1 Microscopy of biofilmsout light, epifluorescence and electron microscopy. The first section of...

Biofouling Methods, First Edition. Edited by Sergey Dobretsov, Jeremy C. Thomason and David N. Williams. © 2014 John Wiley & Sons, Ltd. Published 2014 by John Wiley & Sons, Ltd.

1 Microscopy of biofilms

Abstract

Identification, visualization and investigation of biofouling microbes are not possible with-out light, epifluorescence and electron microscopy. The first section of this chapter presents methods of quantification of microbes in biofilms and Catalyzed Reporter Deposition Fluorescent in situ hybridization (CARD-FISH). The second section provides an overview of Laser Scanning Confocal Microscopy (LSCM) imaging, which focuses mainly on the Fluorescent in situ Hybridization Technique (FISH) technique. This technique is very useful for visualization and quantification of different groups of microorganisms. The third section describes the principles of transmission (TEM) and scanning (SEM) electron microscopy.

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COPYRIG

HTED M

ATERIAL


Section 1 Traditional light and epifluorescent microscopy

Sergey Dobretsov1 and Raeid M.M. Abed2

1 Department of Marine Science and Fisheries, College of Agricultural and Marine Sciences, Sultan Qaboos University, Al Khoud, Muscat, Oman2 Biology Department, College of Science, Sultan Qaboos University, Al Khoud, Muscat, Oman

1.1 Introduction

Light microscopy is among the oldest methods used to investigate microorganisms [1, 2]. Early microscopic observations are usually associated with the name of Antony van Leeuwenhoek, who was able to magnify microorganisms 200 times using his designed microscope [1]. A modern light microscope has a magnification of about 1000× and is able to resolve objects separated by 0.275 μm. This resolving power is limited by the wave-length of the used light for the illumination of the specimens. Several light microscopy techniques, such as bright field, dark field and phase contrast, enhance contrast between microorganisms and background [1]. Fluorescent microscopy takes advantage of the abil-ity of some materials or organisms to emit visible light when irradiated with ultraviolet radiation at a specific wavelength. Phototrophic organisms have a natural fluorescence due to the presence of chlorophyll in their cells [3]. Other organisms require additional dyes in order to become fluorescent.

Light microscopy is a simple and cheap method [2]. It is commonly used for observation of relatively large (>0.5 μm) cells of microorganisms (Figure 1.1). In comparison, epifluo-rescent microscopy provides higher resolution and is generally used for observation of bacteria or cell organelles. The pros and cons of these methods are presented in Table 1.1.

Epifluorescent stains allow quick and automatic counting of bacteria using flow cytometry (discussed later in this chapter). Epifluorescent microscopy is preferable over scanning electron microscopy (SEM) (Chapter 1, section 3) for bacterial size and abundance studies [4]. While direct light microscopy measurements can be highly sensitive to low cell num-bers, electron microscopy methods are not. Light and epifluorescent microscopy has the advantage over electron microscopy that a larger surface area can be assessed for a given amount of time [5]. Two fluorescent stains are widely used to stain microbial cells, namely 4’,6-diamidino-2-phenylindole (DAPI), which binds to DNA [6] (Figure 1.2), and acrydine orange, which binds to DNA and RNA as well as to detritus particles [7]. Therefore, the estimated number of bacteria stained with DAPI is on average 70% of bacterial counts made with acrydine orange [8]. The use of DAPI stain allows a longer period between slide

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Microscopy of biofilms 5

preparation and counting, since DAPI fluorescence fades less rapidly than acrydine orange. DAPI staining does not allow accurate measurement of the size of the bacterial cells, since it could only stain the specific part of the cell containing DNA [8]. Visualization of bacteria in dense biofilms is highly difficult. This problem can be overcome to a certain extent by using confocal scanning laser microscopy (CSLM) (Chapter 1, part 2). DAPI staining has been intensively used for determination of bacterial abundance in water samples [9] as well as in biofilms [10]. This can be useful for the determination of the efficiency of biocides (Chapter 2).

Length = 100.75 µm

Figure 1.1 Microfouling community dominated by different cyanobacteria, diatoms and bacteria under a light microscope. Magnification 100×. Picture by Julie Piraino. For color detail, please see color plate section.

Table 1.1 Pros and cons of light and epifluorescent microscopy.

Method Pros Cons

Light microscopy

• Relatively inexpensive method (<$500) and does not require specialized equipment

• Simple sample preparation. In order to increase contrast, object can be stained

• Visualization of small microorganisms (>0.5 mm) is difficult

• Only large cell organelles (such as nucleus) can be visualized

• Counting of bacteria is difficultEpifluorescent microscopy

• Small microorganisms, such as bacteria, can be visualized and easily counted

• Photosynthetic organisms, such as diatoms and cyanobacteria, do not require staining

• Specialized selective probes allow staining of different cell organelles or different groups of microorganisms

• Require specialized equipment, relatively expensive (>$10 000) equipment (epifluorescent microscope with UV lamp)

• Usually requires staining with fluorescent probes

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6 Biofouling Methods

Fluorescent in situ hybridization (FISH) allows quick phylogenetic identification (phylogenic staining) of microorganisms in environmental samples without the need to cultivate them or to amplify their genes using the polymerase chain reaction (PCR) [11] (Table 1.2, Figure 1.3). This method is based on the identification of microorganisms using short (15–20 nucleotides) rRNA-complementary fluorescently labeled oligonucleo-tide probes (species, genes or group specific) that penetrate microbial cells, bind to RNA and emit visible light when illuminated with UV light [12]. Common fluorescent dyes include Cy3, Cy5 and Alexa®. In comparison with other molecular methods (Chapter 3), FISH provides quantitative data about abundance of bacterial groups without PCR bias [13]. The FISH-based protocol is presented later in this chapter (Chapter 1, section 2); here the modified protocol of catalyzed reporter deposition fluorescent in situ hybridiza-tion (CARD-FISH) is described. CARD-FISH is based on the deposition of a large number of labeled tyramine molecules by peroxidase activity (Figure 1.3), which enhances visualization of a small, slow growing or starving bacteria that have a small amount of rRNA and, thus, give a weak FISH signal [14]. Additionally, CARD-FISH can be used for the visualization and assessment of the densities of microorganisms in the samples that have high background fluorescence, such as algal surfaces, fluorescent paints, phototro-phic biofilms and sediments [14–16]. In this procedure, FISH probes are conjugated with the enzyme (horseradish peroxidase) and after hybridization the subsequent deposition of fluorescently labeled tyramides results in substantially higher signal intensities on target cells [16]. The critical step of CARD-FISH is to ensure probe microbial cell permeability with cellular integrity, especially in diverse, multispecies microbial communities [17]. Recent improvements in CARD-FISH samples preparation, permeabilization and staining techniques have resulted in a significant improvement in detection rates of benthic and planktonic marine bacteria [14, 15].

0.01 mm

Figure 1.2 Bacterial cells stained with DAPI visualized under an epifluorescent microscope. Magnification 1000 ×. For color detail, please see color plate section.

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Table

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0002124997.INDD 7 6/14/2014 9:04:09 AM


1.2 Determination of bacterial abundance

1.2.1 Material and equipment

The materials and equipment necessary for counting bacteria in biofilms using DAPI stain-ing are listed in Table 1.3.

1.2.2 Method

1. Add a few drops of DAPI solution in order to fully cover the biofilm.2. Stain for 15 minutes in the dark. Stained samples should be processed within 2–3 days in

order to avoid loss of bacterial numbers [18].3. Place a cover slip.4. Remove excess water using filter paper.5. Place immersion oil on the top of the cover slip.6. Using 100× objective count bacteria in 20 fields of view selected randomly. In the case

of digital camera coupled with an epifluorescent microscope, an automatic counting of

Epifluorescent microscopy

Environmental sample

Fixation

Hybridization with probes

Washing

DAPI staining

Fish Card-fish

Fixation andembedding

Permeabilization andinactivation of peroxidases

Tyramide signalamplification

Figure 1.3 Outline of fluorescent in situ hybridization (FISH) and catalyzed reporter deposition fluorescent in situ hybridization (CARD-FISH).

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microorganisms is possible using free image processing software ImageJ (http://rsbweb.nih.gov/ij/).

7. Calculate the number of bacteria.8. Slides can be storied frozen at –20 °C in the dark for up to one year.

1.2.3 Troubleshooting hints and tips

Special attention should be paid in the random selection of fields of view for microbial counting. This can be done using MS Excel or other software that allow a table of random X and Y stage coordinates to be generated.

A stock solution of DAPI (1 mg ml–1 in distilled water) can be prepared and stored in a dark cold (+4 °C) place for several months. This stock solution can be diluted with distilled water prior to staining in order to make working solution.

1.3 Catalyzed reporter deposition fluorescent in situ hybridization (CARD-FISH)

1.3.1 Material and equipment

The materials and equipment necessary for CARD-FISH are listed in Table 1.4.

1.3.2 Sample preparation

Microbial samples should be fixed with paraformadehyde (final concentration 2–3%) for 12 h at 4 °C. Short (1–12 h) fixation by 3% formaldehyde is possible. Fixed samples should be washed with PBS buffer for 30–60 minutes and then stored in a 2:3 PBS:ethanol mixture at –20 °C for further use without loss in signal. In the case of cell suspensions, the suspen-sion should be filtered through non-fluorescent black 0.2 μm filters prior to staining.

1.3.3 Method

Embedding

1. Mark slides or filters using pencil or permanent marker that resists alcohol.2. Dip filters in 0.1–0.2% low melting agarose and air dry them at 35 °C.

Table 1.3 Materials and equipment needed for the DAPI-based determination of bacterial abundance in biofilms.

Materials Equipment

Biofilm samples developed on glass slides and fixed with 3% formaldehyde or glutaraldehyde

Epifluorescent microscope with total magnification at least 1000×

Glass slides A blue filter set (excitation 365 nm, splitter 395 nm, barrier filter 420 nm) for DAPI stain

Cover slips An eye piece of known areaImmersion oil4,6-Diamidino-2-phenylindole (DAPI) working solution 50 µg ml–1

Blotting paper

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3. Dehydrate slides or filters in 96% ethanol for one minute at room temperature.4. Air dry slides or filters at room temperature. Samples may be stored at –20 °C for several

weeks without loss in signal.

