USGS

Open-File Report 2004–1446

In cooperation with the U.S. Environmental Protection Agency

Bioaccumulation of Mercury in Riverine Periphyton

By Amanda H. Bell and Barbara C. Scudder

The PDF for the report is 2,294 kb


Table of Contents

Introduction

Methods

Results

Acknowledgments

References Cited

Figures

Figure 1. Location of the three detailed study units and the U.S. Geologica...

Figure 2. Location of the eight study sites for mercury in periphyton, 2003...

Figure 3. A wide range of algal species at 400X magnification found in the ...

Figure 4. A Chlorophyta (green algae) Closterium sp. at 400X magnification ...

Tables

Table 1. Site information for the eight stations sampled for mercury in per...

Table 2. Raw data of all periphyton samples. See table 1 for complete descr...

Table 3. Divisional classification data of all periphyton samples. See tabl...


Introduction

In aquatic ecosystems, algae are the primary producers and the base of the food web. To date, there has been little research on the role of benthic algae (periphyton) in the bioaccumulation of mercury (Hg) in riverine systems —a key step of the process of bioaccumulation from the physical environment (water and sediments) to higher aquatic organisms (invertebrates, fish, and others). Periphyton has been shown to have an important role in the transfer of mercury in wetlands of the Florida Everglades (Cleckner and others, 1999) and in some situations served as the host site for mercury methylation, which is the key process controlling mercury toxicity in the environment. Pickhardt and others (2002) found that algal blooms in lakes resulted in reduced bioaccumulation of mercury in algal-rich eutrophic lake systems due to decreases in the concentration of mercury per algal cell.

In 2003, the United States Geological Survey (USGS) National Water Quality Assessment (NAWQA) and Toxic Substances Hydrology (Toxics) programs initiated a study to assess mercury bioaccumulation and cycling in eight differing stream-ecosystem settings across the U.S. One aspect of this project involved a detailed examination of the role of periphyton in the trophic transfer of methylmercury (MeHg) in indigenous food webs. This periphyton-mercury study was based in three NAWQA study basins chosen for the first intensive mercury project sampling; the Western Lake Michigan Drainages (WMIC), the Willamette Basin (WILL), and the Georgia-Florida Coastal Plain (GAFL), shown in figure 1. Two to three study sites in each of the three basins were chosen for the NAWQA/Toxics study to represent one urban site and one to two reference/non-cultivated (low- and high-percent wetland) sites. Table 1 lists site information for the sampled rivers, and site locations are shown in figure 2. Currently, there are no generally accepted methods for collection of periphyton for mercury or methylmercury determinations. This paper discusses the collection process and analysis for mercury in periphyton.


Figure

Figure 1. Location of the three detailed study units and the U.S. Geological Survey's National Water Quality Assessment Program's study units.


Table 1. Site information for the eight stations sampled for mercury in periphyton, 2003.

[d, degree; m, minute; s, second]

Study unit USGS station ID Station name River code Landscape type Basin area (square miles) Latitude-longitude (ddmmssdddmmss) Percent wetland1
GAFL 02234998 Little Wekiva River near Longwood, FL LW Urban 44.5 2842070812332 4.50
GAFL 02322500 Santa Fe River near Fort White, FL SF Reference, non-cultivated 1020 2950550824255 18.0
GAFL 02231000 St. Marys River near MacClenny, FL SM Reference, non-cultivated 700 3021310820454 48.0
WILL 14206435 Beaverton Creek at SW 216th Ave, near Orenco, OR BT Urban 36.9 4531151225354 0.16
WILL 14161500 Lookout Creek near Blue River, OR LO Reference, non-cultivated 24.1 4412351221520 0.00
WMIC 04075365 Evergreen River below Evergreen Falls near Langlade, WI EG Reference, non-cultivated 64.5 4503570884034 9.30
WMIC 04087204 Oak Creek at South Milwaukee, WI OC Urban 25.0 4255300875212 8.10
WMIC 04066500 Pike River at Amberg, WI PR Reference, non-cultivated 255 4529490881818 18.0

1 Land use data derived from National Land Cover Dataset using 30-meter Thematic Mapper data (Vogelmann and others, 2001).


Figure

Figure 2. Location of the eight study sites for mercury in periphyton, 2003.


