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. Author manuscript; available in PMC: 2022 Jan 30.
Published in final edited form as: Methods Mol Biol. 2014;1147:85–103. doi: 10.1007/978-1-4939-0467-9_7

Experimental Approaches to Investigating the Vaginal Biofilm Microbiome

Marc M Baum 1, Manjula Gunawardana 1, Paul Webster 1
PMCID: PMC8801157  NIHMSID: NIHMS1552111  PMID: 24664828

Abstract

Unraveling the complex ecology of the vaginal biofilm microbiome relies on a number of complementary techniques. Here, we describe the experimental approaches for studying vaginal microbial biofilm samples with a focus on specimen preparation for subsequent analysis. The techniques include fluorescence microscopy, fluorescence in situ hybridization, and scanning and transmission electron microscopy. Isolation of microbial DNA and RNA from these samples is covered along with a brief discussion of chemical analysis methods.

Keywords: Vaginal microbiome, Microbial biofilms, Fluorescence microscopy, Fluorescence in situ hybridization, Scanning electron microscopy, Transmission electron microscopy, Genomic microbial DNA, RNA, Correlative microscopy

1. Introduction

Bacteria colonize surfaces and establish a sessile mode of growth in biofilms, chemically and morphologically heterogeneous matrices of extracellular polymeric substances (EPS). Microbial biofilms are implicated in chronic infections [1] and are more resistant to antimicrobial agents and the immune system than their planktonic counterparts [2, 3]. The normal vaginal microbiota makes up a complex, diverse community that is believed to play an important protective role in maintaining the health of a woman, sexual partner, or newborn [4]. Microorganisms in the vaginal tract are believed to grow predominantly as sessile polymicrobial communities encapsulated in biofilms [5-7]. The Human Microbiome Project [8, 9] is improving our understanding of how vaginal bacteria in healthy individuals maintain a balanced community [10, 11] and the pathogenic capabilities of key species that mediate poor health outcomes. Vaginal bacterial biofilms have been associated with a bacterial vaginosis and mortality resulting from tampon-related toxic shock syndrome in menstruating women. In previous studies, we have demonstrated that polymicrobial biofilms grow in vivo on the surface of intravaginal rings (IVRs) implanted in pig-tailed macaques [12] and women [13].

Gaps in our understanding of medical microbial ecology largely stem from our reliance on culture-dependent microbiological methods, which typically can identify less than 1 % of the bacterial cells in a given ecosystem [14]. This seminal realization, known as the “great plate count anomaly,” has led to tremendous advances over the past 20 years through the development and application of environmental metagenomics, microanalytical methodologies, novel cultivation methods, and the coupling of stable and radiogenic isotopes with molecular analysis of biosignatures [15]. The strict requirement for pure cultures has been alleviated by the refinement of experimental approaches aimed at improving our understanding of the composition, structure, and function of vaginal microbial biofilm communities. The approaches covered here are based on a combination of established imaging techniques using instrumentation accessible to most research laboratories, including:

  • Fluorescence microscopy: Allows community structure to be studied by selectively labeling different components of the biofilm EPS with a range of fluorescent probes [16-19].

  • Fluorescence in situ hybridization (FISH): FISH is an established molecular, cultivation-independent technique that detects nucleic acid sequences by a fluorescently labeled probe, which hybridizes specifically to its complementary target sequence within the intact cell [14].

  • Scanning and transmission electron microscopy (SEM and TEM): Electron microscopy allows the biofilm microbial community structure to be examined at high resolution. We have developed a method to combine FISH and SEM data to label bacteria (FISH) and provide high-resolution information on the reference space (SEM) [20].

  • Molecular analysis: Genomic DNA and RNA isolation, amplification, and sequencing.

  • Chemical analysis: Colorimetry, nondestructive spectroscopy, and chromatography with tandem mass spectrometric detection techniques.

The corresponding protocols described below were validated in our laboratories for vaginal microbial biofilm samples.

2. Materials

All solutions should be prepared with deionized water, unless otherwise specified, using analytical grade reagents. Prepare and store all reagents at room temperature unless otherwise noted. Diligently follow all appropriate regulations and precautions when handling reagents and disposing of waste materials.

