THE EFFECT OF FERRIC CHLORIDE FLOCCULATION ON CYANOBACTERIAL CELLS

PII: S0043-1354(97)00276-5

Wat. Res. Vol. 32, No. 3, pp. 808±814, 1998 # 1998 Elsevier Science Ltd. All rights reserved Printed in Great Britain 0043-1354/98 $19.00 + 0.00

THE EFFECT OF FERRIC CHLORIDE FLOCCULATION ON CYANOBACTERIAL CELLS M C. W. K. CHOW*, J. HOUSE, R. M. A. VELZEBOER, M. DRIKAS* , M. D. BURCH and D. A. STEFFENSEN

Australian Water Quality Centre, Private Mail Bag 3, Salisbury, South Australia 5108, Australia (First received April 1997; accepted in revised form August 1997) AbstractÐTaste, odour and cyanobacterial toxins are not generally removed during conventional water treatment and can sometimes be increased during the treatment process due to cell lysis. Some reasons for this have been documented in the literature, but there appears to be contradictory evidence regarding the e€ect of ¯occulation. In an earlier study, aluminium sulphate did not appear to cause lysis of cells of cultured Microcystis aeruginosa or Anabaena circinalis, nor increase the amount of microcystinLR or geosmin in the water. In this study, the e€ect of ferric chloride on cultured cells of M. aeruginosa and A. circinalis was evaluated at concentrations that would occur in a water treatment plant. The results showed that ferric chloride did not appear to cause cell lysis of cultured M. aeruginosa and A. circinalis. However, ferric chloride seemed to stimulate growth of both M. aeruginosa and A. circinalis in the experiments with South Para Reservoir Water. # 1998 Elsevier Science Ltd. All rights reserved Key wordsÐMicrocystis aeruginosa, microcystin-LR, Anabaena circinalis, geosmin, ferric chloride, water treatment

INTRODUCTION

Cyanobacteria (blue-green algae) are photoautotrophic Gram negative bacteria that are common members of the freshwater phytoplankton community in surface waters. They are of concern in relation to drinking water because of their ability to produce toxins and odours which can signi®cantly impair water quality. The use of chemicals in the ¯occulation stage of the water treatment process may cause cell lysis and release intracellular metabolities. These compounds are not then removed easily by the conventional ¯occulation and ®ltration treatment processes (Monteil, 1983; McGuire and Gaston, 1988; Keijola et al., 1988; Himberg et al., 1989; Ando et al., 1992). The removal of these compounds has been a major reason for the inclusion of additional treatment such as the use of oxidation or activated carbon, which adds signi®cantly to both the cost and complexity. The removal of cyanobacterial cells without damage would signi®cantly reduce the concentration of taste, odour and toxic metabolites present in the ®nished water. Although this would not completely negate the need for further treatment, there is no doubt that the e€ectiveness of treatment would be improved together with a reduction of costs by minimising oxidant/powdered activated *Author to whom all correspondence should be addressed.

carbon doses and/or extension of the e€ective lifetime of granular activated carbon ®lters. Previous studies have indicated that dosing of aluminium sulphate at concentrations used in water treatment, and also lower than optimum level, does not appear to damage cyanobacterial cells of Microcystis aeruginosa and Anabaena circinalis (Velzeboer et al., 1995a; 1995b). This paper reports on similar experiments to determine the e€ect of ferric chloride on cultured cells of the two cyanobacteria, M. aeruginosa and A. circinalis. This study focuses on microscopic measurements (total cell count and cell viability), cell pigments (phycocyanin and chlorophyll) and the release of cell metabolites (microcystin-LR or geosmin).

MATERIALS AND EXPERIMENTAL METHODS

Materials Water. Water used in these experiments was obtained from a Milli-Q puri®cation system or taken from the South Para Reservoir (South Australia). The reservoir water was ®ltered through a 0.22 mm ®lter to remove any natural algae and bacteria. Relevant water quality parameters were; pH: 8.2, dissolved organic carbon: 8.1 mg/l, true colour: 15 HU, turbidity: 0.53 NTU (after ®ltration) and iron: 0.066 mg/l.