Permeabilization and inactivation of peroxidases

1. Incubate in lysozyme at 37 °C for >60 minutes.2. Incubate in achromopeptidase at 37 °C for >30 minutes.3. Wash twice with MQ water (1 minute at room temperature).4. Incubate in 0.01 M HCl for 10 minutes at room temperature in order to bleach endoge-

nous peroxidise.5. Wash twice with MQ water (1 minute at room temperature).6. Wash with 96% ethanol (1 minute at room temperature). Air dry samples at room temperature.

The samples may be stored at –20 °C for several weeks without loss in signal.

Hybridization and washing

1. Place filters or slides in the centrifuge tubes. Use 1.5 ml and 50 ml centrifuge tubes for filters and microscopic slides, respectively. Place a blotting paper in the large centrifuge tube.

2. Prepare hybridization solution containing appropriate amount of formamide (Table 1.5). Mix 400 μl of hybridization solution and 4 μl of probe solution (concentration = 50 ng μl–1) and add to filters or slides. Cover the microscopic slide with a cover slip. Wet the blot-ting paper in the large centrifuge tube with hybridization solution. It should not be dripping wet.

Table 1.4 Materials and equipment needed for CARD-FISH.

Materials Equipment

Fixed biofilm samples on glass slides or non- fluorescent (e.g., black 0.2 µm Millipore®, GTBP02500) filters (see sample preparation)

Epifluorescent microscope with a total magnification of at least 1000×

Glass slides A blue filter set (excitation 365 nm, splitter 395 nm, barrier filter 420 nm) for DAPI stain and specific filter for the labeled probe

Cover slips An eye piece of a known areaImmersion oil Water bath4,6-Diamidino-2-phenylindole (DAPI) working solution (concentration = 50 µg ml–1)

Thermostatic incubator

50 ml or 1.5 ml centrifuge tubesHorseradish-labeled oligonucleotide probe (Table1.2)Reagents: 96% ethanol, 0.2% low-gelling agarose, lysozyme, 0.01 M HCl, achromopeptidase, MilliQ water, phosphate buffer saline (PBS) buffer (NaCl – 8.01 g, KCl – 0.2 g, Na2HPO4 2H2O – 1.78 g, KH2PO4 – 0.27 g, add 1 l of distilled water, adjust to pH = 7.6), 2% paraformaldehyde solution in distilled water, 0.0015% H2O2 solution, hybridization solution prepared according to Table 1.5, washing solution prepared according to Table 1.6.Blotting paper

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3. Incubate filters or slides in the centrifuge tubes at 35 °C for at least two hours. Longer incubation gives better results usually.

4. Prepare appropriate washing solution (Table 1.6). Wash filters or slides in a pre-warmed washing buffer (10 minutes at 37 °C) (Table 1.7). Do not air dry after washing.

Tyramide signal amplification

1. Remove excess of water using blotting paper. Do not let the filters or slides run dry. Incubate them in 1× PBS buffer for 15 minutes with mild agitation.

Table 1.5 Hybridization solutions for horseradish labeled probes. Amount of compounds is enough for one probe hybridization. These solutions are stable for 2 months at –20 °C. Pre-warm the mixtures in water bath (60 °C) until dextran sulfate dissolves.

Compound

Formamide concentration

20% 25% 30% 35%

5 M NaCl (µl) 360 360 360 3601 M Tris-HCl (µl) 40 40 40 40Dextran sulfate (g) 0.2 0.2 0.2 0.2Formamide (µl) 400 500 600 700MilliQ water (µl) 1000 900 800 70010% (w/v) Blocking reagent (Roche #1096176) dissolved in maleic acid buffer (µl)

200 200 200 200

10% Sodium dodecyl sulfate (SDS) (µl) 2 2 2 2

Table 1.6 Washing buffer solutions for horseradish labeled probes. Amount of compounds is enough to prepare 50 ml of washing solution. Pre-warm washing buffer at 37 °C in order to dissolve compounds.

Compound

Formamide concentration

20% 25% 30% 35%

1 M Tris-HCl (µl) 1000 1000 1000 10005 M NaCl (µl) 1350 950 640 4200.5 M EDTA (µl) 500 500 500 50010% SDS (µl) 50 50 50 50MilliQ water (µl) 47 100 47 500 47 810 48 030

Table 1.7 Amplification buffer solution. Can be used for any formamide concentration. p-iodophenylboronic acid (IPBA; 20 mg IPBA per 1 mg tyramide) enhances the CARD-FISH signal of tyramides labeled with Alexa488 and Alexa546 but does not work for tyramides labeled with Alexa350 and Cy3.

Compound Amount

20× PBS buffer 2 ml10% (w/v) Blocking reagent (Roche #1096176) in maleic acid buffer, pH = 7.5. Can be autoclaved and storied at –20 °C

0.4 ml

5 M NaCl solution in MilliQ water 16 mlSterile MilliQ water To a final volume 40 mlDextrane sulfate. Buffer can be heated to 60 °C in order to dissolve it. 4 g

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2. Incubate samples in a substrate mix (1 part fluorescently labeled tyramide, 10–500 parts of amplification buffer and 0.0015% H

2O

2) for 20 minutes in the dark at 46 °C.

3. Remove excess of buffer using blotting paper. Do not let the filters or slides run dry.4. Wash filters or slides in 1× PBS buffer for 5–10 minutes at room temperature.5. Wash twice in 50 ml of MilliQ water at room temperature in the dark.6. Wash in 50 ml of 96% ethanol at room temperature in the dark.7. Air dry samples. These samples can be stored at –20 °C for several weeks without loss

in signal.8. Overlay each slide or filter with 10 μl DAPI solution and incubate for five minutes in the

dark (see determination of bacterial abundance using DAPI stain).9. Mount slide or filter under a cover glass. Rapid loss of fluorescence can be prevented

by using anti-fading reagents.10. Observe sample under an epifluorescent microscope. Count bacterial cells stained with

DAPI (total count) and amount of bacteria stained with probe. It is recommended to count about 600–800 bacterial cells [15].


One of the common problems of CARD-FISH is the high background fluorescence, which might be due to (i) the use of high tyramide concentration, (ii) high probe concentration and (iii) short washing. Possible solutions may include decreasing tyramide concentrations or increasing the blocking reagent concentrations, decreasing the probe concentrations and extended washing in deionized water.

Low signal intensity might be observed and could be due to several reasons:

● Low ribosome content of target cells. In this case it is recommended to increase the tyra-mide concentration or the temperature during the tyramide signal amplification. A pro-longed hybridization time (>4 hours) may also help.

● Too low tyramide concentration. Check the concentration and increase it 1.5–2 times. ● The probes has too low or no activity. In this case, check the probe. Make sure that the probe is thawed only once and is not be stored in the fridge for more than six months. Check the pH of the PBS buffer (should be around 7.6) and H

2O

2 concentration and its age. If neces-

sary, prepare new PBS buffer and H2O

2 solution. Check the reactivity of the tyramide.

● The horseradish peroxidase is not coupled with the probe. In this case, use new horserad-ish peroxidase probe.

● The horseradish peroxidase probe cannot penetrate the cell wall. In this case, try different permeabilization protocols.

1.4 Suggestions, with examples, for data analysis and presentation

The number of bacteria per an eye piece of a known area obtained using DAPI counting can be transformed to a number of bacteria per mm2 [10]. In the case of normally distributed data, densities of bacteria can be compared using a t-test (for the comparison of 2 means) or ANOVA followed by a suitable post hoc (for multiple comparisons). Usually, normality of the data can be improved by taking the natural log.

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Usually, FISH or CARD-FISH data are expressed as percentages of total bacterial num-bers obtained using particular oligonucleotide probe versus total DAPI counts for the same sample. This can be calculated using the following formula:

% DAPI count = n/N × 100

where n = number of bacteria stained with the probe and N = number of bacteria stained with DAPI.

Acknowledgements

The work of SD was supported by a Sultan Qaboos University (SQU) internal grant IG/AGR/FISH/12/01 and by a HM Sultan Qaboos Research Trust Fund SR/AGR/FISH/10/01.

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1.5 Introduction

Laser scanning confocal microscopy (LSCM) imaging offers many advantages over conven-tional light and fluorescence microscopy, including the elimination of out-of-focus signal and the capability to collect images from serial sections of thick specimens. First invented and developed by Marvin Minsky in the 1950s, the confocal microscope was not widely used by researchers until lasers became used in conjunction with confocal microscopes in the 1980s. The result was LSCM, which, in contrast to conventional fluorescence microscopy, illumi-nates and scans specimens with a beam of light from a laser source. This excitation results in the system’s ability to focus exclusively on planes within thick, opaque objects and eliminate signal from out-of-focus planes, producing “optical sections” of a specimen. Optical sections of a sample can, therefore, be obtained without the disturbance from physical sectioning, making confocal microscopy a popular tool for in situ imaging of microbial biofilms and other samples thicker than a few micrometers. Confocal image processing software offers the ability to acquire a series of optical sections that have matching register (to “build a z-stack”) and render those into three-dimensional images. The laser source also provides the user with the ability to expose the sample to a specific or narrow range of excitation wavelengths, resulting in a reduction of autofluorescence and an increase in specific detection of the target. Most confocal image acquisition software has the ability to link multiparameter quantitative data sets of fluorescence intensity to each image, which can translate into measurements including, but not limited to, cell counts, cell size, cell species identification, biofilm thick-ness, and quantitative gene expression within biofilmed surfaces.

The development of LSCM has its origins in the biomedical sciences, where it was used to image cells in vivo, but with the advancement of molecular techniques for the study of microbial communities in the 1980s and 1990s, it quickly became a tool widely used by microbial ecologists. In the past 20+ years, researchers have used confocal microscopy to study in situ physical structure, polysaccharide excretion, and metabolite production in natural and cultured biofilms [1–10] [reviewed in 11, 12]. LSCM has been used exten-sively with fluorescence in situ hybridization (FISH) techniques to assess the taxonomic

Section 2 Confocal laser scanning microscopy

Koty SharpDepartments of Marine Science and Biology, Eckerd College, St. Petersburg, FL, USA

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composition Sequence of environmental biofilms. In situ hybridization was first designed as a technique to target 16S rRNA molecules in fixed bacterial cells and visualize and identify individual bacterial cells among natural samples using radiography and fluorescence-based detection methods [13–15]. The small subunit (16S) rRNA molecule has been well-established as a phylogenetic marker for bacterial phylogeny, and oligodeoxynucleotide probes targeting 16S rRNA molecules in bacterial ribosomes have become so widely used as primers and probes by microbial ecologists (Chapter 1.1) that oligonucleotides with custom sequences are commercially available at extremely low costs, and these oligonu-cleotides can be ordered with fluorophore labels either conjugated to one of the ends of the probe fragment or with a label incorporated throughout the probe.