2.1. Fluorescence Microscopy

  1. Method optimization and imaging were carried out using an EVOS fl digital inverted fluorescence microscope (AMG).

  2. Samples are prepared and viewed in a Lab-Tek II Chamber #1.5 German Coverglass System. We typically use the 8-chamber system (Model 155409, Nalge Nunc International Corp.).

  3. 20 μM SYTO® 63 in deionized H2 O, 0.5 % w/v fluorescein isothiocyanate isomer I (FITC) in PBS, a 5,000× stock solution SYPRO® Orange in DMSO, 10 μg/mL Calcofluor White in sodium phosphate buffer (10 mM, pH 7.5), 500 μg/mL Nile red in acetone, and 10 mM 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI) in deionized H2O.

  4. 1× PBS (pH 7.2), phosphate buffer (0.05 M, pH 7.4).

  5. Aqueous glycerol solution (75 %, v/v).

  6. Aqueous acetic acid solution (7.5 %, v/v).

  7. Aqueous potassium hydroxide solution (10 %, w/v).

  8. Deionized H2O.

  9. Timer.

  10. Kimwipes.

  11. Fluorescence microscope.

2.2. Fluorescence In Situ Hybridization (FISH)

  1. Method optimization and imaging were carried out using an EVOS fl digital inverted fluorescence microscope (AMG).

  2. Samples are prepared and viewed in a Lab-Tek II Chamber #1.5 German Coverglass System. We typically use the 8-chamber system (Model 155409, Nalge Nunc International Corp.).

  3. Published oligonucleotide probe sequences [21] are employed when possible. We typically order our probes from Integrated DNA Technologies, Inc., and request purification by HPLC.

  4. Hybridization buffer: 0.9 M NaCl, 0.02 M Tris-HCl buffer (pH 7.4), and 0.001 % SDS in deionized H2O (see Note 1).

  5. Washing buffer: 0.175 M NaCl, 0.02 M Tris-HCl buffer (pH 7.4), and 0.001 % SDS in deionized H2O.

  6. 1× PBS (pH 7.2).

  7. Aqueous ethanol solution (50 %, v/v).

  8. Deionized H2O.

  9. Hybridization oven.

  10. Timer.

  11. Kimwipes.

  12. Fluorescence microscope.

2.3. Electron Microscopy

  1. Transmission electron microscope.

  2. Scanning electron microscope.

  3. Critical-point dryer.

  4. Ultramicrotome.

  5. 45° diamond knife for routine sectioning.

  6. Formvar-coated metal specimen grids.

  7. Specimen stubs.

  8. 2.5 % v/v glutaraldehyde in 100 mM phosphate buffer (pH 7.2).

  9. 1 % w/v aqueous osmium tetroxide.

  10. 1 % w/v aqueous tannic acid.

  11. Saturated aqueous thiocarbohydrazide (TCH).

  12. Maleate buffer (50 mM, pH 5.2).

  13. 3 % potassium ferricyanide.

  14. Dilution series of ethanol or acetone.

  15. 100 % ethanol or acetone.

  16. Epoxy resin for specimen embedding.

  17. 60 °C oven.

  18. Embedding molds.

  19. Fine jewelers forceps (e.g., Dumont tweezers styles 5 and 7).

  20. Parafilm.

  21. 3 % w/v aqueous uranyl acetate.

  22. Aqueous lead citrate.

  23. Deionized H2O.

3. Methods

3.1. Sample Preservation and Storage

The vaginal biofilm sample collection, preservation, and storage strategy are fundamentally important in determining the range and quality of the subsequent measurements. Typically samples need to be divided upon collection and stored in a variety of preservative media, as described below:

  1. Electron microscopy: 2.5 % v/v glutaraldehyde in phosphate buffer (pH 7.2) (see Note 2).

  2. Fluorescence microscopy/FISH: 50 % v/v ethanol in deionized H2O.

  3. Microbial DNA/chemical analysis: Flash freeze in liquid nitrogen.

  4. Microbial RNA: Aurum lysis buffer (Bio-Rad) [22].

  5. Samples preserved in steps 1 and 2 should be stored and transported at 4 °C.

  6. Samples preserved in steps 3 and 4 should be stored and transported at −80 °C.

3.2. Fluorescence Microscopy

  1. The combination of multiple, selective fluorescent probes allows the principal chemical components to be imaged (see Note 3).

