Environ. Sci. Technol. 1994, 28, 2366-2371
Sunlight4 nduced Formation of Dissolved Gaseous Mercury in Lake Waters Marc Amyot,"lt Greg Mierle,s David R. S. Lean,$ and Donald J. McQueentlll
Department of Biology, Faculty of Pure and Applied Science, York University, 4700 Keele Street, North York, Ontario, Canada, M3J 1P3, Ministry of the Environment, Dorset Research Centre, Bellwood Acres Road, P.O. Box 39, Dorset, Ontario, Canada, POA IEO, and National Water Research Institute, Burlington, Ontario, Canada, L7R 4A6 ~
Formation of dissolved gaseous mercury (DGM) was measured in lake water incubated at midday in Teflon bottles. DGM production was photoinduced as transparent bottles yielded DGM concentrations that were 2.4-9 times higher than dark controls. These results provide the first experimental evidence obtained in the field of a direct link between solar radiation and DGM production. A positive relationship was found between photoinduced DGM production at different times of the year and incident radiation. Removal of UVBlight or addition of hydrogen peroxide during incubation did not result in significant changes in DGM levels. A diel pattern in DGM production was observed, and a depth profile of DGM revealed that most of the production was occuring in the epilimnion. It is concluded that sunlight plays a key role in DGM production in the epilimnion and may alter the fluxes of Hg in aquatic ecosystems. Introduction
Despite the fact that anthropogenic emissions of Hg from chloralkali plants have been drastically reduced since the 1960s and early 1970s (1-3), Hg is still spreading in the environment. In particular, elevated concentrations of mercury in freshwater fish is currently a problem in many northern countries. Even in remote lakes, fishes are regularly found with levels of mercury that often exceed American and Scandinavian guidelines for complete consumption restriction (0.5-1.0 ppm) (4-6). Mercury concentration in any water body is due to the balance between inputs (deposition and runoff) and outputs (volatilization, outflow, and sedimentation). Hg deposition is controlled by the oxidation of HgO to Hg(I1) in the air followed by dissolution in atmospheric water (wet deposition by rain) or adsorption to particulates (dry deposition). Due to its low solubility and favorable Henry's law constant, HgO is the principal form contributing to volatilization and dissolved gaseous mercury (DGM) concentration (7,8). Evasional fluxes of Hg from remote lakes can represent from 7 96 (7)-95 96 (9)of the estimated atmospheric Hg deposition to the lake. Rates of volatilization are controlled by rates of formation of HgO which are a function of biotic and abiotic processes. Abiotic reduction in water of Hg(I1) into HgO can be initiated by humic substances (10). Turner et al. (11) suggested that this abiotic reduction could account for as much as 10-7096 of the volatile Hg released from some contaminated streams. It is assumed that intermediates like semiquinones, which are present in humic substances, are involved. There have been some indica-
* Address correspondence to this author; E-mail address:
[email protected]. + York University. 3 Ministry of the Environment. 8 National Water Research Institute. 11 E-mail address: FS300006@~01.yorku.ca 2366
Environ. Sci. Technol., Vol. 28, No. 13, 1994
Table 1. Abbreviations DGM UVA UVB
PAR
DGM, [DGMla [DGMlb RFD
dissolved gaseous mercury WV radiations between 320 and 400 nm UV radiations between 280 and 320 nm photosynthetically active radiation between 400 and 700 nm (kJ m-2 h-1) photoinduced DGM production rate (fMh-l) DGM concentration in fM measured in transparent bottles DGM concentration in fM measured in black bottles radiant flux densities (kJ m-2 h-l)
tions that, under laboratory conditions, the rate of this abiotic reduction can be photochemically enhanced (12). Among the abiotic photochemical pathways that may be involved, Schroeder et al. (13) have suggested that hydrogen peroxide (HzOz), mainly originating from UVinduced transformations of dissolved organic matter in surface waters (14), could either act as a reducing or oxidizing agent for Hg, depending on the pH. The following equations have been proposed:
+ 2H+ + Hgo + 20H- + Hg2+
H,02 H,O,
-
-
2H20 + Hg2+
0, + 2H20+ Hgo
(1) (2)
