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With the exception of Objective #1 (Biogeographic aspects of land-applied wastes), we have extensive experience with all the soil and rhizosphere methods proposed in the Methods section for Objectives #2 and #3 (e.g., BIOLOG: Oka et al., 2000; glomalin production: Wright and Upadhyaya, 1997; mycorrhizae: Douds and Millner, 1999; organic carbon: Franzluebbers et al., 1999; phytoremediation: Olexa et al., 2000; whole soil FAME: Cattelan et al., 1999). For this reason, the procedures for Objective #1 are given in detail, whereas the procedures for Objectives #2 and #3 are given only generally. Similarly, the experiments outlined for Objective #1 are given for the entire 5-year lifespan of the multistate proposal, whereas the experiments for Objectives #2 and #3 are given in groups.

Biogeographic aspects of land-applied wastes

A. Specific experiments and collaborations

Two experiments are proposed. In the first experiment, 500 isolates of E. coli will be obtained from humans and a variety of agricultural and wild animals common to the participating states (AL, DE, GA, PR, TN, TX, WI, USDA-GA, USDA-ID). The number of isolates obtained will be: humans (100), cattle (50), deer (50), dog (50), duck or goose (50), poultry (50), sheep (50), and swine (50). The remaining 50 isolates will be allocated to an animal species that each investigator believes to be important in that location (e.g., elk in Idaho). To maximize ribotype diversity, each human or animal will contribute no more than 5 isolates. For example, 12 different cattle/dairy operations will be visited in order to obtain the necessary 60 cattle E. coli isolates. This experiment will not only form the basis of a national library of E. coli ribotypes, but also will provide information on the biogeographic distribution of the ribotypes. Because of the cost, isolates will initially be ribotyped at one location (GA). As extramural funding becomes available, isolates will also be ribotyped at two other locations (USDA-GA, USDA-ID).

In the second experiment, the temporal variability of E. coli ribotypes in three different agricultural animal species, cattle, swine, and poultry will be tracked. Specific animals of each species will be identified and at least 150 isolates of E. coli will be isolated from at least 5 animals (30 isolates per animal) periodically, depending on the normal agricultural lifespan of on the animal. In the case of cattle, steers will be sampled every two months for one year, after which the cattle will be replaced with another batch of yearling calves. In the case of swine, animals will sampled biweekly for 12 weeks, and in the case of poultry, broilers will be sampled biweekly for 6 weeks. In this experiment, only the locations actively ribotyping isolates will participate in the experiment (DE, GA, USDA-GA, USDA-ID).

B. Methods

Fresh fecal samples will be obtained as aseptically as possible. With the exception of humans, fresh fecal samples will be collected with an ethanol flame-sterilized spatula and will be transferred to sterile Whirl-Pak bags. Collection information will consist of the animal species, person collecting, and the date and place collected. The bags will be kept on ice until the samples are processed. Because of the restrictions on isolating bacteria directly from humans, human isolates of E. coli will be collected from septic tanks. These septic tanks will be from homes without any outside sources of E. coli (e.g., dogs). Septic tanks will be sampled with sterile dilution bottles and will be treated in the same manner as the fecal samples.

To isolate E. coli from fecal samples, a 20-mL sample of 0.1% peptone will be added to the Whirl-Pak bag containing the feces, and the feces will be blended for 30 seconds with a Stomacher blender. The septic tanks samples will be shaken by hand for 1 minute. A 10-µL sample from the Whirlpak bag or dilution bottle will be streaked onto mTEC agar plates. The plates will be wrapped in quadruple Ziploc bags and will be incubated submerged in a water bath at 44.5± 0.2°C for 24 h. All streakings will be done in duplicate. Yellow colonies growing on mTEC agar (Difco) after a 24-h incubation will be considered as presumptive E. coli.