  2. Numerous fluorescent probes have been reported for microbial biofilm imaging [16-19]. The most useful probes for vaginal biofilm analysis are described below.

  3. We have used confocal laser scanning and inverted fluorescence microscopes to image vaginal microbial biofilms with similar results [12, 13].

  4. Biofilms are stained after hybridization for FISH, directly in the chamber slide, unless otherwise noted.

  5. Vortex agitation is not used to mix the samples. Sample washing is carried out gently using a 1 mL pipet and by blotting with a clean Kimwipes. (See Note 4).

  6. Samples containing the fluorescent stains are incubated in the dark.

  7. Prior to staining, gently remove the residual liquid from the chamber slide sample.

  8. SYTO® 63 (total cells): Add stock solution to just cover the sample.

  9. Incubate for 30 min at room temperature.

  10. Gently remove stain solution and wash sample with 1× PBS (pH 7.2) until no background stain is visible (2–3 washes are typical).

  11. FITC (proteins, amino sugars): Wash sample three times with phosphate buffer (0.05 M, pH 7.4).

  12. Immerse the sample in 0.5 mL phosphate buffer (0.05 M, pH 7.4) and add FITC stock solution (5 μL).

  13. Incubate for 10 min at room temperature.

  14. SYPRO® Orange (proteins) (Fig. 1): Dilute the stock SYPRO Orange solution 1:5,000 with aqueous acetic acid (7.5 %, v/v) with vigorous mixing.

  15. Add diluted stock solution to just cover the sample.

  16. Incubate for 5 min at room temperature.

  17. Gently remove stain solution and wash sample with 1× PBS (pH 7.2) until no background stain is visible (2–3 washes are typical).

  18. Calcofluor White ( cellulose and chitin): Place sample onto a clean glass slide (see Note 5).

  19. Add on drop of Calcofluor White stock solution and one drop of potassium hydroxide solution (10 % w/v).

  20. Place coverslip over the specimen and leave for 1 min prior to visualization.

  21. Nile red (lipids, hydrophobic sites): Dilute the stock solution with aqueous glycerol (75 % v/v) to a final concentration of 2.5 μg/mL.

  22. Add just enough of the diluted stock solution to just cover the sample.

  23. Incubate for 1–2 min at room temperature.

  24. Gently remove stain solution and wash sample with 1× PBS (pH 7.2) until no background stain is visible (2–3 washes is typical).

  25. DAPI (nucleic acids): Dilute the DAPI stock solution to 300 nM in PBS.

  26. Equilibrate the sample briefly in PBS.

  27. Add the diluted DAPI solution (300 μL) to the sample.

  28. Incubate 1–5 min at room temperature.

  29. Gently remove stain solution and wash sample with 1× PBS (pH 7.2) until no background stain is visible (2–3 washes are typical).

Fig. 1.

Fig. 1

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FISH micrograph of Lactobacillus gasseri (ATCC 33323) biofilms grown in vitro. The sample was fixed overnight at 4 °C in 50 % ethanol. The cells were hybridized with universal bacterial probe EUB-338-Cy5 (red), and the biofilm proteins were labeled with SYPRO Orange (blue). The scale bars is 20 μm

3.3. Fluorescence In Situ Hybridization (FISH)

  1. Our validated FISH method is based on a combination of literature procedures [23-25] (Fig. 2).

  2. Fix specimen in 50 % ethanol for at least 18 h at 4 °C (once the sample is fixed, it can be stored at 4 °C for several months without appreciable deterioration).

  3. On the day when hybridization is planned, prepare the hybridization and washing buffers (see Note 1) according to the probe specification and incubate at hybridization temperature (we typically use 47 °C) until needed (see Note 6).

  4. Remove sample from cooler and thaw to room temperature.

  5. Gently remove the liquid with a pipet, taking care not to disturb the fragile biofilm. Remove the residual 50 % ethanol by blotting with a clean Kimwipes (see Note 4).

