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Jacob, J. Sulfur Analogues of Polycyclic Aromatic Hydrocarbons (Thiaarenes); Cambridge University Press: Cambridge, 1990; Chapter 1, pp 70-87. Fedorak, P. M.; Westlake, D. W. S. Can. J. Microbiol. 1983, 29, 291. Fedorak, P. M.; Westlake, D. W. S. Water, Air, Soil Pollut. 1984, 21, 255. Westlake, D. W. S. In Proceedings of the 1982 International Conference on Microbial Enhancement of Oil Recovery; Donaldson, E. C., Clark, J. B., Eds.; Technology Transfer Branch, Bartlesville Energy Technology Center: Bartlesville, OK, 1983; CONF-8205140; pp 102-111. Williams, J. A.; Bjoray, M.; Dolcater, D. L.; Winters, J. C. Org. Geochem. 1986,10, 451. Fedorak, P. M.; GrbiE-GaliC, D. Appl. Enuiron. Microbiol. 1991, 57, 932. Andersson, J. T. J. Chromatogr. 1986, 354, 83. Campaigne, E.; Cline, R. E. J. Org. Chem. 1956, 21, 39. Bordwell, F.; Lampert, B. B.; McKellin, W. H.J. Am. Chem. Soc. 1949, 71, 1702. Hannoun, M.; BlazeviE, N.; Kolbah, D.; MihaliE, M.; Kajfez, F. J. Heterocycl. Chem. 1982, 19, 1131. Cerniani, A.; Modena, G. Gazz. Chim. Ital. 1959,89,843. Dean-Raymond, D.; Bartha, R. Dev. Ind. Microbiol. 1975, 16, 97. Fedorak, P. M.; Westlake, D. W. S. Appl. Environ. Microbiol. 1986, 51, 435. Fedorak, P. M.; Andersson, J. T. J. Chromatogr. 1992,591, 362. Geneste, P.; Grimaud, J.; Oliv6, L.; Ung, S. N. Bull. SOC. Chim. Fr. 1977, 271. Porter, Q.N. Aust. J. Chem. 1967, 20, 103. Shriner, R. L.; Fuson, R. C.; Curtin, D. Y.; Morrill, T. C. The Systematic Identification of Organic Compounds, 6th ed.; John Wiley & Sons: New York, 1980; p 333.
Bohonos, N.; Chou, T.-W.; Spanggord, R. J. Jpn. J . Antibiot. 1977, 30 (suppl), 275. Kodama, K.; Umehara, K.; Shimizu, K.; Nakatani, S.; Minoda, Y.; Yamada, K. Agric. Biol. Chem. 1973, 37, 45. Mormile, M. R.; Atlas, R. M. Can. J . Microbiol. 1989,35, 603.
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Received for review February 20, 1992. Revised manuscript received M a y 15, 1992. Accepted May 18, 1992. Financial support was provided by Canada Centre for Mineral and Energy Technology, Energy Mines and Resources Contract 23440-09163101-SS and by the Natural Sciences and Engineering Research Council of Canada.
Accumulation of Cobalt, Zinc and Manganese by the Estuarine Green Microalga Chlorella salina Immobilized in Alginate Microbeads Geoffrey W. Garnham, Geoffrey A. Codd, and Geoffrey M. Gadd"
Department of Biological Sciences, University of Dundee, Dundee, DD1 4HN, Scotland, U.K. This paper describes cobalt, zinc, and manganese accumulation by Chlorella salina immobilized in calcium alginate microbeads, investigated by use of the radioisotopes 6oCo, 54Mn,and 65Zn. A rapid biosorption of the metals to C. salina cell walls and the alginate matrix, which was independent of light, temperature, or the metabolic inhibitor CCCP, was followed by a slower energy-dependent phase of uptake. Under similar conditions, immobilized cells accumulated greater amounts of Co,Zn, or Mn than free cells due to an increased active phase of uptake. Accumulation was also dependent on cell density in the alginate beads, with a greater uptake of cobalt at the highest cell densities. Desorption of cobalt from loaded beads was increased by decreasing pH and increasing concentrations of the cations, probably due to exchange of cobalt bound to the cell wall/alginate matrix for H+ or cations. H
Introduction Studies with immobilized microalgae and other photosynthetic microorganisms have mainly focused on the use of immobilized cells for biotransformations and biosynthesis of organic materials (1,2).However, several groups have investigated whether living immobilized microor1764 Environ. Scl. Technol., Vol. 26, No. 9, 1992
ganisms could be useful for the removal of metals and radionuclides from contaminated wastewaters and effluents (3, 4). Immobilized cell systems possess several advantages over freely suspended cells in both batch and continuous-flow systems, including better capability of reuse and regeneration of the biomass, easy separation of cells from the reaction mixture, high biomass loadings within a given bioreactor, manipulation of biomass independent of dilution rate, and minimal clogging in continuous-flow systems (5). Work involving the use of immobilized algae for removal of metals from solution has mainly concentrated on dead cells. For example Nakajima et al. (6) used polyacrylamide-entrappedChlorella vulgaris to remove UO$+,Au3+,Cu2+,Hg2+,and Zn2+,while Darnall et al. (7, 8) used C.vulgaris cells immobilized in a silica gel matrix for removal of A13+,Be2+,Cu2+,Pb2+,Ni2+,Zn2+, Cr3+,Co2+,Fe2+,U022+,Ag+, Hg2+,and Mn2+from solution. Such studies have shown that immobilized dead algae can accumulate at least as much metal as freely suspended dead cells. However, relatively few studies have used living algal cells in immobilized systems for metal removal, and to our knowledge, none has examined active uptake, although there are several examples where greater metal uptake by immobilized, as compared with freely suspended, cells has been demonstrated (9-11).
