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Dietary mercury exposure and bioaccumulation in amphibian larvae inhabiting Carolina bay wetlands
Environmental Pollution 135 (2005) 245–253 www.elsevier.com/locate/envpol
Dietary mercury exposure and bioaccumulation in amphibian larvae inhabiting Carolina bay wetlands J.M. Unrine), C.H. Jagoe, A.C. Brinton, H.A. Brant, N.T. Garvin Savannah River Ecology Laboratory, The University of Georgia, P.O. Drawer E, Aiken, SC 29803, USA Received 21 June 2004; accepted 5 November 2004
Tadpoles accumulate significant concentrations of mercury in isolated wetlands. Abstract Inorganic mercury and methylmercury concentrations were measured both in guts and remaining carcasses of southern leopard frog (Rana sphenocephala) larvae from 10 Carolina bay wetlands in South Carolina, USA. Significant variation among bays in methylmercury and inorganic mercury concentrations existed both in guts and carcasses. There was a moderate negative correlation between dissolved organic carbon concentration in bays and mean inorganic mercury concentrations in guts. There was also a weak positive correlation between pH in bays and mean methylmercury concentrations in carcasses. The ratio of methylmercury to inorganic mercury decreased with increasing total mercury concentration in guts and in larvae, but the rate of decrease was highly variable among bays. Ratios of concentrations in carcasses to concentrations in guts were inversely related to gut concentration. Mercury concentrations in carcasses in some bays were within the range of concentrations at which adverse effects have been observed in laboratory studies of R. sphenocephala. Ó 2004 Elsevier Ltd. All rights reserved. Keywords: Amphibian; Mercury; Speciation; Trophic transfer; Wetland
1. Introduction Relatively high concentrations of mercury (Hg) have been measured in biota inhabiting Carolina bays (isolated depression wetlands on the southeastern coastal plain of the United States) that have no obvious local anthropogenic or geologic point sources of Hg contamination, raising concerns about potential risks of Hg exposure to organisms that utilize these habitats (Brant et al., 2002; Snodgrass et al., 2000). Carolina bays have several characteristics often associated with enhanced bioaccumulation of Hg including: low pH
) Corresponding author. Tel.: C1 803 725 5892; fax: C1 803 725 3309. E-mail address: [email protected] (J.M. Unrine). 0269-7491/$ - see front matter Ó 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.envpol.2004.11.003
(4.5–6.0), high dissolved organic carbon (DOC) content (often O15 mg/L and up to nearly 100 mg/L), and highly fluctuating water levels (Snodgrass et al., 2000). As many as 10,000–20,000 bays may currently exist on the Atlantic Coastal Plain of the United States from New Jersey to northern Florida, and they are important breeding sites for many species of amphibians (Sharitz, 2003). Exposure and subsequent bioaccumulation of Hg in amphibian larvae inhabiting Carolina bays has not been systematically investigated. A recent study suggested that dietary Hg exposure at environmentally realistic concentrations may be sufficient to disrupt normal growth and development in amphibian larvae (Unrine et al., 2004). It is therefore critical that Hg exposure, bioaccumulation, and potential risk to amphibian populations utilizing these habitats be evaluated.
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Diet is probably the most important source of Hg exposure in amphibians because concentrations in the diet can be greater than those in water by a factor of 106 (Unrine and Jagoe, 2004). Previous studies investigating dietary Hg exposure in wildlife have involved analyses of stomach contents (e.g. Gariboldi et al., 1997). Due to the small size of anuran larvae, the length of the gut, and the fragility of the gut tissue, the amount of labor required for separation of gut contents from the gut itself prohibits analyses of gut contents from large numbers of tadpoles. It has been suggested that the relationship between metal concentrations in tadpole guts (including contents) and carcasses minus the gut may provide an indication of uptake of metals relative to concentrations in the diet (Burger and Snodgrass, 1998; Sparling and Lowe, 1995). The primary assumption underlying this approach is that metal concentrations in the gut plus contents are related to metal concentrations in the diet. In a previous study with southern leopard frog (Rana sphenocephala) larvae fed three experimental diets of known Hg concentrations ranging from 0.02–1.40 mg Hg/g dry mass for 60 d (Unrine and Jagoe, unpublished results), concentrations in the guts including contents were not significantly different from concentrations in the experimental diets. It was also found that while gut tissue contributed approximately 25% of the dry mass of the gut and contents, it only contributed about 10% of the total amount of Hg present. Considering the small contribution of Hg contained within the gut to the total amount of Hg in the combined gut and contents, we do not expect Hg contained within the gut that was absorbed from prior feeding to heavily influence estimates of Hg in the gut contents. Thus, gut concentrations can provide a consistent estimate of dietary Hg concentrations over a wide range of gut concentrations in this species. Therefore, the ratio of concentration in carcasses to concentration in guts (CGR) provides an index of bioaccumulation from the diet. This approach helps to reduce uncertainties associated with using concentrations in environmental media such as sediment or aufwuchs (periphyton and associated organisms) as an estimate of Hg concentrations in the larval diet. The composition of the diet of anuran amphibian larvae can vary considerably (Hoff et al., 1999); therefore, metal concentrations and bioavailability in individual environmental components may not be reflective of concentrations and bioavailability in the actual larval diet. In this paper, we present a survey of Hg concentrations and speciation in guts and in carcasses (with the gut removed) of R. sphenocephala larvae inhabiting 10 Carolina bays on the upper coastal plain of South Carolina. A principal objective was to test the predictions of our previous work in artificial impoundments in South Carolina (Unrine and Jagoe, 2004) which used Hg concentrations and speciation in aufwuchs to estimate
Hg concentrations and speciation in the anuran larval diet. We accomplished this by using concentrations and speciation of Hg in tadpole guts as an estimator of concentrations in the tadpole diet. We previously predicted that tadpoles could be exposed to total mercury (THg) concentrations as high as 1600 ng Hg/g dry wt., and that Hg would be mostly present as inorganic mercury (Hg (II)) with percent methylmercury (MMHg) decreasing with increasing total mercury (THg) concentration (Unrine and Jagoe, 2004). For both guts and carcasses, we attempt to correlate observed MMHg and Hg (II) concentrations with water chemistry parameters of the bays. We also attempt to correlate MMHg and Hg (II) concentrations in carcasses with concentrations in guts. A second principal objective was to determine whether ratios of bioaccumulated concentrations to concentrations in the diet in natural habitats are inversely related to concentrations in the diet, as shown in the laboratory (Unrine and Jagoe, 2004). Our third objective was to examine variation among wetlands in the assimilation efficiency of MMHg relative to Hg (II).
2. Materials and methods 2.1. Study area and sampling R. sphenocephala larvae were collected from 10 Carolina bays on the United States Department of Energy Savannah River Site (SRS) located in Aiken and Barnwell counties on the upper coastal plain of South Carolina (Fig. 1). The bays selected were a subset of bays from previous studies of mercury in the diet of wood storks (Mycteria americana) (Brant et al., 2002). The numbers used to identify these bays are from Schalles et al. (1989) who provide more specific locations on the SRS. The SRS is a 780 km2 area established by the United States Atomic Energy Commission in the early 1950s for the production of nuclear materials and contains at least 300 isolated depression wetlands. Most of the land area at the SRS is a large security buffer, with a relatively small area dedicated to industrial activities. The bays and associated watersheds in this study have remained relatively undisturbed since the 1950s and have received no known Hg input from local anthropogenic or geologic sources. Since, the bays have minimal surface and ground water input, and are contained in small, protected watersheds on the SRS, atmospheric deposition is likely the major source of Hg input to these wetlands (Snodgrass et al., 2000). R. sphenocephala breed in many of the depression wetlands on the SRS and may successfully complete metamorphosis in wetlands with hydroperiods of at least 75 d (Gibbons and Semlitsch, 1991). Un-baited minnow traps were used to sample larvae between April 15, 2003 and
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Fig. 1. Location of Carolina bays where larval Rana sphenocephala were sampled in the spring of 2003 on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA. Numbers used to refer to bays are from Schalles et al. (1989).