  6. Gently add a predetermined volume of PBS solution (pH 7.2) along the inside walls of the tube to rinse the biofilm taking care not to disturb the fragile structure. Never add solutions directly to the specimen. With our specimens (microfuge tubes and chamber slides), 500 μL is typically sufficient.

  7. Remove the PBS solution as in step 5.

  8. Remove hybridization buffer from the incubator and rapidly (to avoid cooling of the buffer) add 400–500 μL to the sample (see Note 7).

  9. Immediately add the FISH probe(s) (50–200 pmol) to the sample.

  10. Mix probe and hybridization buffer by gently aspirating the supernatant fluid up and down in the pipet, 3–5 times.

  11. In all subsequent steps, protect the sample from light to avoid probe photobleaching.

  12. Incubate at 47 °C for 90 min.

  13. Quickly add deionized H2O (1 mL, room temperature) to the sample.

  14. Using a 1 mL pipet, remove all the liquid from the tube, leaving only the hybridized sample behind.

  15. Repeat steps 13 and 14.

  16. Add washing buffer directly from incubator using the same volume as used in step 8 with the hybridization buffer.

  17. Incubate for 20 min at 47 °C.

  18. Remove the washing buffer with a 1 mL pipet.

  19. Add deionized H2O (1 mL, room temperature) to the sample.

  20. Remove most of the H2O with a 1 mL pipet. Lease enough fluid to just cover the specimen.

  21. Store sample at 4 °C until it is to be imaged (see Notes 8-11).

Fig. 2.

Fig. 2

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FISH micrographs of Lactobacillus gasseri (ATCC 33323) biofilms grown in vitro. The cells were hybridized with (a) universal bacterial probe EUB-338-Cy5 (red) and (b) Lactobacillus-specific probe Lac-158-Cy3 (green) [21]. (c) Cells that hybridized with both probes appear yellow/orange. The images clearly show that universal coverage was obtained with both probes. The scale bars are 50 μm

3.4. Scanning Electron Microscopy (SEM)

  1. Fix specimen in 2.5 % glutaraldehyde in 100 mM phosphate buffer (2.5 % v/v) for 2 h (see Notes 2 and 12-16).

  2. Wash thoroughly with deionized H2O to remove aldehyde and phosphate from the specimens.

  3. Soak in 1 % w/v aqueous tannic acid solution for 1 h.

  4. Wash thoroughly with deionized H2O.

  5. Postfix in 1 % w/v aqueous osmium tetroxide for 1 h.

  6. Wash thoroughly with deionized H2O.

  7. Treat specimens with aqueous saturated TCH for 1 h.

  8. Wash thoroughly with deionized H2O.

  9. Treat with aqueous 1 % w/v osmium tetroxide for 1 h.

  10. Wash thoroughly with deionized H2O.

  11. Dehydrate using the following ethanol series: 30 % ethanol (2 × 10 min), 50 % ethanol (2 × 10 min), 70 % ethanol (2 × 10 min), 95 % ethanol (2 × 10 min), and 100 % ethanol (2 × 10 min).

  12. Finally, change in 100 % dry ethanol (see Note 17).

  13. Dry the specimen using a critical-point dryer or hexamethyldisilazane (HMDS) (see Note 18).

  14. Mount dried specimen on aluminum specimen stub and examine in a scanning electron microscope (see Note 19) (Fig. 3).

  15. Specimens should be stored in the presence of desiccant (see Notes 20-22).

Fig. 3.

Fig. 3

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Day 14 microbial biofilms forming on intravaginal rings, delivering the antiherpetic drug acyclovir, worn by women with recurrent genital herpes [13, 26]. (a) SEM image of epithelial cell monolayer on the ring surface; scale bar = 200 μm. (b) SEM image of nanowire-linked microbial biofilm cluster; scale bar = 20 μm

3.5. Transmission Electron Microscopy (TEM)

  1. For examination of morphology, fix specimen in glutaraldehyde buffered in 100 mM phosphate buffer (2.5 % v/v/, pH 7.2) for up to 3 h (see Note 23).

