Microbial Ecology of the Lake Erie Ecosystem (click here to return)                            

MELEE - Standard Methods Employed in the Analysis of Great Lakes Microbial Communities and Biogeochemistry

This website is a compilation of methods that are commonly used by researchers working in the Microbial Ecology of the Lake Erie Ecosystem (MELEE) Network. While text has been checked for accuracy, it is strongly suggested that researchers interested in employing these approaches see the original citations as well as contact colleagues knowledgeable in this area.

Chlorophyll a estimates

Size-fractionated chlorophyll-a is determined immediately after sample collection from parallel filtration of samples collected on 0.2-µm, 2-µm and 20-µm nominal pore-size polycarbonate filters (47 mm dia.; Osmonics) after extraction (ca 24 h, 4 °C) in 90 % acetone.  Chlorophyll-a retained on the different size class filters is quantified using the non-acidification protocol (Welschmeyer 1994) with either a TD-700 or AU-10 fluorometer (both from Turner Designs). 

Bacterial abundance estimates

Bacterial particles in 2 mL samples are collected onto 25 mm diameter, 0.20-µm nominal pore-size black polycarbonate filters and stained with acridine orange (Hobbie et al. 1977). For all samples, 20 full grids or 200 particles are enumerated using a Leica DMRXA epifluorescence microscope equipped with a Hammamatsu CCD camera system. 

Virus abundance estimates

The abundance of virus-like particles in samples is determined in glutaraldehyde-preserved (2.5% final concentration).  Virus samples are processed to slide stage immediately upon collection and stored at –20° C on the ship (Wen et al. 2004). For viruses, 800 µL aliquots will be collected onto 25-mm diameter, 0.02-µm nominal pore-size Anodisc filters (Whatman) and stained with SYBR Green 1 (Noble and Fuhrman 1998). For all samples, 20 full grids or 200 particles are enumerated using a Leica DMRXA epifluorescence microscope equipped with a Hammamatsu CCD camera system.   

Bacterial production rate estimates

Bacterial production is estimated using the ³H-leucine incorporation microcentrifuge method (Kirchman 2001), which we have demonstrated works well in this system (DeBruyn et al. 2004).  Triplicate 1.5-mL water samples plus 2 trichloroacetic acid-killed blanks are incubated for 1 hour with ³H-(U)-leucine (ca 40 nM final concentration). Cellular protein is subsequently isolated by microcentrifugation and the amount of leucine incorporated determined by liquid scintillation counting. Specific production rates are calculated by dividing the total bacterial production (moles of leucine incorporated L^-1h^-1) by the bacterial abundance of that sample (cells mL^-1) to arrive at the moles L incorporated per cell per hour.  To generate bacterial carbon production estimates, the conversion factor (3.1 kg C per mol leucine) of Wetzel and Likens (2000) are employed.

Primary production and respiration rates.

Primary production and respiration will be determined using the light-dark bottle technique.  Water samples will be placed into BOD bottles, incubated under natural light and temperature, and a comparison of initial and final oxygen concentrations will be used to production-respiration rates.  Concentrations of dissolved oxygen in water samples will be determined using a modified Winkler technique, where whole BOD bottles (300 ml) are titrated using an automated Brinkman Metrohom potentiometric end-point detection system (Fahnenstiel and Carrick 1988; Carrick 2004). For benthic samples, changes in oxygen over time will be measured by inoculating BOD bottles with sediment subsamples and processed as described above. Coefficients of variation among replicate samples are typically < 0.04%. 

The water column oxygen balance in the central basin of Lake Erie will be assessed by direct comparison.  Estimates of production and respiration rates will be derived from the bottle experiments, and balanced to evaluate losses and gains to the system.  This will allow the relative contribution of metalimnetic and benthic algal biomass to be estimated directly. 

Zooplankton inferred mortality rates

The impact of macrozooplankton (organisms > 153-µm in size) grazing on protozoa is determined by experimentally manipulating macrozooplankton concentrations across a series of bottles and evaluating changes in protozoan densities within the bottles over time (Lehman 1980; Carrick et al. 1991).  Collected lake water (150 L) will be screened through a 153-µm mesh size Wisconsin type zooplankton net and is dispensed into a shaded 200-L polyethylene tank, after which an additional 90 L of screened lake water is dispensed into a second 200-L tank.  Epilimnetic macrozooplankton are collected with a 10 m vertical haul using a solid bucket (to avoid excess mechanical damage to the zooplankton) and carefully added to that 90 liter tank by submerging the solid bucket into the collected lake water and allowing the macrozooplankton to escape.