Methods

Collection

Trace-metal clean sampling techniques were used to minimize potential sample contamination (USEPA, 1996; Cleckner and others, 1998; Cleckner and others, 1999). These techniques generally serve to minimize contact between the sample and field crews that employ sampling devices and sample containers that have been stringently cleaned in acid. Prior to use in the field, all glass and Teflon®1 equipment was cleaned by immersing in 4 normal (N) hydrochloric acid (HCl) at 65° celsius (C) for at least 48 hours and then immersed and rinsed at least three times with reagent-grade deionized water (>18 megohms (MOhm)). Equipment other than glass or Teflon® was soaked for at least four hours in a solution of reagent-grade water and Liquinox® (a non-ionic surfactant detergent). This equipment was then triple-rinsed with reagent-grade water, placed in five-percent HCl (Omnitrace) for at least four hours, and finally immersed and triple rinsed with reagent-grade water. After cleaning, all sampling equipment and sample containers were stored by double bagging in hermetically sealed plastic bags.

1 The use of firm, trade, and brand names does not constitute endorsement by the U.S. Government.

At each sampling site, two types of periphyton samples were collected. The (USEPA) Rapid Bioassessment Protocol (Barbour and others, 1999) recommends "single-habitat sampling should be used when biomass of periphyton will be assessed." The single-habitat sampling targets two contrasting habitats that are estimated to be the primary periphyton habitats in the study streams: the depositional-targeted habitat (DTH) and the taxonomically richest-targeted habitat (RTH). The DTH sample may be collected from fine sediment such as silt/clay or sand as appropriate. The DTH and RTH periphyton samples for this study were collected and composited from separate locations in the stream. The NAWQA single habitat sampling method (Moulton and others, 2002) was used for this study because the surface area sampled by this method is quantifiable and those two habitats are generally where periphyton growth dominates in streams.

The overall study design for this project called for seasonal comparison of mercury and methylmercury concentrations and fluxes during spring (high flow) and fall conditions (base flow). For the spring DTH sampling, three areas in each stream were sampled, typically in depositional areas with high organic carbon content in the streambed sediment. These areas were targeted for sampling due to relative abundance of fine sediments and presumably low redox conditions that would promote mercury methylation. Hem (1985) defines redox as the processes of a participating element losing or gaining orbital electrons. The sediment sampling procedure employed by this study seeks to capture the upper 0.5 centimeters (cm) of sediment by employing a Teflon® petri dish (2.54 cm diameter) that is carefully placed open-side down on the streambed sediment to enclose a 19.64 cm2 circle of sediment. A thin sheet of Teflon® was slid under the opening to capture the sediment contained in the petri dish, which is then transferred into a 500 milliliter (mL) Teflon® jar. A more complete description of this method can be found in Porter and others (1993) and Moulton and others (2002) with the addition of trace-metal-clean techniques. For the fall DTH sampling, subsamples were collected at specific areas where methylation rates were found to have the highest methylation potential during the springtime sampling (M. Marvin-DiPasquale, U.S. Geological Survey, Menlo Park, Calif., written commun., August 11, 2003).

The RTH samples were collected from either rock cobbles (epilithic) or woody snags (epidendric) by brushing the algal growth with a stiff-bristled toothbrush-type brush into a Teflon® dish and transferring the slurry into a 500-mL Teflon® jar. Five cobble or woody snags were collected at five locations in each stream for a total of twenty-five composited samples as described in Moulton and others (2002). This method was used with the addition of trace-metal-clean techniques. Rock cobble was the preferred substrate in the WMIC and WILL basins; however, woody snags were used in the GAFL basin because of a general lack of cobble or larger sized rocks at these sites. To determine the surface area of the cobble samples, a section of aluminum foil was placed over the rock and cut to the size and shape of the area scraped. Each sample template was then weighed to determine surface area based on a seven-point curve of mass to surface area from each roll of aluminum foil used. To determine surface area of the woody snags, the length of each snag was measured to the nearest millimeter excluding the first two centimeters from each end. These areas were not scraped to minimize contamination from handling and cutting of the snag.