  2. Take care not to cause handling trauma during any relevant dissection (e.g., no crushing, pulling, squeezing, etc.).

  3. Wash with phosphate buffer (100 mM, pH 7.2), at least 3 × 10 min.

  4. Postfix with 1 % w/v aqueous osmium tetroxide on wet ice for 2 h.

  5. Wash thoroughly with deionized H2O.

  6. Postfix again with 1 % aqueous osmium tetroxide. It is possible to reuse the first solution. For the second fixation, add aqueous potassium ferrocyanide solution (3 % w/v) for a final concentration of 0.3 % reduced osmium. Leave for 2 h.

  7. Wash with deionized H2O 3 × 10 min (see Notes 24-27).

  8. Wash with sodium maleate buffer (50 mM, pH 5.2) (see Note 28).

  9. En bloc stain with 1 % w/v uranyl acetate in maleate buffer (50 mM, pH 5.2) for 1 h (see Note 29).

  10. Wash with sodium maleate buffer (50 mM, pH 5.2), two changes, 10 min each.

  11. Wash with deionized H2O 3 × 10 min (see Notes 30 and 31).

  12. Dehydrate with an ethanol series: 30 % ethanol (2 × 10 min), 50 % ethanol (2 × 10 min), 70 % ethanol (2 × 10 min), 95 % ethanol (2 × 10 min), and 100 % ethanol (2 × 10 min).

  13. Ensure that the final ethanol steps are carried out using dried 100 % ethanol.

  14. Replace ethanol with propylene oxide, and leave for two changes, 10 min each.

  15. Replace with fresh propylene oxide for 10 min.

  16. Replace with epoxy resin/propylene oxide (1:1) and leave overnight. At this stage, the epoxy resin should not contain catalyst (see Note 29).

  17. Replace 1:1 mixture with 1:3 propylene oxide/epoxy resin mix and leave for 3–6 h.

  18. Open caps for 1 h to let propylene oxide evaporate.

  19. Replace with fresh resin for 4 h with rotation on mixing wheel.

  20. Prepare fresh resin containing catalyst and place in embedding molds.

  21. Place specimen in molds with unique identifier labels, orientate for best sectioning position, and transfer to 60 °C oven overnight.

  22. Section the embedded specimen in an ultramicrotome equipped with a 45° angle diamond knife.

  23. Collect the section onto Formvar-coated specimen grid and contrast by floating section-side down on drops of 3 % w/v aqueous uranyl acetate placed on Parafilm.

  24. Wash with deionized H2O.

  25. Repeat the staining procedure using aqueous lead citrate (see Notes 32-34).

  26. Dry the grids and examine in the transmission electron microscope (see Note 35).

3.6. Genomic DNA Extraction

  1. The PowerBiofilm™ DNA Isolation Kit is recommended for genomic DNA extraction (see Notes 36-38).

  2. Follow the manufacturer’s instructions without modifications.

  3. The RNeasy Mini Kit is recommended for total RNA isolation.

  4. Follow the manufacturer’s instructions without modifications.

  5. Include an on-column DNase digestion step with the RNase-free DNase kit (see Note 39).

3.7. Chemical Analysis

A detailed discussion of the chemical analysis of vaginal microbial biofilm samples is outside the scope of this book chapter as these endeavors usually are hampered by the small sample sizes available for study. A number of destructive (colorimetric and fluorometric) methods have been described [27] and can provide semiquantitative biofilm chemical composition. SEM coupled with energy dispersive spectroscopy (SEM-EDS) is a powerful tool to obtain elemental microanalysis overlaid with high-resolution SEM images. Nondestructive methods such as Fourier transform infrared (FTIR) spectroscopy and 2-D nuclear magnetic resonance (NMR) spectroscopy can be useful techniques for determining and comparing the principal components of vaginal microbial biofilms [27-35].

Acknowledgment

Research reported in this publication was supported, in part, by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under Award Number R01AI100744. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Footnotes

1.

Formamide can be added to the hybridization buffer if required.

2.

Rapid freezing, ideally using a high-pressure freezer, is the preferred method of sample preservation for electron microscopy examination but is rarely feasible in practice due to logistical constraints at the collection point and the need for specialized, expensive equipment.

3.

