Handbook of Methods in Aquatic Microbial Ecology
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Handbook of Methods in Aquatic Microbial Ecology

Paul F. Kemp, Jonathan J. Cole, Barry F. Sherr, Evelyn B. Sherr, Paul F. Kemp, Jonathan J. Cole, Barry F. Sherr, Evelyn B. Sherr

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eBook - ePub

Handbook of Methods in Aquatic Microbial Ecology

Paul F. Kemp, Jonathan J. Cole, Barry F. Sherr, Evelyn B. Sherr, Paul F. Kemp, Jonathan J. Cole, Barry F. Sherr, Evelyn B. Sherr

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About This Book

Handbook of Methods in Aquatic Microbial Ecology is the first comprehensive compilation of 85 fundamental methods in modern aquatic microbial ecology. Each method is presented in a detailed, step-by-step format that allows readers to adopt new methods with little difficulty. The methods represent the state of the art, and many have become standard procedures in microbial research and environmental assessment. The book also presents practical advice on how to apply the methods. It will be an indispensable reference for marine and freshwater research laboratories, environmental assessment laboratories, and industrial research labs concerned with microbial measurements in water.

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Publisher
CRC Press
Year
2018
ISBN
9781351442367
Section IV
Activity, Respiration, and Growth
CHAPTER 45
Microautoradiographic Detection of Microbial Activity
Kevin R. Carman
INTRODUCTION
Brock and Brock1 introduced microautoradiography (MA) as a tool for the study of activities by individual aquatic microorganisms from natural samples. The technique has subsequently been used to address a variety of ecological questions regarding bacterial activities in natural aquatic systems.2,3,4,5 In particular, MA can be used in radiotracer studies to determine the proportion of microorganisms in a sample that is metabolizing a given radiolabeled compound. MA is also useful for determining labeling specificity, i.e., whether targeted microorganisms take up labeled substrates at the exclusion of nontargel microorganisms.6,7 MA has also been used to check assumptions of radiotracer-grazing techniques, i. e., whether label is taken up by grazers via consumption of labeled microbes or as a consequence of activity by microorganisms associated with the grazers.8
The MA methods described here are based primarily on the “MARGE-E” procedure described by Tabor and Neihof,9 and secondarily on suggestions from Rogers10 and Meyer-Reil.3 These procedures are designed for use in radiotracer studies that involve the use of compounds labeled with 3H or 14C. With relatively minor modifications (e.g., the use of different emulsions), the basic methods described here should be applicable for studies involving other β-emitting isotopes (e.g., 32P, 33P and 35S). Readers who wish to further investigate practical and theoretical aspects of autoradiography are referred to Rogers‘10 excellent book.
MATERIALS REQUIRED
Sample Preparation
Equipment
• Centrifuge
• 0.2-μm filter-sterilization device
• Waring blender: semi-micro blender container (Eberbach, Ann Arbor, MI)
Supplies
• 50-ml centrifuge tubes
• 5-ml pipettes
Solutions
• 0.01% sodium pyrophosphate solution, filter sterilized (0.2 μm) (in deionized water or appropriate saline solution)
• 3% glutaraldehyde solution in 0.1 M cacodylate buffer
• Sterile, 15-ml, screw-cap test tube or comparable sample containers
• 25 mm, 0.2-μm membrane filters, prestained with Irgalan black (Poretics Corporation, Livermore, CA or Nuclepore Corporation, Pleasanton, CA)
Microautoradiography
Equipment
• Refrigerator set at 4°C
• Safelight with 30-W bulb and Kodak #2 filter
• Fool switch (to control safelight)
• Water bath (to maintain temperature at 43°C)
• Epifluorescence microscope
• Filtration apparatus for 25-mm filters
Supplies
• 10-μl Hamilton syringe
• 25-mm and 0.2-μm membrane filters
• Membrane-filter forceps
• Microscope slides with frosted end
• Autoradiography slide boxes (RPI, Mount Prospect, IL; standard plastic slide boxes may also be used)
• 30-ml beaker (used as a dipping container for emulsion)
• 100-ml beaker (in which 30-ml dipping container is placed)
• Kimwipes (or equivalent)
• Cold metal plate (chilled in freezer compartment of standard refrigerator)
• Paper towels
• Silica-gel desiccant
• Desiccator (large enough to hold slide boxes)
• 60-ml, plastic, screw-cap bottles
• Aluminum foil
• Coplin jars
Solutions
• Chromic acid (dissolve 100 g K2Cr2O7 in 850 ml deionized water, add 100 ml H2SO4)
• Deionized water
• Glycerin
• Autoradiographic emulsion (e.g., NTB-2, Kodak, Rochester, NY)
• Dektol developer (Kodak; 1:2, stock solution:deionized water)
• Kodak fixer solution (NOT Rapid fixer), make up in deionized water
• Glycerin solution, 1% in deionized water
Low-fluorescence immersion oil
