Environ. Sci. Technol. 1990, 24, 1413-1418
Environmental Factors Affecting the Production of Peptide Toxins in Floating Scums of the Cyanobacterium Microcystis aeruginosa in a Hypertrophic African Reservoir Richard J. Wicks**+and Pleter G. Thiel$ Division of Water Technology, CSIR, P.O. Box 395,Pretoria 0001, South Africa and Research Institute for Nutritional Diseases, South African Medical Research Council, P.O. Box 70, Tygerberg 7505, South Africa
The presence of six cyclic, heptapeptide toxins in floating scums of the cyanobacterium Microcystis aeruginosa in Hartbeespoort Dam, South Africa, was monitored over 2.5 years. Separation of the six peptide toxins was achieved by reversed-phase high-pressure liquid chromatography. The six toxins were either not detectable or in very low concentration during the winter (May-August), reaching maximum concentrations during the summer. Combined concentrations of four of the toxins ranged from 5 to 415 wg (g of dry scum)-' and were strongly, positively correlated to primary production per unit of chlorophyll a , solar radiation, surface water temperature, pH, and percent oxygen saturation (r = 0.52-0.67,n = 16-20,p < 0.001)and weakly, negatively correlated to surface chlorophyll a and orthophosphate concentrations (r = -0.33, n = 16-20,p < 0.05).No strong relationships were found between total toxin concentrations in scum samples and surface water organic and inorganic nutrient concentrations. The data from the subtropical reservoir indicated that the specific rate of photosynthesis of M . aeruginosa together with several environmental factors is closely coupled to the concentrations of peptide toxins in this cyanobacterium.
vironmental factors and the presence of more than one strain of M . aeruginosa (17, 18). Defects in the rather crude bioassay methodology may also be responsible for some of the observed variation (17). Only very recently has there been available an analytical procedure capable of detecting and quantifying the individual M . aeruginosa toxins in cyanobacterial cultures and natural blooms in order to assess their potential threat to human and animal health (11,19). A number of laboratory experiments investigating the influence of environmental conditions on the toxicity of M. aeruginosa cultures have been reported. Environmental factors investigated were light intensity, temperature, pH, aeration, culture age, and nutrients (7, 17, 20-30). However, no studies have been published concerning the effect of environmental factors on the toxicity of M . aeruginosa growing in lakes or reservoirs using an analytical technique that can detect and quantify individual M . aeruginosa toxins. We report here a quantitative analysis of the toxin content of M . aeruginosa buoyant scums determined by high-pressure liquid chromatography (HPLC) in relation to various environmental factors. The scum samples were collected during 2.5 years when scum was present part of the time in hypertrophic Hartbeespoort Dam, South Africa (31). This reservoir is enriched with nitrogen and phosphorus (32) and is characterized by high, variable chlorophyll a concentrations [mainly M. aeruginosa (33)], which can reach -3000 mg m-3, and large (1-2ha), thick (up to 1 m), buoyant cyanobacterial hyperscums (34).
Introduction Environmental conditions that favor cyanobacterial (blue-green algal) bloom formation include (A) moderate-to-high levels of nutrients especially phosphorus and nitrate or ammonia, (B)water temperatures between 15 and 30 "C, and (C) a pH of between 6 and 9 or higher ( I ) . Experimental Details The cyanobacterium Microcystis aeruginosa occurs worldwide, the toxic form having been reported in the The study was carried out at a permanent station in the United States, Canada, United Kingdom, Norway, South main basin of Hartbeespoort Dam (33),except solar raAfrica, Australia, China, and Japan ( 2 , 3 ) , and has been diation was measured 7 km southeast of the station with connected to animal deaths and even human illness (I, a Kipp solarimeter (range 320-1500 nm) and M.aerugi4-6). The toxins involved have been designated as fast nosa scums were sampled near the dam wall up to 1 km death factors, microcystins, aeruginosins, and recently as from the permanent station. M . aeruginosa scum samples, cyanoginosins (7). They are peptide hepatotoxins (1). The when present, were collected usually two or three times structures of five different M . aeruginosa toxins were per month between December 1984 and May 1987. Only elucidated only recently (8,9)and were found to be cyclic recently formed, green regions of the scum were sampled. heptapeptides comprised of five amino acids common to Scum samples were stored at 4 "C for up to 24 h and all toxin variants and two variable L-amino acids. Their freeze-dried; the dry material was stored at 4 "C until general structure is cyclo(D-Ala-L-X-erythro-P-methy1-D- analyzed. Samples collected during the same month were isoAsp-L-Y-Adda-D-isoGlu-N-methyldehydroAla) where X combined, except for the periods January-February and and Y represent the two variable amino acids and Adda March-April 1987,each of which were combined. Two or is 3-amino-9-methoxy-2,6,8-trimethyl-lO-phenyldeca-4,6- three portions from each combined sample were analyzed dienoic acid. At least four additional variants have been for toxin content. The combined samples were not mixed detected (10-15). so that an indication of sample variability would be obMany reports have noted the variable toxicity of samples tained. from cyanobacterial water blooms with regard to site, Water samples were collected weekly from December season, week, or even day of collection (e.g., ref 16). The 1984 to March 1986 and biweekly from April 1986 to May variable toxicity in M . aeruginosa is probably due to en1987 with a 6-LVan Dorn water sampler from which all of the following measurements were made. Surface temperature was measured with a Cole-Parmer 8502-20 * Present address: Department of Microbiology, University of thermistor, and a Metrohm Herisau E444 was used to Georgia, Athens, GA 30602. measure surface pH. Surface dissolved oxygen (expressed Division of Water Technology, CSIR. t Research Institute for Nutritional Diseases. as percent saturation and corrected for temperature and 0013-936X/90/0924-1413$02.50/0
0 1990 American Chemical Society
Environ. Sci. Technol., Vol. 24, No. 9, 1990 1413
altitude) was measured by the azide modification of the Winkler method and determined spectrophotometrically (35). Chlorophyll a , minus degradation products, was measured spectrophotometrically after being extracted with boiling ethanol (33). Surface chlorophyll a was measured to determine whether floating M. aeruginosa biomass was related to toxin production whereas integral euphotic zone chlorophyll a is a measure of photosynthetically active M. aeruginosa biomass. Primary production and the production of dissolved organic carbon (EDOC) by photosynthesizing phytoplankton were measured by using the 14C light- and dark-bottle technique (33, 36) a t seven depths covering the euphotic zone. Integral, hourly rates were obtained by planimetrically integrating the depth profiles. Surface dissolved organic carbon (DOC) concentrations were determined by filtering water samples through glass-fiber filters (mean pore size ca. 0.7 pm) and injecting the filtered water into a Beckman 915A TOC Analyzer following K2S208and UV-irradiation digestion (37). For ammonium, nitrate + nitrite, and orthophosphate, surface water samples were filtered through glass-fiber filters and analyzed by Technicon Autoanalyzers according to published methods (38). Ammonium was determined by colorimetry after reaction with alkaline phenol and hypochlorite; nitrate was reduced to nitrite, which was determined colorimetrically; and orthophosphate was determined by colorimetry of the molybdenum blue color. Toxins were extracted from freeze-dried M. aeruginosa scums by use of distilled water instead of the usual butanol-methanol-water solution (11)because recovery of toxin was better (results not shown) and the procedure was quicker due to one less drying step. Freeze-dried material (0.1 g) and 10 mL of distilled water were homogenized in a Turrax homogenizer at medium speed for 30 s. After being gently shaken for 1 h at room temperature, the suspension was centrifuged (16000g; 30 min) and the pellet reextracted as above. The supernatants were pooled and the toxins purified by using a Sep-Pak C18 cartridge as described previously (1I). Dried, purified toxins were stored a t 4 " C and dissolved in 38% (v/v) acetonitrile in distilled water just before analysis. Reversed-phase high-pressure liquid chromatography was carried out as described previously (11, 19) except a Beckman Ultrasphere ODS 5-pm, 15 cm X 4.6 mm column was used and the mobile phase was 38% (v/v) acetonitrile, 0.1% (v/v) trifluoroacetic acid in distilled water. Six M . aeruginosa toxin standards designated YR, LR, FR, YA, LA, and LAba were used. The nomenclature proposed by Botes et al. (8) identifies the individual toxins (referred to as cyanoginosins or microcystins) by a two-letter suffix, which denotes the two variant L-amino acids in the cyclic heptapeptides; Y = tyrosine, R = arginine, L = leucine, F = phenylalanine, A = alanine, and Aba = aminoisobutyric acid. The general structure of the toxins is presented in the Introduction. The toxins were isolated from a culture of M . aeruginosa and purified by the method of Botes et al. (39). Toxins FR and LAba were supplied by the National Chemical Research Laboratory, CSIR, Pretoria, South Africa. Due to the fact that the standards for FR and LAba were not of the required purity, the exact concentrations of these two toxin variants in scums could not be determined. Concentrations of toxin variants YR, LR, YA, and LA in freeze-dried scums were determined by use of the linear relationship between peak area and injected amounts of the toxin standards (11);the peak area of each toxin was compared with that of the corresponding toxin 1414
Environ. Sci. Technol., Vol. 24, No. 9, 1990
0.0075
-
t
A iR
E -5 0.0050 0
W 0
2
a
m ir
2m
0.0025
6
0 IO
20
RETENTION TIME (min) 0 0075
-
E
0
%
v
00050-
W
0 Z Q
m LT 0
2
00025
-
6
Ot' 0
IO
20
RETENTION TIME (min)
Figure 1. Typical reversed-phase HPLC chromatograms of (A) toxln standards YR, LR, FR, YA, LA, and LAba and (B) extract of M. aefugin- obtained from Hartbeespoort Dam, South Africa. The amounts of toxin standards YR, LR, YA, and LA injected were 42.1, 24.8, 35.4, and 29.7 ng, respectivety. FR and LAba were not of the required purity. A Beckman Ultrasphere ODS 5-pm (15 cm X 4.6 mm) column was used with a moblle phase of 38 % (v/v) acetonitrile, 0.1 % (v/v) trifluoroacetic acid in distilled water at a flow rate of 1.0 mL min-'.