Mucins, highly glycosylated large proteins (10–40 MDa) secreted by epithelial cells, make up an entangled viscoelastic gel that forms part of the cervicovaginal secretions [36]. To differentiate these proteins from biofilm EPS, mucins are labeled specifically with commercial monoclonal antibodies in biofilm cryostat sections using standard protocols [37].

4.

Care should be taken to ensure that sample remains hydrated throughout.

5.

Calcofluor White staining can reduce the signal intensity of FISH probes.

6.

The fixation and hybridization times were optimized for our specimens. Due to variability in the vaginal microbial biofilm composition across subjects, it is recommended to vary these times as part of protocol optimization.

7.

Minimize the amount of time the hybridization buffer is out of the incubator to avoid cooling of the solution to room temperature.

8.

When imaging the samples, move from low to high magnification. Use the low magnification to align the sample as much as possible and thereby minimize photobleaching.

9.

Minimize the time spent under high magnification to minimize photobleaching.

10.

Magnification at 60× is usually sufficient to obtain images of single microbial cells. Oil immersion optics and higher magnification provide little benefit due to high photobleaching rates.

11.

While a number of commercial photobleaching suppressants exist, they are not recommended as they can add a haze to the sample image and reduce the probe intensity. It is recommended to practice obtaining high-quality images in a short time period instead.

12.

High-pressure freezing produces immediate immobilization of small specimens in a frozen state. These specimens can be sectioned and examined by electron microscopy while still frozen [38-42]. This approach produces optimal ultrastructural preservation but requires specialized specimen preparation and imaging equipment, as well as considerable technical skills.

13.

High-pressure frozen specimens can be processed for embedding in resin by dehydration at low temperature using a method called freeze substitution [43]. Freeze-substituted specimens can be embedded subsequently in low-temperature resin such as Lowicryl HM20 [43, 44] or warmed for embedding in epoxy resin [45]. Such an approach makes it possible for high-pressure frozen specimens to be examined using routine resin sectioning.

14.

Although chemical fixation is a convenient collection method and dehydration at ambient temperature is a routinely applied protocol in almost all electron microscopy laboratories, it is not an ideal approach for preserving bacterial cells or bacterial biofilms [46, 47].

15.

A feasible approach for preparing bacterial biofilms that does not require high-pressure freezing and avoids chemical fixation is freezing by immersion in a cryogen such as liquid propane or ethane [47]. The cryogen can be prepared in advance and taken to the collection site while frozen in liquid nitrogen. Immediately prior to sample freezing, the cryogen can be warmed by immersion of a warm metal block, and, when liquid, the specimen can be immersed. The frozen specimen is then transferred to 100 % dry ethanol or acetone on dry ice for subsequent processing or transport. Further processing consists of transferring to fresh solvent (optional contents include 2 % glutaraldehyde, 1 % osmium tetroxide, 1 % uranyl acetate, or suitable combinations). Immersion fixation will not produce optimal ultrastructure of bacterial cells or bacterial biofilms, but the preservation will be improved when compared with chemical fixation [47].

16.

Formaldehyde or glutaraldehyde both work well, but glutaraldehyde will make the sample harder [48]. Best results are obtained if specimens are fixed as soon as possible. Handling of the specimen should be kept to a minimum, both before and after fixation. Specimen drying should be avoided at all stages during collecting and subsequent processing.

17.

The second approach for dehydrating chemically fixed material at low temperature requires the specimens to be frozen in the presence of cryoprotectant. Chemically fixed specimens are soaked in dimethylformamide, 30 % v/v glycerol, or 2.3 M sucrose and then immersed in liquid cryogen (propane, ethane, or liquid nitrogen). The frozen specimens are freeze substituted in solvent. If sucrose is used as the cryoprotectant, then methanol must be used as the freeze substitution medium [49]. Methanol is able to solubilize the sucrose in the specimens, whereas if ethanol or acetone is used, the sucrose crystallizes and interferes with subsequent processing.

18.