Processing

Samples were processed on site, or in some instances held in a darkened cooler with wet ice for up to six hours until processing. If the DTH sample contained a large quantity of sand, the slurry was shaken vigorously for 30 seconds and immediately decanted into another 500-mL wide-mouth Teflon® bottle. Fifty milliliters of reagent-grade water was added to the original container, which was shaken again for 30 seconds. The new slurry was immediately decanted into the second 500-mL wide-mouth Teflon® bottle. This elutriation was repeated a third time so that all that remained in the original 500-mL wide-mouth Teflon® bottle were sand particles and the final sample contained little or no sand. The RTH samples contained little or no sand at the time of sampling and were processed without decanting/elutriation.

For total mercury, methylmercury, and stable isotopes, the sample was swirled and shaken to homogenize and suspend algal cells and 5 to 15 mL of the sample was placed on a 47 millimeters (mm) Whatman® quartz fiber filter (QFF) for each subsample. The subsample was filtered by vacuum filtration, using methods of Lewis and Brigham, (2005, in press). Care was taken to ensure that pressure inside the vacuum filtration chamber remained below 10 pounds per square inch (psi) so that the algal cells did not lyse due to high pressures. Each filter was then placed into a petri dish (Teflon® for mercury samples and sterile polystyrene for stable isotopes) and frozen on dry ice for shipment to the respective laboratory for analysis. The chlorophyll a and ash-free biomass subsamples were prepared similarly on 47-mm Whatman® glass fiber filters (GFF). GFF filters were folded in quarters, wrapped in aluminum foil, placed into a sterile polystyrene petri dish, then frozen on dry ice and shipped to the laboratory for analysis.

Two 100-mL subsamples were removed from the remaining sample for gross taxonomic identification and preserved to 5 percent (5 mL addition) with 100 percent formalin buffered to pH 7.

Laboratory Analysis

Total Hg and MeHg analyses were performed by the USGS Wisconsin District Mercury Laboratory (WDML) in Middleton, Wis. For total Hg, the frozen filters were thawed at room temperature for 20 minutes and placed in a 125-mL wide-mouth Teflon® bottle. The petri dish that contained the filter was rinsed three times with five percent bromine chloride (BrCl) into the same bottle. The volume of the bottle was then brought to 100.0 mL with 5-percent BrCl. The bottles were tightly capped, double bagged and allowed to oxidize in an oven at 50° C for five days. The oxidized samples were analyzed with USEPA Method 1631 (USEPA, 2002) using an automated mercury analysis system (Tekran® 2600) with gold trapping, thermal desorption, and cold vapor atomic fluorescence spectrometry detection.

For methylmercury, the extraction method for filtered periphyton was used, which is the same procedure developed by the WDML for the analysis of suspended solids on filters (DeWild and others, 2004). Thawed filters were placed into 125 mL distillation vessels and the petri dish that contained the filter was rinsed three times with reagent-grade water into the vessel. The volume in the vessel was brought up to 50.0 mL by weight, and 2.0 mL of a combined reagent (two parts 8 M sulfuric acid (H2SO4), one part 20-percent potassium chloride (KCl), and two parts copper sulfate (CuSO4)) was added. The distillation vessel was capped and placed in a distillation block, and 50.0 mL of reagent-grade water was added to a receiving vessel. Nitrogen gas (N2) was allowed to flow into the distillation vessel at 60 mL/min. The distillation block was heated to 125 ± 5° C and distillation was allowed to proceed until approximately 25 percent of the original volume remained. The volume was recorded and the solution in the receiving vessel was analyzed for methylmercury according to DeWild and others (2002).

Chlorophyll a and ash-free biomass were analyzed at the National Water Quality Laboratory (NWQL) in Denver, Colo., using a spectrofluorometric method described in USEPA Method 445 (Arar and Collins, 1997).

The USGS National Research Program Isotopic Tracers Laboratory in Menlo Park, Calif., analyzed periphyton samples for stable 13C and 15N isotopes as described in Kendall and others (2001) by determining carbon and nitrogen isotopic and elemental composition on a Carlo Erba 1500® elemental analyzer that is linked in series to a Micromass Optima® mass spectrometer.

Taxonomic identification to algal division was performed using a Bausch and Lomb compound microscope with 400X magnification. Samples were gently swirled to homogenize, and 10 wet-mount slides per sample were prepared using one milliliter aliquots of homogenized sample slurry. Each slide was viewed for identification, and one digital picture was taken for each genus of algal cells occurring more than once and for unique cells encountered, with examples of cells found in figures 3 and 4. Divisional characteristics were determined based on Prescott (1962 and 1970), and Wehr and Sheath (2003). The number of times an algal division was encountered on each slide was recorded per sample site. The divisional composition was rated as very common (≥50 percent of the total cells on the slide), common (50–25 percent), few (≤25 percent), and unique (≤1 percent).