May 22, 2003, trapping for approximately 24 h at each bay. The larvae were immediately transported to the laboratory and stored in polyethylene bags at ÿ4 C until analysis. R. sphenocephala breed throughout the year on the SRS except for the months of December and January, with major periods of breeding activity occurring in February and September (Caldwell, 1986). Prior to mid-February 2003, all of the bays sampled in
this study were dry due to severe drought (A. Lawrence Bryan, personal communication); thus, the larvae sampled were at most 100 days post-hatching (DPH). Water samples (1 L) were collected from each bay in polyethylene sample jars, immediately transported to the laboratory, filtered to 0.45 mm, and analyzed for pH, alkalinity and conductivity. Samples for DOC determinations were acidified and stored at ÿ4 C prior to
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analysis. Conductivity was determined with an Orion model 126 conductivity meter (Thermo-Orion, Beverly, MA, USA) and pH with and Orion model 720A pH meter equipped with a Ross-type electrode. Alkalinity was determined according to standard titration methods (APHA, 1989). Dissolved organic carbon concentration was measured with a Shimadzu TOC-500 carbon analyzer (Shimadzu, Kyoto, Japan). 2.2. Sample preparation and mercury analyses Only tadpoles that were in developmental stages prior to forelimb emergence (FL; Gosner, 1960 stage 42) were analyzed for Hg. The entire gut of each tadpole was removed through an incision in the abdomen. Wet weights were measured to the nearest 0.1 mg with an analytical balance for the gut and carcass separately and each tadpole was classified according to whether or not complete hindlimb development had occurred (HL; Gosner, 1960 stage 39). Guts and carcasses were lyophilized for approximately 48 h and stored in a desiccator for 24 h prior to determining dry weight. Surgical-grade stainless steel dissecting scissors were used to finely mince and homogenize the guts and carcasses. A sub-sample of up to 100 mg of whole carcass or gut homogenate was then weighed into closed, heavy walled, acid washed polytetrafluoroethylene (PTFE) vials containing 2.5–5 ml (depending on sample size) of 25% (w/v) KOH in methanol and allowed to digest at 70 C overnight. Digestates were then diluted to twice their initial volume in methanol. Small aliquots of the diluted digestates were analyzed by aqueous phase ethylation, room temperature pre-collection, isothermal gas chromatography and cold vapor atomic fluorescence spectrophotometry (GC-CVAFS) (Brooks Rand Model III atomic fluorescence spectrophotometer, Brooks Rand, Seattle, WA, USA) using the methods of Liang et al. (1994). Methylmercury and Hg (II) concentrations were determined by direct standardization and THg was determined by summing the concentrations of MMHg and Hg (II). Peaks for other Hg species were not present in the chromatographs indicating that the sum of the MMHg and Hg (II) concentrations was approximately equivalent to the THg concentration in our samples. 2.3. Quality assurance A National Institute of Standards and Technology (NIST) traceable standard was used for Hg (II) and the actual titer of the MMHg standard was determined as described in United States Environmental Protection Agency (EPA) method 1630 (EPA, 2001). Each analytical batch of 24–36 samples included two extraction blanks, one standard reference material sample (TORT2 lobster hepatopancreas, National Research Council of
Canada, Ottawa, ON, Canada), and one sub-sample replicate. Analytical spike recovery was determined in duplicate for each Hg species for each analytical batch as well as relative percent difference (RPD) between replicate analyses. Detection limits were 1.67 ng Hg/g dry wt. and 9.03 ng Hg/g dry wt. for MMHg and Hg (II), respectively, in 100 mg samples. Concentrations in all samples exceeded these detection limits. Recovery of MMHg and THg from TORT-2 was 112 G 6% and 125 G 3%, respectively (mean G standard error of the mean; TORT-2 is not certified for Hg (II) content). Analytical spike recovery was 84 G 6% for MMHg and 97 G 6% for Hg (II). Relative percent difference between replicate extractions and analyses of subsamples was 23 G 5% for MMHg and 26 G 6% for Hg (II). Relative percent difference between replicate analyses of a single sub-sample was 10 G 4% for MMHg and 11 G 6% for Hg (II). A portion of the variation between replicate sub-samples resulted from the difficulty in homogenizing the tadpole carcasses which were very fibrous and difficult to finely divide. However, because mercury concentrations being measured are low, the measurement uncertainty expressed as a percentage of the mean is inherently large. For example, the standard reference material TORT-2, which has a THg concentration similar to tadpoles in this study, has a certified range for THg which varies by G22% (0.27 G 0.06 ng Hg/g dry mass). 2.4. Data analyses SAS 9.00 (SAS Institute, Cary, NC, USA) was used for all statistical analyses. Potential outliers in the data were identified by examining box plots and Dixon’s test was used to determine if such outliers should be excluded from the data set. Two outliers were removed from the data set. Data were normalized by logtransformation after rejecting the null hypotheses of normality and homoscedasticity required for parametric analysis using Shapiro–Wilk’s and Bartlett’s tests, respectively. Concentrations of MMHg and Hg (II) in carcasses and guts were compared among bays using a generalized linear model (GLM) with carcass dry weight and presence or absence of hindlimbs as covariates. Post hoc multiple comparisons between bays were performed using the Student–Newman–Keuls (SNK) procedure. Correlations of conductivity, pH, alkalinity, and DOC with mean MMHg and Hg (II) concentrations within bays for both carcasses and guts were tested using Spearman’s rank-order correlation analyses. Correlations between mean Hg concentrations in carcasses and guts were also tested for each Hg species. To quantify the relationship between [MMHg] and [Hg (II)] in carcasses and guts and examine variation in this relationship among bays, multiple regression analysis was performed on log-transformed
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dry wt. concentrations with bay and [Hg (II)] as the dependent variables and [MMHg] as the independent variable. Carcass gut ratios (CGRs) were calculated by dividing Hg concentration in the carcass by Hg concentration in the gut for each Hg species. Hg concentrations in the carcasses were assumed to approximate steady state based on a previous finding that tadpoles in the laboratory attained steady state prior to 50 DPH (Unrine and Jagoe, 2004). This minimizes any potential confounding effects of temporal variation in bioaccumulation. Simple linear regression was used to test for a relationship between exposure concentration and CGR. Assimilation efficiency of MMHg relative to Hg (II) was estimated by dividing the ratio of MMHg to Hg (II) in the carcasses by the ratio of MMHg to Hg (II) in the guts as in Mason et al. (1996).
of hindlimbs (F1,175 Z 0.17, p Z 0.6837; Fig. 4). Significant variation among bays for gut Hg (II) concentration was also found (F9,175 Z 3.00, p Z 0.0023; Table 1, Fig. 2), but there was no significant effect of carcass mass (F1,175 Z 0.25, p Z 0.6199; Fig. 3) or presence/ absence of hindlimbs (F1,175 Z 1.66, p Z 0.1997; Fig. 4) on gut Hg (II) concentration. Methylmercury concentrations in carcasses ranged from 13 to 265 ng Hg/g dry wt. with an overall mean of 64 G 3 ng Hg/g dry wt. Inorganic mercury concentrations in carcasses ranged from 26 to 314 ng Hg/g dry wt. for Hg (II) with an overall mean of 120 G 5 ng Hg/g dry wt. (Table 1). There was significant variation in MMHg concentration in carcasses among bays (F9,175 Z 38.78, p ! 0.0001; Table 1; Fig. 2) with no significant effects due to carcass mass (F1,175 Z 0.13, p Z 0.7173; Fig. 3) or presence/absence of hindlimbs (F1,175 Z 1.61, p Z 0.2059; Fig. 4). Similarly, for Hg (II) significant variation among bays existed (F9,175 Z 19.09, p ! 0.0001; Table 1, Fig. 2) with no significant effect due to carcass mass (F1,175 Z 1.75, p Z 0.1874; Fig. 3) or presence/absence of hindlimbs (F1,175 Z 0.07, p Z 0.7983; Fig. 4).
3. Results 3.1. Mercury concentrations Concentrations of MMHg in guts ranged from 15 to 292 ng Hg/g dry wt. with an overall mean of 90 G 4 ng Hg/g dry wt. Concentrations of Hg (II) in guts ranged from 202 to 4125 ng Hg/g dry wt. with an overall mean of 1185 G 52 ng Hg/g dry wt. Significant variation existed in gut MMHg concentrations among bays (F9,175 Z 6.08, p ! 0.0001; Table 1, Fig. 2). However, there was no significant effect of carcass mass (F1,175 Z 0.83, p Z 0.3625; Fig. 3) or presence/absence
3.2. Correlation analyses Conductivity of water from the bays ranged from 36.1 to 58.9 mS/cm at 25 C; pH from 4.2 to 5.7; alkalinity from 0 to 6.4 mg/L; DOC from 3.0 to 97.1 mg/L (Table 2). Concentration of Hg (II) in the gut was negatively correlated with DOC in water (rs Z ÿ0.64, p Z 0.0479). A marginally significant positive correlation was also
Table 1 Methylmercury (MMHg) and inorganic mercury (Hg (II)) concentrations in Rana sphenocephala larvae (carcasses and guts) collected from 10 Carolina bays (isolated depression wetlands) from the United States Department of Energy’s Savannah River Site in South Carolina, USA including ratio of MMHg to Hg (II) in guts and carcasses as well as carcass gut ratio (CGR), and assimilation efficiency of MMHg relative to Hg (II) Bay nb a number 5 42 77 83 86 97 136 139 143 147 All bays
Mean carcass Mean carcass Carcass MMHgc Hg (II)c ratiod
15 90 (7) 20 100 (9) 22 37 (8) 15 89 (7) 12 154 (16) 27 23 (2) 24 39 (3) 8 57 (15) 28 45 (4) 15 35 (4) 186
60 (3)
218 72 73 165 150 82 60 112 110 170
(12) (7) (10) (17) (21) (7) (6) (15) (120) (14)
120 (5)
0.41 1.40 0.51 0.54 1.02 0.28 0.64 0.51 0.41 0.21 0.6 (0.1)
Mean gut Mean gut MMHgc Hg (II)c
Gut ratiod
Assimilation efficiency of MMHg MMHg relative to Hg (II)e CGRf
Hg (II) CGRf
62 102 63 99 45 65 115 33 72 92
0.10 0.12 0.06 0.07 0.06 0.08 0.10 0.03 0.08 0.10
4.03 11.31 8.28 7.84 16.54 3.28 6.14 19.29 5.08 2.02
0.4 0.1 0.1 0.1 0.2 0.1 0.1 0.1 0.1 0.2
(8) (16) (12) (20) (7) (5) (9) (6) (12) (9)
75 (4)
600 824 1024 1432 736 769 1100 1243 884 902
(67) (153) (202) (197) (57) (67) (154) (203) (141) (144)
951 (50)
0.10 (0.008) 8.38 (1.81)
1.5 1.0 0.6 0.9 3.4 0.4 0.3 1.7 0.6 0.4
1.1 (0.3) 0.1 (0.03)
Means are followed by standard errors of the mean in parentheses. a Numbers used to identify wetlands are from Schalles et al. (1989). b Number of larvae collected. c All mercury concentrations expressed in ng Hg/g dry wt. d Ratio of MMHg to Hg (II). e Ratio of MMHg to Hg (II) in carcasses divided by ratio of MMHg to Hg (II) in guts. f Carcass to gut ratio (CGR) calculated by dividing the mercury concentration in the carcass by the mercury concentration in the gut.