• Acridine orange, 0.04% in pH 6.6 citrate buffer
• Citrate buffer, 0.004 M, pH 6.6, pH 5.0 and pH 4.0
Citrate buffer recipe:
1. Make 0.1 M stock solutions of citric acid and sodium citrate in 25% methanol (MeOH; prevents microbial growth in stock solution):
citric acid: 2.10 g/100 ml 25% MeOH
sodium citrate: 2.94 g/100 ml 25% MeOH
2. Mix 20 ml of stock solutions in the following ratios to achieve the appropriate pH:
pH
ml citric acid
ml sodium citrate
4.0
13
7
5.0
8
12
6.6
1.5
18.5
3. For working solution, dilute the 20 ml of mixed stock solution with 480 ml deionized water. Final pH can be adjusted by adding small amounts of citric acid or sodium citrate.
PROCEDURES
Sample Preparation
If the study involves the use of plankton samples, microorganisms can be concentrated onto membrane filters of appropriate pore size (e.g., 0.2 μm for bacteria, 1 to 5 μm for eukaryotic algae) using standard filtration procedures. For studies involving sedimentary microorganisms, it is recommended that microorganisms be separated from sediments before they are concentrated on filters. The procedure described below for separating microbes from sediments is derived from Balkwill et al.11 It is suitable for a 1-cm3 sediment sample. If other sediment volumes are used, appropriate adjustments to solution volumes should be employed:
1. Place 1-cm3 sediment in sterile test tube, fix in 6 ml of 3% glutaraldehyde solution. Store at 4°C until use.
2. Transfer entire sample from test tube to a clean, sterile, semi-micro blender container.
3. Add 24 ml of 0.2-μm-filtered (FS), 0.01% sodium pyrophosphate solution to the blender container.
4. Blend sample for 1 minute on, 30 s off, 30 s on, 30 s off, and 30 s on.
5. Transfer blender contents to a clean, sterile 50-ml centrifuge tube.
6. Pelletize sediment and cells by centrifugation at 2350 × g for 10 min.
7. Taking care not to disturb pellet, remove and discard 25 ml of supernatant using pipette.
8. Add 5 ml of FS 0.01% sodium pyrophosphate solution to centrifuge tube, vortex tube for 30 s.
9. Centrifuge at 650 × g for 5 min. This pelletizes sediment but leaves cells in suspension.
10. Decant supernatant into a separate clean, sterile 50-ml centrifuge tube.
11. Add 5 ml 0.01% sodium pyrophosphate solution to tube containing sediment pellet, vortex 30 s.
12. Centrifuge at 650 × g for 5 min.
13. Repeat steps 10 through 12 until 25 ml of supernatant has been collected.
14. Stain an aliquot of the supernatant with acridine orange (or DAPI) and filter onto a 0.2-μm membrane filter prestained with Irgalan black. Determine supernatant volume required to obtain 50 to 100 bacteria per field of view when viewed with a 100 × oil-immersion objective. If MA is to be performed on microalgae, microalgal abundance may be determined using membrane filters of a larger pore size (1 to 5 μm).
Microautoradiography
Microautoradiography is performed on microbes that have been concentrated on membrane filters. Briefly, slides are coated with emulsion and the filter is placed face down on the liquid emulsion. The emulsion is then hardened by cooling and drying. After exposure, development, staining, and redrying, filters are peeled from the emulsion, leaving the cells embedded in the emulsion. Cells and developed silver grains can then be detected with fluorescence and bright-field microscopy. Experience in our lab has been that slides are most conveniently processed in batches of 10. Several batches may be processed in a single day.
Advance Preparations
1. Soak microscope slides and 30-ml beaker in chromic acid overnight, then rinse thoroughly with FS deionized water. Store acid-cleaned slides and glassware so that they will not be exposed to dust.
2. Label (sample #) frosted end of microscope slide with indelible ink.
3. Using a 10-μl syringe, place a 1-μl drop of glycerin on a corner of the frosted end of the slide.
4. Filter an aliquot of the sample supernatant onto a 25-mm, 0.2-μm membrane filter. Rinse twice with deionized water to remove unincorporated radioactivity.
5. Attach filter (filtered side up) to corner of slide by placing an edge of the filter on the glycerin drop. This minimizes the chance that filters will become disassociated from labeled slides (leading to confusion over sample i.d.s), and greatly facilitates handling of slides and filters in the darkroom.
Sample Processing
Darkroom preliminaries:
1. Place emulsion in 43°C water bath. Emulsion should liquify in approximately 1 h. This can be performed in the light if lid of the emulsion container is not removed. Each of the following steps should be performed in total darkness.
2. Dilute emulsion with distilled water, two parts emulsion to one part water (this dilution yields an emulsion layer 1 to 2 μm thick).10
3. Dispense 20-ml aliquots of diluted emulsion into 60-ml screw-cap plastic bottles. Place bottles in a film-develop...

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