Environ. Sci. Technol. 2007, 41, 7363-7369
Bioconcentration Factor Hydrophobicity Cutoff: An Artificial Phenomenon Reconstructed MICHIEL T. O. JONKER* AND STEPHAN A. VAN DER HEIJDEN Institute for Risk Assessment Sciences, Utrecht University; P.O. Box 80177, 3508 TD Utrecht, The Netherlands
The debate on whether highly hydrophobic organic chemicals (with log Kow > 5-6) bioconcentrate less than may be expected from their hydrophobicity is still not settled. The often-observed hydrophobicity “cutoff” might either be explained by artifacts occurring during bioconcentration factor (BCF) measurements or by a true mechanism, i.e., reduced uptake of larger molecules due to decreased membrane permeation. In this paper, we advocate there is no hydrophobicity cutoff, at least not for compounds with log Kow of up to 7.5. Data are presented on the uptake of polycyclic aromatic hydrocarbons (PAHs) in the aquatic worm Lumbriculus variegatus. For this combination of chemicals/organism, BCFs were measured using several approaches, including traditional batch uptake kinetics measurements and alternative ones, involving solid-phase microextraction (SPME), polyoxymethylene solid-phase extraction (POM-SPE), field exposures, and the substitution of living worms by dead worm material or liposomes. A hydrophobicity cutoff was observed at two levels during the traditional approach only, whereas for the other approaches it was absent. The data were used to demonstrate the presence and impact of artifacts due to so-called “third phase effects” and nonequilibrium conditions that can obscure “true uptake”. The experiments suggest that previously observed cutoff effects can be ascribed to artifacts, and that current risk assessment (often incorporating a BCF cutoff) as well as BCF measurement techniques of very hydrophobic chemicals should be revised.
Introduction Bioconcentration of chemicals is a key process in ecotoxicology, because it serves as the link between exposure and effects of most toxicants and thereby generally controls toxicity in organisms. Bioconcentration of nonionic, hydrophobic organic chemicals (HOCs) that cannot be (easily) metabolized (e.g., PCBs, dioxins, chlorobenzenes, brominated flame retardants) is usually considered as a simple partitioning process between water and the organism’s lipid pool, i.e., membranes and storage fat. As such, it is believed to be solely driven by the chemical’s hydrophobicity and to be characterized by the octanol-water partition coefficient (Kow), being a simple measure of this chemical property. Numerous studies have shown that in many cases bioconcentration of HOCs indeed can be described by this parameter (e.g., (1, 2)); however, for highly hydrophobic HOCs with log Kow values * Corresponding author e-mail:
[email protected]; phone: +31 30 2535338; fax: +31 30 2535077. 10.1021/es0709977 CCC: $37.00 Published on Web 09/27/2007
2007 American Chemical Society
greater than 5.5-6, the picture is not so clear. After two decades of bioconcentration research, there is still an active debate on the relationship between bioconcentration and hydrophobicity beyond log Kow of 5.5-6. Logarithmic bioconcentration factors (log BCFs, i.e., the logarithm of the ratio of a chemical’s concentration in organism to that in the water phase) would either (a) level off or decrease (1, 3, 4), or (b) still be proportional with log Kow values (1, 5). Argumentation for possibility (a) is that very hydrophobic chemicals have such large molecular dimensions that they might not be able to (easily) permeate membranes, as the energy for cavity formation is too high (6, 7). This size exclusion phenomenon is generally referred to as the “bioconcentration (factor) hydrophobicity cutoff”, even though it does not involve a true cutoff (i.e., observations concern a leveling-off