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Flocculation of cyanobacterial cells

Cyanobacterial cultures. M. aeruginosa or A. circinalis were grown in 12 litres ASM-1 medium (Gorham et al., 1964) at 258C under continuous illumination (080 mM photons mÿ2sÿ1). Cultures were harvested at the late exponential phase of growth, which corresponded to 10 days after inoculation for M. aeruginosa and 14 days for A. circinalis (Velzeboer et al., 1995a,b). Chemicals. A 20,000 mg/l ferric chloride stock solution was prepared in Milli-Q water using AR grade Ferric Chloride (FeCl3
6 H2O), BDH. Methods Toxicity experiments. A cyanobacterial batch culture (24 l) was concentrated to 900 ml by centrifugation at 8000 rpm for 10 minutes. The concentrate was mixed with 5.4 l of South Para reservoir water to make up a total of 6.3 l. The water was transferred into 9 polythene bottles (700 ml each) and left overnight at 258C under constant illumination. Three samples were retained as controls, and the other 6 samples were dosed with 15 mg/l (half dose) and 30 mg/l (optimum dose) of ferric chloride (as FeCl3). The amount of ferric chloride required to achieve the optimum dose was determined using a standard jar test method. Blanks using 600 ml of natural water with 100 ml of ASM-1 bu€er at the same chemical dose concentrations but without addition of cyanobacteria were used to determine the background concentrations of chlorophyll-a, phycocyanin, microcystin and geosmin. Samples (230 ml) were taken for analysis initially (t = 0), after 4 hours (t = 4) and after 24 hours (t = 24). The experiments were done in triplicate for each species, i.e. a separate batch culture was grown independently for each experiment. Total cell number. Cyanobacterial cells were counted on a compound microscope in a Sedgewick-Rafter Counting chamber after preservation in Lugol's iodine. Cell counts were carried out to a minimum precision of 20%. Cell viability. Cyanobacterial cells were stained with ¯uorescein diacetate (FDA) and propidium iodide (PI). FDA stains cells with an intact cell membrane and active esterases. PI stains cells that do not have an intact cell membrane and non-active esterases (Velzeboer et al., 1995a). A minimum number of 100 cells were counted to achieve a precision greater than 10%. Chlorophyll-a. 100 ml samples were ®ltered through GF/ C ®lter papers and the chlorophylls were extracted using 10 ml ethanol (95%). The optical densities of the extracts at 665 and 750 nm were determined using a GBC UV/VIS 918 spectrophotometer with 4 cm matched cells. The chlorophyll-a concentrations were determined using the equations derived by Wintermans and de Mots (1965). Phycocyanin. 100 ml samples were concentrated by centrifugation at 3750 rpm for 15 minutes. The pellets were then resuspended with 5 ml of 20 mM sodium acetate buffer, pH 5.5. The cells were broken up by sonication for 2 minutes. The cell-free extracts were precipitated with 5 ml of 1% (w/v) streptomycin sulfate. The optical densities of the supernatants at 620 and 650 nm were determined using a GBC UV/VIS 918 spectrophotometer with 4 cm matched cells. The phycocyanin concentrations were calculated using the equations described in Tandeau de Marsac and Houmard (1988). Geosmin. The samples were gravity ®ltered through GF/ C ®lter papers into glass bottles with no air gap. Concentrations of geosmin were determined using a HewlettPackard gas chromatograph-mass spectrometer (GC/MS) after preconcentration by closed loop stripping. Microcystin-LR. 100 ml samples were ®ltered through GF/C ®lter papers, the ®ltrates were concentrated on preprimed Sep-Pak vac /3cc C18 Cartridges. The microcystin