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A 350
ln [Hg] ng/g dry wt.
300
ng Hg/g dry wt.
MMHg Hg (II)
5.5 Hg (II) MMHg
250 200 150 100
5.0 4.5
b
4.0 a
3.5 3.0 2.5
50
2.0 0.0
0 86
5
83
147
139
42
143
77
97
0.2
136
0.4
0.6
0.8
1.0
1.2
gutted tadpole mass (g dry wt.)
bay number
B
2000 1800
ng Hg/g dry wt.
1600
Hg II MMHg
1400
Fig. 3. Natural log of methylmercury (MMHg) and inorganic mercury (Hg (II)) concentrations versus body mass in gutted carcasses of Rana sphenocephala larvae collected from 10 Carolina bays (isolated depression wetlands) on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA. Solid lines are least squares regression lines for MMHg (a) and Hg (II) (b).
1200 1000 800 600 400 200 0 83
139
136
77
147
143
42
97
86
5
bay number Fig. 2. Mercury concentrations (ng Hg/g dry wt.) for methylmercury (MMHg) and inorganic mercury (Hg (II)) in gutted tadpoles (A) collected from 10 Carolina bays on the U.S. Department of Energy Savannah River Site (SRS) in South Carolina, USA and corresponding mercury concentrations in guts (B). Numbers used to refer to bays are from Schalles et al. (1989). Error bars correspond to G1 standard error of the mean. Columns under the same horizontal bar at top are not significantly different from each other (type I experimentwise error rate Z 0.05).
(F1,183 Z 7.94, p Z 0.0054) indicating that significant variation in slopes existed among bays (Table 3). Calculated slopes ranged from ÿ0.03 to 0.91 but were only significantly different from zero in bays 5 (slope Z 0.88), 42 (slope Z 0.91), and 139 (slope Z 0.91). The ratio of MMHg to Hg (II) in carcasses was much higher than in guts ranging from 0.2 to 1.4. A linear relationship between ln [MMHg] and ln [Hg (II)] existed in carcasses where data from all bays were pooled (F1,183 Z 3.82, p Z 0.0522, r2 Z 0.33; Fig. 5B, Table 3) but there was no significant variation in slopes among bays (F1,183 Z 0.93, p Z 0.3360). 160 140 120 ng Hg/g dry wt.
found between pH and carcass [MMHg] (rs Z 0.60, p Z 0.0667). Other water chemistry variables were not well correlated with carcass or gut concentrations of MMHg or Hg (II) ( p O 0.11 in all cases). Mean MMHg, Hg (II), and THg carcass concentrations were not correlated with corresponding concentrations in guts (in all cases p ! 0.22).
100 80 60 40 20 0 HL
3.3. Mercury speciation The ratio of MMHg to Hg (II) in guts was small, varying from 0.03 in bay 139 to 0.12 in bay 42 with an overall average of 0.1 (Table 1). A linear relationship existed between ln [MMHg] and ln [Hg (II)] in guts (Fig. 5A, Table 3) when all bays were pooled (F1,187 Z 24.04, p ! 0.0001, r2 Z 0.16). However, there was a significant interaction between bay and [Hg (II)]
NO HL MMHg
HL
NO HL Hg (II)
Fig. 4. Mercury concentrations for Rana sphenocephala larvae with no hindlimbs or incompletely developed hindlimbs (NO HL) compared to larvae with completely developed hindlimbs (HL) for both methylmercury (MMHg) and inorganic mercury (Hg (II)). Number of larvae Z 90 for NO HL and 96 for HL Error bars correspond to G1 standard error of the mean. Larvae were collected from 10 Carolina bays (isolated depression wetlands) on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA.
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J.M. Unrine et al. / Environmental Pollution 135 (2005) 245–253 Table 2 Conductivity, pH, alkalinity, and dissolved organic carbon concentration (DOC) from 10 Carolina bays (isolated depression wetlands) from the United States Department of Energy’s Savannah River Site in South Carolina, USA Bay numbera
pH
Conductivityb
Alkalinityc
DOCc
5 42 77 83 86 97 136 139 143 147
5.72 5.77 4.51 4.61 4.62 4.47 4.98 4.94 4.26 4.50
50.7 58.9 39.3 23.7 36.1 40.3 39.1 49.1 44.4 43.1
6.19 6.46 0.00 0.00 0.00 0.00 0.00 4.31 0.00 0.00
38.8 45.4 33.8 24.5 54.9 97.1 3.0 32.7 61.4 68.4
a b c
Numbers used to identify wetlands are from Schalles et al. (1989). mS/cm at 25 C. mg/L.
Table 3 Slope, intercept and coefficient of determination (r2) for the linear relationship between log of methylmercury concentration (MMHg) and inorganic mercury concentration (Hg (II)) in Rana sphenocephala larvae (carcasses and guts) collected from Carolina bays on the United States Department of Energy Savannah River Site in Aiken and Barnwell counties, SC, USA Baya
nb
5 42 77 83 86 97 136 139 143 147 All bays
Guts
Carcasses
Slope
Intercept
r2
Slope
Intercept
r2
15 20 22 15 12 27 24 8 28 15
0.88** 0.91** 0.24 0.55 0.77 0.61 ÿ0.03 0.91** 0.11 0.53
ÿ1.54 ÿ1.50 ÿ2.46 0.58 ÿ1.29 0.07 4.98** ÿ2.99 3.47 0.88
0.59 0.72 0.06 0.23 0.21 0.50 0.00 0.79 0.01 0.59
0.35 0.45* 0.47* 0.51* 0.65** 0.38* 0.57** 0.39 0.39** 0.12
2.65 2.70* 1.59 1.87 1.79* 1.46* 1.33 2.20 1.97** 2.90
0.08 0.22 0.24 0.33 0.68 0.19 0.32 0.08 0.25 0.01
186
0.41**
1.47
0.16
0.52**
1.53**
0.20
*Significantly different from zero at a Z 0.05, **significantly different from zero at a Z 0.01. a Numbers used to identify wetlands are from Schalles et al. (1989). b Number of larvae collected.
7
A
ln [MMHg]
6
5
4
3
2 5.0
5.5
6.0
6.5
7.0
7.5
8.0
8.5
ln [Hg (II)] 6
Carcass gut ratios were quite low ranging from 0.4 to 3.4 for MMHg and 0.1 to 0.4 for Hg (II). There was a statistically significant inverse relationship between gut concentration and CGR both for MMHg (F1,8 Z 5.56, p Z 0.0461, r2 Z 0.41) and Hg (II) (F1,8 Z 5.80, p Z 0.0426, r2 Z 0.42; Fig. 6). Slopes for MMHg and Hg (II) were similar (ÿ1.28 and ÿ1.39, respectively), however, the intercepts differed (5.14 and 7.37, respectively). The increase in ratio of MMHg to Hg (II) in carcasses versus guts, which is indicative of the assimilation efficiency of MMHg relative to Hg (II) (Mason et al., 1996), was quite variable and ranged from 2.02 to 19.29-fold (Table 1).
B 5 y = -1.23x + 5.15 r2 = 0.44
1
4
MMHg Hg II Linear (MMHg) Linear (Hg II)
0.5
3
0
ln CGR
ln [MMHg]
1.5
2
-0.5 y = -1.36x + 7.24 r2 = 0.43
-1 -1.5
1 3.0
3.5
4.0
4.5
5.0
5.5
6.0
-2
ln [Hg (II)] -2.5
Fig. 5. Relationship between log of inorganic mercury (Hg (II)) and methylmercury (MMHg) in the guts of Rana sphenocephala larvae collected from 10 Carolina bays on the United States Department of Energy Savannah River Site (SRS) in South Carolina (A). Solid line indicates least squares regression line, dashed lines correspond to 95% confidence intervals and dotted lines correspond to 95% prediction intervals. Relationship for mercury concentrations in corresponding gutted whole carcasses (B) is also shown.
-3 3
3.5
4
4.5
5
5.5
6
6.5
7
7.5
ln ng Hg/g dry wt. in gut
Fig. 6. Relationship between log of mercury concentration in the gut versus carcass gut ratio (CGR) calculated by dividing mean concentration in gutted larvae by mean concentration in guts for each of 10 Carolina bays sampled.