The general bacterial oligonucleotide EUB338 was developed first as a probe that would hit a majority of known bacteria [14, 15,]. As molecular techniques allowed identification of more and more previously uncharacterized bacterial groups, new probes (EUB338II and EUB338III) were developed to be used in combination with the original probe, now named EUB338I, to ensure coverage of diverse bacterial taxa, including the strains from the orders Planctomycetales and Verrucomicrobiales [18]. Over the past decade, important advances have been made on the design and development of sequence-specific 16S oligonucleotide probes, targeting specific bacterial taxa or groups. Sequence-specific probes have been designed targeting hypervariable regions of 16S rRNA from most of the known bacterial taxonomic groups [16, 17]. Another important development in the science of oligonucleotide probe design was the determination of steric accessibility of regions across the 16S rRNA molecule. Studies show that variation in primary sequence across different bacterial taxonomic groups results in differential secondary structure and accessibility of particular regions of the 16S molecule. Patterns of accessibility of particular regions of the 16S molecule have been well characterized and appear to vary across phylogenetic affiliation [16, 17]. Methods have been developed to increase probe binding to less accessible regions of the 16S molecule [17], to amplify the signal from end-labeled oligonu-cleotide probes [19], and to construct probes with higher intensity signal [20]. In situ hybridi-zation with end-labeled oligonucleotide probes (FISH) and amplification of signal from those end-labeled probes (CARD-FISH) are detailed in this chapter (Figure 1.3), but any type of hybridization can be imaged on a confocal microscope with the following imaging protocol.

The data that result from FISH approaches are no longer limited to taxonomic identifica-tion and localization of bacteria. In order to determine which genes are being expressed by bacteria or what proteins and other bioactive molecules are being synthesized, LSCM is now used with variations on FISH, combined with microanalytical and chemical methods, such as immunohistochemistry and Raman spectroscopy, to detect gene expression, protein synthesis, and certain metabolites of interest to single bacterial cells [32–34, 53–58].

Fully equipped LSCM setups across a wide range of costs are available from a variety of major microscopy companies. A typical LSCM package includes a microscope, laser line(s), a laser scan head attached to the microscope via fiber optic cables, and a computer with enough processing speed and memory specifications for image acquisition and processing. A schematic diagram of a typical LSCM setup is shown in Figure 1.4. The epifluorescence microscope is often outfitted with filter sets customized to the user’s needs and a charge-coupled device (CCD) camera to allow the researcher to image single plane images from the epifluorescence scope before switching to confocality. Attached to the epifluorescence scope is a scan head, which contains a series of mirrors and beam splitters that focus the laser beam emission onto the sample and then collect and detect specific wavelengths of light via photomultiplier tubes (PMTs). The light is detected through a series of pinholes, apertures of adjustable diameter that are confocal with each other, which allow exclusive detection of

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specific two-dimensional planes on the Z-axis without noise from other out-of-focus planes. Detection of emitted light, or signal, from the sample is collected point-by-point or line-by-line on a two-dimensional plot. User-defined scan speed results in variation in resolution (typically ranging from 512 × 512 to 1028 × 1028 pixels). Though slow scan speed yields a high image resolution, there is a trade-off in target photobleaching that must be considered by the user and, as a result, optimal image acquisition speed is often determined by the user based on empirical testing of image resolution and target fluorophore bleaching rates.

Several laser lines can be employed in conventional LSCM systems. Multiple lasers, including krypton–argon and helium–neon, can be attached to the epifluorescence scope via a single laser scan head. The lasers, in addition to specialized filter sets, can be controlled by acquisi-tion software for detection of specific wavelength ranges to target fluorophores of interest. This combination offers confocal systems the ability to excite a specimen with light from a narrow range of wavelengths and detect specific wavelengths of emission from the specimen. High performance confocal systems, such as the Zeiss LSM 710 system, have spectral imag-ing, or the capability to detect emission at a 5 nm range and produce a spectral profile of each pixel of an image, yielding increased target signal acquisition specificity and sensitivity.

There is now a wide range of fluorescent molecular probes and labels that have been developed for biological imaging. Among the most commonly used for confocal microscopy applications are fluorophores for labeling oligonucleotide probes. The fluorophores can be

Photomultiplier (PMT)

Pinhole

Beam splitter

Objective lens

xy

z

Z Control

Scanner

Laser

Figure 1.4 General schematic diagram of a typical LSCM setup. Sample (green) is exposed to the laser. source. (Image used with permission of Carl Zeiss MicroImaging).

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detected in the near-UV, visible, and near-infrared wavelengths, including blue (DAPI, Alexa Fluor® 405), green (FITC and fluoroscein derivatives; Alexa Fluor® 488), red (Cy 3; TRITC, rhodamine, and rhodamine derivatives; Alexa Fluor® 568) and far-red (Cy 5 and Alexa Fluor® 635) wavelengths. Alexa Fluor® dyes (Invitrogen Life Technologies), have a higher cost than the Cy or the FITC dyes, but they are also more robust to photobleaching, which is critical when the user is trying to image a faint signal that requires prolonged exci-tation by the laser. Imaging with two or three wavelength ranges (i.e., DAPI, FITC, and Cy 3) is sufficient for the desired goal in many applications. One of the most promising recent developments in FISH imaging of microbial ecology is the use of increased numbers of probes. Combinatorial labeling and spectral imaging FISH (CLASI-FISH) is a recently developed protocol [29] in which the authors used spectral imaging and performed FISH with factorial combinations of fluorophores to detect an unprecedented high number of fluo-rescent probes in a single specimen. In Figure 1.5, images from this study show that the spectral imaging capacity of the Zeiss LSM 710 system and construction of combinatorial probes allowed simultaneous imaging and localization of 15 distinct probes, identifying bacterial taxa important to the initial developmental stages of dental biofilms [29].

LSCM is an effective, nondestructive tool for quantifying, characterizing diversity of, visual-izing the structural characteristics, and determining the activity of organisms in microbial bio-films. LSCM systems are becoming increasingly common in multiuser equipment facilities in research institutions; the methods and sample preparation described here are a summary of basic methods used in for determining the phylogenetic makeup of cells in fixed or live biofilms.

1.6 Materials, equipment, and method

1.6.1 Materials and stock solutions

● Fixed biofilm sample on glass slide or other substrate (see Notes 1, 2) ● Cover slips

Raw spectral image merge Taxon-assigned segmented image

Figure 1.5 Confocal images of CLASI-FISH-labeled human oral biofilm. Color in spectral images (left) represents the merge of six different fluorophore channels. Color in the segmented image (right) represents resulting false coloration of cells from each of the 15 taxa. Scale bar: 10 μm. Source: From Valm et al. [29] and reproduced with the permission of Proceedings of the National Academy of Sciences. For color detail, please see color plate section.

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● Immersion oil ● 50 ml tubes ● Stock solutions (Table 1.8) ● Oligonucleotide probe (Chapter 1.1, Table 1.2).

1.6.2 Equipment

● Microscope ● Laser line(s) ● Laser scan head attached to the microscope via fiber optic cables ● Computer.

1.6.3 Methods

Localizing specific phylogenetic groups of bacteria in a sample (FISH)

1. Submerge sample in fixative of choice (see Notes 1, 2, 6–8).2. Remove fixative from sample.3. Perform a dehydration series (50, 80, 96% ethanol) on sample, three minutes per ethanol

concentration.4. Remove ethanol and air dry the biofilm sample.5. The sample is ready for FISH or microscopy or can be stored at room temperature for

months [30]. Cells can be imaged and enumerated in fixed or live cells via nucleic acid stains 4′,6-diamidino-2-phenylindole (DAPI) and/or acridine orange (see Chapter 1.1 for method). DAPI, which emits fluorescence in the blue wavelengths, is an ideal counterstain to fluorophores commonly used for FISH, such as those in the yellow, green, orange, red, and far red wavelengths.

6. Prepare 2 ml of hybridization buffer (Table 1.9), containing appropriate concentration of formamide per specimen (see Note 9).

7. Add probe(s) of interest to a final concentration of 5 ng/ul each probe (see Note 10).8. Overlay specimen with 10–200 ul probe/hybridization buffer.9. Place clean tissue or filter paper along the side of a 50 ml tube and wet the paper with

remaining hybridization buffer (this is a hybridization chamber).

Table 1.8 Solutions for fixing and fluorescence in situ hybridization of biofilm samples for LSCM (see Notes 3–5).

Reagent/Solution

Fixation 2.5 % glutaraldehyde*Note4 % paraformaldehyde*Note

Hybridization 5 M NaCland Washing 1 M Tris-HCl, pH 7.4

0.5 M EDTAMilliQ water10% Sodium dodecyl sulfate (SDS)Molecular grade formamide*Note

Mounting VectaShield or Citifluor*Note

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10. Place the slide, keeping it flat, into the 50 ml tube, cap it tightly, and incubate it, lying sideways, at 46 °C in dark for at least two hours.

11. Remove hybridization solution from specimen by tipping specimen, decanting solu-tion, or rinsing in wash buffer (Table 1.10).

12. Incubate specimen in wash buffer at 48 °C for a time according to hybridization incuba-tion time (see Notes 11, 12).

13. Remove wash buffer.14. Air dry the specimen at room temperature.15. Mount in VectaShield or Citifluor for imaging.

Table 1.9 Hybridization buffer and wash buffer recipes for hybridization buffer containing 35% formamide.