Macrozooplankton treatments are administered by filling 10- L carboys with screened lake water (from the 150-liter sample), while subsequently inoculating the carboys with subsamples from the 90-L zooplankton sample.  Carboys are inoculated with concentrations of zooplankton that approximate 1-, 1.5-, and 3 x ambient macrozooplankton concentrations.  Some bottles will have no macrozooplankton added to them and served as the 0x treatment. To minimize the effects of nutrient recycling via zooplankton excretion to protozoa, phosphate (230 nM final conc.) is added to all bottles.  We choose to add phosphorus to our bottles because this nutrient limits phytoplankton growth in Lake Erie (Schelske et al. 1986) and this concentration will saturate requirements (Wilhelm et al. 2003).  To explore the possibility that macro-zooplankton might be supplying protozoa with organic compounds which could enhance growth by either direct uptake (Haas and Webb 1979) or by augmenting bacterial prey densities (Taylor and Lean 1981), glucose (0.09 µM final conc) is added to one 0x and one 1x bottle. 

All bottles are incubated for 24 h at ambient light and temperature in a shipboard incubator equipped with rotating racks. Initial and final subsamples for nano- and micro-protozoa are removed from the bottles, preserved, and enumerated (as described previously) to estimate exponential growth as follows:

r = ln [Nt / No] / t

where r is the rate of population growth (d^-1), No and Nt are initial and final cell densities, and t is the duration of incubation. At the end of each experiment, macrozooplankton abundance is determined by passing the entire contents of each carboy through a 153-µm mesh size screen and enumerating the retained animals as above. 

The relationship between protozoan growth (dependent variable) and zooplankton biomass (independent variable) is assessed using simple linear regression.  The slope of this relationship provides an estimate of the weight-specific zooplankton clearance rate on protozoa (µg dry wt L^-1 d^-1) and the y-intercept is an estimate of the exponential growth rate (d^-1) of the protozoa (Lehman and Sandgren 1985).  We calculate the flux of carbon from protozoa to macrozooplankton (µg C L^-1 d^-1) by multiplying the clearance rate for the protozoan group under question by the ambient macrozooplankton biomass and, in turn, multiplying this product by the ambient biomass of the protozoan group itself.

Virus-induced mortality rates

Virus production rates are determined using the dilution approach of Wilhelm et al. (2002). For the dilution assay the native microbial community is collected over an acid clean 0.2-µm polycarbonate filters or spiral cartridge that retains the infected members of the native community. This community is maintained at ambient concentrations by the introduction of virus free (< 30,000 Da) seawater from the specific station of interest during the filtration (generated by ultrafiltration – Wilhelm and Poorvin 2001).  The population is then maintained under ambient conditions for up to 12 hours and the reoccurrence of viruses in the sample denoted at 2-3 hour increments to determine the production rate in true triplicate samples. Incubations are kept short during these experiments in order to avoid the products to reinfection in the sample.  This approach has worked well in several environments. Virus induced mortality (VIM) is calculated as follows:

VIM = (viral production rate / burst size) / bacterial abundance

To determine burst size, whole water (40 mL) samples preserved with glutaraldehyde (2.5% v/v) are collected and stored in the dark at 4°C. Upon returning to the lab samples are collected onto carbon-coated Formvar films atop 400-mesh electron microscope grids by centrifugation and stained with 0.75% uranyl formate.  Frequency of visibly infected bacteria cells (FVIC) as well as burst size will be determined by using the Hitachi H-600 and H-800 transmission electron microscopes at UTK as previously described (Weinbauer and Suttle 1996, Wilhelm and Smith 2000).  For each sample, three grids are prepared and at least 1,000 bacteria cells per grid for infection in each.  Burst size (BS) is defined as the average number of viral particles in all visibly infected cells (VIC). As this is likely the minimum BS, we  also keep track of the BS in cells that appear to be completely filled with viral particles (as per Weinbauer and Suttle 1996) to determine a maximal burst size. In total, the results provide an estimate of the total abundance of bacteria destroyed per day by viral activity.

Measurement of alkaline phosphatase activity (APase; EC. 3.1.3.1)

APase activity will be measured in coordination with experiments and survey samples, and used to determine the nutrient stature of plankton in this P-limited system. Unfiltered water collected is dispersed to triplicate methacrylate cuvettes (2.5-mL) and incubated with 40-µM 4-methyl umbelliferyl phosphate (Sigma) at ambient laboratory temperature (~ 21 °C) for 4 - 5 h in darkness.  Sodium bicarbonate (4 mM) is substituted for unfiltered water in substrate controls whereas quench standards are prepared using unfiltered water and 1 µM of 4-methylumbelliferone (Sinsabaugh et al. 1997). APase-catalyzed fluorescence is determined using a TD-700 laboratory fluorometer (Turner Designs,) equipped with a near UV lamp and a methylumbelliferyl filter set (λex: 300-400 nm; λem: 410-610 nm). Enzyme activities are calculated using a 4-methylumbelliferone reference standard. Activities are normalized to total chlorophyll a (> 0.2 mm) in each bottle.

