ELECTIVES > MICROBIAL METHODS

The usual methods of microbiology were developed to study microbes affecting human health (Lynch and Hobbie 1988). When these methods are used for field studies they have many problems; to begin with, less than one percent of the bacteria found in a sample of ocean water will grow on laboratory petri plates. Recent methodological advances, such as the application of fluorescently-labeled immunochemical or nucleic acid probes, provide a new way to taxonomically identify individual cells in the mixed populations of soils and waters. Yet, the application of these methods is limited to the detection of physiologically active cells with a large number of ribosomes, a rare occurrence in nature. Despite these and other problems, ecological microbiologists have developed a number of methods for enhancing our ability to measure the activities, abundances, and types of microbes found in soils, waters, and sediments of natural systems (e.g., the methods book of Kemp et al. 1993).

In this course, we will present the scientific rationale behind a number of methods suitable for determining the role of microbes in ecosystems. The methods will be described and students will carry out the procedures in a series of hands-on laboratories. One theme will be choosing methods and temporal and spatial scales appropriate to the question being asked (e.g., the review by Paerl and Pinckney 1996). The course will be taught, under the leadership of Joseph Vallino, in collaboration with scientists from the Ecosystems Center, the permanent staff of the Marine Biological Laboratory, and other Woods Hole institutions.

Lectures will describe the biology and logic that underlie the various methods used by microbial ecologists. In the laboratory, students will work with the latest techniques to measure microbial biomass, activity, extracellular enzymes, biogeochemistry and species diversity. These include epifluorescence microscopy, radioisotopic tracers for bacterial production, fluorescent substrates, hydrogen sulfide and methane production, and molecular probes for classes of bacteria. Students will be required to turn in answers and calculations to problem sets that are associated with each lab topic. These problem sets will form the basis of the grade. In addition, students will be asked to present their laboratory results and participate in group discussions on the findings by the entire class. Students will be encouraged to use the methods developed in this course for their individual research projects associated with the SES course.

Syllabus:

Session 1: Introduction

Session 2: Construct Winogradsky column.
Field trip to Little Sippewisset Marsh

Session 3: Bacterial Abundance

Session 4: Prepare dilution plates, Fix samples for direct counts

 Session 5: DAPI staining and counts; examine plate

Session 6: Bacterial production. Lecture on method; count dilution plates

Session 7: Measure bacterial production

Session 8: Measure filter 14C activity, Scintillation counter demonstration, Explain calculations, Chemolithotrophy

Session 9: Lecture on Winogradsky column

Session 10: Measure methane and redox gradients

Session 11: Measure hydrogen sulfide gradient

Session 12: Extracellular Enzyme Assays

Session 13: Lecture on extracellular enzymes. Fluorometry

Session 14: Measure enzyme activities

Session 15: Microbial food webs: flagellate and ciliate grazing on bacterial

Session 16: Lecture and demonstration

Session 17: Lecture and demonstration

Session 18: Microbial food webs: bacteria phytoplankton competition

Session 19: Lecture. Microcosm startup and sample

Session 20: Sample microcosm

Session 21: Sample microcosm

Session 22: Sample microcosm

Session 23: Sample microcosm

Session 24: Sample microcosm

Session 25: Sample microcosm

Session 26: Sample microcosm

Session 27:  Process microcosm samples. Molecular techniques. Bahr

Session 28:  PCR

Session 29:  Electrophoresis

 

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