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4. Discussion These results support our previous conclusion (Unrine and Jagoe, 2004) that, in general, dietary THg exposures for tadpoles would range from 50 to 1600 ng Hg/g dry wt. in sites with no local anthropogenic or geologic sources of Hg. However, we predicted a range of average exposure concentration on the SRS of about 60 to 120 ng Hg/g dry wt. THg, which underestimated THg concentrations actually observed in the guts (599 to 1432 ng Hg/g dry wt.) (Unrine and Jagoe, 2004). The discrepancy between our predicted and observed Hg concentrations likely stems from differences in dietary components and the relative partitioning of Hg species within these components. Aufwuchs from artificial substrates used to estimate dietary Hg concentrations in our previous work (Unrine and Jagoe, 2004) probably differed from the diet of tadpoles in the bays both in organic composition (autotrophic, heterotrophic, and detrital components) and inorganic composition (sediment content and composition). The prediction from our previous work (Unrine and Jagoe, 2004) that the ratio of MMHg to Hg (II) in the tadpole diet would decrease with increasing THg concentration was supported by the observed relationship between ln [MMHg] and ln [Hg (II)]. In these plots, a slope !1 indicates that the ratio of MMHg to Hg (II) decreases with increasing THg concentration because ln [MMHg] does not increase as rapidly as ln [Hg (II)]. The slope was less than one in all bays, and not significantly different from zero in most bays. The overall slope for the pooled data in this experiment was 0.41, which agrees well with our predictions based upon aufwuchs samples (0.31). However, significant variation in slopes among individual bays (ÿ0.03 to 0.91) demonstrates that Hg speciation in the tadpole diet can vary greatly among bays. Such differences in slope may relate to differences in net methylation rate among bays and/or differences in dietary composition among bays. For example, in bays where the relationship between ln [Hg (II)] and ln [MMHg] had a large slope, a large proportion of the Hg (II) entering the system may have been methylated. Components of the tadpole diet may also differ in their capacities to concentrate MMHg and Hg (II) from the water column, which could explain differences in the relationship between Hg (II) and MMHg among bays. Considerable variation in assimilation efficiency of MMHg relative to Hg (II) probably existed among bays. In the laboratory (Unrine and Jagoe, 2004), we observed an approximately 5.7-fold increase in the ratio of MMHg to Hg (II) in tadpoles relative to experimental diets. Hg content in experimental diets was varied by combining proportions of control and Hg-enriched aufwuchs grown in experimental mesocosms; the mesocosms aufwuchs was primarily composed of filamentous
algae. Mason et al. (1996) showed that the assimilation efficiency of MMHg for zooplankton consuming phytoplankton was approximately 3.5–4.5 times greater than for Hg (II). They related this to the observation that percent of MMHg found in the cytoplasm of diatoms is approximately four times greater than percent Hg (II), and concluded that zooplankton assimilate Hg primarily from the cytoplasm of diatoms. In our previous work, the assimilation efficiency of MMHg relative to Hg (II) was probably reflective of differences in sub-cellular fractionation of MMHg and Hg (II) in the algal cells used as a source of Hg in the experimental diets. The present study shows that assimilation efficiency of MMHg relative to Hg (II) for amphibian larvae in the field can be quite variable. Although experimental diets in the laboratory (Unrine and Jagoe, 2004) may have had concentrations and speciation of Hg reflective of natural environments, relative assimilation efficiencies of MMHg and Hg (II) from the experimental diets were slightly different from what we observed in nature (this study). In both the laboratory (Unrine and Jagoe, 2004) and the field (current study), we found an inverse relationship between exposure concentration and BAF or CGR both for MMHg and Hg (II). A decrease in the rate of bioaccumulation with increasing dietary exposure concentration would tend to dampen the relationship between dietary concentration and bioaccumulated concentration. This at least partially explains the lack of correlation between Hg concentrations in carcasses and concentrations in guts. Differences in dietary composition among bays that resulted in differences in bioavailability could have also contributed to the lack of relationship. Carcass gut ratios observed in this study were similar to dietary bioaccumulation factors (BAFs) observed in the laboratory (Unrine and Jagoe, 2004) for Hg (II) but were slightly higher than laboratory BAFs for MMHg. Bioaccumulation factors for Hg are currently used under the assumption that they are independent of exposure concentration; however, the results of our work using BAFs and CGRs and the work of McGeer et al. (2003) concerning bioconcentration factors (BCFs) for Hg, show that predictions of Hg concentrations in biota based upon either BAFs, CGRs or BCFs are confounded by exposure concentration. An alternative approach might utilize a bioaccumulation factor that is corrected for exposure concentration using a slope factor derived from experimental data. Observed total mercury concentrations in guts in bay 83 exceeded dietary concentrations (approximately 1400 ng Hg/g dry wt.) at which adverse effects on development and decreased metamorphic success were observed in R. sphenocephala in the laboratory (Unrine et al., 2004). Bioaccumulated concentrations in larvae from the bays exceeded bioaccumulated concentrations (approximately 240 ng Hg/g dry wt.) at which effects
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were observed in the laboratory (Unrine et al., 2004) in three bays (5, 83, and 86). This suggests that amphibian larvae inhabiting Carolina bays may be exposed to and bioaccumulate concentrations of Hg within the range of Hg concentrations at which changes in development and reduced metamorphic success might be observed. The conclusion that wildlife utilizing Carolina bays are potentially exposed to toxicologically significant quantities of mercury was reached in a previous investigation concerning dietary exposure in nestling wood storks (Brant et al., 2002). Carolina bays have been described as ‘‘islands’’ of species richness on the Atlantic Coastal Plain and their degradation due to draining could result in a major loss of biodiversity in the southeastern United States (Sharitz, 2003). Contamination by atmospheric mercury deposition should be further investigated to determine if it is an additional threat to the ecological integrity of these important ecosystems. 5. Conclusions Ranid larvae inhabiting relatively pristine Carolina bay wetlands may be exposed to and bioaccumulate toxicologically significant quantities of mercury. This study confirmed the predictions of our previous work (Unrine et al., 2004) that Hg concentrations in the diet of larval anuran amphibians would be as high as 1600 ng Hg/g dry wt. THg in sites with little or no local anthropogenic or geologic mercury input. It also confirmed that Hg is mostly present as Hg (II) in the diet, and that percent MMHg decreases with increasing THg concentration. Even if Hg concentrations and speciation in experimental diets may reflect what is observed in nature, the relative bioavailabilities of MMHg and Hg (II) may not. Future studies should focus on the relationship between composition of gut contents (autotrophic, heterotrophic, detrital, and inorganic components) with MMHg and Hg (II) concentrations as well as the influence of dietary composition on relative bioavailabilities of MMHg and Hg (II). Finally, further investigation into the potential risks of Hg exposure to amphibians and other biota inhabiting Carolina bays is warranted. Acknowledgements This research was supported by the Environmental Remediation Sciences Division of the Office of Biological and Environmental Research, U.S. Department of Energy, through Financial Assistance Award No. DEFC09-96-SR18546 to the University of Georgia Research Foundation. J. Unrine was supported by a doctoral assistantship provided by the Savannah River Ecology Laboratory while conducting this research. Earlier versions of the manuscript benefited from the comments
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of J. Snodgrass, W. Hopkins, M. Smith, I.L. Brisbin, Jr., P. Williams, M. Black and two anonymous reviewers. The authors are also grateful for the advice and assistance of A.L. Bryan and the technical support of T. Paulin. References APHA, 1989. 2320 Alkalinity/titration method. Standard Methods for the Examination of Water and Waste Water. American Public Health Association, Baltimore, MD, USA, pp. 2-35–2-39. Brant, H.A., Jagoe, C.H., Snodgrass, J.W., Bryan, A.L., Gariboldi, J.C., 2002. Potential risk to wood storks (Mycteria americana) from mercury in Carolina Bay fish. Environ Pollut 120, 405–413. Burger, J., Snodgrass, J., 1998. Heavy metals in bullfrog (Rana catesbeiana) tadpoles: effects of depuration before analysis. Environ Toxicol Chem 17, 2203–2209. Caldwell, J.P., 1986. Selection of egg deposition sites: a seasonal shift in southern leopard frog. Copeia 1986, 249–253. EPA, 2001. Method 1630: Methylmercury in Water by Distillation, Aqueous Ethylation, Purge and Trap, and CVAFS. United States Environmental Protection Agency. Gariboldi, J.C., Jagoe, C.H., Bryan, A.L., 1997. Dietary exposure to mercury in nestling wood storks (Mycteria americana) in Georgia. Arch Environ Contamin Toxicol 34, 398–405. Gibbons, J.W., Semlitsch, R.D., 1991. Guide to the Reptiles and Amphibians of the Savannah River Site. University of Georgia Press, Athens, GA, USA. Gosner, K.L., 1960. A simplified table for staging anuran embryos and larvae with notes on identification. Herpetologica 16, 183–190. Hoff, K., Blaustein, A.R., McDiarmid, R.W., Altig, R., 1999. Behavior: interactions and their consequences. In: McDiarmid, R.W., Altig, R. (Eds.), Tadpoles: The Biology of Anuran Larvae. The University of Chicago Press, Chicago, IL, USA, pp. 215–239. Liang, L., Bloom, N.S., Horvat, M., 1994. Simultaneous determination of mercury speciation in biological-materials by GC/CVAFS after ethylation and room-temperature precollection. Clin Chem 40, 602–607. Mason, R.P., Reinfelder, J.B., Morell, F.M.M., 1996. Uptake, toxicity, and trophic transfer of mercury in a coastal diatom. Environ Sci Technol 30, 1835–1845. McGeer, J.C., Brix, K.V., Skeaff, J.M., DeForest, D.K., Brigham, S.I., Adams, W.J., Green, A., 2003. Inverse relationship between bioconcentration factor and exposure concentration for metals: implications for hazard assessment of metals in the aquatic environment. Environ Toxicol Chem 22, 1017–1037. Schalles, J.F., Sharitz, R.R., Gibbons, J.W., Leversee, G.J., Knox, J.N., 1989. Carolina Bays of the Savannah River Plant. Savannah River Plant National Environmental Research Park Program, Aiken, SC, USA. Sharitz, R.R., 2003. Carolina bay wetlands: unique habitats of the southeastern United States. Wetlands 23, 550–562. Snodgrass, J.W., Jagoe, C.H., Bryan, A.L., Brant, H.A., Burger, J., 2000. Effects of trophic status and wetland morphology, hydroperiod, and water chemistry on mercury concentrations in fish. Can J Fish Aquat Sci 57, 171–180. Sparling, D.W., Lowe, T.P., 1995. Metal concentrations of tadpoles in experimental ponds. Environ Pollut 91, 149–159. Unrine, J.M., Jagoe, C.H., 2004. Dietary mercury exposure and bioaccumulation in southern leopard frog (Rana sphenocephala) larvae. Environ Toxicol Chem 23, 2956–2963. Unrine, J.M., Jagoe, C.H., Hopkins, W.A., Brant, H.A., 2004. Adverse effects of ecologically relevant dietary mercury exposure in southern leopard frog (Rana sphenocephala) larvae. Environ Toxicol Chem 23, 2964–2970.
