm and funnel with rubber stoppers to contain the culture in the funnel. Remove the fixative by gentle vacuum on the filtration flask, after removing both stoppers.
      Note: If the cell density on the polycarbonate filter is not optimal, adjust the amount of starting culture used.
  2. Prepare single cell algae (euglenoids).
    1. Grow unialgal cultures in AF-6 media26 with 150 mL L-1 of commercially-available soil-water medium added to the media, at a temperature of 22 °C on a 14:10 h light dark cycle. Transfer 2 mL of low density (OD600 < 0.5) culture with a glass pipette into a 10 mL filtration rig (Figure 1) containing a polycarbonate 25 mm filter with 8 µm pores. Seal the side arm and funnel with rubber stoppers to contain the culture in the funnel.
      Note: If the cell density on the polycarbonate filter is not optimal, adjust the amount of starting culture used.
    2. Add three drops of 4% osmium tetroxide (OsO4) directly to the culture and allow to incubate for 30 min. Remove the fixative by gentle vacuum on the filtration flask, after removing both stoppers.
      Note: OsO4 is an acute toxin (dermal, oral, and inhalation routes), corrosive to the skin, damaging to the eyes, and a respiratory sensitizer. OsO4 should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.
  3. Prepare Drosophila melanogaster (fruit fly)23.
    1. Anesthetize adult flies using 100% carbon dioxide. Place anesthetized adults (about 10 to 30 flies) in a small plastic screw cap vial or 1.5 mL centrifuge tube.
    2. Immerse anesthetized flies in 1 mL of fixative (1.25% glutaraldehyde, 0.1 M phosphate buffer pH 7.2) for 2 h (or overnight) at 4 °C. If the flies float to the surface of the fixative, add a few drops of 2.5% polyethylene glycol tert-octylphenyl ether to weaken the surface tension of the fixative allowing for total submersion of the tissue. Remove the fixative using a glass pipet.

2. Washing and Dehydration

  1. Wash and dehydrate cyanobacteria.
    1. Wash the fixed sample three times with 5 mL of 0.1 M phosphate buffer pH 7.0 at room temperature for 10 min each. Keep the flask and funnel stoppered to hold the wash. Remove each wash by gentle vacuum on the filtration flask, after removing both stoppers.
    2. Dehydrate the sample while maintaining continuous immersion using a graded ethanol series, for 10 min in a volume of 5 mL in the funnel. The graded ethanol concentrations are: 25%, 50%, 75%, 95%, and 2 changes of 100% ethanol. Keep the flask and funnel stoppered to contain the ethanol in the funnel, preventing loss both via evaporation and passive flow through the filter.
    3. Remove ethanol by gentle vacuum on the filtration flask, after removing both stoppers. Transfer the filter/sample to a disposable aluminum weighing dish containing just enough 100% ethanol to cover the sample before drying.
  2. Wash and dehydrate single cell algae (euglenoids).
    1. Wash the fixed sample three times with 5 mL of deionized water at room temperature for 10 min each. Keep the flask and funnel stoppered to contain the wash in the funnel. Remove each wash by gentle vacuum on the filtration flask, after removing both stoppers.
    2. Dehydrate the sample while maintaining continuous immersion using a graded ethanol series for 10 min in a volume of 5 mL in the funnel. The graded ethanol concentrations are: 25%, 50%, 75%, 95%, and 2 changes of 100% ethanol. Keep the flask and funnel stoppered to contain the ethanol in the funnel, preventing loss both via evaporation and passive flow through the filter.
    3. Remove the ethanol by gentle vacuum on the filtration flask, after removing both stoppers. Transfer the filter/sample to a disposable aluminum weighing dish containing just enough 100% ethanol to cover the sample before drying.
  3. Wash and dehydrate Drosophila melanogaster (fruit fly).
    1. Wash the fixed sample three times with 1 mL of 0.1 M phosphate buffer pH 7.2 at room temperature for 10 min in a 1.5 mL microcentrifuge tube. Remove each wash with a glass pipette, being careful not to remove the flies.
    2. Dehydrate the sample using a graded ethanol series, for 10 min in a volume of 1 mL in a microfuge tube. The ethanol concentrations are: 25%, 50%, 75%, 80%, 95%, 100%. Remove the ethanol with a glass pipette, being careful not to remove the flies.
    3. Retain the sample in the 1.5 mL microcentrifuge tube with just enough 100% ethanol to cover the sample before drying.