standard. Product-moment correlation coefficients were calculated by the method of Karl Pearson (40).
Results and Discussion The six toxin standards eluted a t different retention times by reversed-phase HPLC using a mobile phase of 38% acetonitrile, 0.1 % trifluoroacetic acid (Figure 1A). We found that the same mobile phase with a 5-pm Novapak CIScolumn (Waters, 15 cm X 3.9 mm) produced a different elution order of toxin standards: YA eluted between YR and LR instead of between FR and LA (data not shown). This is an improvement on the reversed-phase HPLC elution profile obtained previously ( I l ) , where LR and YR coeluted when a mobile phase of 26% acetonitrile in 10 mM ammonium acetate was used. Extracts of freeze-dried M. aeruginosa often produced several peaks in the HPLC elution profile in addition to peaks corresponding to the six standards (Figure 1B; retention times of 4.4,9.1, and 16.8 min). These additional peaks may be produced by more toxin variants, several of which have been detected in South African strains (10-12), but are not available in sufficient quantity for their chemical structure to be determined, or by unknown
Table I. Yearly Variations in the Relative Levels of Peptide Toxins FR and LAba in Freeze-Dried Scums of M. aeruginosa from Harbeespoort Dam, South Africa" vear and month toxin
1984 12
1985 3
4
5
6
1986 7
11
2
1
3
+ + + - - - - ++ ++ ++ ++ ++ - - - - - + + + + + + + indicates not detectable; + and ++ indicate relative levels of toxin.
FR LAba
1987
4
5
+ +
-
6
7
8
9
- - - - - - -
1
3
4
+
+
-
+
-
5
-
-
-
Table 11. Summary of Correlation Coefficients between Peptide Toxin Concentration in Surface Scums of M . aeruginosa [fig (g of dry scum)-1] and Various Environmental Parameters in Hartbeespoort Dam, South Africa" toxin total
YR LR YA LA
solar radiatn
water temp
PH
% oxygen
saturatn
0.56* 0.40 0.48 0.51* 0.72*
0.60* 0.56* 0.49 0.59* 0.62*
0.67* 0.41 0.64* 0.49 0.75*
0.52* 0.31 0.47 0.47 0.65*
surface Chl a
PPIZeu Chl a
nitrate + nitrite
-0.33
0.57* 0.43 0.49 0.43 0.72*
NS NS NS
-0.33 -0.34
0.34
-0.34
NS
NS
NS NS NS -0.37
orthophosphate
NS
"Number of cases is 20 except for surface and euphotic zone (ZeJ chlorophyll a (chl a ) and primary production (PP), where it is 16. NS, not significant. No asterisk indicates a significant correlation at D < 0.05: one asterisk indicates sienificance at I) < 0.001.