Figure

Figure 3. A wide range of algal species at 400X magnification found in the sediment of the St. Marys River, Fla.


Figure

Figure 4. A Chlorophyta (green algae) Closterium sp. at 400X magnification found in the sediment of the Santa Fe River, Fla.


Results

All raw periphyton data collected are given in table 2 and table 3. Quality control procedures for the collection and processing included collection of approximately 17 percent replicate samples. Replicate values for all analytical parameters were found to be within 5 percent of targeted values.

Table 2. Raw data of all periphyton samples. See table 1 for complete description of sites and river codes.

[First two characters of the sample code denote the river code; the next charater denotes the season (S, spring; F, fall); and the last character denotes the habitat (R, rock; S, sediment; W, wood). Number of samples, n, is 32.]

Sample code Total mercury (nanograms per square meter) Methylmercury (nanograms per square meter) Biomass, ash free dry mass (grams per square meter) Chlorophyll a (milligrams per square meter) Total mercury/AFDM (nanograms per gram) Methylmercury/ CHL A (nanograms per milligram) Percent methyl/total mercury Stable isotopes
d13C d15N
LWFW 589.8 25.75 8.00 4.20 73.73 6.13 4.37 -28.29 10.07
LWSW 603.7 30.91 8.40 4.30 71.87 7.19 5.12 -29.28 11.15
LWFS 10,800 265.3 29.30 18.60 368.7 14.26 2.46 -27.03 5.77
LWSS 27,400 534.6 70.40 67.40 389.3 7.93 1.95 -28.02 6.99
SFFW 861.8 54.94 238.4 12.30 3.62 4.47 6.38 -27.90 4.05
SFSW 637.0 51.43 4.70 5.00 135.5 10.29 8.07 -28.78 6.22
SFFS 89,360 856.2 7.40 7.40 12,080 115.7 0.96 -27.67 4.08
SFSS 127,200 3,798 240.3 20.50 529.4 185.3 2.99 -28.20 3.44
SMFW 1,530 184.1 26.50 23.10 57.74 7.97 12.03 -28.13 3.69
SMSW 1,267 35.34 10.20 <0.1 124.1 353.4 2.79 -28.86 -0.01
SMFS 6,519 218.5 8.70 8.90 749.3 24.55 3.35 -27.23 5.74
SMSS 6,206 181.3 15.20 0.70 408.3 258.9 2.92 -27.70 1.69
BTFS 247,800 2,458 335.2 7.70 739.1 319.3 0.99 -27.68 3.58
BTSS 21,000 132.2 28.70 2.30 731.6 57.48 0.63 -27.49 -6.31
BTFR 1,024 18.54 3.30 2.60 310.3 7.13 1.81 -27.81 3.87
BTSR 556.1 13.63 1.80 2.60 308.9 5.24 2.45 -37.52 12.46
LOFS 18,750 432.3 122.8 33.60 152.7 12.87 2.31 -25.64 -1.21
LOSS 3,024 18.54 7.10 0.80 425.9 23.17 0.61 -27.13 5.93
LOFR 74.35 1.43 2.60 0.60 28.60 2.39 1.93 -18.60 -1.21
LOSR 38.13 1.33 1.00 1.40 38.13 0.95 3.48 -25.85 3.59
EGFS 27,280 3,653 53.40 41.00 510.8 89.10 13.39 -27.05 3.20
EGSS 44,770 2,857 209.0 93.40 214.2 30.59 6.38 -27.43 2.42
EGFR 2,253 191.4 19.10 40.60 117.9 4.71 8.50 -27.47 2.98
EGSR 321.0 35.23 1.80 10.00 178.3 3.52 10.98 -31.00 5.80
OCFS 50,200 665.5 82.10 59.90 611.5 11.11 1.33 -28.85 4.85
OCSS 110,000 2,589 152.3 275.0 722.2 9.42 2.35 -28.42 4.11
OCFR 3,009 77.79 25.10 77.30 119.9 1.01 2.59 -26.66 10.05
OCSR 3,163 213.0 22.30 126.0 141.8 1.69 6.73 -30.42 13.24
PRFS 39,720 1,284 277.3 61.40 143.2 20.92 3.23 -27.09 1.51
PRSS 24,580 1,751 114.2 51.70 215.3 33.87 7.12 -28.03 1.03
PRFR 1,165 73.35 12.60 25.30 92.46 2.90 6.30 -29.50 2.20
PRSR 314.2 14.73 1.20 2.40 261.8 6.14 4.69 -27.97 5.97
Maximum 257,500 3,798 358.6 275.0 12,080 353.4 13.39 -18.60 13.24
Minimum 38.13 0.91 1.00 <0.10 3.62 0.65 0.61 -38.32 -6.31
Mean 30,070 684.8 67.73 30.32 590.0 53.89 4.17 -28.22 4.39
Median 3,024 132.2 15.20 9.03 215.3 9.42 2.99 -27.90 3.69