or gradual decrease, not a sudden drop to zero). Obvious implication of this cutoff is that very hydrophobic chemicals would not, or only to a lesser extent, exert toxic effects. In many risk assessment models, a cutoff has been implemented and bioconcentration is modeled by a nonlinear (polynomial, bilinear, parabolic) equation (8, 9). Advocates of possibility (b) presume that the cutoff phenomenon results from artifacts occurring during BCF measurements (10) and that actual bioconcentration should be proportional with hydrophobicity, even beyond log Kow of 5.5-6. Recently, a number of papers have been published that support this second possibility. For instance, Kraaij et al. (11) and Van der Wal et al. (12) observed log BCFs of several chemicals taken up by worms to be proportional with the chemicals’ log Kows up to log Kow of about 7.5-8. Jabusch and Swackhamer (13) found that PCBs with log Kow > 6 could be taken up by algal cells and that uptake was only kinetically restricted. Later on, these authors reported membrane-water partition coefficients for PCBs that were proportional with log Kow values up to log Kow of about 8 (14). Finally, Sobek et al. (15) showed that plankton-water partitioning was proportional with hydrophobicity up to a log Kow of about 7.5. All in all, there is still no consensus on whether and how very hydrophobic organic chemicals accumulate in organisms. This paper aims at contributing to the debate by arguing that the bioconcentration hydrophobicity cutoff often being observed at log Kow of 5.5-6 is not caused by a true biological or physicochemical mechanism, but by experimental artifacts instead, being nonequilibrium conditions and so-called “third-phase effects” (i.e., overestimation of aqueous HOC concentrations by the presence of and HOC binding to dissolved organic matter). In contrast to previous attempts based on assumptions and/or models (10, 16), we will actually reconstruct these artifacts on the basis of experimental bioconcentration measurements.
Experimental Design ‘Bioconcentration factors’ were determined for 13 polycyclic aromatic hydrocarbons (PAHs; 4.6 < log Kow < 7) and the aquatic worm Lumbriculus variegatus, following six different experimental approaches. These included (1) a traditional batch-shake setup, in which concentrations in worms and concentrations in water (as determined by solvent extractions) were measured as a function of time (0-11 days). Eleven days were set as maximum exposure time, as the worms started to decay beyond this point; (2) a batch-shake setup similar to the one used in approach 1, but with a fixed exposure duration of 11 days and quantification of the freely dissolved aqueous concentrations by solid-phase microexVOL. 41, NO. 21, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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traction (SPME); (3) a batch-shake setup similar to the one used in approach 2, but with the addition of cellulose pulp as substrate for the worms to increase the organisms’ survival time, allowing a prolonged exposure duration of 4 weeks; (4) exposures of worms and SPME fibers for 4 weeks in enclosures inserted at two field locations, investigating bioaccumulation in situ; (5) a batch-shake setup in which dead worm homogenate was exposed for 4 weeks, testing the hypothesis that bioconcentration is a passive partitioning process. Aqueous concentrations were determined by either SPME or polyoxymethylene solid-phase extraction (POM-SPE; (17)) in order to obtain data for different phase (worm material: water:solid-phase extraction material) distribution ratios; (6) a batch-shake setup similar to the one used in approach 5, but with liposomes instead, investigating whether these artificial membranes can be used as a surrogate phase in bioconcentration studies.