substances. However, it is likely that the six toxin standards represent the majority of the peptide toxin content of the M. aeruginosa samples analyzed in this study because the six standards are the most commonly occurring toxins in M . aeruginosa scums produced in South Africa. Toxins LA and LR have previously been isolated from a natural bloom of M . aeruginosa in Hartbeespoort Dam and were characterized by amino acid analysis (39). It is, therefore, not surprising to find these toxins in samples taken during the present study. All six peptide toxins were either not detected or in very low concentrations in M . aeruginosa scums collected during the winter months (May-August) but were generally in high concentrations during the summer (Table I and Figure 2). Samples collected during the same month sometimes showed substantial variability in individual toxin content (Figure 2), which is a common characteristic of cyanobacterial water blooms (16). Individual toxin concentrations were usually, in decreasing order, LR I YR I LA = YA for the four toxins for which accurate concentrations could be determined. Judging from the size of the chromatographic peaks, FR was a major toxin in most samples while LAba was a minor contaminent. The total toxin concentrations (Figure 3), which varied from 5 to 415 pg (g of dry scum)-*,were the sums of the four individual peptide toxin concentrations that could be determined accurately (Figure 2). Kungsuwan and coworkers (41) found two toxins, YR and LR, in cultured M. aeruginosa M-228; YR was in much higher concentration than LR. In contrast to the Japanese sample, YR was not the major toxin in any of the Hartbeespoort Dam scum samples collected during the present study (Figure 2), in a sample collected from the same source in November 1974 ( 3 9 ) , or in a culture of M . aeruginosa originating from Witbank Dam, South Africa (39). However, YR was the major toxin in M . aeruginosa WR-70, which originates from South Africa (11). Twelve environmental variables were measured on a regular basis during the period of the study (Figure 3). Mean daily solar radiation was averaged for each month while the other variables were monthly means of weekly or biweekly measurements. For every month scum samples were analyzed, except June and July 1985, September 1986, and May 1987, M . aeruginosa dominated (>80% by volume) the phytoplankton species in the euphotic zone
400
LR I
-
1 4 0 m m 20
a
-
IJ
A
J
1985
0
IJ
A
J
0
IJ
A
1986
Figure 2. Changes in the concentrations with time of four peptide toxins in freeze-dried scums of M . aeruginosa from Hartbeespoort Dam. Error bars represent SEM.
[mean depth 3.5 m (42,43)]. Correlation analyses therefore excluded these 4 months' data for chlorophyll a, primary production, and EDOC (Table 11). Primary production per unit of chlorophyll a , solar radiation, surface water temperature, pH, and percent oxygen saturation were strongly, positively correlated to total toxin concentration while surface chlorophyll a and orthophosphate concentrations were weakly, negatively correlated. All four individual toxins were positively correlated to the first five variables listed above (Table 11). Harris and Gorham (20,21) compared the toxin content of M. aeruginosa NRC-1 at two light intensities of 220 and 1600 foot-candles [37 and 270 peinsteins &E) m-2 s-l] using mouse bioassays and found little difference. Similarly, Watanabe and Oishi (29) found no significant difference in the toxicity of M. aeruginosa M-228 grown at either 30 or 75 p E m-2 s-l (LD, 9.65 and 9.97 mg kg-', respectively; LD,, is the median lethal dose in milligrams of dry algae per kilogram of mice), but a lower light intensity of 7.5 p E m-2 s-l produced a culture -4 times less toxic (LDm36.9 mg kg-'). Van der Westhuizen and Eloff (26) used a range Environ. Sci. Technol., Vol. 24, No. 9, 1990
1415
-
4000
L
5
7 3000 2000 1000
TEMPERATURE 20
0