3. Drying

  1. Perform chemical drying using t-butyl alcohol (TBA)27.
    1. Replace the 100% ethanol solution with a 1:1 solution of TBA and 100% ethanol for 20 min. Replace the 1:1 solution with 100% TBA for 20 min. Repeat twice. Keep the solution at 37 °C so that TBA does not freeze.
      Note: 100% TBA has a freezing point of 25.5 °C; the 1:1 solution will not freeze at room temperature. TBA is flammable, causes serious eye irritation, is harmful if inhaled, and may cause respiratory irritation, drowsiness, or dizziness. TBA should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.
    2. Transfer the sample in TBA, if in a 1.5 mL microcentrifuge tube, into a disposable aluminum weighing dish. Once in the aluminum weighing dish, remove TBA and replace with just enough fresh 100% TBA to cover the sample. Freeze the TBA at 4 °C for 10 min.
    3. Transfer to a vacuum desiccator (bell jar) with frozen gel packs to keep TBA frozen. Evacuate and maintain vacuum with a rotary pump to allow the sample to dry by vacuum sublimation of the frozen TBA for at least 3 h or overnight.
  2. Perform chemical drying using hexamethyldisilazane (HMDS)20.
    1. Replace the 100% ethanol solution with a 1:2 solution of HMDS and 100% ethanol for 20 min. Replace the 1:2 solution with a 2:1 solution of HMDS and 100% ethanol for 20 min. Replace the 2:1 solution with 100% HMDS for 20 min. Repeat once.
      Note: HMDS is flammable and an acute toxin (dermal route). HMDS should be handled in a chemical fume hood using appropriate personal protective equipment including gloves, lab coat, and eye protection.
    2. Transfer the sample in HMDS, if in a 1.5 mL microcentrifuge tube, into a disposable aluminum weighing dish. Once in the aluminum weighing dish, replace the 100% HMDS with just enough fresh 100% HMDS to cover the sample.
    3. Transfer the sample to a plastic or glass non-vacuum desiccator with fresh desiccant (5-6 cm deep) and place into in a chemical fume hood. Alternatively, place the sample directly in a chemical fume hood to dry with a loose lid, such as a box, to prevent debris from falling on the sample. Allow the sample to dry for 12 to 24 h.

4. Mounting

  1. Mount cyanobacteria and euglenoids.
    1. Label the bottom of either a 12- or 25-mm aluminum mounting stub to indicate what is being placed on top. Cut the polycarbonate filter with the dried cells into quarters with a clean razor blade or scalpel if using a 12 mm stub, or place the entire filter if using a 25 mm stub.
    2. Place each filter on adhesive or a carbon adhesive tab secured to the top of a stub.
  2. Mount Drosophila melanogaster (fruit fly).
    1. Label the bottom of the aluminum mounting stub to indicate what is being placed on top.
      Place the dried flies in the desired position on adhesive or carbon adhesive tab secured to the top of a stub under a dissecting microscope with precision tweezers.
    2. Apply silver conductive adhesive, i.e., silver paint, around the outer edges of the stubs. Connect the silver paint to the flies using a toothpick to ensure conductivity. Do not allow the paint to touch the desired imaging area.
    3. Place the stubs in a stub holder box and place the open stub holder box in a desiccator. Allow the silver paint to dry at least 3 h or overnight for best results.

5. Sputter Coating

  1. Prepare the sputter coating apparatus by performing four to five flushes of argon gas. If the humidity is high (i.e., the room air feels moist, usually about 50% relative humidity), perform six to seven flushes. Sputter coat the stubs following the manufacturer’s instructions.
  2. Coat samples based on specimen used:
    1. For filters with cyanobacteria, set the timer to sputter coat for 180 s straight on.
    2. For filters with eukaryotic algae, i.e., euglenoids, set the timer to sputter coat for 120 s straight on, then 20 s on three sides at a 45° angle.
    3. For large samples, i.e., fly heads, set the timer to sputter coat for 60 s straight on, then 30 s on each side at a 45° angle.
      Note: After coating is complete, stubs should be light gray in color.