Dietary mercury exposure and bioaccumulation in amphibian larvae inhabiting Carolina bay wetlands J.M. Unrine), C.H. Jagoe, A.C. Brinton, H.A. Brant, N.T. Garvin Savannah River Ecology Laboratory, The University of Georgia, P.O. Drawer E, Aiken, SC 29803, USA Received 21 June 2004; accepted 5 November 2004
Tadpoles accumulate significant concentrations of mercury in isolated wetlands. Abstract Inorganic mercury and methylmercury concentrations were measured both in guts and remaining carcasses of southern leopard frog (Rana sphenocephala) larvae from 10 Carolina bay wetlands in South Carolina, USA. Significant variation among bays in methylmercury and inorganic mercury concentrations existed both in guts and carcasses. There was a moderate negative correlation between dissolved organic carbon concentration in bays and mean inorganic mercury concentrations in guts. There was also a weak positive correlation between pH in bays and mean methylmercury concentrations in carcasses. The ratio of methylmercury to inorganic mercury decreased with increasing total mercury concentration in guts and in larvae, but the rate of decrease was highly variable among bays. Ratios of concentrations in carcasses to concentrations in guts were inversely related to gut concentration. Mercury concentrations in carcasses in some bays were within the range of concentrations at which adverse effects have been observed in laboratory studies of R. sphenocephala. Ó 2004 Elsevier Ltd. All rights reserved. Keywords: Amphibian; Mercury; Speciation; Trophic transfer; Wetland
1. Introduction Relatively high concentrations of mercury (Hg) have been measured in biota inhabiting Carolina bays (isolated depression wetlands on the southeastern coastal plain of the United States) that have no obvious local anthropogenic or geologic point sources of Hg contamination, raising concerns about potential risks of Hg exposure to organisms that utilize these habitats (Brant et al., 2002; Snodgrass et al., 2000). Carolina bays have several characteristics often associated with enhanced bioaccumulation of Hg including: low pH
) Corresponding author. Tel.: C1 803 725 5892; fax: C1 803 725 3309. E-mail address: [email protected] (J.M. Unrine). 0269-7491/$ - see front matter Ó 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.envpol.2004.11.003
(4.5–6.0), high dissolved organic carbon (DOC) content (often O15 mg/L and up to nearly 100 mg/L), and highly fluctuating water levels (Snodgrass et al., 2000). As many as 10,000–20,000 bays may currently exist on the Atlantic Coastal Plain of the United States from New Jersey to northern Florida, and they are important breeding sites for many species of amphibians (Sharitz, 2003). Exposure and subsequent bioaccumulation of Hg in amphibian larvae inhabiting Carolina bays has not been systematically investigated. A recent study suggested that dietary Hg exposure at environmentally realistic concentrations may be sufficient to disrupt normal growth and development in amphibian larvae (Unrine et al., 2004). It is therefore critical that Hg exposure, bioaccumulation, and potential risk to amphibian populations utilizing these habitats be evaluated.
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Diet is probably the most important source of Hg exposure in amphibians because concentrations in the diet can be greater than those in water by a factor of 106 (Unrine and Jagoe, 2004). Previous studies investigating dietary Hg exposure in wildlife have involved analyses of stomach contents (e.g. Gariboldi et al., 1997). Due to the small size of anuran larvae, the length of the gut, and the fragility of the gut tissue, the amount of labor required for separation of gut contents from the gut itself prohibits analyses of gut contents from large numbers of tadpoles. It has been suggested that the relationship between metal concentrations in tadpole guts (including contents) and carcasses minus the gut may provide an indication of uptake of metals relative to concentrations in the diet (Burger and Snodgrass, 1998; Sparling and Lowe, 1995). The primary assumption underlying this approach is that metal concentrations in the gut plus contents are related to metal concentrations in the diet. In a previous study with southern leopard frog (Rana sphenocephala) larvae fed three experimental diets of known Hg concentrations ranging from 0.02–1.40 mg Hg/g dry mass for 60 d (Unrine and Jagoe, unpublished results), concentrations in the guts including contents were not significantly different from concentrations in the experimental diets. It was also found that while gut tissue contributed approximately 25% of the dry mass of the gut and contents, it only contributed about 10% of the total amount of Hg present. Considering the small contribution of Hg contained within the gut to the total amount of Hg in the combined gut and contents, we do not expect Hg contained within the gut that was absorbed from prior feeding to heavily influence estimates of Hg in the gut contents. Thus, gut concentrations can provide a consistent estimate of dietary Hg concentrations over a wide range of gut concentrations in this species. Therefore, the ratio of concentration in carcasses to concentration in guts (CGR) provides an index of bioaccumulation from the diet. This approach helps to reduce uncertainties associated with using concentrations in environmental media such as sediment or aufwuchs (periphyton and associated organisms) as an estimate of Hg concentrations in the larval diet. The composition of the diet of anuran amphibian larvae can vary considerably (Hoff et al., 1999); therefore, metal concentrations and bioavailability in individual environmental components may not be reflective of concentrations and bioavailability in the actual larval diet. In this paper, we present a survey of Hg concentrations and speciation in guts and in carcasses (with the gut removed) of R. sphenocephala larvae inhabiting 10 Carolina bays on the upper coastal plain of South Carolina. A principal objective was to test the predictions of our previous work in artificial impoundments in South Carolina (Unrine and Jagoe, 2004) which used Hg concentrations and speciation in aufwuchs to estimate
Hg concentrations and speciation in the anuran larval diet. We accomplished this by using concentrations and speciation of Hg in tadpole guts as an estimator of concentrations in the tadpole diet. We previously predicted that tadpoles could be exposed to total mercury (THg) concentrations as high as 1600 ng Hg/g dry wt., and that Hg would be mostly present as inorganic mercury (Hg (II)) with percent methylmercury (MMHg) decreasing with increasing total mercury (THg) concentration (Unrine and Jagoe, 2004). For both guts and carcasses, we attempt to correlate observed MMHg and Hg (II) concentrations with water chemistry parameters of the bays. We also attempt to correlate MMHg and Hg (II) concentrations in carcasses with concentrations in guts. A second principal objective was to determine whether ratios of bioaccumulated concentrations to concentrations in the diet in natural habitats are inversely related to concentrations in the diet, as shown in the laboratory (Unrine and Jagoe, 2004). Our third objective was to examine variation among wetlands in the assimilation efficiency of MMHg relative to Hg (II).
2. Materials and methods 2.1. Study area and sampling R. sphenocephala larvae were collected from 10 Carolina bays on the United States Department of Energy Savannah River Site (SRS) located in Aiken and Barnwell counties on the upper coastal plain of South Carolina (Fig. 1). The bays selected were a subset of bays from previous studies of mercury in the diet of wood storks (Mycteria americana) (Brant et al., 2002). The numbers used to identify these bays are from Schalles et al. (1989) who provide more specific locations on the SRS. The SRS is a 780 km2 area established by the United States Atomic Energy Commission in the early 1950s for the production of nuclear materials and contains at least 300 isolated depression wetlands. Most of the land area at the SRS is a large security buffer, with a relatively small area dedicated to industrial activities. The bays and associated watersheds in this study have remained relatively undisturbed since the 1950s and have received no known Hg input from local anthropogenic or geologic sources. Since, the bays have minimal surface and ground water input, and are contained in small, protected watersheds on the SRS, atmospheric deposition is likely the major source of Hg input to these wetlands (Snodgrass et al., 2000). R. sphenocephala breed in many of the depression wetlands on the SRS and may successfully complete metamorphosis in wetlands with hydroperiods of at least 75 d (Gibbons and Semlitsch, 1991). Un-baited minnow traps were used to sample larvae between April 15, 2003 and
J.M. Unrine et al. / Environmental Pollution 135 (2005) 245–253
247
Fig. 1. Location of Carolina bays where larval Rana sphenocephala were sampled in the spring of 2003 on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA. Numbers used to refer to bays are from Schalles et al. (1989).