of 21-205 p E m-2 s-l and found toxicity was lowest at the minimum light intensity and declined a t the highest light intensity. The above studies show that light intensity has little effect on toxin content except at the lower end of the range 7.5-270 p E m-2 s-l, where culture growth rate is light limited. Light is a limiting factor for growth of M . aeruginosa in Hartbeespoort Dam (33). Therefore it is not surprising that solar radiation correlates to toxin concentration of M. aeruginosa scums. Harris and Gorham (20, 21) found that toxin concentration of M. aeruginosa NRC-1 increased approximately 10-fold when cultures were incubated at 25 "C instead of 32.5 "C. Microcystis isolates were 4 times more toxic when grown a t 18 "C than at 29 "C (23). In contrast, Van der Westhuizen and Eloff (26) found less pronounced changes in toxicity in the range 16-32 "C. M . aeruginosa UV-006 was twice as toxic when cultured at 20 "C (LD,, 25.4 mg kg-') than at 32 "C, but at 16 "C the toxicity was reduced by -35% compared to 20 "C. M . aeruginosa M-228 had qualitative characteristics common to UV-006 with similar toxicities at 18 and 25 "C (LD,, ca. 11mg kg-') decreasing by -40% at 32 "C (29). The above studies indicate that maximum toxicity of M . aeruginosa cultures is achieved at a temperature between 18 and 25 "C. In Hartbeespoort Dam, the greatest toxin concentration in M. aeruginosa scums was found during the summer with surface water temperatures up to 27 "C, but negligible toxin concentrations were always found during the winter with a minimum water temperature of 13 "C. The reservoir data therefore only agree with the culture studies where maximum toxicity was found around 25 "C. The effect on toxicity of M . aeruginosa UV-006 by a pH range from 6.5 to 10.5 was studied by controlled addition of carbon dioxide to the culture suspension (25). Maximum toxicity (LD, ca. 35 mg kg-') was found at the lower and upper ends of the range of pH studied, which corresponded with the lowest growth rates. The authors concluded that the slower the cells grew the more toxic they were. In Hartbeespoort Dam, during the present study period, the pH ranged from 7.7 to 9.4 and toxicity generally increased with increasing pH. This agrees well with the above culture study where the minimum toxicity (LD, 52 mg kg-') was found at pH 8. Harris and Gorham (20,21) found that increasing the aeration rate from 100 to 1000 mL min-' in M. aeruginosa NRC-1 more than doubled the toxicity of the culture under otherwise optimum conditions. Similarly, in the present study, increased percent oxygen saturation during the summer correlated with greater toxin concentrations in M . aeruginosa scums. For most of the year Hartbeespoort Dam is characterized by dense populations of M. aeruginosa (33,43). During calm weather the buoyant cyanobacterium accumulated a t lee shores to form scums. However, the amount of surface M . aeruginosa biomass measured as chlorophyll a was weakly, negatively correlated to toxin concentration in the cells (Table I). Therefore, large concentrations of M. aeruginosa on the surface of Hartbeespoort Dam, which often occurred even during winter when the buoyant cells maintained themselves within shallow diurnal mixed layers (43),tended to indicate that toxin was not present in high concentration in the cells. The low levels of toxin in scum samples taken in the winter is probably due to the dominance of a nontoxic strain of M. aeruginosa, whereas toxic strains dominate in the summer (44). Hartbeespoort Dam is somewhat unusual in that blooms are often present year round, making it possible to determine toxin content on a yearly basis rather than just a seasonal one.
5 SOLAR RADIATION
10.0
9.0 8 .O
t
DH
t
~
X OXYGEN
200
tI
loo
I
SURFACE CHL I
EUPHOTIC ZONE CHL
r?
3367
I
I
400
F PRIMARY PRODUCTION
2000
I
9.0
E
6 .O 400
-
m 200
-
-
L
a
7
AMMONIUM I
NITRATE
4000
+
NITRITE
g 2000
I
cn
1
PHOSPHATE
500
TOTAL TOXINS
A
400
J
A
J
1985
O
J
A
J
O
J
A
1986
Figure 3. Changes in the total toxin concentration in freeze-dried scums of M . aeruginosa and a number of environmental factors with time in Hartbeespoort Dam. Monthly means of measurements were plotted only when scums occurred. Data sources: present study and calculated from Robarts ( 4 5 ) . 1416
Environ. Sci. Technol., Vol. 24, No. 9, 1990