6. Imaging

  1. Image samples using a scanning electron microscope that includes a secondary electron (SE) detector.
    1. For cyanobacteria, apply the following settings: 3-5 kV accelerator voltage (AC), 20-30 probe current (PC), and 5 mm working distance (WD). For euglenoids, use 3 kV AC, 30 PC, and 5 mm WD. For flies, use 5 kV AC, 30 PC, and 5 to 10 mm WD.
  2. Adjust magnification depending on the size of the detail or object to be visualized. Adjust the stigmators, apertures, and focus until a clear image is produced. Capture the image using high-resolution settings according to the manufacturer’s instructions of the SEM.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

Cyanobacteria are a prokaryotic group of organisms that are critical to the global carbon, oxygen, and nitrogen cycles28,29. Of the estimated 6000 species of cyanobacteria30, most have a mucilaginous sheath that cover and connect the cells together and to other structures31, which along with the shape, can be resolved microscopically32. Cell size, shape, and pigments present distinguish individual species and in aquatic habitats, can be visualized as cyanobacterial "blooms"28. The modern cyanobacterial classification combines all morphological features possible using light microscopy, TEM, and SEM, as well as molecular data33. Similar to the mucilage sheath of cyanobacteria, pellicle strips of euglenoid species aid in phylogeny determination. Euglenoids are aquatic flagellates that possess a great deal of morphological and behavioral diversity, and SEM analysis has proven useful in classifying distinct species. Specifically, euglenoids possess novel structures beneath their plasma membrane that are composed of parallel proteinaceous strips and microtubules, called the pellicle34,35,36. The number, arrangement, and size of pellicle strips are critical morphological comparators, and together with molecular phylogenetic data, are used to construct the phylogeny for Euglenozoa37.

The first step in preparing both cyanobacteria and euglenoids requires filtering the organisms from their growth medium and is accomplished by assembling a filtration rig (Figure 1A). Aspiration is used to pull the medium through a polycarbonate filter, leaving the organisms on the surface of the filter. The pore size of the polycarbonate filter should be selected such that the intact organisms will not pass through (compare Figure 1B and 1C). Figure 2 shows representative results of three different genera of cyanobacteria. Two are freshwater and one is marine. Details of the cell, including shape and texture of the surface are visible, as well as a sheath of mucilage or projections from the cells. Figure 3 illustrates how SEM is used to visualize the pellicle strips of two different euglenoid species. The helical arrangement of the pellicle strips that merge at the posterior end to form a tail aid in phylogeny determination when comparing Monomorphina aenigmatica (Figure 3A and 3B) with Monomorphina pseudopyrum (Figure 3C and 3D).

In addition to morphological characteristics that identify species, the external morphology of model organisms such as Drosophila melanogaster can be used to identify genetic modifiers using the photoreceptors neurons that comprise the compound Drosophila eye38. Since the eye is a non-essential tissue in flies, it is possible to express toxic proteins in retinal cells and use SEM to examine the morphological changes that genetic modification has on the eye. Normal fly eyes are characterized by an ordered array of bristles and lenses (Figure 4A), however expression of proteins associated with Alzheimer's disease create what is called the 'rough eye' phenotype (Figure 4B). Genes that suppress (Figure 4C) or enhance (Figure 4D) the rough eye phenotype may play a physiological role in disease progression and have potential for drug targeting39. Also shown in Figure 4 are examples of potential pitfalls to avoid including an example of the collapsed structure of the eye from improper drying technique (Figure 4E) and electron charging from insufficient sputter coating that appears during image acquisition (Figure 4F and 4G).

Figure 1
Figure 1: Schematic representation of the filtration rig and SEM of a polycarbonate filter. (A) This apparatus is used to separate small or single celled organisms such as cyanobacteria and euglenoids away from their growth medium. Application of a vacuum to the side arm of the flask will pull the contents of the funnel through the polycarbonate filter and into the base of the filtration flask, leaving the organisms on the surface of the filter. (B) SEM of a 0.8 µm pore filter with no sample as the sample was lost because of mishandling. Scale bar = 20 µm. (C) SEM of a 0.8 µm pore filter with cyanobacteria present. Scale bar = 20 µm. Please click here to view a larger version of this figure.