0013-936X/92/0926-1764$03.00/0
0 1992 American Chemical Society
In this study, we describe the accumulation of cobalt, manganese, and zinc by Chlorella salina cells immobilized in alginate microbeads and compare metal accumulation and some basic aspects of their physiology to freely suspended cells. These elements are required for normal growth of Chlorella species and other microalgae (12)but also occur in the aquatic environment as the neutron activation products 6oCo,54Mn,and 65Znarising from the wastestreams of nuclear installations (4, 13, 14). The ability of microalgae, including C. salina, to accumulate metals from aqueous solution is well documented (5,15, 16). However, relatively little is known about the accumulation of these elements by algae in an immobilized state, which could be important in the movement of such pollutants in aquatic food chains because in natural aquatic environments large numbers of microorganisms, including microalgae, are deposited on and/or attached to solid surfaces as an immobilized biofilm (17,18).
Experimental Section Organism and Growth Conditions. Axenic cultures of Chlorella salina Kufferath (obtained from Dr. G. Russell, Department of Evolutionary & Environmental Biology, University of Liverpool, P.O. Box 147, Liverpool, L69 3BX England) were grown at 23 "C in 10 L of MN culture medium which comprised the following: 0.04 g, MgS04.7HzO;0.02 g, CaC12.2H20;0.75 g, NaNO,; 0.02 g, K,HP04; 0.03 g, citric acid; 0.003 g, ferric ammonium citrate; 0.0005 g, ethylenediaminetetraacetic acid (disodium salt) (EDTA); 0.02 g, Na2C0,; 10 mL, trace metal mix A5 [2.86 g, H,BO,; 1.81 g, MnC12*4H20; 0.079 g, CuS04.5H20; 0.0494 g, Co(N03),.6H20; 0.222 g, ZnS04.7H20; 0.039 g, NazMo04.2H20in 1L of distilled water] in 1L of filtered seawater diluted 3:l with distilled water (16). The medium was adjusted to pH 8 with 1M NaOH and autoclaved (120 "C, 15 min) before being inoculated to approximately 2 X lo5 cells mL-'. Cultures were incubated in 12-L flasks, at 23 "C, on magnetic stirrers, sparged with sterile air and with a photon fluence rate incident on the surface of the flask of 12 peinsteins m-2 s-l provided by white fluorescent light tubes. Production of Microbeads. A microbead maker (obtained from Dr.Gudmund Skjak-Braek, Institute of Biotechnology, University of Trondheim, Trondheim, Norway) was used to produce calcium alginate beads of approximately 0.5-mm diameter. This apparatus uses a high air flow to dislodge drops from the needle of the bead maker where the alginate gel is extruded. Air is forced into a chamber surrounding the needle, escaping from a small opening encircling the needle. It then flows down the outside of the needle, striking and prematurely dislodging gel droplets forming at its end. Bead size can be varied by changing the rate of air flow striking the droplets. The pressure at the top of the chamber is kept to a minimum as gravity alone draws the gel at a sufficient rate for bead formation which can exceed 240 beads min-l. A mixture of 25 mL of sterile sodium alginate and 25 mL of a C. salina suspension from a logarithmic culture (at an appropriate cell density and washed once with 10 mM Ntris(hydroxymethyl)methyl-3-aminopropanesulfonicacid (Taps) buffer, pH 8, to remove Co2+,Zn2+, and Mn2+ present in the growth media) or sterile distilled water was placed in the sterilized bead maker chamber and the top sealed. A rate of filter-sterilized air flow of -30 mL m i d was applied to the top of the bead maker, with the bottom chamber being subjected to an air pressure of 0.25 bar. The beads (0.5-mm diameter) were allowed to drop into cold (-4 "C), sterile 0.2 M CaCl2.2H,O. After 1h in this solution the beads were hard; they were then rinsed three