Neither photosynthetically active biomass (measured as integral, euphotic zone chlorophyll a ) nor integral primary production and EDOC were correlated to M. aeruginosa toxin concentration in Hartbeespoort Dam. However, primary production per unit of chlorophyll a was strongly, positively correlated to cell toxin concentration (Table 11). This result agrees quite well with findings of other researchers, who have reported that with increasing light intensity, temperature (up to 18-25 "C), and pH (range from 8 to 9), growth rate and toxicity of M. aeruginosa cultures increase although not at the same rate (20,21,25, 26,29). EDOC per unit of chlorophyll a showed no correlation to toxin concentration. Brown (22) investigated the effect of different nutrient concentrations on the toxicity of M . aeruginosa NRC-1. The toxicity of cells grown in the standard BG-11 culture medium was LD50 45.3 mg kg-'. Toxicity more or less doubled (LD50 ranging from 19.8 to 25.6 mg kg-') when cells were grown at the following concentrations: 0.5X NaNO,, K2HP04,CaC12and microelements; 2.0X K2HP04, MgSO,, ferric citrate, and microelements; and 3.0X K2HPO4 and ferric citrate. There was also an increase in toxicity (LDbOranging from 30.1 to 35.9 mg kg-') for the 0.5X ferric citrate, the 2.0X NaN03 and CaC12, and the 3.0X MgSO, and CaC12treatments. The only treatment that did not produce any increase in toxicity was the half-strength MgSO,. In contrast, decreases in toxicity of -25% with and 1/20 the normal concentration of phosphate in culture medium MA were found in M. aeruginosa M-228 (29). Toxicity was decreased 150% at nitrate concentration but by only 50% at ',Im of the normal concentration of nitrate. In the present study, neither organic nor inorganic nutrients showed strong correlations with total toxin concentration in M. aeruginosa although several individual toxin concentrations were weakly correlated to nitrate nitrite and orthophosphate (Table 11). The lack of strong correlations between M . aeruginosa toxin concentration and nitrogen and phosphorus concentrations in Hartbeespoort Dam is not surprising because these inorganic nutrients are always in excess of phytoplankton requirements (33). Van der Westhuizen and co-workers (7,27),using ranges of light intensity from 21 to 205 p E m-2 s-', temperature from 16 to 36 "C, and pH from 6.5 to 10.5, found the ratio of toxins LR to LA remained relatively constant in M. aeruginosa UV-006 (derived from Hartbeespoort Dam) at around 1:1, whereas in the present study the ratio varied from 2:l to 8:l with no seasonal trend. At temperatures of 16 and 20 "C they found a third peptide of unknown amino acid composition at up to 45% of the total toxin concentration, but at 28 "C and above, only toxins LR and LA were found. In the present study, none of the six toxins or unknown peaks on the HPLC chromatogram were prominent when the surface water temperature was below 20 "C. These differences in toxin compositions between cultured isolates and naturally occurring scums consisting of variable mixtures of toxic and nontoxic strains of M . aeruginosa are probably due to the numerous, variable environmental conditions affecting Hartbeespoort Dam compared to the controlled environment and genetic homogeneity of laboratory cultures.
+
Conclusion Our results of strong, positive correlations between total toxin concentrations in M.aeruginosa scums and primary production per unit of chlorophyll a , solar radiation, surface water temperature, pH, and percent oxygen saturation in a hypertrophic African reservoir agree well with published M . aeruginosa laboratory culture studies using
mouse bioassays. However, greater variation in the relative proportions of the individual toxins was found in the present study when compared to the very limited published data available from toxic isolates. Laboratory culture studies, therefore, can give an indication of the environmental factors affecting toxin concentrations in naturally occurring cyanobacterial scums that pose a health hazard.
Acknowledgments We thank R. D. Robarts for critical assessment of the draft manuscript and W. E. Scott for the supply of M. aeruginosa scums. We gratefully acknowledge the chlorophyll a data of T. Zohary, the nitrogen data of P. J. Ashton, the phosphorus data of J. A. Thornton, the supply of standards of toxins FR and LAba by F. H. H. Carlsson, and the technical assistance of W. Kekana.