Presumptive isolates of E. coli will be transferred onto tryptic soy agar (TSA) and will be incubated overnight at 35°C. This step will be repeated to ensure pure cultures. After two streakings, each isolate will be streaked a third time onto TSA as well as on urea agar and Simmons citrate agar. In addition to the E. coli isolates, type cultures from the American Type Culture Collection will be used as appropriate controls. These organisms represent almost all the bacteria that can be found on mTEC agar plates. The type cultures include Escherichia coli #11775 (urease, citrate), Klebsiella pneumoniae subspecies pneumoniae #13883 (urease+, citrate+), Citrobacter freundii #8090 (urease+, citrate+), and Enterobacter aerogenes #13048 (urease, citrate+). After overnight incubation at 35°C, presumptive E. coli isolates that can grow on TSA but are urease and citrate will be subjected to an oxidase test. If the isolate is also oxidase, then the isolate will be considered as confirmed E. coli and will be frozen. All other isolates will be autoclaved and discarded. To freeze each E. coli isolate, a loopful of each isolate will be transferred from the third streaking of the TSA plate into two labeled cryovials, each containing a 3.5:1 mixture of saline/phosphate buffer and cryoprotectant. The two cryovials, one representing the working stock and the other the reserve stock, will be kept in separate -80°C freezers.

Confirmed isolates of E. coli will be streaked from the -80°C frozen stocks onto TSA and incubated overnight at 35°C. A single clone will be inoculated into 10 mL of Luria-Bertani broth (Maniatis et al., 1982) and incubated on a rotary shaker at 100 rpm overnight at 35°C. The DNA from a turbid, 1-mL sample will be obtained with a commercial kit (Qiagen DNeasy tissue kit). The DNA will be quantified with a UV spectrophotometer at 260 nm (DNA) and 280 nm (protein). Samples with an acceptable 260:280 ratio (i.e., >1.75) will be used for ribotyping.

A digoxigenin (DIG)-labeled probe will be prepared to yield a reverse transcribed, DIG-labeled probe of E. coli 5S, 16S, and 23S rRNA. The DIG label will be quantified against a kit standard (Roche). To perform a restriction digest of the genomic DNA, a sample of DNA will be added to each of two microfuge tubes and each brought to a specific volume with distilled water. A sample of EcoRI or PvuII with the appropriate restriction enzyme buffer will be added and the mixture will be incubated at 37˚C.

After overnight incubation, loading dye will be added to each tube of restricted DNA. A portion of the DNA will be added to each well of a 0.7% agarose gel. Additional wells will be set aside for DNA ladders of -DNA cleaved with EcoRI and HindIII (molecular weight marker), no DNA control, and DNA from type culture E. coli ATCC #11775. The gel will be submerged in 1X Tris acetate EDTA buffer and electrophoresed at 55 volts for approximately 3 h.

Once the DNA separation is complete, the gel will be placed on a Nytran nylon membrane contained in a vacuum blotting assembly (VacuGene blotting assembly, ). The gel will be sequentially washed with denaturing solution, neutralizing buffer, and 20X transfer buffer. After transfer, the gel will be discarded and the membrane washed in 2X transfer buffer before the DNA on the membrane is fixed with UVirradiation.

The membrane will be hybridized with preheated DIG-labeled probe overnight at 42°C. The membrane will be washed in a series of stringency washes before equilibrating in washing buffer for 1 min. The membrane will be incubated in blocking solution for 60 minutes at room temperature. The blocking solution will be discarded and a new batch of blocking solution containing anti-DIG-alkaline phosphatase will be added. After 30 minutes incubation at room temperature, the membrane will be washed twice for 15 min, and detection buffer will be added for 2 min. The membrane will be removed from detection buffer and will be treated with a chemiluminescent substrate. The chemiluminescence will be quantified with an Alpha Innotech FluorChem 8000 imager and the image saved as a TIFF file.

All banding patterns contained in the image files will be sent to one location (GA) to be quantified with gel analysis software (GelCompar II). The host origin, temporal, and geographic relationships among the isolates will be examined by cluster analysis, and cluster dendrograms will be plotted with the same gel analysis software. Ribotype patterns with respect to geography, time, and host origin will be made publicly available on the multistate project website, <http://eclass.ifas.ufl.edu/dmsa/msp>.