Figure 2
Figure 2: SEM micrograph of cyanobacteria. (A, B) Toxifilum mysidocida, a filamentous freshwater species from Colorado with a visible sheath of mucilage (M). Scale bars = 5 µm. (C, D) A freshwater Golenkina sp. from Ohio. Individual cells sometimes form colonies held together by a thin layer of mucilage (M). Filament-like protrusions (white arrows) extend for the individual cells. Scale bars = 10 µm. (E, F) Unknown filamentous species from a high salinity estuary in Texas coastal waters. Individual cell (black arrows) and dividing cells (white arrowheads) are visible. Scale bar in panel E = 2 µm. Scale bar in panel F = 1 µm. Please click here to view a larger version of this figure.

Figure 3
Figure 3: SEM micrograph of the pellicle strips from two species of euglenoids. (A, B) Monomorphina aenigmatica. (A) Low magnification images of the entire cell. Scale bar = 5 µm. (B) High magnification of the posterior end. Scale bar = 3 µm. (C, D) Monomorphina pseudopyrum. (C) Low magnification images of the entire cell. Scale bar = 10 µm. (D) High magnification of the posterior end. Scale bar = 5 µm. Please click here to view a larger version of this figure.

Figure 4
Figure 4: SEM analysis of phenotypic modification of the Drosophila eye. Compared to the normal external appearance of the ordered array of bristles and lenses of the fly eye (A), expression of the toxic protein tau causes the death of the photoreceptor neurons, generating the 'rough eye' phenotype (B). Expression of genetic modifiers that either suppress (C) or enhance (D) the tau rough eye provides evidence of genes that may play a role in tau neurotoxicity. Improper technique during drying steps can lead to a collapse of the structure of the eye (E) while insufficient coating of the sample will cause charging, which appears as either a white (F) or black (G) band during image acquisition, as shown by the arrow. Scale bar = 400 µm. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

Here we described a protocol using SEM to obtain detailed information about external morphological characteristics of three types of organisms that others can apply to examine features of many types of organisms or tissues. Within each step of the protocol, there are potential points of error that may arise and are discussed in detail below.

While the volumes for fixative and washes given here are specific, in general the fixative and washes should be 5-10x the volume of the specimen. All fixation, washing, and dehydration times are based on our experience, if problems arise, e.g., shriveling of the sample, you may need to increase the times at each step of dehydrating or drying. Depending on your single cell sample, you may need to do both a glutaraldehyde fixation and a post-fixation in osmium tetroxide. If a duel fixation is needed after step 1.1.2 of the cyanobacteria procedure cover with an equal amount 2% OsO4 in the same buffer and proceed from step 1.2.2 of the single cell procedure. From work not shown here, we have found that Drosophila can either be directly placed into fixative following anesthetization or can be frozen indefinitely at -20 °C in a microfuge tube and placed into fixative at a later time with no difference in finished image quality.

During the washing and dehydration steps, it is imperative to move through the graded ethanol series slowly and without skipping a noted concentration. If samples are dehydrated too quickly, it negatively impacts the underlying structural components in the tissue and specimen will shrivel, wither, or collapse. An example of tissue collapse is shown in Figure 4E. In addition, to prevent the introduction of artifacts such as clumping and flattening of morphological features, it is critical to leave just enough ethanol to cover the sample between steps during dehydration and drying.

When using the filtration rig, a drop of water on the base will hold the filter in place when assembling. Also, the filter should be placed with the shiny side up and all liquids should be added gently, being careful to handle the filter with care to prevent washing away the sample. Although we have found the results of drying using either HMDS or TBA to be of equal quality, we recommend using TBA: while both HMDS and TBA are flammable, TBA is about one third the cost and less toxic than HMDS.