Hybridization Buffer

Stock Volume Final Concentration

5 M NaCl 360 µl 900 mM1 M Tris-HCl pH 7.4 40 µl 20 mMFormamide (molecular grade) 700 µl 35%dH2O 1.9 ml —10% SDS 2 µl 0.01%

Wash Buffer

Stock Volume Final Concentration

5 M NaCl 800 µl 80 mM1 M Tris-HCl pH 7.4 1 ml 20 mM500 mM EDTA 500 µl 5 mMdH2O add to 50 ml —10% SDS 50 µl 0.01% SDS

Table 1.10 Concentrations of NaCl in washing buffer (48°C) at different concentrations of formamide in hybridization buffer (46°C).

% Formamide in hybridization buffer mM NaCl in washing buffer

0 9005 636

10 45015 31820 22525 15930 11235 8040 5645 4050 2855 2060 1465 1070 775 580 3.5

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1.7 Image acquisition

There is a general sequence of steps for obtaining a confocal image of a specimen. Each brand’s software program will differ slightly but the following broad guidelines should guide a beginning user through any LSCM acquisition and analysis.

1. Turn on the mercury (HBO) lamp.2. Turn on the computer and run the LSCM software.3. View the specimen on the microscope (without the confocal scan head on).4. Using the microscope in epifluorescence mode, view specimen under the filter cube of

choice and adjust the specimen/magnification so that the region of interest is centered in the field of view.

5. Select a configuration of lasers, filters, and mirrors, according to which fluorophore is being detected.

6. Select the option on the LSCM software to scan the specimen with the laser scan head, and scan the image using a short-duration scan to “find” the specimen or optimize speci-men orientation.

7. Once a low-resolution image of the sample is visible, adjust the software settings to take a longer scan with increased resolution.

The above steps detail a sequence for taking a single-plane image; for generation of z-stacks, series of images in the x-y plane, see your software user manual. In general, taking a z-stack includes defining the upper and lower limit of the z-position within the specimen, choosing an interval or “thickness” of each section slice, optimizing the resolution and image acquisition settings, and running the acquisition software so that it collects an image across each section within the “stack.”

1.8 Presentation

For publication-quality images, it is best to use the LSCM software for acquisition of 12-bit images with a pixel depth of 1024 × 1024 pixels. However, while exploring the sam-ple or scanning for an appropriate portion specimen, it is best practice to minimize scan speed and use low-resolution imaging until generation of an image for presentation. In software from most of the leading brands of microscope software, the resulting image file is a proprietary file format. The files, which average around 0.5 MB to 5 MB in size depending on resolution, are organized via a small (<1 MB) database file. There is an option to export the files into compatible file formats from image acquisition software, including jpg or tiff, which can then be annotated and saved in a separate location from the raw image files.

1.9 Troubleshooting hints and tips

When beginning confocal imaging efforts on samples, it is useful to image unhybridized samples in the different detection channels available on the system. This is important for two reasons: (i) it will help the researcher determine in which wavelengths the specimens emit the least autofluorescence, so that the detection channels and, subsequently, fluorophores that will produce the highest signal:noise ratio can be selected; (ii) inherent autofluorescence

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(such as photosynthetic pigments, bioactive compounds) has often been used to characterize general structural arrangement or other parameters of the specimen. Note that fixation of a sample will alter the fluorescent characteristics of a specimen. It is, therefore, necessary to assess the baseline fluorescence of a specimen after fixation if the hybridization or staining is being done on fixed samples. Once the baseline fluorescence of a sample is assessed, the user can select the fluorophore label or stain to be used for the application.

If samples are on slides or an apparatus that allows movement of the slide, submerge the slides in a 50 ml plastic tube full of wash buffer, with a maximum of two slides per tube, back to back so the samples do not touch.

One of the most significant challenges in imaging natural biofilms is the shape of the surface. Use of LSCM to image biofilms cultivated on flat surfaces, such as Robbins devices or other flow-through reactors, is more straightforward than microscopy on wild surfaces, such as pieces of rocks, algae, wood, or plastic. Irregular substrates can be placed in sterile seawater in small petri dishes or depression slides with a water immersion lens on an upright microscope. Substrate opacity also is most easily overcome by imaging on an upright system.

Whenever possible, perform hybridizations in a hybridization oven with accurate digitally measured temperature control. A single degree difference in temperature may affect probe hybridization stringency.

Formamide concentration in the hybridization buffer destabilizes DNA–DNA hybrids, therefore controlling stringency of probe-target binding [31]. When formamide concentration is increased in the hybridization buffer, the level of monovalent cations (NaCl in this case) is decreased in order to maintain the low stability of DNA–DNA binding. When testing new probes and new hybridization buffers, the NaCl concentration in the wash buffer should be adjusted to accommodate the altered formamide concentration. Table 1.10 shows the concen-tration adjustments at hybridization temperature of 46 °C and wash temperature of 48 °C.

Unprobed specimens often exhibit autofluorescence from photosynthetic pigments and polysaccharide fluorescence properties, and this noise is a significant problem that often requires additional protocol adjustment and testing. Techniques developed to overcome high sample autofluorescence, such as CARD-FISH for signal enhancement [19] and DOPE-FISH [20], which constructs brighter probes with more signal molecules, are two methods that help to increase the ratio of probe-conferred fluorescence to autofluorescence in probed specimens. Some researchers find it useful to photobleach the pigments in samples with the laser line or UV lamps before hybridization, but those protocols are at the expense of run-ning the laser source and can potentially do damage to the cellular structure in specimens.

Optimal stringency conditions promoting probe specificity are adjusted most easily by changing formamide concentration in the hybridization buffer. Hybridization conditions for optimal stringency of previously tested probes are available in probeBase (http://www.microbial-ecology.net/probebase/), a searchable catalog-like database of published rRNA-targeted oligonucleotide probes for FISH and microarray technology [21] maintained by the University of Vienna Department of Microbial Ecology. A list of some of the most com-monly used probe sequences are listed in Table 1.2. probeCheck is another useful online tool that links to ribosomal 16S rRNA sequence databases, such as SILVA, RDP-II, and Greengenes, for checking probe or primer coverage and specificity [22]. In testing a novel FISH probe, it is best to test the stringency across a range of formamide concentrations empirically.

When scanning images, it is best practice taking images of the specimen at low magnifi-cation and then move to higher magnification. Depending on the fluorophore used and

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the laser intensity setting, the scan may leave a faded square where the scan head bleached the specimen. This is unfortunately impossible to reverse once it is done, and it can scar an otherwise informative image.

1.10 Notes

Note 1: Though glutaraldehyde strongly cross-links nucleic acids and preserves cells extremely well, it exhibits more autofluorescence than paraformaldehyde. As a result, paraformaldehyde is often chosen for fluorescence and confocal imaging.

Note 2: Fixation can be done in a well-buffered, 2–5% paraformaldehyde solution that has similar osmolarity to the specimen itself. In biomedical applications, phosphate-buffered saline and 3-(N-morpholino)propanesulfonic acid (MOPS) paraformaldehyde solutions are common. In marine research, especially in field research, liquid paraform-aldehyde stocks are particularly useful, because hazardous paraformaldehyde in its powder form can be avoided. These stocks can be diluted directly in sterile filtered (0.22 μm) seawater.

Note 3: It is helpful to sterile filter all solutions that go into the hybridization buffer to exclude any autofluorescent particles from the hybridization.

Note 4: These reagents can be stored at room temperature.Note 5: In addition to the user-made solutions, commercially available molecular grade

deionized formamide, VectaShield (Vector Labs, Burlingame, CA) and Citifluor (Citifluor, Ltd, London, UK), are used in this protocol. These reagents should be stored at 4 °C.

Note 6: Prolonged exposure to fixatives should be avoided because it decreases cell perme-ability to probes, and for bacterial biofilms and samples should not be fixed for more than about four hours.

Note 7: Biofilms should be fixed in a small volume; often this can be done by submerging the biofilmed surface in a small petri dish or vessel with an area enclosed by inert aquar-ium silicone adhesive.

Note 8: In order to study the fully hydrated, intact biofilm for in situ visualization, skip the fixation step altogether and add stains or perform hybridizations on unfixed samples if possible.

Note 9: Hybridization solution volume will vary based on the sample/vessel used, but a volume of 10–200 μl is typical. For example, if the specimen is a biofilm on a glass microscopy slide, use enough hybridization solution to cover the specimen to ensure that the entire specimen is exposed to the solution and that none of the specimen dries during hybridization.

Note 10: for use of multiple probes (such as EUB338I, EUB338II, and EUB338III), add probes in equimolar concentration at 5 ng/μl final concentration each.

Note 11: 20 min wash after a two hour hybridization, 30 min after a three hour hybridization, an so on.

References

1. Parsek, M.R. and Greenberg, E.P. 1999. Quorum sensing signals in development of Pseudomonas aeruginosa biofilms. Biofilms, 310: 43–55.

2. Schmidt, M., Cavaco, A., Gierlinger, N., et al. 2009. In situ imaging of barnacle (Balanus amphitrite) cyprid cement using confocal Raman microscopy. Journal of Adhesion, 85(2–3): 139–151.

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3. Bjorkoy, A. and Fiksdal, L. 2009. Characterization of biofouling on hollow fiber membranes using confocal laser scanning microcscopy and image analysis. Desalination, 245(1–3): 474–484.

4. Bester, E., Kroukamp, O., Wolfaardt, G.M., et al. 2010. Metabolic differentiation in biofilms as indicated by carbon dioxide production rates. Applied and Environmental Microbiology, 76(4): 1189–1197.

5. Chen, M.Y., Lee, D.J. and Tay, J.H. 2006. Extracellular polymeric substances in fouling layer. Separation Science and Technology, 41(7): 1467–1474.

6. Chen, M.-Y., Lee, D.-J., Yang, Z., et al. 2006. Fluorecent staining for study of extracellular polymeric substances in membrane biofouling layers. Environmental Science and Technology, 40(21): 6642–6646.

7. Norton, T.A., Thompson, R.C., Pope, J., et al. 1998. Using confocal laser scanning microscopy, scanning electron microscopy and phase contrast light microscopy to examine marine biofilms. Aquatic Microbial Ecology, 16(2): 199–204.

8. White, D.C., Arrage, A.A., Nivens, D.E., et al. 1996. Biofilm ecology: On-line methods bring new insights into MIC and microbial biofouling. Biofouling, 10(1–3): 3–16.