May 22, 2003, trapping for approximately 24 h at each bay. The larvae were immediately transported to the laboratory and stored in polyethylene bags at ÿ4 C until analysis. R. sphenocephala breed throughout the year on the SRS except for the months of December and January, with major periods of breeding activity occurring in February and September (Caldwell, 1986). Prior to mid-February 2003, all of the bays sampled in
this study were dry due to severe drought (A. Lawrence Bryan, personal communication); thus, the larvae sampled were at most 100 days post-hatching (DPH). Water samples (1 L) were collected from each bay in polyethylene sample jars, immediately transported to the laboratory, filtered to 0.45 mm, and analyzed for pH, alkalinity and conductivity. Samples for DOC determinations were acidified and stored at ÿ4 C prior to
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analysis. Conductivity was determined with an Orion model 126 conductivity meter (Thermo-Orion, Beverly, MA, USA) and pH with and Orion model 720A pH meter equipped with a Ross-type electrode. Alkalinity was determined according to standard titration methods (APHA, 1989). Dissolved organic carbon concentration was measured with a Shimadzu TOC-500 carbon analyzer (Shimadzu, Kyoto, Japan). 2.2. Sample preparation and mercury analyses Only tadpoles that were in developmental stages prior to forelimb emergence (FL; Gosner, 1960 stage 42) were analyzed for Hg. The entire gut of each tadpole was removed through an incision in the abdomen. Wet weights were measured to the nearest 0.1 mg with an analytical balance for the gut and carcass separately and each tadpole was classified according to whether or not complete hindlimb development had occurred (HL; Gosner, 1960 stage 39). Guts and carcasses were lyophilized for approximately 48 h and stored in a desiccator for 24 h prior to determining dry weight. Surgical-grade stainless steel dissecting scissors were used to finely mince and homogenize the guts and carcasses. A sub-sample of up to 100 mg of whole carcass or gut homogenate was then weighed into closed, heavy walled, acid washed polytetrafluoroethylene (PTFE) vials containing 2.5–5 ml (depending on sample size) of 25% (w/v) KOH in methanol and allowed to digest at 70 C overnight. Digestates were then diluted to twice their initial volume in methanol. Small aliquots of the diluted digestates were analyzed by aqueous phase ethylation, room temperature pre-collection, isothermal gas chromatography and cold vapor atomic fluorescence spectrophotometry (GC-CVAFS) (Brooks Rand Model III atomic fluorescence spectrophotometer, Brooks Rand, Seattle, WA, USA) using the methods of Liang et al. (1994). Methylmercury and Hg (II) concentrations were determined by direct standardization and THg was determined by summing the concentrations of MMHg and Hg (II). Peaks for other Hg species were not present in the chromatographs indicating that the sum of the MMHg and Hg (II) concentrations was approximately equivalent to the THg concentration in our samples. 2.3. Quality assurance A National Institute of Standards and Technology (NIST) traceable standard was used for Hg (II) and the actual titer of the MMHg standard was determined as described in United States Environmental Protection Agency (EPA) method 1630 (EPA, 2001). Each analytical batch of 24–36 samples included two extraction blanks, one standard reference material sample (TORT2 lobster hepatopancreas, National Research Council of
Canada, Ottawa, ON, Canada), and one sub-sample replicate. Analytical spike recovery was determined in duplicate for each Hg species for each analytical batch as well as relative percent difference (RPD) between replicate analyses. Detection limits were 1.67 ng Hg/g dry wt. and 9.03 ng Hg/g dry wt. for MMHg and Hg (II), respectively, in 100 mg samples. Concentrations in all samples exceeded these detection limits. Recovery of MMHg and THg from TORT-2 was 112 G 6% and 125 G 3%, respectively (mean G standard error of the mean; TORT-2 is not certified for Hg (II) content). Analytical spike recovery was 84 G 6% for MMHg and 97 G 6% for Hg (II). Relative percent difference between replicate extractions and analyses of subsamples was 23 G 5% for MMHg and 26 G 6% for Hg (II). Relative percent difference between replicate analyses of a single sub-sample was 10 G 4% for MMHg and 11 G 6% for Hg (II). A portion of the variation between replicate sub-samples resulted from the difficulty in homogenizing the tadpole carcasses which were very fibrous and difficult to finely divide. However, because mercury concentrations being measured are low, the measurement uncertainty expressed as a percentage of the mean is inherently large. For example, the standard reference material TORT-2, which has a THg concentration similar to tadpoles in this study, has a certified range for THg which varies by G22% (0.27 G 0.06 ng Hg/g dry mass). 2.4. Data analyses SAS 9.00 (SAS Institute, Cary, NC, USA) was used for all statistical analyses. Potential outliers in the data were identified by examining box plots and Dixon’s test was used to determine if such outliers should be excluded from the data set. Two outliers were removed from the data set. Data were normalized by logtransformation after rejecting the null hypotheses of normality and homoscedasticity required for parametric analysis using Shapiro–Wilk’s and Bartlett’s tests, respectively. Concentrations of MMHg and Hg (II) in carcasses and guts were compared among bays using a generalized linear model (GLM) with carcass dry weight and presence or absence of hindlimbs as covariates. Post hoc multiple comparisons between bays were performed using the Student–Newman–Keuls (SNK) procedure. Correlations of conductivity, pH, alkalinity, and DOC with mean MMHg and Hg (II) concentrations within bays for both carcasses and guts were tested using Spearman’s rank-order correlation analyses. Correlations between mean Hg concentrations in carcasses and guts were also tested for each Hg species. To quantify the relationship between [MMHg] and [Hg (II)] in carcasses and guts and examine variation in this relationship among bays, multiple regression analysis was performed on log-transformed
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dry wt. concentrations with bay and [Hg (II)] as the dependent variables and [MMHg] as the independent variable. Carcass gut ratios (CGRs) were calculated by dividing Hg concentration in the carcass by Hg concentration in the gut for each Hg species. Hg concentrations in the carcasses were assumed to approximate steady state based on a previous finding that tadpoles in the laboratory attained steady state prior to 50 DPH (Unrine and Jagoe, 2004). This minimizes any potential confounding effects of temporal variation in bioaccumulation. Simple linear regression was used to test for a relationship between exposure concentration and CGR. Assimilation efficiency of MMHg relative to Hg (II) was estimated by dividing the ratio of MMHg to Hg (II) in the carcasses by the ratio of MMHg to Hg (II) in the guts as in Mason et al. (1996).
of hindlimbs (F1,175 Z 0.17, p Z 0.6837; Fig. 4). Significant variation among bays for gut Hg (II) concentration was also found (F9,175 Z 3.00, p Z 0.0023; Table 1, Fig. 2), but there was no significant effect of carcass mass (F1,175 Z 0.25, p Z 0.6199; Fig. 3) or presence/ absence of hindlimbs (F1,175 Z 1.66, p Z 0.1997; Fig. 4) on gut Hg (II) concentration. Methylmercury concentrations in carcasses ranged from 13 to 265 ng Hg/g dry wt. with an overall mean of 64 G 3 ng Hg/g dry wt. Inorganic mercury concentrations in carcasses ranged from 26 to 314 ng Hg/g dry wt. for Hg (II) with an overall mean of 120 G 5 ng Hg/g dry wt. (Table 1). There was significant variation in MMHg concentration in carcasses among bays (F9,175 Z 38.78, p ! 0.0001; Table 1; Fig. 2) with no significant effects due to carcass mass (F1,175 Z 0.13, p Z 0.7173; Fig. 3) or presence/absence of hindlimbs (F1,175 Z 1.61, p Z 0.2059; Fig. 4). Similarly, for Hg (II) significant variation among bays existed (F9,175 Z 19.09, p ! 0.0001; Table 1, Fig. 2) with no significant effect due to carcass mass (F1,175 Z 1.75, p Z 0.1874; Fig. 3) or presence/absence of hindlimbs (F1,175 Z 0.07, p Z 0.7983; Fig. 4).