If immersion fixation is not an option, perhaps due to the hazardous properties of gaseous propane and ethane, an aldehyde fixation is the only option for preserving bacteria and biofilms. The ultrastructure can be improved by dehydration at low temperature. Two approaches are possible. The first, called the progressive lowering of temperature (PLT) method, gradually takes chemically fixed specimens through washing and dehydration steps while lowering the temperature. Fully dehydrated specimens can be held at low temperature, infiltrated with low-temperature embedding resin, and polymerized in resin by ultraviolet light [50]. Alternatively, cold, dehydrated specimens can be gradually warmed to ambient temperature and either embedded in epoxy resin for examination by transmission electron microscopy or further dried using a critical-point drier for examination by scanning electron microscopy. If a critical-point drier is not available, then specimens can be dried in the presence of HMDS, a reasonable substitute for critical-point drying [51].

19.

Metal coating of specimens using a sputter coater may be required to reduce charging in the microscope. Coating with platinum produces a finer layer, but any metal coating over the specimen surface can potentially cover essential fine structural details.

20.

For electron microscopy, specimen preparation is a compromise between applying ideal preparation protocols and practical considerations. For example, the best approach for preserving subcellular morphology usually is high-pressure freezing [47]. However, such an approach is challenging to apply to specimens removed from human subjects due to logistical considerations. Specimens have to be transferred immediately to a high-pressure freezer for immediate freezing by a trained operator.

21.

In extreme circumstances, specimens collected for SEM can be air-dried immediately after collection. The dry specimens can be attached to specimen stubs and examined in the SEM, either with or without a metal coating. The specimen ultrastructure will not be optimal, and damage caused by air-drying will be present. However, the specimen can be observed at low magnification to document structures that might not be obvious by light microscopy. Specimens that have been processed for FISH can also be simply air-dried for CLEM, and if fine, ultrastructural detail is not important.

22.

Specimens processed for FISH (Subheading 3.3) subsequently can be imaged by SEM [20]. Once light microscopy or confocal images have been obtained, the specimens are carefully removed from the glass slide or coverslip. The fully hydrated specimens then are processed by fixation in buffered glutaraldehyde (2.5 %), dehydrated, and critical-point dried. When mounted onto the specimen stub, the orientation is noted so that the same fields examined by light microscopy are imaged in the SEM. This approach to correlative light and electron microscopy (CLEM) is not common, but offers an opportunity for imaging large specimens where the fluorescent signal is overlaid on an SEM image of the intact specimens (or reference space).

23.

Fix specimens immediately after collection and avoid drying. If specimens are to be used for immunolabeling, then fixation in phosphate-buffered formaldehyde (4 %, v/v, pH 7.2) alone may be more useful. However, specimens should then be embedded in acrylic resins (e.g., Lowicryl HM20 or LR White). Dehydration and resin infiltration times can be reduced with the assistance of a microwave processor [52]. Microwave processing is useful for embedding in epoxy resins and LR White resin. Semi-thin sections of embedded biofilms mounted on glass substrate can be immunolabeled for light microscopy examination, and thin sections of embedded biofilms labeled for TEM examination.

24.

The protocol has been written to give regular contrast to the specimens and can be modified in many ways to affect contrast. Some suggested changes are described below.

25.

Different fixatives will affect the final appearance of the specimens. Substituting sodium cacodylate (100 mM, pH 7.2) for the phosphate buffer will produce less extraction of cell cytoplasm and thus result in decreased contrast in the TEM. The use of Good’s buffers (TRIS, HEPES, PIPES, etc.) [53] will result in decreased extraction and thus reduced contrast.

26.

Shorter fixation times, or the use of selective detergents, also can be used to manipulate retention of cellular contents and thus affect contrast.

27.

Shorten the postfixation step in osmium tetroxide. In step 4 (Subheading 3.5), incubate for 3–4 h in 1 % osmium tetroxide. Jump to step 7.

28.

Steps 9–11 (Subheading 3.5): Omit the sodium maleate buffer and use aqueous uranyl acetate for slightly less contrast. Alternatively, leave the specimens overnight at 4 °C in saturated uranyl acetate in 70 % methanol for more contrast.

29.

Step 12 (Subheading 3.5): Instead of using ethanol, dehydrate in graded acetone series. The propylene oxide steps then can be omitted, substituting acetone for the propylene oxide. If acetone is used, ensure it is completely removed from the resin before polymerization starts.

30.

The incubation times given here are only suggested guidelines. They can be changed to fit different protocols and specimen types. Small pellets of cells will require less incubation at each step than will tissues with tightly packed cells such as nerve or muscle.