Some biotic processes leading to the reduction of Hg have also been identified. The primary mercury-detoxifying mechanism used by mercury-resistant bacteria is the reduction of mercuric ion to volatile HgO (15). Photosynthesis may also produce Hg(I1) reductants, the highest rates of Hgo production having been observed during algal blooms (7). While photochemical and photobiological processes are thought to alter the rate of volatile mercury production in lakes, there is no clear experimental evidence obtained under field conditions of a direct link between solar radiation and in situ DGM production. In this context, we determined the effects of sunlight and hydrogen peroxide on dissolved gaseous Hg (DGM) production in lake water (see Table 1 for a list of abbreviations). The effect of UVBlight (280-320 nm) was also assessed because many photochemical processes are triggered by this very energetic form of radiation. With the recent depletion of the ozone layer and the resulting increase in UVB light, these processes could increase in importance and could significantly alter the biogeochemical cycle of some compounds. Also, the presence of a diel pattern of DGM production in surface water was investigated. Finally, depth profiles of DGM concentrations were also obtained in order to identify the site of maximum DGM production in the water column. Experimental Section
Water samples were taken in the pelagic zone of Ranger Lake (45'09' N, 78'51' W), a small (11.2 ha, 13 m deep) acidic (pH 6.1) oligotrophic (total phosphorus 6 pg L-l) 0013-936X/94/0928-2366$04.50/0
0 1994 American Chemical Society
softwater lake and Jack Lake (44'41' N, 78'02'W), alarge oligotrophic (total phosphorus 12 pg L-l) headwater lake of neutral pH (pH 7.2). Samples were collected from a plastic canoe by filling 1-L Teflon bottles with surface water. These bottles, as all the glasswaresubsequently in contact with the samples, were acid-washed and thoroughly rinsed with doubledistilled water. In addition, the Teflon bottles were rinsed with surface water downwind from the sampling site. Polyethylene gloves were worn at all times. To assess the effect of sunlight on DGM production, experiments were conducted in Ranger Lake on August 21, September 8, and November 13, 1993. Different combinations of treatments were applied to the samples: (1)absence of light (black bottles); (2) presence of sunlight (transparent bottles); (3)absence Of UVBlight (transparent bottles wrapped in Mylar). To determine whether the observed photoinducedDGM production could be mainly attributed to the reduction by hydrogen peroxide of Hg(I1) to the volatile species HgO (see eq 2), experiments were conducted in both Ranger Lake and Jack Lake. Half of the transparent and black bottles were spiked with 500 nM H202 (Ranger Lake: August 27) or 5000 nM H202 (Ranger Lake: November 14; Jack Lake November 15). These levels of H2Oz are higher than natural levels, which usually range from 60 to 180 nM at the surface of these lakes at midday (16). After sampling, the bottles were incubated for 4.5 h, at midday, on a platform (Ranger Lake) or a small island (Jack Lake). During incubation, the bottles were placed horizontally with their caps pointing north. The samples were maintained half submerged in a bath of lake water. Temperature of the water bath was kept close to the temperature of surface waters. An experiment was conducted on September 7 in the pelagic zone of Ranger Lake to determine diel changes in DGM concentrations in surface water. Samples were collected in Ranger Lake, a t 6-h intervals using transparent bottles. Since changes in wind speed can alter the volatilization rate of Hg and the mixing of surface water so that diel patterns in dissolved [HgOI cannot be observed (17),samples were incubated for a short period (2 h) after each sampling. Note that, due to time constraints, no black controls were included during these incubations. Consequently, the rates obtained after incubation are not true DGM production rates because they include the amount of DGM already present in the water before incubation. For depth profiles of DGM and total Hg concentrations, water samples were taken with a peristaltic pump. Freshly acid-cleaned Teflon and silicon tubing were used to collect samples from 0,2,4,6,8,10, and 12 m. The depth of the sampling site was 13 m. Oxygen and temperature were measured using a Yellow Spring Instrument. The pH was measured using a pH-meter, Fisher Tetramatic Model 150. At the end of each experiment, the bottles were bagged and stored in a dark cooler for shipment. Upon arrival at the clean laboratory, the samples were processed within 12 h. Total mercury analysis was based on a method described by Gill and Bruland (18). Approximately 100 mL of sample was placed in a water-jacketed sparging vessel at 70 'C, and 5 mL of 4 M NaOH and 1mL of 1% NaBH4 (w/v) were added through an injection port. The sample was purged for 2 min with Hg-free air at a flow rate of approximately 300 mL min-l, and the released Hg vapor