Specific substrate utilization.

Heterotrophic bacteria acquire carbon from a variety of sources.  Classified according to general preference, these sources are identified as labile, semilabile, recalcitrant and refractory.  Labile compounds, because of their suitability to a broad range of heterotrophs, are often rapidly depleted from the pool of dissolved organic matter (DOM). Of the compounds remaining, those contained in the semilabile DOM pool are preferred since they can be hydrolyzed into labile components by bacterial ectoenzymes. Rates of hydrolysis are inferred from use of small soluble substrate proxies, commonly monosaccharides or amino acids covalently linked to fluorophores. These fluorogenic substrates are added to the water sample and rates of hydrolysis calculated from measuring increases in fluorescence intensity over time.

Serving as representative carbohydrate and polypeptide moieties, respectively, we will use methylumbelliferyl-(MUF)-glucoside (both a- and b-linked) and L-leucine 7-amido-4-methylcoumarin (MCA). By examining the ratio of leucine aminopeptidase:b-glucosidase activity, we can infer the relative importance and bioavailability of proteins and polysaccharides as carbon sources to the endemic heterotrophic community (Christian and Karl 1995, Arnosti 2003). These ratios may change seasonally or even across basins depending on inputs of allochthonous and autochthonous carbon sources. Rates of hydrolysis measured using these substrates can be used in our mass balance modeling of carbon consumption.

Experimental procedures will be identical to those outlined for the measurement of alkaline phosphatase activity (see above). At the beginning of each cruise, a kinetic analysis will be conducted where substrate concentrations will be varied. Subsequent assays will be conducted using the lowest concentration deemed saturating with respect to enzyme activity. Attempts will be made to regulate incubation temperatures to coincide with the temperature of the water column at the time of sampling. Maximum potential activity will be calculated by incubating samples at 37 °C. Recognizing that MUF-glucoside is not representative of all potential carbohydrate carbon sources, we will supplement our assays using additional MUF-monosaccharides (e.g. MUF-mannosides, MUF-galactosides).

Spatial Distribution of EEA

Extracellular enzyme activity (EEA) and substrate induced respiration (SIR) assays have been described by Sinsabaugh and Foreman (2001).  Sediment slurries for each sampling station are made by mixing two grams of sediment with 400 mL of sterile water.  Both EEA and SIR assays are measured on 96-well microplates.  For hydrolytic enzymes, 200 mL of slurry is mixed with 50 mL of methylumbelliferyl (MUF)-linked substrate solution for a final substrate concentration of 40 mM.  EEA measured were alkaline phosphatase (EC 3.1.3.1), b-1, 4-glucosidase (EC 3.2.1.21), b-N-acetylglucosaminidase (EC 3.2.1.30), b-1, 4-galactosidase (EC 3.2.1.23) and sulfatase (EC 3.1.6.1).  Each enzyme assay was replicated eight times.  Appropriate MUF reference standards, quenching and substrate controls were measured as well.   Microplates were then incubated at room temperature in the dark and read at 0.5, 1, 2, 3 and 4 hours in order to determine the V_max for each enzyme.  Fluorescence is measured using a Wallac Victor 1420 Multipurpose Counter at an excitation wavelength of 365 nm and an emission wavelength of 450 nm.

Oxidases are measured spectrophotometrically with L-3, 4-dihydroxyphenylalanine (DOPA) as the enzymatic substrate.  Phenol oxidase (EC 1.14.18.1) activity was quantified by adding 50 mL of 25 mM DOPA to 200 mL slurry samples.  Horseradish peroxidase (EC 1.11.1.7) activity is measured similarly, with the addition of 10 mL of 3% H₂O₂ to each well.  Treatments were assayed in replicates of 16, while controls are performed in replicates of eight.  Plates were incubated at room temperature in the dark and read regularly for 11 hours in order to determine V_max.  Enzyme activities are measured spectrophotometrically at 450 nm using a Wallac Victor 1420 Multipurpose Counter.