During mounting when using silver conductive adhesive (also called silver paint), be careful when applying as your sample can easily be covered (buried) in the paint, thereby obscuring fine details in morphology. Sputter coating times may vary based on your sample, and values given here are based on our experience. One potential negative outcome of insufficient sputter coating is a phenomenon called charging, which often appears as a white or black line (sometimes a flash) in the final image (see Figure 4F and G for examples). If the amount of charging observed is minimal, it may be possible to mitigate the effects by adjusting various microscope parameters, such as lowering the accelerating voltage or beam current, increasing the condenser lens strength, and/or using a smaller objective aperture. Most SEMs used for biological samples have secondary electron detectors, however some SEMs may also be equipped with backscatter detectors or environmental secondary electron detectors, all of which can be adjusted to diminish or eliminate the effects of minimal charging. If the charging effects are more pronounced, it is likely that there is poor grounding of the sample or too little sputter coating to produce the secondary electrons, causing the charged electrons to build up in the specimen and be spontaneously released, producing distortion in the image during acquisition. Pronounced charging is most often resolved with a thicker coating or better grounding with silver paint. Because both too much and too little sputter coating can negatively impact the quality of the image, it is best to optimize the amount of coating applied by adjusting the variables of time (10 to 180 s) and pressure (70 to 150 mTorr) of the sputter coater to achieve an optimal coating. As the coating cannot be removed once applied to the sample, a test run is recommended with a single sample before coating all of the samples.

SEM parameters such as accelerating voltage (in kV), probe current (PC), and working distance (WD) must be adjusted to get the best image possible, as the settings given here are suggested starting parameters based on our experience. While the quality of the sample is dependent upon the care taken in preparation, the quality of the final image relies upon the experience and skill of the SEM operator. Thus, we recommend working with an SEM technician or spending time developing your own SEM skills. All of the data (including sample preparation and imaging) shown in this paper were generated by undergraduate students in the Biology Microscopy program at CMU, with guidance from faculty and our microscopy technician.

The protocols described here include the use of potentially harmful chemicals, and the appropriate safety measures should always be observed to protect users, particularly students who may be handling these chemicals. While the protocols specify safety concerns associated with each chemical upon first use, users should always: (1) use a chemical fume hood, (2) have access to an emergency eye wash and safety shower, (3) use appropriate personal protective equipment including nitrile gloves, lab coat, and eye protection, and (4) know where to find written safety procedures inclusive of handling procedures, designated use areas, spill procedures, decontamination procedures, and waste disposal procedures.

In conclusion, these organisms illustrate how information about the three-dimensional topography of the external surfaces of each is useful for phylogenetic grouping or modifier analysis. For cyanobacteria, features determined from the SEM can be used in combination with additional data from light microscopy, TEM, and molecular studies to identify, characterize, and name the cyanobacteria. For euglenoids, the number, width, arrangement (helical or straight), and ornamentation, if present, of the pellicle strips can be used with other morphological data, and if necessary molecular data, to identify, characterize, and name euglenoids. Moreover, other types of single-celled algae and protozoans or micro-invertebrates can be prepared and examined in a similar manner to determine external morphological features such as flagella, feeding structure, external ornamentation, or details of appendages in the case of micro-invertebrates. Finally, SEM has broad applications for examining morphological details, such as the eye of the model system Drosophila, to characterize genetic modifiers of disease pathology. The protocols described here utilize organisms that vary greatly with respect to complexity, yet all are amenable to SEM analysis to gather useful morphological information that is critical for scientific discovery spanning many different subdisciplines of biology.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have nothing to disclose.

Acknowledgments

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

Materials

Name Company Catalog Number Comments
gold–palladium Ted Pella, Inc. 91212
Silver conductive adhesive 503 Electron Microscopy Sciences 12686-15
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 8 μm, polycarbonate Sigma WHA110614
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.8 μm, polycarbonate, black Sigma WHA110659
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.2 μm, polycarbonate Sigma WHA110606
aluminum weighing dish  Fisher Scientific 08-732-100
aluminum mounting stubs (12 mm) Electron Microscopy Sciences 75210
aluminum mounting stubs (25 mm) Electron Microscopy Sciences 75186
Adhesive tabs  Electron Microscopy Sciences 76760
conductive carbon adhesive tabs Electron Microscopy Sciences 77825
F/2 media Culture Collection of Algae at the University of Texas at Austin https://utex.org/products/f_2-medium No Catalog number given - see link 
AF6 media Bigelow - National Center for Marin Algae and Microbiota https://ncma.bigelow.org/media/wysiwyg/Algal_recipes/NCMA_algal_medium_AF6_1.pdf No Catalog number given - see link 
Soil water medium Carolina Biological Supply Company 153785
Polyethylene glycol tert-octylphenyl ether (Triton X-100) VWR 97062-208
Hummer 6.2 Sputter Coater Anatech USA http://www.anatechusa.com/hummer-sputter-systems/4 No Catalog number given - see link 
Hitachi 3400N-II SEM  Hitachi https://www.hitachi-hightech.com/us/product_list/?ld=sms2&md=sms2-1&sd=sms2-1
-2&gclid=EAIaIQobChMIpq_jtJfj3A
IVS7jACh2mdgkPEAAYASAAEgKA
nfD_BwE
The company doesn't appear to sell this model any longer 