was collected on a 4 X 15 mm (diameter X length) goldcoated sand column in a 6 mm 0.d. quartz tube. The trapped Hg was thermally desorbed by heating with a 25-turn, 22-gauge Chrome1 A wire energized at 18 V for 15 s and flushed into an LDC mercury monitor Model 1255 for quantification at a flow rate of 30 mL min-'. The detection limit (three times the standard deviation of the blanks) was 150 fM.The relative precision at concentrations of 5-10 pM was about 1.5%. For DGM analysis, 500 mL of water was decanted slowly into a 1-L bubbler and sparged for 15 min with Hg-free argon at a flow of 1L min-l. The volatile Hg compounds were trapped on a gold-coated sand column. The bubbler was then bypassed to allow dry gas to pass through the column for an additional 10 min (flow 500 mL min-l). The Hg trapped on the column was then thermally desorbed, and readings were made at low flow (less than 100 mL min-1) with a LDC mercury monitor Model 1255. The limit of detection for the complete procedure (including sampling, storage, and analysis), defined as three times the standard deviation on replicates of lake water samples with low DGM levels (48 f 9 fM,N = 6), was 27 fM.The limit of detection of the analytical procedure, defined as three times the mean standard deviation on the procedural blanks, was 11 fM. In the incubation experiments, black bottles were used as controls. To assess the temporal stability of DGM levels in black bottles, Ranger Lake surface water samples were taken and kept in the dark on two dates (September 1and November 16). In September, four replicates were analyzed at 3-h intervals, for a total length of 13.5 h, which was representative of the longest delays encountered between sampling and analysis. In November, six replicates were analyzed at 40-min intervals, for a total length of 4 h, which simulated what happens during the incubation of black bottles in the field. DGM concentrations did not increase or decrease in a consistent way during both experiments. Coefficients of variation in DGM concentrations were 18% (49 f 9 fM;N = 6) for the short experiment and 12% (179 f 22 fM;N = 4) for the long one. We concluded that there is no detectable DGM production in the dark under our experimental conditions. The absorption of light by the Teflon bottles was determined by comparingspectral irradiance (280-800 nm) at a depth of 10 cm in lake water with and without the presence of a Teflon bottle covering the sensor. We used an Optronics 752 spectroradiometer connected with an underwater optical sphere with a 3-m fiber optic cable. During our evaluation, we compared values collected every 10 min on an overcast day (December 3, 1993). Values were similar with Teflon bottles absorbing less than 2.5%. An example is shown in Figure 1. The wavelength cutoff of Mylar was also assessed with the spectroradiometer. Cutoff values ranged from 315 to 323 nm. Light measurements were obtained from the weather station of the Ontario Ministry of the Environment and Energy (Dorset Research Centre). Total incident radiation (A = 285-2800 nm) and photosynthetically active radiation (PAR, X = 400-700 nm) were measured with an Eppley precision pyranometer model PSP, Results and Discussion
Effect of Sunlight and UVB on DGM Production. Samples incubated in transparent bottles yielded DGM Envlron. Scl. Technot., Vot. 28, No. 13, 1994
2367
...
. . . . .. . With bottie
Without bottle
;...:
.. ,
.
I
i 200 300
400 500 600 700 800 900 Waveiength (nm)
Flgure 1. Spectral energy distribution of the global radiation on Dec 3, 1993,at Jacks Lake. Measurements were taken with the sensor: (1)in a Teflon bottle filled with lake water (at 15:lO);(2)under lake water, at a depth equal to the diameter of a bottle (at 15:20).
1000
,-.
800
z
Vylar 9iack ;3 Transparent
v
v)
s
.-
+
600
These results indicate that sunlight induces DGM production in unfiltered lake water. When compared to dark DGM production, photoinduced DGM production is quantitatively the only process of significance under midday conditions. These results provide the first experimental evidence obtained under field conditions of a direct link between solar radiation and in situ DGM production. Such a link has been suggested by others (17, 19).But confounding factors (e.g., variations in temperature, wind speed, and loss of DGM by volatilization) were not kept constant. Consequently, a direct effect of sunlight could only be hypothesized. Samples in bottles wrapped in Mylar (treatment without UVBlight) did not yield DGM concentrations significantly different (t-test,p > 0.05; N = 3 per treatment) from those in transparent unwrapped bottles (Figure 2). Thus, most of the observed DGM production was induced by other wavelengths. However, these results do not rule out a potential contribution of UVB light on Hg volatilization from lakes, since our experiments were conducted during periods of relatively low levels of UVg radiation. It is possible that at other times of the year (e.g., spring and summer) or a t other locations (near the poles or in the equatorial zone),UVBlight could have a significant impact on DGM production.