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toxins were eluted by MeOH. The microcystin concentrations were determined by HPLC using standard microcystin-LR as a reference. pH. The pH of the samples were measured using a portable Hanna Instruments H18424 pH meter with a combined pH electrode. Calibration was carried out with pH 6.86 and pH 9.18 NBS bu€ers. Iron. Total and soluble iron concentrations of the samples were determined using inductively coupled plasma-atomic emission spectrometry (ICP-AES). Speciation calculation. The iron speciation calculation was carried out using GEOCHEM-PC, a multi-purpose chemical speciation program (Parker et al., 1990). The required input parameters were calcium, magnesium, sulphate, alkalinity and chloride. Calcium, magnesium and sulphate concentrations were determined using ICP-AES, alkalinity by acid titration using a glass pH electrode and chloride by a colorimetric method. RESULTS AND DISCUSSION

In this study, total cell number, cell viability, chlorophyll-a, phycocyanin, microcystin release (M. aeruginosa) and geosmin release (A. circinalis) were measured to evaluate the e€ect of ferric chloride on the two selected cyanobacteria. In order to account for the variability associated with using cultured organisms and to provide reliable results, the experiments were repeated three times on three separate occasions for each cyanobacterium. Results for one of the triplicate experiments are presented here, as very similar results were obtained on all occasions. The e€ect of ferric chloride dosage on pH and soluble iron concentration The optimum ferric chloride dose required for the natural water used in these experiments was found to be 30 mg/l as FeCl3. Our previous work showed that underdosing of aluminium sulphate released more available aluminium in the samples (Velzeboer et al., 1995b), therefore samples in this experiment were also dose at half the optimum concentration based on the assumption that ferric chloride behaves similarly to aluminium sulphate. In general, all ferric coagulants are used over a wide range of pH from 4.0 to 11.0 (Bratby, 1980; Smethurst, 1988). The pH was reduced from 8.1 to 7.6 and 8.2 to 7.3, with 15 and 30 mg/l, FeCl3 dosages respectively for samples without cells (Table 1). For water with cyanobacterial cells, the pH was reduced from pH 9.0 to 8.1 and 9.0 to 7.7 with 15 and 30 mg/l FeCl3 dosages respectively (Table 1). A calculation of the iron speciation for the natural water used in these experiments, indicated that solid ferric hydroxide should be the major species (99% at pH above 5) present in these samples after addition of ferric chloride. The soluble iron concentration of the water samples both with and without cyanobacterial cells are given in Table 1. In all cases, the soluble iron concentrations were less than 1% of the added iron which is consistent with the speciation calculation. The soluble iron concentrations were higher in the samples

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C. W. K. Chow et al. Table 1. The e€ect of pH and soluble iron with two di€erent dosages of ferric chloride

No Algal Cells Control 15 mg/l FeCl3 30 mg/l FeCl3 No Algal Cells Control 15 mg/l FeCl3 30 mg/l FeCl3

0 Hrs

pH 4 Hrs

24 Hrs

Soluble Iron Concentration2s.d. (mg/l) 0 Hrs 4 Hrs 24 Hrs

8.0 8.1 8.2

8.1 7.6 7.3

8.1 7.6 7.3

0.023 2 0.006 0.019 2 0.011 0.033 2 0.018

0.013 2 0.001 0.044 2 0.005 0.077 2 0.015

0.0192 0.001 0.1192 0.047 0.1452 0.050

9.1 9.0 9.0

9.2 8.1 7.7

9.9 9.4 8.2

0.015 2 0.003 0.015 2 0.007 0.015 2 0.003

0.019 2 0.007 0.031 2 0.014 0.044 2 0.026

0.0122 0.005 0.0262 0.007 0.0492 0.008

Note: 15 mg/l FeCl3=5.2 mg/l Fe; 30 mg/l FeCl3=10.4 mg/l Fe.