3. Results 3.1. Mercury concentrations Concentrations of MMHg in guts ranged from 15 to 292 ng Hg/g dry wt. with an overall mean of 90 G 4 ng Hg/g dry wt. Concentrations of Hg (II) in guts ranged from 202 to 4125 ng Hg/g dry wt. with an overall mean of 1185 G 52 ng Hg/g dry wt. Significant variation existed in gut MMHg concentrations among bays (F9,175 Z 6.08, p ! 0.0001; Table 1, Fig. 2). However, there was no significant effect of carcass mass (F1,175 Z 0.83, p Z 0.3625; Fig. 3) or presence/absence
3.2. Correlation analyses Conductivity of water from the bays ranged from 36.1 to 58.9 mS/cm at 25 C; pH from 4.2 to 5.7; alkalinity from 0 to 6.4 mg/L; DOC from 3.0 to 97.1 mg/L (Table 2). Concentration of Hg (II) in the gut was negatively correlated with DOC in water (rs Z ÿ0.64, p Z 0.0479). A marginally significant positive correlation was also
Table 1 Methylmercury (MMHg) and inorganic mercury (Hg (II)) concentrations in Rana sphenocephala larvae (carcasses and guts) collected from 10 Carolina bays (isolated depression wetlands) from the United States Department of Energy’s Savannah River Site in South Carolina, USA including ratio of MMHg to Hg (II) in guts and carcasses as well as carcass gut ratio (CGR), and assimilation efficiency of MMHg relative to Hg (II) Bay nb a number 5 42 77 83 86 97 136 139 143 147 All bays
Mean carcass Mean carcass Carcass MMHgc Hg (II)c ratiod
15 90 (7) 20 100 (9) 22 37 (8) 15 89 (7) 12 154 (16) 27 23 (2) 24 39 (3) 8 57 (15) 28 45 (4) 15 35 (4) 186
60 (3)
218 72 73 165 150 82 60 112 110 170
(12) (7) (10) (17) (21) (7) (6) (15) (120) (14)
120 (5)
0.41 1.40 0.51 0.54 1.02 0.28 0.64 0.51 0.41 0.21 0.6 (0.1)
Mean gut Mean gut MMHgc Hg (II)c
Gut ratiod
Assimilation efficiency of MMHg MMHg relative to Hg (II)e CGRf
Hg (II) CGRf
62 102 63 99 45 65 115 33 72 92
0.10 0.12 0.06 0.07 0.06 0.08 0.10 0.03 0.08 0.10
4.03 11.31 8.28 7.84 16.54 3.28 6.14 19.29 5.08 2.02
0.4 0.1 0.1 0.1 0.2 0.1 0.1 0.1 0.1 0.2
(8) (16) (12) (20) (7) (5) (9) (6) (12) (9)
75 (4)
600 824 1024 1432 736 769 1100 1243 884 902
(67) (153) (202) (197) (57) (67) (154) (203) (141) (144)
951 (50)
0.10 (0.008) 8.38 (1.81)
1.5 1.0 0.6 0.9 3.4 0.4 0.3 1.7 0.6 0.4
1.1 (0.3) 0.1 (0.03)
Means are followed by standard errors of the mean in parentheses. a Numbers used to identify wetlands are from Schalles et al. (1989). b Number of larvae collected. c All mercury concentrations expressed in ng Hg/g dry wt. d Ratio of MMHg to Hg (II). e Ratio of MMHg to Hg (II) in carcasses divided by ratio of MMHg to Hg (II) in guts. f Carcass to gut ratio (CGR) calculated by dividing the mercury concentration in the carcass by the mercury concentration in the gut.
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A 350
ln [Hg] ng/g dry wt.
300
ng Hg/g dry wt.
MMHg Hg (II)
5.5 Hg (II) MMHg
250 200 150 100
5.0 4.5
b
4.0 a
3.5 3.0 2.5
50
2.0 0.0
0 86
5
83
147
139
42
143
77
97
0.2
136
0.4
0.6
0.8
1.0
1.2
gutted tadpole mass (g dry wt.)
bay number
B
2000 1800
ng Hg/g dry wt.
1600
Hg II MMHg
1400
Fig. 3. Natural log of methylmercury (MMHg) and inorganic mercury (Hg (II)) concentrations versus body mass in gutted carcasses of Rana sphenocephala larvae collected from 10 Carolina bays (isolated depression wetlands) on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA. Solid lines are least squares regression lines for MMHg (a) and Hg (II) (b).
1200 1000 800 600 400 200 0 83
139
136
77
147
143
42
97
86
5
bay number Fig. 2. Mercury concentrations (ng Hg/g dry wt.) for methylmercury (MMHg) and inorganic mercury (Hg (II)) in gutted tadpoles (A) collected from 10 Carolina bays on the U.S. Department of Energy Savannah River Site (SRS) in South Carolina, USA and corresponding mercury concentrations in guts (B). Numbers used to refer to bays are from Schalles et al. (1989). Error bars correspond to G1 standard error of the mean. Columns under the same horizontal bar at top are not significantly different from each other (type I experimentwise error rate Z 0.05).
(F1,183 Z 7.94, p Z 0.0054) indicating that significant variation in slopes existed among bays (Table 3). Calculated slopes ranged from ÿ0.03 to 0.91 but were only significantly different from zero in bays 5 (slope Z 0.88), 42 (slope Z 0.91), and 139 (slope Z 0.91). The ratio of MMHg to Hg (II) in carcasses was much higher than in guts ranging from 0.2 to 1.4. A linear relationship between ln [MMHg] and ln [Hg (II)] existed in carcasses where data from all bays were pooled (F1,183 Z 3.82, p Z 0.0522, r2 Z 0.33; Fig. 5B, Table 3) but there was no significant variation in slopes among bays (F1,183 Z 0.93, p Z 0.3360). 160 140 120 ng Hg/g dry wt.
found between pH and carcass [MMHg] (rs Z 0.60, p Z 0.0667). Other water chemistry variables were not well correlated with carcass or gut concentrations of MMHg or Hg (II) ( p O 0.11 in all cases). Mean MMHg, Hg (II), and THg carcass concentrations were not correlated with corresponding concentrations in guts (in all cases p ! 0.22).
100 80 60 40 20 0 HL
3.3. Mercury speciation The ratio of MMHg to Hg (II) in guts was small, varying from 0.03 in bay 139 to 0.12 in bay 42 with an overall average of 0.1 (Table 1). A linear relationship existed between ln [MMHg] and ln [Hg (II)] in guts (Fig. 5A, Table 3) when all bays were pooled (F1,187 Z 24.04, p ! 0.0001, r2 Z 0.16). However, there was a significant interaction between bay and [Hg (II)]
NO HL MMHg
HL
NO HL Hg (II)
Fig. 4. Mercury concentrations for Rana sphenocephala larvae with no hindlimbs or incompletely developed hindlimbs (NO HL) compared to larvae with completely developed hindlimbs (HL) for both methylmercury (MMHg) and inorganic mercury (Hg (II)). Number of larvae Z 90 for NO HL and 96 for HL Error bars correspond to G1 standard error of the mean. Larvae were collected from 10 Carolina bays (isolated depression wetlands) on the United States Department of Energy Savannah River Site (SRS) in Aiken and Barnwell Counties, SC, USA.
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J.M. Unrine et al. / Environmental Pollution 135 (2005) 245–253 Table 2 Conductivity, pH, alkalinity, and dissolved organic carbon concentration (DOC) from 10 Carolina bays (isolated depression wetlands) from the United States Department of Energy’s Savannah River Site in South Carolina, USA Bay numbera
pH
Conductivityb
Alkalinityc
DOCc
5 42 77 83 86 97 136 139 143 147
5.72 5.77 4.51 4.61 4.62 4.47 4.98 4.94 4.26 4.50
50.7 58.9 39.3 23.7 36.1 40.3 39.1 49.1 44.4 43.1
6.19 6.46 0.00 0.00 0.00 0.00 0.00 4.31 0.00 0.00
38.8 45.4 33.8 24.5 54.9 97.1 3.0 32.7 61.4 68.4
a b c
Numbers used to identify wetlands are from Schalles et al. (1989). mS/cm at 25 C. mg/L.
Table 3 Slope, intercept and coefficient of determination (r2) for the linear relationship between log of methylmercury concentration (MMHg) and inorganic mercury concentration (Hg (II)) in Rana sphenocephala larvae (carcasses and guts) collected from Carolina bays on the United States Department of Energy Savannah River Site in Aiken and Barnwell counties, SC, USA Baya
nb
5 42 77 83 86 97 136 139 143 147 All bays
Guts
Carcasses
Slope
Intercept
r2
Slope
Intercept
r2
15 20 22 15 12 27 24 8 28 15
0.88** 0.91** 0.24 0.55 0.77 0.61 ÿ0.03 0.91** 0.11 0.53
ÿ1.54 ÿ1.50 ÿ2.46 0.58 ÿ1.29 0.07 4.98** ÿ2.99 3.47 0.88
0.59 0.72 0.06 0.23 0.21 0.50 0.00 0.79 0.01 0.59
0.35 0.45* 0.47* 0.51* 0.65** 0.38* 0.57** 0.39 0.39** 0.12
2.65 2.70* 1.59 1.87 1.79* 1.46* 1.33 2.20 1.97** 2.90
0.08 0.22 0.24 0.33 0.68 0.19 0.32 0.08 0.25 0.01
186
0.41**
1.47
0.16
0.52**
1.53**
0.20
*Significantly different from zero at a Z 0.05, **significantly different from zero at a Z 0.01. a Numbers used to identify wetlands are from Schalles et al. (1989). b Number of larvae collected.