9. Decho, A.W. and Kawaguchi, T. 1999. Confocal imaging of in situ natural microbial communities and their extracellular polymeric secretions using Nanoplast (R) resin. Biotechniques, 27(6): 1246–1252.

10. Pope, R., Little, B., and Ray, R. 2000. Microscopies, spectroscopies and spectrometries applied to marine corrosion of copper. Biofouling, 16(2–4): 83.

11. Lawrence, J.R. and Neu, T.R. 1999. Confocal laser scanning microscopy for analysis of microbial biofilms. Biofilms, 310: 131–144.

12. Neu, T.R., Manz, B., Volke, F., et al. 2010. Advanced imaging techniques for assessment of structure, composition and function in biofilm systems. FEMS Microbiology Ecology, 72(1): 1–21.

13. Delong, E.F., Wickham, G.S., and Pace, N.R. 1989. Phylogenetic stains – ribosomal RNA-based probes for the identification of single cells. Science, 243(4896): 1360–1363.

14. Giovannoni, S.J., DeLong, E.F., Olsen, G.J., and Pace, N.R. 1988. Phylogenetic group-specific oligodeoxynucleotide probes for identification of single microbial cells. Journal of Bacteriology, 170(2): 720–726.

15. Amann, R.I., Krumholz, L., and Stahl, D.A. 1990. Fluorescent oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. Journal of Bacteriology, 172(2): 762–770.

16. Behrens, S., Rühland, C., Inácio, J., et al. 2003. In situ accessibility domains of small-subunit rRNA of members of the domains Bacteria, Archaea, and Eucarya to Cy3-labeled oligonucleotide probes. Applied and Environmental Microbiology, 69(3): 1748–1753.

17. Fuchs, B.M., Wallner, G., Beisker, W., et al. 1998. Flow cytometric analysis of the in situ accessibility of Escherichia 16S rRNA for fluorescently labeled oligonucleotide probes. Applied and Environmental Microbiology, 64: 4973–4982.

18. Daims, H., Brühl, A., Amann, R., et al. 1999. The domain-specific probe EUB338 is insufficient for the detection of all Bacteria: development and evaluation of a more comprehensive probe set. Systematic and Applied Microbiology, 22(3): 434–444.

19. Pernthaler, A., Pernthaler, J., and Amann, R., 2002. Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Applied and Environmental Microbiology, 68(6): 3094–3101.

20. Stoecker, K., Dorninger, C., Daims, H., and Wagner, M. 2010. Double labeling of oligonucleotide probes for fluorescence in situ hybridization (DOPE-FISH) improves signal intensity and increases rRNA accessibility. Applied and Environmental Microbiology, 76(3): 922–926.

21. Loy, A., Maixner, F., Wagner, M., and Horn, M. 2007. probeBase – an online resource for rRNA-targeted oligonucleotide probes: new features 2007. Nucleic Acids Research, 35: D800–D804.

22. Loy, A., Arnold, R., Tischler, P., et al. 2008. probeCheck – a central resource for evaluating oligonucleotide probe coverage and specificity. Environmental Microbiology, 10(10): 2894–2898.

23. Moraru, C., Lam, P., Fuchs, B.M., et al. 2010. GeneFISH - an in situ technique for linking gene presence and cell identity in environmental microorganisms. Environmental Microbiology, 12(11): 3057–3073.

24. Huang, W.E., Stoecker, K., Griffiths, R., et al. 2007. Raman-FISH: combining stable-isotope Raman spectroscopy and fluorescence in situ hybridization for the single cell analysis of identity and function. Environmental Microbiology, 9(8): 1878–1889.

25. Lechene, C.P., Luyten, Y., McMahon, G., and Distel, D.L., 2007. Quantitative imaging of nitrogen fixation by individual bacteria within animal cells. Science, 317(5844): 1563–1566.

26. Neu, T.R. and Lawrence, J.R. 1997. Development and structure of microbial biofilms in river water studied by confocal laser scanning microscopy. FEMS Microbiology Ecology, 24(1): 11–25.

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27. Neu, T.R., Swerhone, G.D.W., Böckelmann, U., and Lawrence, J.R. 2005. Effect of CNP on composition and structure of lotic biofilms as detected with lectin-specific glycoconjugates. Aquatic Microbial Ecology, 38(3): 283–294.

28. Neu, T.R., Woelfl, S., and Lawrence, J.R. 2004. Three-dimensional differentiation of photo-autotrophic biofilm constituents by multi-channel laser scanning microscopy (single-photon and two-photon excitation). Journal of Microbiological Methods, 56(2): 161–172.

29. Valm, A.M., Mark Welch, J.L., Riekena, C.W., et al. 2011. Systems-level analysis of microbial community organization through combinatorial labeling and spectral imaging. Proceedings of the National Academy of Sciences of the United States of America, 108(10): 4152–4157.

30. Manz, W. 1999. In situ analysis of microbial biofilms by rRNA-targeted oligonucleotide probing. Biofilms, 310: p. 79–91.

31. Schwarzacher, T. and Heslop-Harrison, J. 2000. Practical In Situ Hybridization. , BIOS, Oxford, UK.

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Section 3 Electron microscopy

Omar Skalli, Lou G. Boykins, and Lewis CoonsIntegrated Microscopy Center and Department of Biological Sciences, The University of Memphis, Memphis, TN, USA

1.11 Introduction

Shortly after the invention of the transmission and scanning electron microscopes, in the 1930s, biologists realized the potential that these instruments had to revolutionize their field by enabling scrutinizing biological structures beyond the 200 nanometers resolution of compound light microscopes [1–4]. To explore this new resolution landscape, ingenious methods and tools were rapidly devised to fix, stain and process cells and tissues for obser-vation with high-energy electron beams [1–4]. The investigations carried out by electron microscopy pioneers revealed novel organelles and provided morphological and functional information about biomolecules such as nucleic acids and proteins [5, 6]. These studies also revealed that bacteria, which at the time were thought of as “bags of enzymes”, had a cellular architecture and internal organelles [7]. Ernst Ruska, one of the electron microscope’s fathers, also used his invention to investigate viruses, which had so far eluded morphological characterization as their size was smaller than the light microscopes resolu-tion limit [8, 9]. By the 1970s, electron microscopy had produced a “bountiful harvest of new structural information that had culminated in the unified new field of science called cell biology” [10] and electron microscopes had become a staple of most research- oriented biology departments.

It is thus perhaps surprising that, to the best of our knowledge, one had to wait for the late 1960s for investigators to add electron microscopy to the arsenal of light microscopy and microbiological approaches used to study biofilms. Jones and colleagues, in 1969, developed transmission electron microscopy (TEM) methods which, they predicted “will prove useful in studying the ecology of naturally occurring microbial films in aquatic systems”, and employed these methods to characterize slime layers growing in polluted streams [11]. Their observations provided a structural characterization of the extracel-lular slime matrix and established its relationship with the microorganisms in the slime. A few years later, the structure and nitrification capacity of bacteria in the slime of aqua-culture systems was further examined by TEM [12]. From that time on, TEM became often used to obtain information on the types of microorganisms present in biofilms and

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on the nature, abundance, and organization of the extracellular matrix [13–15]. Scanning electron microscopy (SEM) was introduced in biofilm investigations in the 1980s and first served to characterize the growth of sessile Sphaerotilus natans in a continuous flow recycle system [16] and bacterial adherence to catheter lumen [17–19]. Since then, SEM has remained a widely used method for investigating biofilms of medical and environ-mental interest [15, 20–23].

TEM generates two-dimensional structural information with a resolution 10–100 times better than that of light microscopes [24]. Sample preparation for TEM involves sectioning samples into thin (~40–70 nm) sections and staining them with uranyl acetate and lead citrate. When sections are illuminated by the electron beam of a TEM, an image is formed by the electrons passing through the sample unabsorbed by the stain. With SEM, the electron beam scans the surface of a three-dimensional sample and an image of the surface is created by collecting the secondary and/or backscattered electrons bounc-ing off the sample surface. SEMs enable magnification ranging from 20× to 30 000× and spatial resolution up to 50 nm [24–26]. SEMs can also be equipped with an attachment to perform energy dispersive X-ray spectroscopy (EDS or EDX). EDS is an important tool for biofilm analysis because it enables mapping the elemental composition of spe-cific regions on a sample [24–26].

Nowadays, morphological assessment of biofilms frequently combines TEM and/or SEM with confocal laser scanning microscopy (Chapter 1, part 2). In the following, the principal methods for biofilm preparation for TEM or SEM observations are described. Instructions on how to operate TEM and SEM are beyond the scope of this chapter and are best left to the manufacturers of these instruments.

1.12 Transmission electron microscopy (TEM)

1.12.1 Purpose of TEM

TEM affords morphological observations at a resolution of up to 0.2 nm, but a resolution of 2–20 nm is usually sufficient for most biofilm studies [24]. TEM allows identification of the types of microorganisms present in biofilms because its high resolution enables the visualization of organelles specific to various types of microorganisms. For example, nuclei and mitochondria are present in algae and protozoa, but not in bacteria. Algae can be further distinguished from protozoa by chloroplasts. It may also be possible to dif-ferentiate between different types of bacteria based on shape and/or features of the cell wall and the presence of specific organelles such as pili, flagella and cytoplasmic inclusions.

TEM observations of a fish tank biofilm revealed different types of prokaryotes, some of which appeared as individual cells while others formed clusters. The cytoplasm of these prokariotes contained highly electron-dense inclusions similar to those found in nitrifying bacteria (Figure 1.6). TEM studies also demonstrated the complex organismal complexity of waste water biofilms [27] and of intracellular infection in otitis media biofilms [28]. However, in recent years, molecular methods such as fluorescence in situ hybridization (FISH) and polymerase chain reaction (PCR) have proven more effective than TEM in ana-lyzing the taxonomic complexity of biofilms [29, 30].