Materials and Methods Chemicals and Solid-Phase Extraction Materials. Solvents used (hexane, acetone, methanol, and acetonitrile) were obtained from Lab-Scan (Dublin, Ireland) and were of Pestiscan or HPLC grade. Testing chemicals (phenanthrene, anthracene, fluoranthene, pyrene, benz[a]anthracene, chrysene, benzo[e]pyrene, benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[a]pyrene, benzo[ghi]perylene, dibenz[ah]anthracene, and indeno[1,2,3-cd]pyrene; all >98%) were from Aldrich (Steinheim, Germany). Other chemicals used were 2-methylchrysene (99.2%; BCR, Geel, Belgium), sodium azide (extra pure; Merck, Darmstadt, Germany), calcium chloride dihydrate (analytical grade; Merck), and aluminum oxide (Super I; ICN Biomedicals, Eschwege, Germany). Prior to use, aluminum oxide was deactivated with 10% (w/w) of Millipore water. Polydimethylsiloxane (PDMS)-coated, disposable SPME fiber (glass fiber core diameter 110 µm, PDMS coating thickness 28.5 µm) was obtained from Poly Micro Industries (Phoenix, AZ). Prior to use, the fiber was cut into pieces of 3 or 5 cm length, which were washed three times with methanol and three times with Millipore water, respectively. Polyoxymethylene (POM) was obtained from Vink Kunststoffen BV (Didam, The Netherlands) as a sheet. The sheet was cut into strips of desired dimensions, which were extracted by shaking with hexane (30 min) and methanol (3 × 30 min), after which they were air-dried (17). Organisms. Aquatic oligochaete worms (Lumbriculus variegatus) were reared in the laboratory in 25 L aquaria at 24 °C. Pulverized, chlorine-free cellulose sheets served as substrate and the aquaria were continuously flushed with preheated, copper-free water. Once a week, the organisms were fed with pulverized flake fish food. Prior to the accumulation experiments, organisms were separated from the substrate and allowed to clear their guts overnight in gently flowing water. For experiments with dead organisms, gut-purged worms were frozen, freeze-dried, and finally ground in a mortar. L. variegatus was chosen as test organism, as it is a widely used species that has been recommended by the US EPA for bioaccumulation studies. The worms probably do not or only minimally metabolize PAHs (18, 19). Liposomes. Small unilamellar liposomes made up of synthetic 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine (POPC) and with a diameter of around 125 nm were used as an artificial model membrane phase. Preparation procedure (freeze-thawing in liquid nitrogen and membrane extrusion) and characteristics of these liposomes are described in (20). Liposomes were prepared in a 0.01 M CaCl2/ 50 mg/L NaN3 solution at EAWAG (Switzerland) and transported (cooled) to Utrecht (The Netherlands) by courier. Liposome content of the solution was determined gravimetrically and by a total phosphate determination (20). Both methods resulted in identical values. 7364
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Determination of BCFs. The six different experimental approaches that were applied to measure ‘bioconcentration factors’ are described below. To prevent PAH photolytic degradation, all laboratory equilibrations were performed in the dark, and brown/covered glassware was used during sample treatment (extractions, cleanup, analyses). Organisms were not fed during exposures. Approach 1: Batch-Shake Method, Including Kinetics. Thirteen 500 mL glass-stoppered Erlenmeyer flasks were filled with water and spiked with 50 µL of a cocktail solution of 13 PAHs in acetone. Resulting aqueous concentrations did not exceed the chemicals’ solubilities. After thoroughly homogenizing the solutions on an orbital shaker for 18 h, small quantities of worms (ca. 1.4 g wet weight) were added to each flask. Then, all flasks were closed again and placed on the orbital shaker at 90 rpm and 20 °C. Systems were allowed to equilibrate for either 0.16, 0.25, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, or 11 days. From each system, a 100 mL water sample was extracted by shaking with three subsequent batches of hexane (4, 3, 3 mL) for several hours on a mechanical shaker. The pooled extract was then concentrated under nitrogen and exchanged to acetonitrile, after which an internal standard solution (2-methylchrysene) was added. Recoveries for this water extraction procedure were determined to be 91-96% (depending on the chemical). Final results were all adjusted for both blanks and extraction recoveries. Worms were collected from the Erlenmeyer flasks, frozen, freeze-dried, and homogenized in a mortar. Approach 2: Batch-Shake Method Combined with SPME. Three 500 mL Erlenmeyer flasks were prepared as described above. Together with the worms, two 5-cm SPME fibers were added, and equilibration time was 11 days (no kinetics determined). After these 11 days, SPME fibers were collected, wiped with wet tissue, and transferred to amber-colored HPLC vials filled with 180 µL of acetonitrile in a 250 µL insert. Then, 20 µL of internal standard solution was added, and the vials were vortexed for 1 min and stored at -18 °C until