Relationship between DGM Production Rate and Incident Radiation. Using eq 3, photoinduced DGM production rates were calculated for six dates (August 19, 21, and 27; September 8; November 13 and 14,1993) and were plotted against corresponding total radiant flux densities (RFD, in kJ m-2 h-l, = 285-2800 nm). The distribution of points (Figure 3) can be described by the following quadratic equation (corrected R = 0.995, p < 0.001):
F Y 7
z0 5 400 0
from 182 fM h-1 in August to 77 fMh-l in September and 17 fM h-' in November.
I
3 u n
200
DGM, = 1.412
+ 0.109 (RFD) - 1.58 X
(RFD)' (4)
0 Aug21
SeptB
Nov13
Dates Flgure 2. Dissolvedgaseous mercury (DGM) concentrationsin surface water samples from Ranger Lake, incubated during 4.5 h at midday on Aug 21, Sept 8,and Nov 13, 1993. Samples were incubated in transparent bottles, black bottles, or transparent bottles wrapped in Mylar. Error bars correspond to the standard deviation of three replicates.
concentrations that were 2.4-8.9 times higher (t-tests, p < 0.001;N = 3 per treatment) than those kept in black bottles (Figure 2). DGM production rate was calculated by the following equation: DGM,= ([DGMl,- [DGMIb)/t
(3)
where DGM, is the photoinduced DGM production rate, [DGMI. and [DGMlb are the DGM concentration measured in transparent and black bottles, respectively, and t is the incubation time. Calculated DGM production rate at midday displayed strong seasonal differences, ranging 2388
Environ. Sci. Technol., Vol. 28, No. 13, 1994
If only photosynthetically active radiant flux densities (PAR, in kJ m-2 h-l, X = 400-700 nm) are considered, the following equation is obtained (corrected R = 0.995, p < 0.001):
DGM, = 0.289 + 0.200 (PAR) - 5.02 X
(PAR)2 (5)
The strength of these relationships suggests that, for a given lake, the light budget of this lake could be used as an efficient and inexpensive means to estimate DGM production in surface waters. The nonlinearity of the function at high light intensities may be due to (1) a limitation of available substrate, i.e., mainly Hg(I1); (2) a saturation of the chromophores involved; or (3) an inhibition of photosynthetic processes. Other covariates of solar radiation, such as day of year and water tem-
200
I
0" 0
1000
2000
3000
Total radiant flux densities (kJ rV2 hr? mure 3. RBlanonshipbetweenthetotairadiantfluxdens~esesnmated for a 4.54 perkd at midday for different days and the corresponding photoinduced DGM production rate in surface waters from Ranger Lake.
1
pH = 6.16
H Black + H,O,
700 -
--2 2 -
600
-
500
-
400
-
300
-
Transparent + HO ,, Black 0 Transparent
pH
=
761
200 100
0
1
L Aug27
Nov 14
Ranger L.
Nov15 -
Jack L
Flgure4. Dissohredgaseowmercury(ffiM)concentratlansin surface water samples from Ranger Lake and Jacks Lake, incubated during 4.5 h at midday. Samples were incubated in transparent and black bottles. On Aug 27, 1993, hall of the samples were spiked wkh 500 nM H2O2.On Nov 14 and 15, 1993. half of the samples were spiked wim 5000 nM H202.Mean pH of the samples are shown for each date. Enor bars correspond to the standard deviation on two replicates. The asterisk indicates the presence 01 only one replicate.
perature, explained a smaller portion of the observed variation in DGM,, yielding adjusted R2of 0.79 (N = 6; p = 0.011) and 0.76 (N = 6;p = 0.015),respectively. Effect of Hydrogen Peroxide on DGM Production. DGM levels in the transparent bottles were 1.5-4.3 times higher than those in black bottles, suggesting strong sunlight-induced DGM production (Figure 4). If Hg reduction by HzOz were the main process for this DGM production, we wouldexpect black and transparent bottles spiked with HzOzto show significant increases in DGM levels after incubation when compared to unspiked treatments.