Rhizosphere-enhanced biotreatment of organic contaminants

A. Specific experiments and collaborations

Three groups of experiments are proposed. In the first group of experiments, collaborative lightroom studies with comparable protocols will determine the effect of interactions associated with different plant species, soils, and nutrients on remediation of several unlabelled organic compounds (e.g., PAHs) in soil and rhizosphere. The experiments will incorporate a) a model system of specific organic contaminants, b) whole soil FAME or PLFA (phospholipid fatty acid) and DNA-based techniques to describe changes in microbial community structure, c) BIOLOG plates to assess functional diversity, d) staining techniques to evaluate mycorrhizal colonization, and e) gas chromatography (GC) to determine the concentrations of the remaining parent compound. In this manner, it may be possible to predict the general conditions under which the rhizosphere will stimulate biodegradation of an organic contaminant (AL, AR, DE, FL, NC, SC, WI, Other Coop.-NH, Other Coop.-FL, Other-Coop.-Canada).

In the second group of laboratory experiments, collaborative lightroom studies with comparable protocols will be conducted with labeled organic compounds. These compounds will be selected on the basis of their different degradative patterns from the set of organic compounds in the first group of experiments. The second group of experiments will determine the effect of rhizosphere processes on contaminant bioavailability, with a specific emphasis on plant- or microbially produced biosurfactants. By following the fate of 14C-labelled organic compounds, the degree to which toxic organics may be degraded to intermediates covalently bound to the humic soil fraction (i.e., not to CO2 and H2O) will be studied. The experiments will be conducted with and without plants to assess the role of plants in bioavailability (AL, AR, DE, FL, NC, SC, WI, Other Coop.-NH, Other Coop.-FL, Other-Coop.-Canada).

In the third group of experiments, the results of the second group of experiments will be conducted under field conditions with unlabelled compounds to determine the rhizosphere effects on remediation and bioavailability of organic contaminants in aged and newly contaminated soils (AL, AR, DE, FL, NC, SC, WI, Other Coop.-NH, Other Coop.-FL, Other-Coop.-Canada).

B. Methods

Soils will be amended with toxic organic compounds. Plant species, including grasses and legumes, will be evaluated for germination percentage, growth rate, and root development in these soils under greenhouse, growth chamber, or lightroom conditions. Following plant selection, cooperative greenhouse studies will determine bioremediation in the rhizosphere. The soil will be amended with appropriate amounts of inorganic N, P, and K as indicated by soil test analysis. At each sampling time, pots will be harvested and the quantity of contaminant and degradation products in the plant top and roots and remaining in the rhizosphere and non-rhizosphere soil will be determined. Plant dry matter production and rooting characteristics will be quantified (Bohm, 1979). The microbial population (i.e., bacteria, fungi, degraders) in the rhizosphere and non-rhizosphere (i.e., R/S ratio), will be evaluated at selected sample times. At each sampling time, isolates will be cultured or microbial community structure will be characterized or both.

The soils will be characterized for physical, chemical, and biological properties by standard methods. The physical properties will be texture and water content at -0.03 MPa, the chemical properties will be pH, NH4-N, NO3-N, total N, % organic C, extractable P, K, Ca, Mg, Na, electrical conductivity, and chloride, and the biological properties will be total viable bacteria and fungi.

Other biological properties will be individual and whole soil FAME analyses. Phospholipid fatty acid (PLFA) analyses will be used in instances when the organic contaminant interferes with the whole soil FAME analyses. Ecotypic variations among arbuscular mycorrhizae will be evaluated with a nested PCR approach (Kjøller and Rosendahl, 2000).

Contaminant concentrations will be determined with GC and gas chromatography–mass spectroscopy (GC–MS). Soils will be initially extracted with an appropriate organic solvent using modified EPA methods.

Disturbed lands and urban landscapes

A. Specific experiments and collaborations

Two groups of experiments will be conducted. In the first group of experiments, collaborative studies with comparable protocols will characterize taxonomic and functional diversity of bacteria and mycorrhizae in disturbed soils (e.g., road construction). Lands will be selected with a range of ages and varying amounts of disturbance and revegetation (e.g., coastal bermudagrass to trees) and will be compared to undisturbed sites (DE, MD, PR, TN, TX, WV, USDA-GA, USDA-ID).