Fig. 1. The e€ect of ferric chloride on Microcystis aeruginosa. (a) total cell number, (b) cell viability, (c) chlorophyll-a and (d) phycocyanin. control; w, half ferric chloride dosage (15 mg/l FeCl3); r and optimum ferric chloride dosage (30 mg/ll FeCl3); q.

Flocculation of cyanobacterial cells

dosed with 30 mg/l FeCl3 than in the samples dosed with 15 mg/l FeCl3. From these results, underdosing of ferric chloride did not contribute to higher soluble iron. In addition, the soluble iron concentrations in samples containing cyanobacteria were lower than the blank (no cells), indicating complexation of FeCl3 by cyanobacteria. The cell metabolic activity of cyanobacteria also caused pH variation during the experiment. Cyanobacteria, as photoautotrophic organisms, use

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carbon dioxide during photosynthesis, the uptake of carbon dioxide by cells led to an increase in the pH of the solution. At time 0, the pH of the samples containing the cyanobacterial cells were higher than the samples without the cells. During the experimental period (24 hours), the pH of the samples with cyanobacterial cells increased after an initial decrease caused by the addition of ferric chloride. However, these increases were not observed for samples without cyanobacterial cells (Table 1).

Fig. 2. The e€ect of ferric chloride on Anabaena circinalis. (a) total cell number; (b) cell viability; (c) chlorophyll-a and (d) phycocyanin. Control: w, half ferric chloride dosage (15 mg/l FeCl3); r and optimum ferric chloride dosage (30 mg/l FeCl3); q.

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The e€ect of ferric chloride dosage on total cell number and cell viability Total cell number measures the total number of cells present while cell viability measures cells with an intact cell membrane. To evaluate the e€ect of ferric chloride dosages, the total cell number of the ferric chloride dosed samples compared with the control are important. For both cyanobacteria, the total cell number for all three series (control, half dose and optimum dose), were increased over the experimental period (Fig. 1a, 1b, 2a and 2b) in the order of optimum dose1half dose > control. It appears that both cyanobacteria grew better in the ferric chloride dosed samples. This may indicate that iron was a limiting nutrient in South Para reservoir water. Under iron limiting conditions, addition of iron has been found to enhance cellular growth, photosynthesis and chlorophyll-a content (Morton and Lee, 1974; Hann, 1984). Both cyanobacteria remained 90% viable in control and ferric chloride dosed samples after 4 and 24 hours in all the experiments. The e€ect of ferric chloride dosage on photosynthetic pigments Cyanobacteria have only one form of chlorophyll, chlorophyll-a, and they also have characteristic biliprotein pigmentsÐphycobilins, which function as accessory pigments in photosynthesis. Phycocyanin, a blue phycobilin, together with the green chlorophyll-a are responsible for the blue green colour of the cyanobacteria. In this study, the concentration of these pigments in the cyanobacterial cells was used as an additional measure of cell integrity.

In Fig. 1c and 2c, the chlorophyll-a concentrations increased with time for both cyanobacteria in all samples. However, the chlorophyll-a results of A. circinalis also showed the same trend as observed with total cell number and cell viability (Fig. 2a and 2b) where optimum dose > half dose > control after 24 hrs. For M. aeruginosa the trend was less clear. The phycocyanin results for both cyanobacteria showed a slight deviation compared with the other three measured parameters (Figs 1d and 2d). In Fig. 1d (M. aeruginosa), the initial concentrations of phycocyanin (t = 4) were lower in the ferric chloride dosed samples than in the control and this di€erence remained constant throughout the duration of the experiment. In Fig. 2d (A. circinalis), the phycocyanin concentration in the control cells increased initially (t = 4) but decreased over 24 hours. For the half dose situation, the phycocyanin concentration decreased throughout the experiment. For the optimum dose, the phycocyanin concentration was constant over the experimental period. This appears to suggest that ferric chloride may have some e€ects on the growth, or even damage, the cyanobacterial cells for both M. aeruginosa and A. circinalis. Phycocyanin is located on the outside of the thylakoid membrane and may be more readily released than, for example, the chlorophyll-a which is localised within the thylakoid membrane (Golecki and Drews, 1982). However, an initial comparison of the standard deviation for replicated measurements showed that the coecient of variation for chlorophyll-a measurement was approximately 5% while phycocyanin was approximately 30%. Thus, it is also possible that the contradictory