31.

If at any time the protocol needs to be halted and continued the next day, it is possible to store the samples at any of the washing steps. Place the samples in the appropriate washing solution and store it at 4 °C.

32.

Reynold’s lead citrate [54]: Dissolve lead nitrate (1.33 g) in 30 mL of deionized H2O in a 50 mL volumetric flask. Add sodium citrate (1.76 g) and mix. Shake the suspension for 1 min and then leave to stand for 30 min with intermittent shaking. The lead nitrate is being converted to lead citrate during this time. Add NaOH solution (4 g in 100 mL, 1 N, 8 mL) and mix well. Filter before use.

33.

Venable and Coggeshall’s lead citrate [55]: Weigh out portions of lead citrate in 10 mL tubes. The amounts can vary between 0.1 and 0.4 g, with the larger amounts producing stronger staining. When needed for staining, add 1 mL of 1 N NaOH to a tube containing the aliquot of lead citrate and dissolve the solid. Then add 9 mL of deionized H2O, filter, and use. Use carbonate-free NaOH and fresh deionized H2O or H2O that has been boiled to remove dissolved CO2.

34.

Staining protocol for lead citrate: Prepare CO2-free, deionized H2O by boiling and cooling H2O or by sparging nitrogen gas through the H2O. Place drops of the lead citrate stain on clean Parafilm. Float grids, section-side down, on the drops of stain (one grid per drop). Incubate for recommended, or optimal, time. Fill four 10 mL beakers with CO2-free deionized H2O; in the first, add one drop of 1 N NaOH solution. Remove each grid from the drop of stain and wash by rapidly immersing, sequentially, in the four beakers of deionized H2O. Start with the beaker containing H2O with 1 drop of NaOH. Dry and examine in the TEM.

35.

If the vaginal biofilms under study are on the surface of a medical device (e.g., intravaginal ring, intrauterine device) and can be embedded in epoxy resin to produce thin sections for examination in the TEM, they should produce high-quality images. However, the epoxy resin will most likely not infiltrate the medical device, causing difficulties in sectioning the polymer at ambient temperature. A convenient alternative approach is to remove the embedded biofilm from the medical device substrate, mark the interface on the device (e.g., using nail polish), and replace the device with fresh resin that is subsequently polymerized. The embedded biofilm can be sectioned easily using routine methods. If the medical device is required to be in the section, remove as much of the material as possible, leaving the biofilm-device interface intact, and either reembed in fresh resin or attempt to section the interface. Soft polymers can be successfully sectioned at low temperatures using a cryoultramicrotome.

36.

The PowerBiofilm™ DNA Isolation Kit (MO BIO Laboratories, Inc.) can accommodate the low sample masses (50–200 mg) typically available from vaginal specimens.

37.

The above DNA isolation kit embodies three important features relevant to vaginal microbial biofilms: Mechanical shearing breaks up the cross-linked biofilm, heat-activated enzymatic digestion dissolves most of the polysaccharides, and the protein/inhibitor removal step usually affords high-quality genomic microbial DNA.

38.

The removal of PCR inhibitors sometimes may require a different DNA isolation strategy. Under these circumstances, the InstaGene Matrix (Bio-Rad Laboratories, USA) kit is recommended.

39.

A discussion of the methods for analyzing DNA and RNA isolated from the vaginal specimens is beyond the scope of this chapter. The first report of culture-independent characterization of vaginal microbial communities dates to 2004 [56]. A number of subsequent studies have surveyed the vaginal microbiome using culture-independent approaches to census bacterial community composition, usually to improve our understanding of the etiology underlying bacterial vaginosis (BV) [11, 57-62]. Analyzing the metatranscriptomes of vaginal microbial communities is more challenging and not as well developed as methods for determining community composition and structure. McNulty and colleagues have described methods for microbial RNA-Seq analysis of the fecal metatranscriptomes from gnotobiotic mice and humans [63]. Total RNA was purified and converted into the corresponding barcode-ligated double-stranded cDNA (dscDNA) [63, 64]. Multiplex sequencing was performed using the Illumina platform according to published protocols [63].

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