In the second group of experiments, collaborative studies with comparable protocols will characterize taxonomic and functional diversity of bacteria and mycorrhizae in urban landscapes (e.g., intensively managed turfgrass systems like golf courses). Experiments will include different grass species. For example, in the case of putting greens, samples from the sand-based root zones will be analyzed for bacterial numbers, BIOLOG carbon-source utilization profiles, whole-soil FAME profiles, water-extractable organic carbon, mycorrhizal infection and possibly quantification of glomalin. If sufficient arbuscular mycorrhizal spores are collected, then they will be submitted to the University of West Virginia for classification. Soils will be sampled periodically during the year to capture seasonal effects on community structure (AL, DE, FL, NC, NE, SC, TX, WV).

B. Methods

Soils from the collaborators will be analyzed for functional diversity (i.e., community level physiological profiles or CLPP) using the commercially available BIOLOG system. This system has been widely adapted for these studies. Soil samples will be collected fresh from the field and refrigerated until analyzed by the BIOLOG system. Ideally, soils will be analyzed immediately upon return to the lab. Appropriate dilutions (i.e., 10-3) will be inoculated into BIOLOG GN plates. The plates will be incubated for 24 to 72 h during which time they will be analyzed with a plate reader to record well-color development. Data from the BIOLOG plates will be analyzed by principal component analysis and other methods to determine the CLPP of the microbial populations.

Taxonomic diversity of the soil microbial communities will be assessed with whole soil FAME profiles. Selected fatty acids will be used as biomarkers for specific groups of microorganisms. The FAME profiles will be analyzed with the software program CANOCO to determine the similarities in microbial communities.

Chemical and physical analysis of the soils will be conducted at Texas A&M University. Microbiological analyses will include enumeration by standard plate count methods, analysis of soil microbial biomass with chloroform-fumigation extraction methods (Brookes et al., 1985), and analysis of microbial communities using the BIOLOG and FAME analyses described above.

Soil samples will be collected, dried in air, and stored at –80oC until they can be extracted for FAME analyses. Extraction will be done under standard protocols developed in our previous regional project research (Franzluebbers et al., 1999) with the exception that prior to the last step of resuspending the extracts in the ether solvent, the GC vials will be sealed and capped under a stream of nitrogen. This eliminates the hazards of mailing flammable solvents. We have shown in earlier research that this method does not adversely affect our ability to determine FAME profiles (Peach et al., 1999). The extracts will be sent to University of Delaware for analysis with the MIDI system (MIDI, Newark, DE).

With regards to mycorrhizae, standard methods (e.g., Sylvia, 1994) will be used to quantify propagules, estimate root colonization, and detect active hyphal lengths of hyphae in soil. Root samples will be collected from vegetation (primarily coastal bermudagrass) at the sampling sites and will be sent to the University of Florida for analysis. The effect of arbuscular mycorrhizae on interplant competition and plant aggressivity will be determined with replacement and additive designs within mesocosm field plots (Snaydon, 1991). Glomalin assays will be conducted to separate the activity of mycorrhizal hyphae from other root-inhabiting fungi. As a specific and persistent glycoprotein (Wright and Upadhyaya, 1997), glomalin will be quantified with an immunofluorescence assay. Detailed methods are available online at <da.gov/nri/smsl/glomalin.htm>.


Biogeographic aspects of land-applied wastes

The proposed research will:

  • identify the degree of geographic variability of E. coli ribotypes among humans, cattle, deer, dog, Canada goose, poultry (broilers), domestic sheep, and swine.

  • identify the degree of temporal variability of E. coli ribotypes in cattle, swine, and poultry (broilers).

  • generate a publicly accessible database of ribotypes of E. coli on the worldwide web for use by resource managers.

Rhizosphere-enhanced biotreatment of organic contaminants

The proposed research will:

  • provide a database of biodegradative strains consisting of organic contaminant degraders isolated from contaminated soils from a diversity of sites across the United States. Such a database will be publicly available on our website and will be similar to the Canadian database at <ak.ca.departments/scsr/department/research/index/html>.

  • create a culture collection of organic contaminant degraders available to researchers upon request.

  • provide guidelines for calculating nutrient amendments for phytoremediating hydrocarbon-contaminated soils.

  • provide an improved knowledge of pollutant bioavailability.

Disturbed lands and urban landscapes

The proposed research will:

  • provide knowledge regarding the effects of disturbed lands and urban landscapes on soil microbial taxonomic and functional diversity.

  • enable land managers to adopt practices that conserve or protect the taxonomic and functional diversity of soil microbes.