times with 50 mL of sterile distilled water and transferred to 250-mL Erlenmeyer flasks containing 100 mL of MN medium. Flasks were then placed on an orbital shaker (150 rpm), at 25 "C, with a photon fluence rate incident on the surface of the flasks of 12 peinsteins m-2 s-l provided by white fluorescent tubes for at least 72 h prior to experimental use. Metal Uptake by Microbeads. A 5-mL volume of packed microbeads (containing C. salina cells dispersed throughout the matrix) was obtained by allowing beads to settle out in a 10-mL measuring cylinder and then removing the MN medium by use of a Pasteur pipet. The beads were then resuspended in 10 mL of 10 mM Taps buffer, containing 0.5 M NaC1, and adjusted to pH 8 with 1 M NaOH in an acid-washed 50-mL polythene beaker. This was then placed in the light (30 peinsteins m-2 s-l) at 23 "C, unless stated otherwise, and the contents were mixed. Aliquots from one of 50 or 500 pM CoC12*6H20, MnC12.4H20,or ZnC12.7H20solutions, containing 17 Bq mL-' 6oCo(Amersham International, C0C12concentration 0.012 nM), 185 Bq mL-l 54Mn(Du Pont, MnC12 concentration 0.8 nM), or 50 Bq mL-l 65Zn (Amersham International, ZnC12 concentration 24 nM), respectively, were added to give final metal concentrations in the range 0.5-100 pM. Triplicate samples (200 pL) of suspending buffer were taken at time intervals from the mixed suspension after the addition of metal isotope and placed in 5 mL of Ecoscint A scintillation fluid (National Diagnostics, Maville, NJ). Samples were measured for radioactivity (at an efficiency of a least 30%) using a Packard Minaxi Tricarb 4000 scintillation counter. For calculation of specific activity, three replicate samples of each isotope solution were counted. The amount of metal taken up by the beads was calculated from the reduction in radioactivity in the buffer, after taking into account metal/isotope binding to the plastic beaker. When desired, beads were treated 30 min prior to transfer to buffer with 100 pM carbonylcyanide n-chlorohydrazone (CCCP), incubated for at least 12 h in the dark, or brought to a temperature of 4 "C with melting ice. Desorption of Cobalt. Bead suspensions were prepared in 10 mM Taps buffer, pH 8, with 0.5 M NaCl as described above and loaded with a 'Wo/CoC12 solution to give a final CoC1, concentration of 25 pM. Bead suspensions were then incubated for at least 24 h in the light, as described. Beads were then separated by using fine muslin, and any excess loading solution was removed carefully from the beads with tissue. The separated beads were then washed off the muslin with 10 mL of a specific desorbing solution into a 50-mL plastic beaker. Triplicate samples (200 pL) from the desorption medium were then taken at time intervals after the addition of the beads, and the radioactivity present was measured as above. Measurement of Respiration, Photosynthesis, and Chlorophyll B Content. Rates of respiration and photosynthesis were calculated from measurement of oxygen evolution in the light and oxygen utilization in the dark by free or immobilized C. salina. Changes in oxygen concentration were measured using a Rank oxygen electrode connected to a Servoscribe 1s potentiometric chart recorder. Where appropriate, a light source (1000 Meinsteins m-2 s-l) was provided by use of a slide projector. Chlorophyll a content of free and immobilized cells was determined by placing a 2-mL volume of packed beads or 2 mL of a cell suspension in 5 mL of methanol overnight. The beads were then homogenized in a ground-glass Potter homogenizer, after which homogenized beads and cell samples were both centrifuged (SOOOg, 1 min), and the Environ. Sci. Technol., Vol. 26, No. 9 , 1992
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Flgure 1. Uptake of (a) zinc, (b) manganese, and (c) cobalt by immoblllzed C.salin8 [2 X 10' cells (mL of beads)-'] in 10 mM Taps buffer containing 0.5 M NaCI, pH 8, with a metal concentration of 25 pM at 23 "c: (0)ilght Incubation; (0)dark incubation; (0)incubation at 4 "C; (W) beads pretreated prlor to Incubation In the ilght with 100 fiM CCCP; (A)beads wlthout a ceii loading. Each point is a mean of three repllcates; bars Indicate standard error of mean and when not shown were smaller than the dimensions of the symbols.
absorbance of the liquid phase was read at 663 nm using a Pye Unicam SP600 spectrophotometer. Chemicals and Reagents. All chemicals used were of analytical grade. Inhibitors, buffers, and sodium alginate were supplied by Sigma.