SIR profiles are attained using Ecolog plates (Biolog Inc.), which consisted of 96-well microplates with 31 different carbon sources and a negative control in triplicate.  Ecolog plates were inoculated with 150 mL of sediment slurry and incubated at room temperature in the dark for 72 hours.  Most of the color change occurred between 48 and 72 hours, with little color change after 72 hours.  SIR profiles were quantified by noting whether or not color change had taken place.  Absorbance measurements could not be obtained due to the growth of a film over many wells.

Nutrient analysis

Nutrient concentrations (total P (filtered and unfiltered), SRP, total N (filtered), NH₄⁺, NO₂, NO₃ + NO₂, SiO₂, and CHN) in both lake water and mesocosm experiments are determined for our work at the National Laboratory for Environmental Testing (NLET) associated with the National Water Research Institute. NLET is a world leader in the development of standards for environmental samples. Bourbonniere will supervise the submission and documentation of nutrient samples collected by all PIs.  Given this significant amount of work, funds for a co-operative education student / summer student are included in the Environment Canada component.  This student will participate in all of our cruises and be responsible for the collection of samples for nutrients. Filtered (<0.2 µm) and total nutrient samples will be collected and measured as previously described in numerous publications (e.g., NLET 1994, Wilhelm et al. 2003, DeBruyn et al. 2004).

Carbon Analysis (DOC/POC/DIC)

DOC is isolated into different size fractions analytically using high performance size exclusion chromatography or preparatively by a combination of ultrafiltration and reverse osmosis.  It can also be fractionated according to physico-chemical character (acidity, hydrophobicity) in analytical or preparative modes using various resins (e.g. DAX-8) and this is carried out as necessary. Fractions are analyzed at Environment Canada for total carbon and nitrogen content.

DOC and any subfractions are quantified on an Apollo 9000HS Carbon Analyzer (Tekmar-Dohrmann), a high temperature catalytic combustion instrument. Inorganic carbon is first removed from DOC samples by the carbon analyzer through acidification with 20% H₃PO₄ and by purging with carrier gas.  DIC is determined on either the Apollo instrument above or on a Dohrmann DC-190 instrument.  In either case the sample is injected into an acid chamber and the CO₂ released is detected.  Standards and blanks are run daily to determine the system blank which is used to correct all sample results.  The CO₂ released by acid or combustion on the Apollo or Dohrmann instruments is detected by non-dispersive infrared detector.

Particulate organic material (POM which includes POC and PON) is measured on dried pre-weighed glass fiber disks, after acid treatment also using a high temperature catalytic instrument at the NLET laboratory.  As well POC only concentrations in the 1-5 mg C L^-1 range can be determined on the unfiltered and acid treated sample on the Apollo instrument with using its particulate sampling option.  C & N are determined on the CHN analyzer by gas chromatography.

Bioreporter determinations of Fe, N, and P During the last several years our group has actively explored new techniques, including the use of bioluminescent reporters of nutrient availability. While development of these tools is ongoing, we have already reported on the use of tools to assess nitrogen and phosphorus (Wilhelm et al. 2003) as well as iron (Mioni et al. 2003, Porta et al. 2003)

References

Caron, D.A. 1983. Technique for enumeration of heterotrophic and phototrophic nanoplankton, using epifluorescence microscopy and comparison with other procedures. Appl. Environ. Microb. 46: 491-498.

Carrick, H.J. and G.L. Fahnenstiel 1989. Biomass, size structure, and composition of phototrophic and heterotrophic nanoflagellate communities in lakes Huron and Michigan. Canadian Journal of FIsheries and Aquatic Sciences 46: 1922-1928.

Carrick, H. J. and Fahnenstiel, G. L. Common planktonic protozoa in the upper Great Lakes: An illustrated guide.  95. Ann Arbor, MI, White Pine Press .

Carrick, H.J. and G.L. Fahnenstiel 1990. Planktonic protozoa in Lakes Huron and Michigan: Seasonal abundance and composition of ciliates and dinoflagellates.  Journal of Great Lakes Research  16: 319-329.

Carrick, H.J. , G.L. Fahnenstiel, E.F. Stoermer, and R.G. Wetzel 1991. The importance of zooplankton-protozoan couplings in Lake Michigan. Limnology and Oceanography 36: 1335-1345.

Choi, J.W. and D.K. Stoecker 1989.  Effects of fixation on cell volume of marine planktonic protozoa. Applied and Environmental Microbiology 55: 1761-1765.

DeBruyn, J.M. , J.A. Leigh-Bell, R.M.L. McKay, R.A. Bourbonniere, and S.W. Wilhelm 2004. Microbial distributions and the impact of phosphorus on bacterial activity in Lake Erie. Journal of Great Lakes Research 30: 166-183.