TEM, however, remains unrivaled for characterizing the fine structure of the extracel-lular matrix of biofilms and evaluating its relationship with the microorganisms in the

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2 µm

Figure 1.6 Transmission electron micrograph of biofilm from a catfish tank. This low-magnification image demonstrates that the biofilm consists of prokaryotes with different morphologies. Two types of microorganisms, however, appear to be predominant, one forming clusters (arrowheads) whereas the other one (arrows) appears as individual cells surrounded by an electron-dense material, which may be a thick cell wall or secreted material. Both prokaryotes contain highly electron-dense inclusions (V-shaped arrowheads). These microorganisms are surrounded by a loose fibrillar extracellular material (asterisks). Note that ruthenium red was added to the glutaraldehyde and osmium tetroxide fixatives to enhance preservation and contrast of extracellular glycoproteins.

100 µm

Figure 1.7 Transmission electron micrograph (30 000×) demonstrating the fibrillar nature of the extracellular matrix (asterisk) in biofilms from catfish tanks. Some of the fibrils appear to associate with a denser, homogenous matrix (arrows) apparently binding the prokaryotes (arrowheads) together. Note that ruthenium red was added to the glutaraldehyde and osmium tetroxide fixatives to enhance preservation and contrast of extracellular glycoproteins.

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biofilm [31–33], as illustrated in Figure 1.7 for a biofilm from a catfish tank. It is thus very likely that TEM will remain a relevant tool to understand the biology of biofilms because the extracellular biofilm matrix is the key component that binds various organ-isms into a film and that binds this film to a substratum [32].

1.12.2 Material and equipment for TEM

Sample preparation for TEM is time consuming and requires highly skilled technicians or investigators experienced in thin sectioning. TEM also necessitates expensive and maintenance-demanding instrumentation, the centerpiece of which is, of course, the TEM apparatus itself. For these reasons, many academic institutions have created core facilities staffed with personnel specialized in processing samples for TEM and in main-taining and operating the TEM equipment. These facilities also house material and equip-ment necessary for sample processing for TEM, as listed in Table 1.11

Investigators using core TEM facilities, however, are usually responsible for fixation of the sample. These investigators should also have an understanding of the processing steps following fixation, should troubleshooting become necessary.

1.12.3 Sample preparation for TEM

Fixation

1. Mince sample into 1 mm3 pieces in the fixation solution consisting of 2% glutaraldehyde plus 2% paraformaldehyde in 0.1 M sodium cacodylate buffer (pH 7.3). Allow fixation to proceed for at least two hours at 4 ºC, with agitation.

All reagents should be electron microscopy grade. Ready to use fixative solutions can be purchased from specialized vendors of electron microscopy products and this is highly recommended for consistent results. Keeping the size of the sample pieces small is critical as glutaraldehyde cross-links proteins and cross-linked proteins eventually act as barrier slowing down the penetration of glutaraldehyde. Paraformaldehyde infiltrates into tissue faster than glutaraldehyde but tissue preservation is not as good as with gluta-raldehyde. Therefore, the combination of these two chemicals is an optimal primary fixative.

Some protocols also call for adding 1% (final) ruthenium red to the fixative described above to enhance the contrast of the bacterial cell wall glycocalyx [34–36]; it is normal for ruthenium red to turn the fixative bright red. Figures 1.6 and 1.7 were obtained with samples fixed with ruthenium red added to glutaraldehyde/formaldehyde and to the osmium tetroxide in the post fixation step described below.

2. Rinse 3 × 10 minutes with 4 ºC sodium cacodylate buffer (0.1 M, pH 7.3) (SCB).3. Post-fix sample at 4 ºC in 2% osmium tetroxide in SCB for two hours. This step is

important for membrane preservation. If 1% ruthenium red was added in the glutaral-dehyde fixation step above, it should also be added to the osmium tetroxide solution, which will then turn bright red.

Alternatively, post-fixation may be performed with 3% potassium ferricyanide and 0.8% OsO

4 in SCB at 4 ºC for two hours; this solution is considered to be optimal for

membrane preservation and may improve the membrane contrast as well.4. Rinse 3 × 10 minutes in cold SCB at 4 ºC.

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Table 1.11 Material and equipment needed for the different steps of specimen preparation for TEM and SEM.

TEM SEM

Fixation Standard laboratory equipment to prepare solutions including: precision balances, pH meter, magnetic stirrer, hot plates, graduated cylinders, beakers, and MilliQ water system.Chemical fume hood to handle noxious fixatives and embedding chemicals.Degreased, clean razor blades, No. 10 scalpel blades, and pink dental wax for mincing samples prior to fixation.Dissecting microscope to identify sample parts of interest.Fine point tweezers to handle minced tissue pieces.Urine cups as vessels for sample fixation.4 ºC refrigerator to store solutions and samples in fixative.

Same as for TEM

Tissue embedding

Borosilicate glass or polyethylene vials with caps.Baskets for carrying tissue pieces from vial to vial.Pre-embedding tissue processing can be either manual or with an automatic, dedicated tissue processor for TEM.Embedding molds or BEEM capsules. Oven (up to 80 ºC) for curing.

Not applicable

Tissue sectioning

Ultramicrotome placed on an anti-vibration table.Diamond knives.Glass knife breaker.Standard light microscopy equipment to stain and observe semi-thin sections. Dissecting microscope and blades to trim the blocks.Copper grids to hold thin sections; grids may be either uncoated or formvar coated.

Not applicable

Staining Standard laboratory equipment for preparing solutions.Fine point tweezers to handle grids during staining.Filter paper.Grid storage boxes.

Standard laboratory equipment for preparing solutions.

Critical point drying

Not applicable. Critical point dryer apparatus for SEM and liquid CO2

Sputter coating

Not applicable. Sputter coater instrument for SEM.Sputtering targets such as gold or gold/palladium (60:40).SEM stubs.Tape or paste to attach samples to stub.SEM stubs storage box.

Observation Transmission electron microscope capable of generating at least 60 kV.

Scanning electron microscope (1–30 kV) or Environmental SEM (ESEMs are capable of functioning in both ESEM and regular SEM modes). Attachment module for EDS if necessary.

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5. Rinse 3 × 10 minutes with distilled water at room temperature.6. En bloc stain samples with 2% (w/v) aqueous uranyl acetate for one hour at room

temperature.7. Rinse samples 3 × 10 minutes with distilled water at room temperature.8. Dehydrate at room temperature in graded ethanol series 10%, 20%, 40%, 50%, 60%,

70%, 80%, 90%, absolute ethanol, 3 × 10 minutes for each ethanol dilution.

Embedding

Embedding of biological material for TEM is routinely performed in Eponate, which is a viscous epoxy resin. Note that low viscosity, hydrophilic media are also available for TEM embedding. These media include Spurr, LR White and LR Gold. They are useful for immu-nogold labeling of proteins because they preserve antigenicity better than Eponate and because immuno-labeling reagents penetrate these hydrophilic media more easily than Eponate [37]. Spurr is also sometimes used for samples that are difficult to infiltrate such as chitinous material and bone [38].

Unless stated otherwise, embedding in Eponate is performed at room temperature and with constant agitation, which is best achieved with a rotary shaker. The following steps are involved:

1. Rinse sample 3 × 10 minutes in propylene oxide.2. Infiltrate sample with 1:1 propylene oxide:eponate mixture for 30 minutes.3. Infiltrate sample with Eponate (2 × 1 hour). Thirty minutes prior to transferring sample to

an embedding mold, place sample in Eponate under vacuum to degas Eponate and to improve infiltration of viscous Eponate into the sample.

4. Place a small drop of Eponate in the area of the flat embedding mold or the BEEM cap-sule where the sample will be transferred. Remove air bubbles from the drop of Eponate with dissecting needles. With applicator sticks or toothpicks, transfer the sample into the drop of Eponate and orient sample such that it will be close to the surface of the final block, which is toward the small end of the flat embedding mold or toward the extremity of the Beem capsule. Finish filling the mold or Beem capsule with Eponate.

5. Cure the Eponate in an oven at 70 ºC for 18–48 hours. Cured Eponate should be hard.6. Remove the hard Eponate block containing the sample from the mold.

Semi-thin section preparation and staining

Prior to carrying out thin sectioning, semi-thin (0.5 μm thick) sections are obtained from the Eponate blocks containing the samples. These sections are examined by light microscopy to determine whether the sample in the block will be useful for TEM observation [39]. This step is necessary because the pieces minced off the initial sample are small and thus may represent a portion of the sample containing our interest. For instance, some of the 1 mm3 pieces minced off of a biofilm may contain debris material without interest for the study, while other pieces actually contain the microorganisms and their matrix. Only blocks containing such sample pieces should be selected for further processing. In addition, light microscopic observation of semi-thin sec-tions enables the investigator to select areas of interest for further ultra-thin sectioning [39].

Semi thin sections are prepared and stained as follows:

1. Excessive epoxy resin is shaved off the tissue block with a clean razor blade.2. Semi-thin sections are obtained with an ultramicrotome fitted with either glass knives or

diamond knives. Operating the ultramicrotome is delicate and should be taught or per-formed by a skilled operator.

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3. Transfer the semi-thin sections onto a glass slide for light microscopy and let them dry and adhere to the slide.

4. Stain the semi-thin sections with filtered aqueous 0.5% toluidine blue plus 1% borax, for five minutes on a hot plate at 100 ºC.

5. Rinse excess stain with running tap water, and remove excess stain with 95% ethanol containing a drop of acetic acid per 50 ml ethanol. The slides are then further rinsed with tap water, air dried, and mounted with a glass cover slip for light microscopy observation.

Preparation of thin sections

Blocks selected for thin sectioning may require additional trimming around the sample to minimize the area being sectioned. Part of the sample than may not be useful for observation may also be trimmed off. The smaller the area sectioned, the higher the chance to obtain quality thin sections.

An ultramicrotome fitted with a diamond blade is used to thin section selected blocks. Sections should be 50–70 nm thick for unsupported grid or ~30 nm for forvar-coated grids, as the forvar film contributes to the overall thickness and will reduce the brightness and contrast of the image during TEM observation. Thin sectioning with the ultramicrotome and picking up thin sections onto copper grids are skills requiring patience and experience and should be taught by an experienced operator.

The grids with the sections are then air dried in a clean dish covered by a lid.Focused ion beam (FIB) microscopes have recently emerged as an alternative to thin

sectioning with ultamicrotomes [40, 41]. However, FIBs are much more costly than ultramicrotomes.