Results Cobalt, Zinc, and Manganese Uptake by Immobilized C.salina. Uptake of each of the three metals from a 25 pM concentration in 10 mM Taps buffer, pH 8, containing 0.5 M NaC1, appeared to be biphasic (Figure 1). An initial phase, in which approximately 10 nmol of Co2+, 6 nmol of Mn2+,and 9.5 nmol of Zn2+ (lo6 cells)-l was accumulated within 15 min, was followed by a second slower phase in which a further 6 nmol of Co2+,8 nmol of Mn2+,and 8 nmol of Zn2" (lo6 cells)-' was accumulated over 5 h (Figure 1). The initial phase of uptake appeared independent of light, temperature, or pretreatment of the beads with the metabolic inhibitor CCCP, indicating this process to be passive. In contrast, the second phase of uptake was clearly dependent on light and could be inhibited by CCCP and by low temperature, indicating that this uptake was an energy-dependent process. Greater uptake of all three metals occurred in beads in the light with a cell loading 2 X lo6cells (mL of packed beads)-' as compared with uptake of the metals by unloaded beads (Figure 1). However, there was little difference between metal uptake by unloaded and loaded beads incubated in the dark, in the presence of CCCP, or at 4 "C. Metal uptake by immobilized cells [calculated by subtracting values for alginate binding (obtained from uptake of metals by beads without cells) from values of binding by beads containing cells] from an initial concentration of 25 pM 1766 Envlron. Scl. Technol., Voi. 26, No. 9, 1992
Time (h)
Figure 2. Uptake of (a) zinc, (b) manganese, and (c) cobalt by (0) lmmobillzed cells and (0)freely suspended cells of C. salina , both at a density of 1 X lo6 cells mL-l In 10 mM Taps buffer containing 0.5 M NaCI, pH 8, wlth a metal concentratlon of 25 pM at 23 OC. Each point is a mean of three repllcates; bars indicate standard error of mean and when not shown were smaller than the dimensions of the symbols.
was greater than metal uptake by an equal amount of free cells in an equal volume of buffer [data for uptake by free cells taken from Garnham, Codd, and Gadd (19)] (Figure 2). Influence of Concentration and Bead Cell Density on Cobalt Accumulation. At all cobalt concentrations examined, the initial, metabolism-independent phase of uptake increased with increasing cobalt concentration in both unloaded beads and those loaded to a density of 2 X lo6 cells (mL of packed beads)-'. The amount of cobalt taken up during this phase was similar in both unloaded and loaded beads at each concentration (Figure 3). A typical straight-line Freundlich adsorption isotherm (20) was obtained for both loaded and unloaded beads (Figure 3). The amount of cobalt taken up in the metabolismdependent phase increased with increasing cobalt concentration. When the initial rates of uptake, taken to be the maximal initial rate of uptake after the initial binding was complete (after -5 min), were plotted according to Lineweaver and Burk (21),a straight line was obtained from which a K, value of 60 pM and a V,, of 55 pmol m i d (lo6cells)-l was calculated (Figure 4). The kinetics of cobalt uptake by free-living cells of C. salina (19)describe a different biphasic uptake system, as shown in Figure 4. As the cell packing of the beads waa increased to greater than 1 X lo6 cells (mL of packed beads)-', the amount of cobalt taken up over a 4-h period also increased (Figure 5). However, at a density of 1 X lo5 cells (mL of packed beads)-l, uptake was less than that for unloaded beads over a 4-h period. The greatest uptake occurred with a cell density of 1 X lo8 cells (mL of packed beads)-l, and it was difficult to produce beads of a cell density higher than this.
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Log c Figure 9. Freundllchplot of cobalt blosorptlon by (0)CelCfree and (0) C. sallna loaded alginate microbeads In 10 mM Taps buffer Containing 0.5 M NaCI, pH 8, at 23 OC. Each point Is a mean of three replicates with a standard error not greater than 0.05. qe, the quantity of cobalt adsorbed per mass of adsorbant at a fixed temperature; c, the concentration of cobalt remaining In solution at equilibrium.
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Figure 4. Llneweaver-Burk plot of cobalt uptake by C . salina at a cell density of 1 X lo6 ml-' In 10 mM Taps buffer containing 0.5 M NaCI, pH 8, at 23 OC: (0)cells Immobilized In calcium alglnate; (0)freely suspended cells. Values are means of three replicates with a standard error of