Fahnenstiel G.L., Carrick HJ 1988. Primary production in lakes Huron and Michigan in vitro and in situ comparisons. Journal of Plankton Research 101273-1283

Haas, L.W. and K.L. Webb 1979. Nutritional mode of several non-pigmented microflagellates from the York River estuary, Virginia. Journal of Experimental Marine Biology and Ecology. 39: 125-134.
Notes: Patty.

Hobbie, J.E., R.J. Daley, and Jasper S 1977. Use of Nuclepore filters for counting bacteria by fluorescence microscopy. Applied and Environmental Microbiology 33: 1225-1228.

Kirchman, D.L. 2001. Measuring bacterial biomass production and growth rates from leucine incorporation in natural aquatic environments. Methods in Microbiology 20: 227-236.

Lehman, J.T. 1980. Nutrient recycling as an interface between algae and grazers in freshwater communities. Chapt. 23 in: Kerfoot 1980.

Lehman, J.T.  and C.D. Sandgren 1985. Species-specific rates of growth and grazing loss among freshwater algae. Limnology and Oceanography 30: 34-46.

Mioni, C.E., A.M. Howard, J.M. DeBruyn, N.G. Bright, M.R. Twiss, B.M. Applegate, and S.W. Wilhelm 2003. Characterization and field trials of a bioluminescent bacterial reporter of iron bioavailability. Marine Chemistry 83 31-46.

NLET (The National Laboratory for Environmental Testing). Manual of Analytical Methods; Volume 1.  Major Ion and Nutrients.  94. Burlington, Ontario, Environment Canada.

Noble, R.T. and J.A. Fuhrman 1998. Use of SYBR Green I for rapid epifluorescence counts of marine viruses and bacteria. Aquatic Microbial Ecology 14: 113-118.

Porta, D., G.S. Bullerjahn, K.A. Durham, S.W. Wilhelm, M.R. Twiss, and R.M.L. McKay 2003. Physiological characterization of a Synechococcus bioreporter (Cyanophyceae) strain PCC 7942 iron-dependent bioreporter for freshwater environments. Journal of Phycology 39 64-73.

Schelske, C.L., E.F. Stoermer,  G.L. Fahnenstiel, and M. Haibach 1986. Phosphorus enrichment, silica utilization, and biogeochemical silica depletion in the Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 43: 407-415.

Sinsabaugh, R.L. and C.M. Foreman 2001. Activity profiles of bacterioplankton in a eutrophic river . Freshwater Biology 46: 1239-1249.

Utermohl, H. 1958. Zur Vervolkommung der quantitativen phytoplankton.  Mitt. Int. Verein. Limnol. 9: 1-13.

Verity, P.G. and M. Vernet 1992. Microzooplankton grzing, pigments, and composition of plankton community during late spring in 2 Norwegian Fjords. Sarisa 77: 263-274.

Weinbauer, M.G. and C.A. Suttle 1996. Potential significance of lysogeny to bacteriophage production and bacterial mortality in coastal waters of the Gulf of Mexico. Appl. Environ. Microbiol. 62: 4374-4380.

Welschmeyer, N.A. 1994.  Fluorometric analysis of chlorophyll a in the presence of chlorophyll b and pheopigments. Limnology and Oceanography 39: 1985-1992.

Wen, K., A.C. Ortmann, and C.A. Suttle 2004. Accurate estimation of viral abundance by epifluorescence microscopy. Applied and Environmental Microbiology 70: 3862-3867.

Wetzel, R. G. and Likens, G. E. Limnological analyses.  2000. New York, Springer.

Wilhelm, S.W., S.M. Brigden, and C.A. Suttle 2002. A Dilution Technique For The Direct Measurement Of Viral Production: A Comparison In Stratified And Tidally Mixed Coastal Waters. Microbial Ecology 43: 168-173.

Wilhelm, S.W., J.M. DeBruyn, O. Gillor, M.R. Twiss, K. Livingston, R.A. Bourbonniere, L.D. Pickell, C.G. Trick, A.L. Dean, and R.M.L. McKay 2003. The effect of phosphorus amendments on present day plankton communities in pelagic Lake Erie. Aquatic Microbial Ecology 32: 275-285.

Wilhelm, S.W. and L. Poorvin 2001. Quantification of algal viruses in marine samples, p. 53-66. In  [ed.], J. Paul Methods in Microbiology.   Academic Press.

Wilhelm, S.W. and R.E.H. Smith 2000. Bacterial carbon production in Lake Erie is influenced by viruses and solar radiation. Canadian Journal of Fisheries and Aquatic Sciences 57: 317-326.