Table 3. Divisional classification data of all periphyton samples. See table 1 for complete description of sample codes.

[First two characters of the sample code denote the river code; the next charater denotes the season (S, spring; F, fall); and the last character denotes the habitat (R, rock; S, sediment; W, wood).

Sample code Habitat Chrysophyta (number of cells encountered) Cyanophyta (number of cells encountered) Chlorophyta (number of cells encountered) Other (number of cells encountered) Total (number of cells encountered)
BTFR Rock 58 148 265 2 473
BTSR Rock 62 159 287 3 511
LOFR Rock 49 77 152 2 280
LOSR Rock 39 98 164 1 302
EGFR Rock 68 168 369 10 615
EGSR Rock 86 156 264 6 512
OCFR Rock 66 168 359 9 602
OCSR Rock 98 192 364 13 667
PRFR Rock 67 148 426 5 646
PRSR Rock 115 234 379 5 733
LWFS Sediment 435 256 121 15 827
LWSS Sediment 521 302 168 19 1010
SFFS Sediment 398 188 156 20 762
SFSS Sediment 415 234 109 11 769
SMFS Sediment 365 219 126 16 726
SMSS Sediment 531 354 150 22 1057
BTFS Sediment 617 316 286 19 1238
BTSS Sediment 486 206 111 16 819
LOFS Sediment 346 194 125 13 678
LOSS Sediment 289 183 213 23 708
EGFS Sediment 582 318 194 16 1,110
EGSS Sediment 617 349 167 22 1,155
OCFS Sediment 423 216 194 13 846
OCSS Sediment 359 168 168 17 712
PRFS Sediment 522 326 124 19 991
PRSS Sediment 456 267 138 23 884
LWFW Wood 123 352 159 3 637
LWSW Wood 131 289 154 4 578
SFFW Wood 99 223 116 7 445
SFSW Wood 111 258 136 6 511
SMFW Wood 86 207 109 1 403
SMSW Wood 119 291 159 8 577
LOSW Wood 114 264 139 9 526

Acknowledgments

Funding for this project was provided by the U.S. Environmental Protection Agency, Office of Research and Development, National Center for Environmental Assessment (USEPA/ORD/NCEA), and the U.S. Geological Survey National Water-Quality Assessment and Toxic Substances Hydrology programs. We thank Keith G. Sappington, USEPA/ORD/NCEA, for providing technical input and support for this study; and Mark E. Brigham, USGS Minnesota District, for assistance in planning and guidance throughout the project. We are grateful for the guidance from Dr. N. Earl Spangenberg, Dr. Robert A. Bell, and Dr. Bryant A. Browne, professors at the University of Wisconsin-Stevens Point, Amanda Bell's graduate committee. Special thanks to David P. Krabbenhoft, Mark L. Olson, Shane D. Olund, and John F. DeWild of the USGS Wisconsin District Mercury Laboratory staff for their dedication and direction in processing and interpretation of the data. We also recognize the help in guidance, preparation, and sample processing from Dennis A. Wentz, Lia S. Chasar, Richard L. Marella, Kurt D. Carpenter, Michelle A. Lutz, Rebecca H. Woll, Krista A. Stensvold, Jennifer L. Hogan, David O. Bratz, Mark C. Marvin-DiPasquale, Jeffery J. Steuer, and others who assisted during this project.

References Cited

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