analysis. Prior to the analysis, vials were defrosted, vortexed again, and left at room temperature for 1 day. PAH extraction recoveries (from fibers) using this method were previously determined to be 99.6 ( 0.1% (21). Worms were collected and treated as described above. Water phase extractions were not performed, as aqueous concentrations were derived from SPME data (see Results section). Approach 3: Batch-Shake Method with SPME, Substrate, and Prolonged Exposure Time. Chlorine-free cellulose sheets were shredded, extracted with hexane by intensive shaking with two subsequent batches of hexane on a horizontal shaker for 5 h, and dried overnight in an oven at 80 °C. Subsamples (3.5 g) of the resulting material were suspended in water, pulverized with a blender, and quantitatively transferred to four 500 mL Erlenmeyer flasks, which were then filled up to contain 350 mL of water. The water-cellulose systems were spiked with 100 µL of a cocktail solution of 13 PAHs in acetone and shaken on a horizontal shaker for 3 days. Then, worms (ca. 3.0 g wet weight) and SPME fibers (2 × 5 cm) were added and the closed systems were equilibrated on an orbital shaker for 4 weeks at 90 rpm and 20 °C. After two weeks, the headspace of the bottles was purged with oxygen to ensure worm survival. Upon finishing the 4-week equilibration period, worms were separated from the cellulose and kept in clean water for 8 h to allow gut clearance, after which they were frozen, freeze-dried, and homogenized. SPME fibers were collected and treated as described above. Approach 4: In Situ Exposures. In the summer of 2004, laboratory-reared worms were exposed in situ in stainless steel, open-bottom, 1 mm-mesh covered enclosures (16 × 16 × 25 cm [L × W × H]) at two Dutch field locations (22): a rural ditch, with PAHs most probably originating from creosote-impregnated sheetpiling (location 1), and a branch
of the river “Hollandsche IJssel” (location 2), receiving PAHs mainly from a large diesel-powered pumping-station located 100 m upstream. In addition, SPME fibers were statically exposed at the locations, by vertically inserting stainless steel mesh devices containing the SPME fibers in the upper 3-cm sediment layer (22). Both worm and SPME exposures were performed in quadruplicate at each location and lasted for 4 weeks. These 4 weeks appeared sufficiently long for the SPME fibers to reach equilibrium, as demonstrated by additional sorption tests in the lab (22). Postexposure activities included manual retrieval of worms from the collected sediments, and worm and SPME fiber treatment as described above. Approach 5: Dead Worm-Water Partitioning. Freezedried, homogenized worm material (25-100 mg) was weighed into 250 mL amber-colored all-glass bottles, which were subsequently filled with water containing 50 mg/L of sodium azide and 0.01 M of calcium chloride. Then, SPME fibers (3 × 5 cm) or POM strips (0.3 g) were added and the bottles were spiked with a cocktail solution of 13 PAHs in acetone. Resulting systems were shaken for 4 weeks at 100 rpm and 20 °C, after which SPME fibers or POM strips were removed. Handling of the SPME fibers was as described above; POM strips were treated as described in ref 17. Experiments were performed for different worm material:water ratios (1.0 and 0.4 g/L for the POM-SPE setup, and 0.09 g/L for the SPME setup). Original, unexposed worm material was analyzed for PAHs, and resulting concentrations were adjusted for during data analysis (i.e., included in the mass balance, which only for phenanthrene and pyrene resulted in a significant contribution of 1-3%). Every measurement was replicated 4-6 times. Approach 6: Liposome-Water Partitioning. Setup for this approach was basically identical to that of approach 5, except for the fact that a liposome solution was weighed into the bottles. Again, both SPME and POM-SPE were applied, equilibration time was 4 weeks, and tests were conducted at two different liposome:water ratios per method (factor of 5 or 10 difference; 0.02-0.2 g liposomes/L). Each determination was replicated 4 or 5 times. Extractions of Worms for PAH and Lipid Analyses. Subsamples of approximately 60-100 mg of freeze-dried worm material were Soxhlet-extracted with a hexane:acetone (3:1) mixture for 16 h. Extracts were concentrated, cleanedup over aluminum oxide columns, and exchanged to acetonitrile. Finally, 2-methylchrysene was added as internal standard. Results were all corrected for extraction/cleanup recoveries and blanks. Lipid contents of the worms were determined gravimetrically by Soxhlet-extraction of 30-100 mg of sample with a hexane/acetone (3:1) mixture for 16 h, and evaporation of the extracts to dryness. Results were corrected for procedural blanks. Instrumental Analysis. PAHs were analyzed on a HPLC system, consisting of a Varian Prostar 420 autosampler, a Gynkotec P580HPG HPLC pump, a Jasco FP-920 fluorescence detector, and a Vydac 201TP54 C18 column (kept at 25.0 °C). The mobile phase consisted of a mixture and flow gradient of methanol and water, being degassed by a Shimadzu DGU14A degasser.