Ononehand,onAugust27 (RangerLake)andNovember 15 (Jack Lake), spiked samples yielded mean DGM concentrations that were slightly higher than their unspiked counterparts, suggestingthat HzOzreduced a small portion of the available Hg(I1) (Figure 4). In these two cases,weconcludethatthereductionofHg(II)byhydrogen peroxide does not significantly contribute to the observed photoinduced DGM production. On the other hand, on November 14 (Ranger Lake), spiked samplesin transparent bottles yielded DGM levels 33% lower than those unspiked. Here HzOz acted as an oxidizing agent (see eq l), decreasing the level of volatile Hg (mainly H e ) . No similar decrease was observed in the black bottle after spiking, probably because the DGM levelswere already very low. The oxidizing action of H,Oz in transparent bottles is probably promoted by the low pH of the water (pH = 5.77)during this experiment. In fact, according to Schroeder et al. (13), the Nernst equation applied to eqs 1and 2 predicts that the oxidation and reduction processes will be balance at pH = 5.5 in distilled water (with oxidation predominating below 5.5 and reduction above that value). However, experimental observationsmadeby Brosset (20)put the transition point somewhere between 5.9 and 7.9 for a solution of 0.7 M NaC1. When considering lake waters, it is likely that this transition point will be different for different lakes, and even for different depths within a lake, depending on variations in water chemistry. For Ranger Lake, it seems that the transition point in pelagic suface waters is somewhere between pH 5.77 and pH 6.16. Since the addition of high levels of H202 did not change DGM production, it is concluded that the photochemical pathways involving the reduction of Hg by H2O2 are probably ofsecondary importance in theselakes. However, rainwater is known to contain high levels of H202. For instance, Cooper and Lean (16) and Lean et al. (21)have reported values up to 55 000 nM at Jack Lake and Lake Erie. Consequently, it cannot be ruled out that, during stormevents,HzOzmaytemporarilyaltertheDGM budget in the first meter of lake water. Diel P a t t e r n s of DGM Production. A clear diel pattern in DGM levels in surface waters was observed, with the highest DGM levels being observed at noon and the lowest ones at 600 AM,just before sunrise (Figure 5). The energy fluence pattern for total incident radiation followed asimilarpattern. Discrepancies between the two patterns are probably due to the fact that we measured total DGM concentrations, including DGM present at the start of the incubation, instead of DGM production rates. For instance, at sunset, the decrease in DGM levels lagged behind decreases in ambiant light. Temperatures in the water column were quite constant, reinforcing the hypothesis that DGM production is mainly caused by a sunlight-induced process and not by a simple effect of temperature on bacterial activity or chemical reactions. ThedielpatterninDGMlevelsinsurfacewaterspredicta that a similar pattern would be observed in the levels of Hg above lake surfaces. Indeed, Schroeder et al. (19) have reported the presence of a diel cycle for Hg emissions from five lakes, with daytime volatilization rates significantly larger than nighttime rates. Our results suggest that these variations in Hg emissions are primarly due to photoinduced DGM production in surface waters. Depth Profile of DGM Concentrations. DGM levels of up to 256 fMwere found in the epilimnion, followed by Envkon. Sol. Tschnol., Vol. 28. No. 13, 1994 23w
01
t
1
& I m
73
+
130
I
0
-
300
3
'2
'8 24 Tim
6
12
0'
d ~ y
16
2d
12
Flgure 5. Diurnal variations in surface water DGM levels (after a 2-h incubation). water temperature, and incident radiation for Sep 7 and 8,1993,at Ranger Lake. Error bars correspondto thestandarddeviation of two replicates.
,E-. v
5 a a,
n 0 IDGMl 0 Temperature
12
C IDGVl (x
6
12
18
24
30
10 fV) TeTperature ("C) Oxygen ( r g L-')
Flgure 6. Depth profiles for dissolved gaseous mercury (DGM) levels, temperature, and dissolved oxygen on Aug 22, 1993,in the pelagic zone of Ranger Lake.