Staining thin-sections with uranyl acetate and lead citrate

Staining should be performed at room temperature using only dry sections. The staining steps can be carried out by depositing a drop of staining solution on a clean surface (e.g. the bottom of tissue culture Petri dish) and by placing the face of the grid supporting the section in contact with the drop.

1. Stain dry thin sections with 4% (w/v) aqueous uranyl acetate for 30 minutes, protected from light. This will enhance contrast of the sections.

2. Rinse with distilled water several times.3. Freshly prepare Reynolds lead citrate. For 50 ml of Reynolds lead citrate add 1.33 g of

lead nitrate and 1.76 g of sodium citrate to 30 ml of distilled water; dissolve with constant stirring (may take up to 1 h). To the milky white suspension add 8 ml of freshly prepared 1 M (1 N) sodium hydroxide until mixture clears. Add distilled water up to 50 ml. The solution may be filtered or passed through a Millipore filter to remove any undissolved material.

4. Stain for two minutes with Reynolds lead citrate, protected from light. Also protect from ambient carbon dioxide by placing dry sodium hydroxide pellets in the staining container (carbon dioxide may dissolve in the staining solution and form insoluble lead carbonate).

5. Rinse with distilled water several times.

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6. Air dry the grids under the lid of a clean petri dish to protect the sections from dust.7. Store the grids at room temperature in a TEM grids storage box kept in a dry and dust-

free environment. Under these conditions, stained sections on grids can be stored indefinitely.


Inadequate fixation

Poorly fixed organisms display mitochondria with swollen or blown out cristae, breaks in the plasma membrane or cell wall, disruption of the general cellular architecture, and cellular lysis. Poor fixation cannot be remedied and new samples need to be collected. The key factors in effective fixation include: using electron microscopy grade fixative and chemicals, mincing the tissue in primary fixative into pieces no bigger than 1–2 mm3 to ensure rapid penetration of the fixative, using a volume of fixative about 20 times that of the tissue pieces. In addition, it is imperative to fix the tissue immediately after sampling to avoid autolysis. Samples frozen either before or after fixation should never be used because the ice crystals forming during freezing will mangle biological structures.

Poor plastic embedding

This problem may result in air bubbles or excessive softness of the plastic block or tissue and eventually hinder sectioning. In addition, incomplete plastic infiltration will cause the tissue to pull off the plastic block during sectioning and/or in the formation of holes in the sections. Poor plastic embedding may occur when samples have not been thoroughly dehydrated after fixation or when sample agitation during the embedding steps was insuf-ficient. The use of an 1:1 epoxy:propylene oxide mixture is important in helping the vis-cous epoxy resin infiltrate the sample. Ideally, epoxy infiltration should be carried out under vacuum with constant rotation or agitation. Processing time with propylene oxide during infiltration of viscous epoxy resin should be minimized to prevent lipid removal from membranes.

Improper embedding with epoxy resins cannot be remedied. The only alternative is to repeat the embedding and pre-embedding steps with spare sample pieces; it is therefore strongly advised that a sample of interest should be minced into many small pieces that can be used as backup should problem arise during the embedding.

Poor quality thin sections

Major problems encountered with thin sections include excessive thickness, shatter, holes, streaks, and folding of sections onto themselves. Assuming proper sample fixation and plas-tic embedding, the quality of thin sections is highly dependent on the skills of the technician performing the ultramicrotome sectioning. In addition, care should be taken to section with ultramicrotomes that are regularly serviced by the vendor and to avoid diamond knifes that are at the end of their lifetime.

Sections that are too thick or folded onto themselves will appear opaque during TEM observation. Chatter manifests itself as a periodic variation in electron density of thin

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sections, akin to a compression wave. Chatter is usually the result of improper clearance angle vibrations occurring during sectioning and because of thinness of the sections even the slightest vibration may cause chatter. Chatter can be minimized by placing the ultrami-crotome on an anti-vibration table and away from air drafts. Holes and streaks in sections may be caused by diamond knives that are at the end of their lifetime.

Low contrast

Poor contrast in thin sections observed by TEM may be due to insufficient fixation or staining. Insufficient fixation results in tissue autolysis and degradation of biomolecules capable to binding uranium and/or lead; this cannot be remedied. Poor contrast due to insufficient staining with uranyl acetate may be remedied by re-staining the sections with a fresh solution of uranyl acetate; in addition, make sure to include an en block staining step with uranyl acetate prior to dehydration. Reduced structural detail and contrast of glycogen, ribosomes, and membrane lipoprotein may also be caused by excessive lead citrate staining time (over two minutes).

Precipitation artifacts

Osmium tetroxide may cause black peppery precipitation throughout the sample. The color of osmium tetroxide is normally straw yellow but it will turn black when con-taminated with incompatible substances. This could be due to improper washing between the glutaldehyde and osmium steps, dirty dissecting instruments or processing containers, or insufficient washing in 0.1 M sodium cacodylate buffer after the osmium tetroxide fixation step.

Fine needle crystals precipitation may occur when phosphate buffer saline is present dur-ing en bloc uranyl acetate staining [42]. Precipitation may also occur when using uranyl acetate solutions that are milky or cloudy rather than clear and crisp yellow in color. To avoid precipitation, always keep uranyl acetate solutions protected from direct light by stor-ing them in amber colored or aliminum foil wrapped bottles.

Improperly prepared Reynold lead acetate can be responsible for precipitate on grids. With time, Reynold lead acetate solution will form precipitate due to reaction with ambient carbon dioxide.

Finally, precipitation may be caused by impurities in the copper grids reacting with heavy metal stains. This problem may be avoided by cleaning the grids by sonication in ethanol.

Dirt and dust contamination

During TEM observations, dirt and dust will appear as filamentous or aggregated material above the section focal plane and will obscure part of the section while the remaining sec-tion is in focus. Dirt and dust may look very similar to staining precipitates (see above) but precipitates tend to be widely spread over sections whereas dirt and dust are more ran-domly distributed. Dirt and dust can be avoided by keeping sections covered at all times when staining, washing and drying, and by using only filtrated solutions and very clean staining and wash containers. In addition, staining and washing thin sections should not be performed in dusty areas or in areas with air drafts. Dirt and dust contamination can be further avoided by avoiding storing TEM sample grids in the open air and by keeping them

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in storage boxes dedicated for TEM grids. Evidently, these boxes should be very clean and if necessary new TEM grids box may be washed with a mild detergent and rinsed with tap and distilled water before being thoroughly dried.

1.12.5 Data analysis and presentation

TEM images are recorded digitally and should be saved as grayscale TIF files. If neces-sary, contrast and gamma point adjustments can be performed with specialized software such as ImageJ or Photoshop. Digital manipulations should be applied to the whole image and not to selected image parts to avoid generating misleading information with respect to contrast in different parts of the image. Scale bars should be added digitally when taking the image and should be used to calculate the dimensions of structures of interest in the image.

Morphometric and stereological quantitative analysis of TEM images can be performed [43, 44] and may be useful in estimating the fraction of the biofilm volume occupied by microorganisms and extracellular matrix components. These methods, however, are very involved and sometime require serial thin sectioning the sample.

1.13 Scanning electron microscopy (SEM)

1.13.1 Purpose of SEM

SEM and TEM are complementary in biofilm high-resolution characterization because TEM provides a two-dimensional image of intracellular organelles while SEM yields a three-dimensional rendering of the surface of the biofilm, thereby revealing the overall shape of the organisms composing the biofilm as well as their organization relative to each other and to the extracellular matrix. For example, SEM examination of fish tank biofilms revealed rod-shaped bacteria densely packed at the surface of a granular extracellular matrix (Figure 1.8A–C). High-magnification SEM observations further demonstrated the presence of two types of bacteria differing by their size (Figure 1.8C), which was in agreement with TEM observations of the same biofilm (Figures 1.6 and 1.7).

SEM necessitates thorough sample dehydration prior to sputter coating with a con-ductive metal. The dehydration steps inevitably lead to some shrinkage, which may alter the size and shape of the organisms present in the biofilm and is also detrimental to the structure of the extracellular matrix [45, 46]. Environmental SEM (ESEM) is a modality of SEM than can be used to minimize sample shrinkage as it does not involve dehydration (Figure 1.8D). However, due to their high moisture content, samples for ESEM cannot be coated with metals. Due to the lack of metal coating of the sample, ESEM images are less contrasted than their SEM counter parts (compare Figures 1.8C and 1.8D).

SEM may also be fitted with optional equipment for energy-dispersive X-ray spectroscopy (EDS or EDX). EDS is a method that can provide highly informative characterization of the elemental composition of biofilms either by determining rela-tive percentage of specific elements or by mapping specific elements to particular fea-tures of the sample [47]. EDS, for example, was instrumental in showing that bacterial biofilms collected in arctic waters were sulfur-rich [48] and in determining the response of bacterial biofilms to silver [49]. EDS also suggested that some dinosaurian soft tis-sues recovered in fossils were actually iron-laden bacterial biofilms [50] and was

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instrumental in revealing the role of biofilms in the deposition of gold nanoparticles in natural gold deposits [51].

Two fairly recent SEM modalities, field emission gun (FEG) and focused ion beam (FIB) SEM bring the resolution of SEM close to that of TEM [52] and also enable intracellular features to be visualized by varying the energy of the incident electron beam and using the secondary electrons emitted by the specimen to form an image [53, 54]. In addition, the backscattered electron detector of FEG SEM can serve to identify gold particles conjugated to antibodies against specific proteins [55].

While the fixation and staining steps for SEM are similar to those of TEM, SEM sample preparation does not require plastic embedding and thin sectioning. SEM sample preparation is thus less labor intensive and less technically challenging than TEM. This makes SEM advantageous for routine, rapid analysis of biofilms. It has been suggested, however, that TEM may be more suited than SEM to characterize the extra-cellular matrix of biofilms, as the matrix structure may be optimally preserved after embedding in plastic resin [47].

1 µm

1 µm2 µm

(C) (D)

10 µm

(A) (B)

Figure 1.8 Scanning electron micrographs of a catfish tank biofilm. (A) Low magnification demonstrates that the biofilm consists of aggregated grains covered by filamentous-structures, which at higher magnification appear to be rod-shaped bacteria (B). (C) In addition, high magnification observations reveal heterogeneity in the size of the microorganisms present in the biofilm, with some of these organisms (arrows) larger than others (arrowheads). (D) Sample processed for ESEM display features similar to those of samples processed for SEM (A–C) but the contrast of ESEM images is lower than that of SEM images due to the absence of metal coating during sample preparation for ESEM. Note that ruthenium red was added to the glutaraldehyde and osmium tetroxide fixatives to enhance preservation and contrast of extracellular glycoproteins.