Fig. 3. The e€ect of ferric chloride on Microcystis aeruginosa. (a) microcystin release, (b) microcystin release per cyanobacterial cell. control; w, half ferric chloride dosage (15 mg/l FeCl3); r and optimum ferric chloride dosage (30 mg/l FeCl3); q.

Flocculation of cyanobacterial cells

813

Fig. 4. The e€ect of ferric chloride on Anabaena circinalis. (a) geosmin release. (b) geosmin release per cyanobacterial cell. control; w, half ferric chloride dosage (15 mg/l FeCl3); r and optimum ferric chloride dosage (30 mg/l FeCl3); q.

result of the phycocyanin measurement was caused by lack of precision in the analysis. The e€ect of ferric chloride dosage on microcystin and geosmin release The parameters discussed above relate to the condition of the cyanobacterial cells. The release of microcystin and geosmin are major concerns in this study. No microcystin or geosmin was present in the blank solutions, the concentrations of both microcystin and geosmin present at t = 0 (Fig. 3a and 4a) were either the residual levels in the medium which cannot be removed completely by centrifugation, or produced during the overnight stabilising period prior to the experiment. It is clear that the amount of microcystin (M. aeruginosa) or geosmin (A. circinalis) released into the solution increased over the experimental period in the general order of optimum dose > half dose 1 control. However, as mentioned earlier, the increase in total cell number and chlorophyll-a concentration indicated that the population of the cyanobacteria was increasing during the experimental period. This may certainly have contributed to the increase in microcystin and geosmin production and to the release of these compounds into the solution. Thus microcystin and geosmin production and to the release of these compounds into the solution. Thus microcystin and geosmin release per cell was calculated. For M. aeruginosa (Fig. 3b), the microcystin release per cell decreased over the experimental period. This clearly indicates that the addition of ferric chloride did not cause any cell damage which led to the release of microcystin. However, the

results shown in Fig. 4b were extremely dicult to interpret, as the trends were rather irregular. The same irregular trends were also found in the repeat experiments. Figure 4b shows that after 4 hours there was a slight increase in geosmin release per cell (optimum dose > half dose > control). However, after 24 hours, the geosmin release per cell increased for the control, decreased for the half dose and stabilised for optimum ferric chloride dosed samples. From these results, it appears that each A. circinalis cell released more geosmin over the experimental period (even for the controls). Based on this observation, there is no conclusive evidence to show that ferric chloride has contributed to the increase in geosmin release in the experiments. It may be concluded that A. circinalis is a sensitive cyanobacteria and disturbances associated with treatment processes may lead to the release of geosmin. CONCLUSIONS

In this study, ferric chloride concentrations used in water treatment practice did not appear to cause cell lysis but seemed to stimulate growth in South Para reservoir water of cultured A. circinalis and M. aeruginosa, as measured by total cell number, cell viability and chlorophyll-a. However, there did appear to be some decrease in phycocyanin concentration of both cyanobacterial species following ferric chloride addition, although the lack of precision in this analysis makes the result inconclusive. There was no increase in concentration of microcystin in the water following treatment of M. aeru-

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ginosa with ferric chloride. However, it appears that A. circinalis may be more susceptible to damage from chemicals although there was no conclusive evidence to suggest that geosmin was released following treatment with ferric chloride. AcknowledgementsÐThe ®nancial assistance of the Urban Water Research Association of Australia is gratefully acknowledged. REFERENCES

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