an abrupt decrease in the metalimnion with a minimum of 59 fM at 6 m and a small increase in the hypolimnion to around 100 fM (Figure 6 ) . This pattern suggests that most of the DGM production occurs in the upper part of the water column. The fact that lower levels were measured at 0 m than at 2 m may be due to a loss of Hg by volatilization in the first meter of water. It is likely that high DGM levels in the upper part of the water column are the result of the same photoinduced processes that produced high DGM levels during our incubation experiments. In particular, Mason et al. (22) have suggested that photosynthetic organisms (mainly cyanobacteria and smaller picoplankton) could reduce Hg(I1) by cell surface enzymatic processes. If these photobiological processes are indeed important, they may explain in part the lower levels of DGM encountered at the surface, where photo2370
~
0
Environ. Sci. Technol., Vol. 28, No. 13, 1994
I
5
10
I
I
15
20
Total l g (pM) and Oxygen (mg L-') Flgure 7. Depth profiles for total mercury levels, temperature, and dissolved oxygen on Oct 5, 1993,in the pelagic zone of Ranger Lake.
synthetic inhibition often occurs. The slight increase in DGM concentrations in the hypolimnion is probably caused by the action of microorganisms in the sediments. Many aerobic and facultatively anaerobic bacteria are known to reduce ionic Hg(I1) to volatile HgO as a detoxification mechanism (15, 23, 24). A depth profile similar to ours was published by Vandal e t al. (17) for a circumneutral dimictic seepage lake in Wisconsin. The surface was depleted in HgO when compared to the epilimnion. In addition, epilimnetic HgO levels exceeded metalimnetic and hypolimnetic levels, suggesting that the epilimnion was the region of maximum HgO production. The range of DGM concentrations were similar to those reported here. If this kind of depth profile is typical of what happens in most boreal forested lakes, photochemical and/or photobiological DGM production in the epilimnion could play a key role in the removal of Hg from lakes. Depth Profile of Total Mercury Concentrations. Total mercury levels were relatively constant through the epilimnion, ranging from 8.4 to 8.7 pM (Figure 7). These levels doubled in the metalimnion and reached a maximum of 21 pM in the hypolimnion. This hypolimnetic increase may be attributed to the downward transport and recycling of particulate Hg prior to incorporation in the sediments (25). Alternatively, Hg may be released from the sediments as a result of the low redox potential usually found in 02-depleted waters. This latter mechanism would also favor the abiotic reduction of Hg and can partly explain the second peak in DGM levels found in the hypolimnion (Figure 6). Conclusion Sunlight had a direct and quantitatively important effect on DGM production in surface waters from lakes. This effect was mainly driven by biological or photochemical processes induced by visible and UVA light, with UVB light responsible for less than 25 % of the DGM production. Furthermore, the presence of hydrogen peroxide did not play a significant role. A clear diel pattern was found which paralleled that for total solar radiation. Most DGM
Vandal, G. M.; Mason, R. P.; Fitzgerald, F. Water Air Soil
production occurred in the epilimnion. We hypothesize that the primary process for DGM production in lakes is the biological or photochemical reduction of Hg in the epilimnion by visible light or UVA light. A possible mechanism would be the cell surface enzymatic reduction of Hg by algae (22). According to the Hg(I1) substrate hypothesis (261, aquatic biological and chemical production processes for volatile HgO and highly bioavailable CH3Hg are in competition with one another for the reactant Hg(I1). In this context, lakes with limnological conditions favoring Hgo production (and thus DGM production) are less likely to have elevated levels of Hg in their fish stock (26). We suggest that the densities and composition of photosynthetic epilimnetic communities may be of major importance in the understanding of Hg accumulation in fish from boreal forested lakes. For instance, oligotrophiclakes have been shown to be more sensitive to Hg contamination in fish (27). It is usually believed that this is a result of an increased biomagnification along the food chain caused by a decrease in the biodilution effect (28). However, our results suggest that this Hg contamination in oligotrophic lakes may be partly due to low DGM production. Also, we hypothesize that lakes of similar trophy and atmospheric inputs but having different light budget (because of latitude or climate) could display differences in the Hg levels of their fish populations as a result of differences in DGM production.
Pollut. 1991, 56, 791-803.
Kim, P. J.; Fitzgerald, W. F. Science 1986, 231, 1131. Xiao, 2. F.; Munthe, J.; Schroeder, W. H.; Lindqvist, 0. Tellus 1991,43B (3), 267-279.
Alberts, J. J.; Schindler, J. E.; Miller, R. W. Science 1974, 184,895-897.
Turner, R. R.; VandenBrook, A. J.; Barkay, T.; Elwood, J. W. Proceedings of the International Conferenceon Heavy Metals in the Environment, Geneva; CEP Consultants Ltd.: Edinburgh, 1989; p 353. Allard, B.; Arsenie, I. Water Air Soil Pollut. 1991,56,457464.