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1.13.2 Material and equipment for SEM

The equipment listed in Table 1.11 should enable individual investigators to prepare samples for SEM or ESEM in their laboratory. Sample preparation for EDS is carried out as for SEM or ESEM. Similar to TEMs, SEMs are capital pieces of equipment and, therefore, often belong to institutional research core facilities rather than to individual laboratories.

1.13.3 Sample preparation for SEM

1. Carefully wipe off razor or scalpel blades with alcohol to remove any greasy substance that the manufacturer might have used to prevent rusting. With a clean blade, mince the sample submerged in primary fixative into 1–2 mm3 pieces; clean, pink dental wax is a common support to mince the sample.

Immediately place pieces into vials that can be capped to avoid evaporation and minimize changes in pH of the fixative, which contain about 20 times the sample volume of 2% (v/v) glutaldehyde and 2% (v/v) paraformaldehyde in 0.1 M sodium cacodylate buffer (SCB) (pH = 7.3). Several pieces of a given sample may be placed into the same vial provided that there is ample space between them. Fix at 4 ºC for a minimum of two hours. Samples may be kept under these conditions until ready for further steps. It is normal for the sample to appear darker in color after fixation.

As described above for TEM, 1% (final) ruthenium red may be added to the glutaraldehyde/paraformaldehyde fixative to enhance the contrast of the bacterial cell wall glycocalyx [56–58]. Figure 1.8 was obtained with samples fixed this way.

2. Rinse 3 × 10 minutes in 4 ºC SCB.3. Post-fix the sample with 2% osmium tetroxide in SCB for two hours at 4 ºC. Note:

osmium tetroxide will form a black precipitate if phosphate-containing buffers are used (hence the utility of SBC). Osmium tetroxide is critical for optimal preservation of lipid-containing structures.

If 1% ruthenium red was added in the glutaraldehyde/paraformaldehyde fixation step above, it should also be added to the osmium tetroxide solution, which will then turn bright red.

4. Rinse 3 × 10 minutes with SCB at 4 ºC.5. Rinse 3 × 10 minutes with distilled water at 25 ºC.6. Stain samples with 2% (w/v distilled water) uranyl acetate for one hour (this is some-

times referred to as “en bloc” staining as opposed to staining of sections). This step increases the contrast of the sample during SEM observation.

7. Rinse samples 3 × 10 minutes with distilled water at 25 ºC.8. Dehydrate sample in graded series of ethanol including 20%, 40%, 50%, 60%, 70%,

80%, 90% and absolute ethanol. Perform each step 3 × 5 minutes at 25 ºC.9. Thoroughly dry the samples either by critical point drying (CPD) or with hexamethyl-

disialzane (HMDS).For critical point drying, the sample is placed in the chamber of the CPD apparatus

and is covered with absolute ethanol. The CPD chamber is closed securely and cooled down to 0 ºC or lower with liquid carbon dioxide (CO

2) as per the manufacturer’s

instructions. Next, cold, liquid carbon dioxide is introduced into the chamber and allowed to mix with the absolute ethanol. The chamber is then purged and refilled with liquid CO

2 several times until all of the ethanol is replaced with CO

2. Complete

removal of alcohol is achieved when a tissue paper placed at the purge vent does not get

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wet. The CPD chamber is then filled with cold liquid carbon dioxide. The CPD fill switch and purge switch are turned off sequentially and the CPD heat switch is then turned on. When the CPD chamber temperature and pressure rise to 32 ºC and 1250 psi, respectively, the carbon dioxide becomes gaseous and transparent. Samples should then be equilibrated at this critical point for several minutes. The gas in the chamber should then be bled off very slowly until the pressure becomes less than 150 psi, at which point the chamber can be vented and the sample removed.

HMDS drying is an inexpensive and rapid alternative to CPD. After the last absolute ethanol bath, samples are incubated for five minutes each in the following: 1:1 ethanol:HMDS, 1:2 ethanol:HMDS, and, finally, pure HMDS. Samples are then air dried in a clean area at room temperature or in a desiccators under low vacuum. Caution: HMDS is very toxic; use with caution under a fume hood following MSDS guidelines.

10. Mount CPD or HMDS dried sample onto SEM stubs. The simplest method is to attach the sample on the SEM stub with regular double-sided adhesive tabs. Other adhesives that can be used include: double-sided adhesive tabs containing carbon, silver or copper to improve conductivity and various gluing pastes, including colloidal silver paste, isopropanol-based colloidal graphite paste, or water-basde colloidal graphite paste. These adhesive pastes and tapes can be purchased from electron microscopy products suppliers.

11. Sputter coat samples with metal or carbon. The purpose of this step is to make non-conductive biofilm samples conductive by applying a thin metallic layer with a sputter coater apparatus dedicated to SEM (Table 1.11). Gold, gold–palladium (60:40), palladium, or platinum can be used to coat samples with low vacuum sputter coater. Gold–palladium was used by us to prepare fish tank biofilm for SEM (Figure 1.8). Note that newer sputter coaters are equipped with turbo pumps that can achieve high vacu-ums permissive to coating with additional targets, including carbon, chromium, silver, and tungsten, which could be useful for some applications.

The thickness of the metal layer should be determined by trial and error, but usually for biological samples and with a gold–palladium target, 60 nm thickness is a good start. It is preferable to err on the thin layer side, because adding additional metal coat-ing can be easily done but a thick metal coating layer cannot be made thinner.

The sputter coater apparatus should be used as directed by the manufacturer. Generally, this requires installing the target of interest and selecting the desired thick-ness of the metal coating. Afterwards, the mounted specimen is transferred onto the sputter coater table, the lid of the chamber is closed and the chamber is then pumped to required vacuum. Next, argon is allowed into the chamber, and coating and sample rotation can start. When the process is over, the chamber is vented and the sample can be removed with tweezers.

12. The stubs with mounted samples can then be stored in a dedicated box, which should be kept in a dessicator until ready for SEM observations.

1.13.4 Sample preparation for ESEM

For ESEM, steps 1–7 are the same as above for SEM. It is important to thoroughly rinse with distilled water in step 7, such that no buffer salt remains on the samples. After the water washes, the samples are mounted on SEM stubs as described in step 10

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above and are then ready for ESEM observation. Sputter coating is not to be performed on samples for ESEM.


Fixation artifacts

These are evidenced by membrane blebbing or disruption. It cannot be stressed enough that samples need to be fixed quickly and that the volume of fixative needs to be at least 20 times that of the sample. In addition, the fixative needs to be of the proper concentra-tion, buffer molar concentration and pH. Freezing the sample should never be performed prior to or after fixation. If necessary, samples can be stored for several weeks in primary fixative at 4 ºC.

Gross sample damageSmall and fragile samples might have been damaged during handling, resulting in cracks, cuts, and holes. Using applicator sticks (or tooth picks) and careful sample handling may remedy the problem.

Gross precipitationGross precipitation may be caused by: (i) insuffisient rinsing between glutaraldehyde fixation and uranyl acetate en bloc staining; (ii) the presence of phosphate ions during osmium tetroxide fixation and/or uranyl acetate staining; (iii) poor water purity – when-ever possible use 800 mOhms MilliQ water; (iv) improperly prepared solutions – osmium tetroxide should be straw yellow, a black color means gross contamination; uranyl acetate solution should be crisp, clear yellow in color and should not appear milky or cloudy. Uranyl acetate precipitation appears as a white surface precipitate on the sample.

Visible moisture in the CPD chamber after ventingThis will result in improper sample drying, which will lead to inadequate metal coating, as a moisture layer will form between the sample and the metal coating. To prevent condensa-tion of ambient room air rushing into the CPD chamber, vent the chamber slowly after the sample has been dried at the critical point temperature.

Crystal forming on the sample during ESEMThis is caused by residual salt left on the sample due to insufficient washing with distilled water.

Excessive sample shrinkingThis may occur when some of the lower alcohol concentration steps are skipped during the dehydration procedure. Excessive shrinking may also occur when tissue is dehydrated with HMDS rather than by CPD. In addition, HMDS may cause cracking artifacts. Therefore, if possible, CPD is preferred to HMDS. Excessive CPD may also create shrinking artifacts; consequently, samples should not be held at the critical point and temperature longer than necessary.

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CrackingSamples may crack if subjected to an electron beam of too high a voltage, or if drying is insufficient (which may occur if the sample is too large) or has been performed with HMDS, or if metal coating is poor. The only alternative is to start over with an unprocessed piece of fixed sample and to optimize the conditions.

Poor surface detailsToo much sputter metal coating may mask fine detail; excessive coating cannot be removed without damaging the sample. Too little metal coating decreases image detail and causes charge artifacts that result in instability of the electron beam during SEM observation. In the case of insufficient coating, the sample can be recoated.

Improper sample orientationIf the samples are not oriented properly for observation, they can be remounted in the desired orientation on another SEM stub. A magnifying binocular may be used to ensure that sam-ples are mounted in the desired orientation. Samples might need to be recoated after remounting for reorientation.

1.13.6 Data analysis and presentation

SEM data consist of digital pictures of representative sample fields. Image files should be in TIF format rather than in formats involving compression algorithms, such as jpeg, because this may result in pixilation. In addition, images should be grayscale rather than indexed color or bitmap. Scale bars should be included during picture collection and should be used to calculate the dimensions of structures of interest and/or to obtain quantitative information on number of organisms present per unit surface and/or percentage of biofilm occupied by microorganisms.

Stereoscopic views can be acquired by taking two pictures, one that is not tilted and the other one that is titled at the eucentric working distance of the SEM. The two images should then be merged with image processing software and the resulting image can be evaluated with stereoscopic glasses if needed.

EDS analysis can be documented as percentage of relative elemental abundance, which can be represented visually with spectrum where the different elements of interest are rep-resented by peaks which height is proportional to the elemental relative abundance. Depending on the study, normalization to specific element(s) may be desirable.

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