Literature Cited (1) Skulberg, 0. M.; Codd, G. A.; Carmichael, W. W. Ambio 1984, 13, 244-247. (2) Carmichael,W. W. In Handbook ofNatural Toxins,Marine Toxins and Venoms;Tu, A. T., Ed.; Marcel Dekker: New York, 1988; Vol. 3, pp 121-147. (3) Carmichael, W. W.; Juan, Y. M.; Rong, H. Z.; Wan, H. J.; Lu, Y. J. Arch. Hydrobiol. 1988, 114, 21-30. (4) Steyn, D. G. South Afr. J . Sci. 1945, 41, 243-244. (5) Schwimmer, M.; Schwimmer, D. In Algae, Man and the Environment; Jackson, D. F., Ed.; Syracuse University Press: Syracuse, NY, 1968; pp 279-358. (6) Carmichael, W. W. In The Water Environment, Algal Toxins and Health; Carmichael, W. W., Ed.; Plenum Press: New York, 1981; pp 1-13. (7) Van der Westhuizen, A. J.; Eloff, J. N.; Kruger, G. H. J. South Afr. J. Bot. 1988,54, 372-374. (8) Botes, D. P.; Tuinman, A. A,; Wessels, P. L.; Viljoen, C. C.; Kruger, H.; Williams, D. H.; Santikarn, S.; Smith, R. J.; Hammond, S. J. J . Chem. SOC.,Perkin Trans. 1 1984, 2311-2318. (9) Botes, D. P.; Wessels, P. L.; Kruger, H.; Runnegar, M. T. C.; Santikarn, S.; Smith, R. J.; Barna, J. C. C.; Williams, D. H. J . Chem. SOC.,Perkin Trans. 1 1985, 2747-2748. (10) Kfir, R.; Johannsen, E.; Botes, D. P. Toxicon 1986, 24, 543-552. (11) Gathercole, P. S.; Thiel, P. G. J . Chromatogr. 1987,408, 435-440. (12) Van der Westhuizen, A. J.; Eloff, J. N.; Kruger, G. H. J. South Afr. J. Sci. 1988, 84, 70-71. (13) Krishnamurthy, T.; Szafraniec, L.; Hunt, D. F.; Shabanowitz, J.; Yates, J. R., 111; Hauer, C. R.; Carmichael, W. W.; Skulberg, 0.;Codd, G. A.; Missler, S. Proc. Natl. Acad. Sci. U.S.A. 1989, 86, 770-774. (14) Meriluoto, J. A. 0.;Sandstrom,A.; Eriksson, J. E.; Remaud, G.; Craig, A. G.; Chattopadhyaya, J. Toxicon 1989, 27, 1021-1034. (15) Watanabe, M. F.; Oishi, S.; Harada, K.-I.; Matsuura, K.; Kawai, H.; Suzuki, M. Toxicon 1988, 26, 1017-1025. (16) Carmichael, W. W.; Gorham, P. R. In The Water Enuironment, Algal Toxins and Health;Carmichael,W. W., Ed.; Plenum Press: New York, 1981; pp 161-172. (17) Eloff, J. N.; Van der Westhuizen, A. J. In The Water Environment, Algal Toxins and Health; Carmichael, W. W., Ed.; Plenum Press: New York, 1981; pp 343-364. (18) Scott, W. E.; Barlow, D. J.; Hauman, J. H. In The Water Environment, Algal Toxins and Health; Carmichael, W. W., Ed.; Plenum Press: New York, 1981; pp 49-69. (19) Brooks, W. P.; Codd, G. A. Lett. Appl. Microbiol. 1986,2, 1-3. (20) Harris, R. E.; Gorham, P. R., unpublished data, 1956. (21) Gorham, P. R. In Algae and Man; Jackson, D. F., Ed.; Plenum Press: New York, 1964; pp 307-336. (22) Brown, P. J. Ph.D. Thesis, Texas A&M University, College Station, TX, 1974. Environ. Sci. Technol., Vol. 24, No. 9, 1990
1417
Environ. Sci. Technol. 1990, 24. 1418-1427
Runnegar, M. T. C.; Falconer, I. R.; Jackson, A. R. B.; McInnes, A. Toxicon 1983, Suppl. 3, 377-380. Van der Westhuizen, A. J. Ph.D. Thesis, University of the Orange Free State, South Africa, 1984. Van der Westhuizen, A. J.; Eloff, J. N. 2. Pflantenphysiol.
Van Steenderen, R. A.; Lin, J. S. Anal. Chem. 1981, 53, 2157-2 158.
Standard Methods for the Examination of Water and Wastewater, 16th ed.; American Public Health Association,
Van der Westhuizen, A. J.; Eloff, J. N. Planta 1985, 163,
American Water Works Association and Water Pollution Control Federation: New York, 1985. Botes, D. P.; Kruger, H.; Viljoen, C. C. Toxicon 1982,20,
55-59.
945-954.
Van der Westhuizen, A. J.; Eloff, J. N.; Kruger, G. H. J.
Sokal, R. R.; Rohlf, F. J. Biometry, 2nd ed.; W. H. Freeman:
Arch. Hydrobiol. 1986, 108, 145-154. Watanabe, M. F.; Oishi, S. Bull. Jpn. SOC.Sci. Fish. 1983,
New York, 1981; pp 565-591. Kungsuwan,A,; Noguchi, T.; Matsunaga, S.; Watanabe, M. F.; Watabe, S.; Hashimoto, K. Toxicon 1988,26,119-125.
1983, 110, 157-163.
49, 1759.
Watanabe, M. F.; Oishi, S. Appl. Environ. Microbiol. 1985, 49, 1342-1344.
Watanabe, M. F.; Harada, K.-I.;Matsuura, K.; Watanabe, M.; Suzuki, M. J . Appl. Phycol. 1989, I , 161-165. Robarts, R. D.; Ashton, P. J.; Thornton, J. A,; Taussig, H. J.; Sephton, L. M. Hydrobiologia 1982, 97, 209-224. Ashton, P. J. J . Limnol. SOC.South. Afr. 1985, 11, 32-42. Robarts, R. D.; Zohary, T. J. Ecol. 1984, 72, 1001-1017. Zohary, T. J . Plankton Res. 1985, 7, 399-409. Ashton, P. J.; Twinch,A. J. J. Limnol. Soc. South. Afr. 1985, 11, 62-65.