DOWNLOAD MATERIALS LIST

References

  1. Bozzola, J. J., Russell, L. D. Electron microscopy, principles and techniques for biologists. 2nd ed, Jones and Bartlett Learning. Boston. (1999).
  2. Goldstein, J., et al. Scanning electron microscopy and X-ray microanalysis. Springer. New York. (2003).
  3. Egerton, R. F. Physical principles of electron microscopy: an introduction to TEM, SEM, and AEM. Springer. New York. (2005).
  4. Mukherjee, K., et al. Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy. Journal of visualized experiments. (113), (2016).
  5. Bello, M. A., Ruiz-Leon, Y., Sandoval-Sierra, J. V., Rezinciuc, S., Dieguez-Uribeondo, J. Scanning Electron Microscopy (SEM) Protocols for Problematic Plant, Oomycete, and Fungal Samples. Journal of visualized experiments. (120), (2017).
  6. Ramirez-Esquivel, F., Ribi, W. A., Narendra, A. Techniques for Investigating the Anatomy of the Ant Visual System. Journal of visualized experiments. (129), (2017).
  7. Endres, B., et al. A Protocol to Characterize the Morphological Changes of Clostridium difficile in Response to Antibiotic Treatment. Journal of visualized experiments. (123), (2017).
  8. Chan, V. B., et al. Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm. Journal of visualized experiments. (120), (2017).
  9. Miquel Guennoc,, C,, et al. A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy. Journal of visualized experiments. (119), (2017).
  10. Bucher, T., Kartvelishvily, E., Kolodkin-Gal, I. Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors. Journal of visualized experiments. (116), (2016).
  11. Thompson, Z. S., Rijal, N. P., Jarvis, D., Edwards, A., Bhattarai, N. Synthesis of Keratin-based Nanofiber for Biomedical Engineering. Journal of visualized experiments. (108), e53381 (2016).
  12. Ponce, A., Mejia-Rosales, S., Jose-Yacaman, M. Scanning transmission electron microscopy methods for the analysis of nanoparticles. Methods in Molecular Biology. 453-471 (2012).
  13. Wang, X. S., Tang, H. P., Li, X. D., Hua, X. Investigations on the mechanical properties of conducting polymer coating-substrate structures and their influencing factors. International Journal of Molecular Sciences. 10, (12), 5257-5284 (2009).
  14. Hortola, P. SEM examination of human erythrocytes in uncoated bloodstains on stone: use of conventional as environmental-like SEM in a soft biological tissue (and hard inorganic material). Journal of Microscopy. 218, (Pt 2), 94-103 (2005).
  15. Joy, D. C. Scanning electron microscopy for materials characterization. Current Opinion in Solid State and Materials Science. 2, (4), 465-468 (1997).
  16. Tsikouras, B., Pe-Piper, G., Piper, D. J. W., Schaffer, M. Varietal heavy mineral analysis of sediment provenance, Lower Cretaceous Scotian Basin, eastern Canada. Sedimentary Geology. 237, (3-4), 150-165 (2011).
  17. Goldberg, M. W., Fiserova, J. Immunogold Labeling for Scanning Electron Microscopy. Methods in Molecular Biology. 1474, 309-325 (2016).
  18. Hall, D. H., Hartwieg, E., Nguyen, K. C. Modern electron microscopy methods for C. elegans. Methods in Cell Biology. 107, 93-149 (2012).
  19. Heegaard, S., Jensen, O. A., Prause, J. U. Hexamethyldisilazane in preparation of retinal tissue for scanning electron microscopy. Ophthalmic Research. 18, (4), 203-208 (1986).
  20. Nation, J. L. A new method using hexamethyldisilazane for preparation of soft insect tissues for scanning electron microscopy. Stain Technology. 58, (6), 347-351 (1983).