7
A
ln [MMHg]
6
5
4
3
2 5.0
5.5
6.0
6.5
7.0
7.5
8.0
8.5
ln [Hg (II)] 6
Carcass gut ratios were quite low ranging from 0.4 to 3.4 for MMHg and 0.1 to 0.4 for Hg (II). There was a statistically significant inverse relationship between gut concentration and CGR both for MMHg (F1,8 Z 5.56, p Z 0.0461, r2 Z 0.41) and Hg (II) (F1,8 Z 5.80, p Z 0.0426, r2 Z 0.42; Fig. 6). Slopes for MMHg and Hg (II) were similar (ÿ1.28 and ÿ1.39, respectively), however, the intercepts differed (5.14 and 7.37, respectively). The increase in ratio of MMHg to Hg (II) in carcasses versus guts, which is indicative of the assimilation efficiency of MMHg relative to Hg (II) (Mason et al., 1996), was quite variable and ranged from 2.02 to 19.29-fold (Table 1).
B 5 y = -1.23x + 5.15 r2 = 0.44
1
4
MMHg Hg II Linear (MMHg) Linear (Hg II)
0.5
3
0
ln CGR
ln [MMHg]
1.5
2
-0.5 y = -1.36x + 7.24 r2 = 0.43
-1 -1.5
1 3.0
3.5
4.0
4.5
5.0
5.5
6.0
-2
ln [Hg (II)] -2.5
Fig. 5. Relationship between log of inorganic mercury (Hg (II)) and methylmercury (MMHg) in the guts of Rana sphenocephala larvae collected from 10 Carolina bays on the United States Department of Energy Savannah River Site (SRS) in South Carolina (A). Solid line indicates least squares regression line, dashed lines correspond to 95% confidence intervals and dotted lines correspond to 95% prediction intervals. Relationship for mercury concentrations in corresponding gutted whole carcasses (B) is also shown.
-3 3
3.5
4
4.5
5
5.5
6
6.5
7
7.5
ln ng Hg/g dry wt. in gut
Fig. 6. Relationship between log of mercury concentration in the gut versus carcass gut ratio (CGR) calculated by dividing mean concentration in gutted larvae by mean concentration in guts for each of 10 Carolina bays sampled.
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J.M. Unrine et al. / Environmental Pollution 135 (2005) 245–253
4. Discussion These results support our previous conclusion (Unrine and Jagoe, 2004) that, in general, dietary THg exposures for tadpoles would range from 50 to 1600 ng Hg/g dry wt. in sites with no local anthropogenic or geologic sources of Hg. However, we predicted a range of average exposure concentration on the SRS of about 60 to 120 ng Hg/g dry wt. THg, which underestimated THg concentrations actually observed in the guts (599 to 1432 ng Hg/g dry wt.) (Unrine and Jagoe, 2004). The discrepancy between our predicted and observed Hg concentrations likely stems from differences in dietary components and the relative partitioning of Hg species within these components. Aufwuchs from artificial substrates used to estimate dietary Hg concentrations in our previous work (Unrine and Jagoe, 2004) probably differed from the diet of tadpoles in the bays both in organic composition (autotrophic, heterotrophic, and detrital components) and inorganic composition (sediment content and composition). The prediction from our previous work (Unrine and Jagoe, 2004) that the ratio of MMHg to Hg (II) in the tadpole diet would decrease with increasing THg concentration was supported by the observed relationship between ln [MMHg] and ln [Hg (II)]. In these plots, a slope !1 indicates that the ratio of MMHg to Hg (II) decreases with increasing THg concentration because ln [MMHg] does not increase as rapidly as ln [Hg (II)]. The slope was less than one in all bays, and not significantly different from zero in most bays. The overall slope for the pooled data in this experiment was 0.41, which agrees well with our predictions based upon aufwuchs samples (0.31). However, significant variation in slopes among individual bays (ÿ0.03 to 0.91) demonstrates that Hg speciation in the tadpole diet can vary greatly among bays. Such differences in slope may relate to differences in net methylation rate among bays and/or differences in dietary composition among bays. For example, in bays where the relationship between ln [Hg (II)] and ln [MMHg] had a large slope, a large proportion of the Hg (II) entering the system may have been methylated. Components of the tadpole diet may also differ in their capacities to concentrate MMHg and Hg (II) from the water column, which could explain differences in the relationship between Hg (II) and MMHg among bays. Considerable variation in assimilation efficiency of MMHg relative to Hg (II) probably existed among bays. In the laboratory (Unrine and Jagoe, 2004), we observed an approximately 5.7-fold increase in the ratio of MMHg to Hg (II) in tadpoles relative to experimental diets. Hg content in experimental diets was varied by combining proportions of control and Hg-enriched aufwuchs grown in experimental mesocosms; the mesocosms aufwuchs was primarily composed of filamentous
algae. Mason et al. (1996) showed that the assimilation efficiency of MMHg for zooplankton consuming phytoplankton was approximately 3.5–4.5 times greater than for Hg (II). They related this to the observation that percent of MMHg found in the cytoplasm of diatoms is approximately four times greater than percent Hg (II), and concluded that zooplankton assimilate Hg primarily from the cytoplasm of diatoms. In our previous work, the assimilation efficiency of MMHg relative to Hg (II) was probably reflective of differences in sub-cellular fractionation of MMHg and Hg (II) in the algal cells used as a source of Hg in the experimental diets. The present study shows that assimilation efficiency of MMHg relative to Hg (II) for amphibian larvae in the field can be quite variable. Although experimental diets in the laboratory (Unrine and Jagoe, 2004) may have had concentrations and speciation of Hg reflective of natural environments, relative assimilation efficiencies of MMHg and Hg (II) from the experimental diets were slightly different from what we observed in nature (this study). In both the laboratory (Unrine and Jagoe, 2004) and the field (current study), we found an inverse relationship between exposure concentration and BAF or CGR both for MMHg and Hg (II). A decrease in the rate of bioaccumulation with increasing dietary exposure concentration would tend to dampen the relationship between dietary concentration and bioaccumulated concentration. This at least partially explains the lack of correlation between Hg concentrations in carcasses and concentrations in guts. Differences in dietary composition among bays that resulted in differences in bioavailability could have also contributed to the lack of relationship. Carcass gut ratios observed in this study were similar to dietary bioaccumulation factors (BAFs) observed in the laboratory (Unrine and Jagoe, 2004) for Hg (II) but were slightly higher than laboratory BAFs for MMHg. Bioaccumulation factors for Hg are currently used under the assumption that they are independent of exposure concentration; however, the results of our work using BAFs and CGRs and the work of McGeer et al. (2003) concerning bioconcentration factors (BCFs) for Hg, show that predictions of Hg concentrations in biota based upon either BAFs, CGRs or BCFs are confounded by exposure concentration. An alternative approach might utilize a bioaccumulation factor that is corrected for exposure concentration using a slope factor derived from experimental data. Observed total mercury concentrations in guts in bay 83 exceeded dietary concentrations (approximately 1400 ng Hg/g dry wt.) at which adverse effects on development and decreased metamorphic success were observed in R. sphenocephala in the laboratory (Unrine et al., 2004). Bioaccumulated concentrations in larvae from the bays exceeded bioaccumulated concentrations (approximately 240 ng Hg/g dry wt.) at which effects
J.M. Unrine et al. / Environmental Pollution 135 (2005) 245–253
were observed in the laboratory (Unrine et al., 2004) in three bays (5, 83, and 86). This suggests that amphibian larvae inhabiting Carolina bays may be exposed to and bioaccumulate concentrations of Hg within the range of Hg concentrations at which changes in development and reduced metamorphic success might be observed. The conclusion that wildlife utilizing Carolina bays are potentially exposed to toxicologically significant quantities of mercury was reached in a previous investigation concerning dietary exposure in nestling wood storks (Brant et al., 2002). Carolina bays have been described as ‘‘islands’’ of species richness on the Atlantic Coastal Plain and their degradation due to draining could result in a major loss of biodiversity in the southeastern United States (Sharitz, 2003). Contamination by atmospheric mercury deposition should be further investigated to determine if it is an additional threat to the ecological integrity of these important ecosystems. 5. Conclusions Ranid larvae inhabiting relatively pristine Carolina bay wetlands may be exposed to and bioaccumulate toxicologically significant quantities of mercury. This study confirmed the predictions of our previous work (Unrine et al., 2004) that Hg concentrations in the diet of larval anuran amphibians would be as high as 1600 ng Hg/g dry wt. THg in sites with little or no local anthropogenic or geologic mercury input. It also confirmed that Hg is mostly present as Hg (II) in the diet, and that percent MMHg decreases with increasing THg concentration. Even if Hg concentrations and speciation in experimental diets may reflect what is observed in nature, the relative bioavailabilities of MMHg and Hg (II) may not. Future studies should focus on the relationship between composition of gut contents (autotrophic, heterotrophic, detrital, and inorganic components) with MMHg and Hg (II) concentrations as well as the influence of dietary composition on relative bioavailabilities of MMHg and Hg (II). Finally, further investigation into the potential risks of Hg exposure to amphibians and other biota inhabiting Carolina bays is warranted. Acknowledgements This research was supported by the Environmental Remediation Sciences Division of the Office of Biological and Environmental Research, U.S. Department of Energy, through Financial Assistance Award No. DEFC09-96-SR18546 to the University of Georgia Research Foundation. J. Unrine was supported by a doctoral assistantship provided by the Savannah River Ecology Laboratory while conducting this research. Earlier versions of the manuscript benefited from the comments
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