Robarts, R. D.; Zohary, T. Appl. Environ. Microbiol. 1986, 51, 609-613.
Zohary, T., personal communication,CSIR,Pretoria, 1989. Zohary, T.; Robarts, R. D. J. Plankton Res. 1989,11,25-48. Scott, W. E. In Mycotoxins and Phycotoxins; Steyn, P. S., Vleggaar, R., Eds.; Elsevier Science Publishers: Amsterdam, 1986; pp 41-50.
Robarts, R. D. Hydrobiologia 1988, 162, 97-107. Received for review January 22, 1990. Revised manuscript received May 18, 1990. Accepted May 21, 1990. This work was supported by the Department of National Health and Population Development. Contribution no. 80 to the Hartbeespoort Dam Ecosystem Program.
Polycyclic Aromatic Hydrocarbon Emissions from the Combustion of Crude Oil on Water Bruce A. Benner, Jr.," Nelson P. Bryner, Stephen A. Wise, and George W. Mulholland National Institute of Standards and Technology, Gaithersburg, Maryland 20899
Robert C. Lao and Mervin F. Fingas Environment Canada, Ottawa, Canada K 1A OH3
This work involved an investigation of some of the factors necessary to assess the environmental impact of an in situ burn: the fraction of an oil layer that can be burned, the quantity of smoke, and the concentrations of 18 polycyclic aromatic hydrocarbons (PAHs) in the smoke, crude oil, and burn residue. Alberta Sweet crude in 2-, 3-, 5-, lo-, and 30-mm layers on water was burned and smoke samples were collected at elevated and ambient temperatures and analyzed by two independent laboratories. While burning the crude oil produced less total PAHs than were in the original crude oil, the concentrations of PAHs with five or more rings were 10-20 times greater in the smoke than in the oil. The organic carbon fraction of the smoke was in the range of 14-21 5%. As the fuel layer thickness was increased from 2 to 10 mm, the smoke yield increased from 0.035 to 0.080 g of smoke/g of fuel, and the percentage of oil residue decreased from 46 to 17%. By consuming much of the oil spill and reducing the amount of PAHs in the water, and by dispersing the combustion products over a larger area, in situ burning can mitigate the local environmental impact of an oil spill. There appears to be a range of conditions, such as in Arctic ice fields, where in situ burning might be the most viable cleanup method. Introduction Since 1970 over 4000 off-shore oil wells ( I , 2 )have been drilled, and off-shore drilling continues to venture into more remote locations, such as the Arctic Ocean. Oil drilling in remote areas and large tankers transporting the crude oil from these wells have increased the possibility of a major oil spill occurring in a remote location. Con1418
Environ. Sci. Technol., Vol. 24, No. 9, 1990
ventional oil spill countermeasures can be effective when the cleanup equipment and ships quickly reach the oil spill, but remote off-shore drilling locations, such as the Canadian Arctic, would be difficult to reach quickly and the hostile weather could seriously impede cleanup operations. The Exxon Valdez oil spill off the Alaskan coast demonstrated some of the difficulties of quickly transporting conventional cleanup equipment to an oil spill. To overcome implementation difficulties with conventional cleanup techniques, in situ burning, igniting the oil and allowing it to burn off, has been suggested. Many aspects of an in situ burn, including oil slick ignition techniques (3),burning rates (4-7),weathering (8, 9), effects of ice (IO, I I ) , and the dispersion of the smoke plume (12),have received attention. Thompson et al. (3) have extensivelydefined the conditions under which in situ burning might be used. However, the environmental implications of such a burn have not been quantified. The present study is directed at clarifying some aspects of the environmental impact of in situ burning, including the fraction of an oil layer that can be burned, the composition of unburned residue, and the quantity and composition of smoke generated. Examining the levels of polycyclic aromatic hydrocarbons (PAHs) in the crude, residue, and smoke is critical for assessing the environmental impact, since some PAH species are believed to be carcinogenic (13-16). It is known that there are some PAHs in the crude oil itself and also that PAHs are produced by the burning of hydrocarbon fuels, but there are no quantitative data on the relative amount of PAHs in the crude oil versus the amount emitted from burning the oil. In this study involving the
0013-936X/90/0924-1418$02.50/0
@ 1990 American Chemical Society