Schroeder, W. H.; Yarwood, G.; Niki, H. Water Air Soil Pollut. 1991, 56, 653-666.
Cooper,W. J.; Shao, C.; Lean, D. R. S.; Gordon, A. S.; Scully, F. E. In Environmental Chemistry of Lakes and Reservoirs; Baker, L. A., Ed.; Advances in Chemistry Series; American Chemical Society: Washington, DC, 1994; pp 391-422. Steffan,R. J.;Korthals,E. L.; Winfrey,M. R.App1. Environ. Microbiol. 1988, 54, 2003-2009.
Cooper,W. J.;Lean,D. R. S.Encyclopediaof Earth Science; Academic Press: New York, 1994 (in press). Vandal, G. M.; Fitzgerald, W. F.; Lamborg, C. H.; Rolfhus, K. R. In Proceedings of the International Conference on Heavy Metals in the Environment, Toronto; CEP Consultants Ltd.: Edinburgh, 1993; Vol2, pp 297-299. Gill, G.; Bruland,K. Environ. Sci. Technol. 1990,24,13921400.
Schroeder,W.;Lindqvist,0.;Munthe,J.; Xiao, 2. Sci. Total Environ. 1992, 125, 47-66. Brosset, C. Water Air Soil Pollut. 1987, 34, 145-166. Lean, D. R. S.; Cooper, W. J.; Pick, F. R. In Aquatic and Surface Photochemistry; Zepp, R. G., Crosby D. G., Eds.;
Acknowledgments
We would like to thank N. D. Yan and two anonymous reviewers for constructive criticisms. This research was supported by Environment Canada's Green Pian Program for evaluating aquatic impacts of UVB light, an NSERC Operating Grant to D.R.S.L. and D.J.M., and an NSERC postgraduate scholarship to M.A. Equipment and logistic support was provided by Ontario Ministry of Environment and Energy and by the National Water Research Institute of Environment Canada.
Lewis Publishers: Ann Arbor, MI, 1993; pp 207-214. Mason,R. P.;Morel,F. M. M.; Hemond,H. F. InProceedings of the International Conference on Heavy Metals in the Environment, Toronto;CEP ConsultantsLtd.: Edinburgh, 1993; Vol. 2, pp 293-296. Lovley, D. R. Annu. Rev. Microbiol. 1993, 47, 263-290. Barkay, T.; Turner, R. R.; VandenBrook, A.; Liebert, C. Microb. Ecol. 1991,21, 151-161. Hurley, J. P.; Watras, C. J.; Bloom, N. S. Water Air Soil Pollut. 1991, 56, 543-551. Fitzgerald, W. F. In Proceedings of the International Conferenceon Heavy Metals in the Environment, Toronto;
Literature Cited
Parks, J. W.; Lutz, A.; Sutton, J. A. Can. J.Fish. Aquat.
CEP Consultants Ltd.: Edinburgh, 1993; Vol. 1, pp 320-
Sci. 1989, 46, 2184-2202. Martinelli, L. A.; Ferreira, J. R.; Forsberg, B. R.; Victor, R. L. Ambio 1988, 17 (4), 252-254. Nriagu, J. 0. Science 1989, 338, 4749.
Meili, M. Comprehensive Summaries of Uppsala Dissertations from the Faculty of Science, Acta Universitatis Upsaliensis. 1991, Vol. 336, 36 pp. Winfrey, M. R.; Rudd, J. W. M. Environ. Toxicol. Chem. 1990, 9, 853-869.
323.
Lindqvist, 0.;Johansson, K.; Aastrup, M.; Andersson, A.; Bringmark, L.; Hovenius, G.; Hakanson, L.; Iverfeldt, A.; Meili, M.; Timm, B. Water Air Soil Pollut. 1991,55,1-261. Meili, M. Water Air Soil Pollut. 1991, 56, 333-348. Received for review April 18, 1994. Revised manuscript received August 29, 1994. Accepted September 14, 1994."
Hakanson, L.; Nilsson, A.; Andersson, T. Enuiron. Pollut. 1988,49, 145-162.
@
Abstractpublishedin Advance ACSAbstracts,October15,1994.
Environ. Scl. Technol., Vol. 28, No. 13, 1994
2371