  21. Wolff, T. Preparation of Drosophila eye specimens for scanning electron microscopy. 2011, (11), Cold Spring Harbor. 1383-1385 (2011).
  22. Fischer, E. R., Hansen, B. T., Nair, V., Hoyt, F. H., Dorward, D. W. Scanning electron microscopy. Current Protocols in Microbiology. Chapter 2 Unit 2B 2 (2012).
  23. Trotter, M. B., Stephens, T. D., McGrath, J. P., Steinhilb, M. L. The Drosophila model system to study tau action. Methods in Cell Biology. 141, 259-286 (2017).
  24. Guillard, R. R., Ryther, J. H. Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt, and Detonula confervacea (cleve) Gran. Canadian Journal of Microbiology. 8, 229-239 (1962).
  25. Guillard, R. R. L. Culture of Marine Invertebrate Animals. Plenum Press. New York. (1975).
  26. Watanabe, M. M., Kawachi, M., Hiroki, M., Kasai, F. National Institute for Environmental Studies. Tsukuba, Japan. 159 (1997).
  27. Inoue, T., Osatake, H. A new drying method of biological specimens for scanning electron microscopy: the t-butyl alcohol freeze-drying method. Archives of Histology and Cytology. 51, (1), 53-59 (1988).
  28. Zurawell, R. W., Chen, H., Burke, J. M., Prepas, E. E. Hepatotoxic cyanobacteria: a review of the biological importance of microcystins in freshwater environments. Journal of Toxicology and Environmental Health. Part B, Critical Reviews. 8, (1), 1-37 (2005).
  29. Berman-Frank, I., Lundgren, P., Falkowski, P. Nitrogen fixation and photosynthetic oxygen evolution in cyanobacteria. Research in Microbiology. 154, (3), 157-164 (2003).
  30. Silva Rocha, B., Carneiro, F. M. How many species of Cyanobacteria are there? Using a discovery curve to predict the species number. Biodiversity and Conservation. (22), 2907-2918 (2013).
  31. Robins, R. J., Hall, D. O., Shi, D. J., Turner, R. J., Rhodes, M. J. C. Mucilage acts to adhere cyanobacteria and cultured plant cells to biological and inert surfaces. FEMS Microbiology Letters. (34), 155-160 (1986).
  32. Diestra, E., et al. Characterization of an oil-degrading Microcoleus consortium by means of confocal scanning microscopy, scanning electron microscopy and transmission electron microscopy. Scanning. 27, (4), 176-180 (2005).
  33. Komárek, J. Modern taxonomic revision of planktic nostocacean cyanobacteria: a short review of genera. Hydrobiologia. 639, (639), 231-243 (2010).
  34. Leander, B. S., Farmer, M. A. Comparative morphology of the euglenid pellicle. I. Patterns of strips and pores. The Journal of Eukaryotic Microbiology. 47, (5), 469-479 (2000).
  35. Leander, B. S., Farmer, M. A. Comparative morphology of the euglenid pellicle. II. Diversity of strip substructure. The Journal of Eukaryotic Microbiology. 48, (2), 202-217 (2001).
  36. Leander, B. S., Witek, R. P., Farmer, M. A. Trends in the evolution of the euglenid pellicle. Evolution. 55, (11), 2215-2235 (2001).
  37. Leander, B. S. Euglenida. euglenids or euglenoids. Available from: http://tolweb.org/Euglenida/97461/2012.11.10 (2012).
  38. Shulman, J. M., Feany, M. B. Genetic modifiers of tauopathy in Drosophila. Genetics. 165, (3), 1233-1242 (2003).
  39. Reinecke, J. B., et al. Implicating calpain in tau-mediated toxicity in vivo. PLoS One. 6, (8), e23865 (2011).

Comments

0 Comments


    Post a Question / Comment / Request

    You must be signed in to post a comment. Please or create an account.

    Usage Statistics