  • produce knowledge needed by managers of intensively maintained agroecosystems to make science-based decisions on efficacy of alternative management inputs (e.g., microbial formulations).



A project chairperson and secretary will be elected at the first meeting of the technical committee. The secretary is automatically the project chairperson-elect for the next year. Therefore, only a new secretary is elected annually after the first year. All voting members of the technical committee are eligible for office, regardless of their affiliation or sponsoring agency. Each of the three multistate objectives will have a chairperson. In addition, there will be two chairpersons, one for the next edition of our textbook and one for the publication of an accompanying laboratory manual. (Because publications may not be listed as objectives, they are not listed in the body of this proposal.) These chairpersons have already been chosen for the multistate project and will be the chairperson for that objective for the entire 5-year-life of the multistate project. However, chairpersons for the objective may be replaced on a vote of the technical committee at the annual meeting. Finally, the technical committee will elect a host chairperson for the annual meeting the next year and designate a general site for the annual meeting.


The project chairperson, in consultation with the administrative advisor, will: a) notify all technical committee members of the time and place for the formal and informal annual meetings (see below), b) prepare a meeting agenda, c) preside at the meetings, and d) prepare an annual and other reports for the multistate project. The last chairperson of the 5-year project will also prepare the final termination report. The chairperson will distribute the annual report to the technical committee members. The chairperson will also consult with the administrative advisor in the conduct of his or her duties.

The chairpersons for each objective will: a) coordinate research within each objective, b) report on objective progress at the annual meeting, and c) prepare publication of results. Therefore, each chairperson is responsible for achieving the overall objective goal.

The secretary will: a) record the minutes of the annual meeting, and b) prepare and distribute copies of the minutes to all members. In addition, the secretary will perform any other duties as needed by the project chairperson.

The host chairperson will make all arrangements for the technical committee at the formal designated annual meeting site (e.g., lodging). The host chairperson is under the direction of the project chairperson.


There will be two meetings per year, one formal meeting at a designated site and another informal meeting at the Soil Science Society of America annual meeting. The formal meeting will be primarily to discuss the three multistate objectives, whereas the Soil Science Society of America annual meeting will be primarily to discuss the two special educational objectives.


The technical committee will maintain a website of the multistate project. Such a website will contain: a) the multistate project with all attachments (if approved), b) minutes of all meetings, c) annual reports, d) progress on the two objectives, and e) hyperlinks to all the technical committee members for further information. The URL for the website is <http://eclass.ifas.ufl.edu/dmsa/msp> and will be maintained by D. Sylvia at the University of Florida. The website is now open.

Organizational chart



Title: Soil Microbial Taxonomic and Functional Diversity as Affected by Land Use and Management

Principal Leader



Area of Specialization

Y. Feng


Auburn University

Soil Microbiology

D. C. Wolf


University of Arkansas

Soil Microbiology

J. J. Fuhrmann


University of Delaware

Soil Microbiology

D. M. Sylvia


University of Florida

Mycorrhizal Ecology

J. Graham


University of Florida

Mycorrhizal Agroecology

K. Jayachandran


Florida International University

Soil Microbiology

A. Franzluebbers


USDA–ARS (Watkinsville)

Soil Microbiology

P. G. Hartel


University of Georgia

Soil Microbiology

M. Jenkins


USDA–ARS (Watkinsville)

Soil Microbiology

J. Entry


USDA–ARS (Kimberly)

Soil Microbiology

J. S. Angle


University of Maryland

Soil Microbiology

R. Klucas


University of Nebraska

Plant Pathology

A. G. Wollum


North Carolina State University

Soil Microbiology

C. M. Reynolds


U. S. Army/CRREL

Soil Microbiology

E. C. Schröder


University of Puerto Rico

Soil Microbiology

H. D. Skipper


Clemson University

Soil Microbiology

M. D. Mullen


Univ. of Tennessee

Soil Microbiology

D. A. Zuberer


Texas A&M University

Soil Microbiology

K. Hatzios


Virginia Polytechnic Institute & State University

Administrative Advisor

J. B. Morton


West Virginia University

Environmental Microbiology

W. J. Hickey


University of Wisconsin

Soil Microbiology

J. J. Germida


University of Saskatchewan

Soil Microbiology

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