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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

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.

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.

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.

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 p...

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 fixa...

Disclosures

The authors have nothing to disclose.

Acknowledgements

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

NameCompanyCatalog NumberComments
 gold–palladiumTed Pella, Inc.91212
Silver conductive adhesive 503Electron Microscopy Sciences12686-15
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 8 μm, polycarbonateSigmaWHA110614
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.8 μm, polycarbonate, blackSigmaWHA110659
Whatman Nuclepore Track-Etched Membranes; diam. 25 mm, pore size 0.2 μm, polycarbonateSigmaWHA110606
aluminum weighing dish Fisher Scientific08-732-100
aluminum mounting stubs (12 mm)Electron Microscopy Sciences75210
aluminum mounting stubs (25 mm)Electron Microscopy Sciences75186
Adhesive tabs Electron Microscopy Sciences76760
conductive carbon adhesive tabsElectron Microscopy Sciences77825
F/2 mediaCulture Collection of Algae at the University of Texas at Austinhttps://utex.org/products/f_2-mediumNo Catalog number given - see link 
AF6 mediaBigelow - National Center for Marin Algae and Microbiotahttps://ncma.bigelow.org/media/wysiwyg/Algal_recipes/NCMA_algal_medium_AF6_1.pdfNo Catalog number given - see link 
Soil water mediumCarolina Biological Supply Company153785
Polyethylene glycol tert-octylphenyl ether (Triton X-100)VWR97062-208
Hummer 6.2 Sputter CoaterAnatech USAhttp://www.anatechusa.com/hummer-sputter-systems/4No Catalog number given - see link 
Hitachi 3400N-II SEM Hitachihttps://www.hitachi-hightech.com/us/product_list/?ld=sms2&md=sms2-1&sd=sms2-1-2&gclid=EAIaIQobChMIpq_jtJfj3AIVS7jACh2mdgkPEAAYASAAEgKAnfD_BwEThe company doesn't appear to sell this model any longer 

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