Results In Figure 1, results of the PAH uptake kinetic measurements (i.e., the conventional batch method; approach 1) are presented for three representative PAHs. In all cases, the ratio Cb/Cw (i.e., lipid-normalized concentration in biota/ concentration in water) gradually increased but appeared to reach a plateau after approximately one week. In uptake studies, a similar plateau is usually considered to represent equilibrium (23), even though this condition may be apparent
FIGURE 1. Uptake kinetics of PAHs in Lumbriculus variegatus as determined with approach 1. Graph shows the logarithm of the ratio of lipid-normalized PAH concentration in worms (Cb): concentration in water (Cw; as determined by hexane extraction) as a function of time (h) for phenanthrene (open circles), benzo[e]pyrene (solid squares), and indeno[1,2,3-cd]pyrene (solid triangles). (see below). Following the standard approach, however, ratios obtained after one week were assumed to be appropriate for BCF calculations. Hence, they were averaged and plotted as lipid-normalized log BCFs against log Kows in Figure 2a. Lipidnormalized BCF values resulting from approaches 2-4 are plotted in Figures 2b-d. Aqueous concentrations in these cases were obtained by combining measured PAH concentrations in SPME fibers (Cf) and SPME fiber-water partition coefficients (Kf; see Table S1: Cw ) Cf/Kf). Lipid-normalized dead worm-water partition coefficients (hereafter referred to as necroconcentration factors, NCFs, as opposed to bioconcentration factors) and liposome-water partition coefficients (Klip) are presented in Figures 2e and 2f, respectively. Both NCF and Klip values were calculated using the mass-balance approach presented in ref 17, i.e., the aqueous concentration of a chemical was determined by means of SPME or POM-SPE, and the accompanying concentration in worm material or liposomes was calculated by subtracting the sum of the chemical mass in water and in POM or SPME fiber from the spiked (total) chemical mass. All BCF, NCF, and Klip values are presented in Table S1. Figure 2 shows a clear BCF cutoff for approach 1 (Figure 2a): log BCF values increase up to log Kow of about 5.5 but then level off, and beyond log Kow of about 6, they decrease. For approach 2 (Figure 2b), a cutoff is observed at log Kow of about 6, but in this case, log BCFs only level off: all values beyond the cutoff measure approximately 6. Finally, for all other approaches (3-6), no cutoff is observed and relationships between log BCF and log Kow are linear (see Figures 2c-f).
Discussion Reconstruction and Explanation of the Cutoff. In Figure 3, data from Figures 2a-f are pooled. This figure clearly shows the above-mentioned three different log BCF-log Kow relationships, as emphasized by manually projected lines: (1) a cutoff involving both a leveling-off and a decrease around log Kow of 5.5-6, as observed for approach 1. BCFs measured with this approach were obtained after a short equilibration time (7-11 days) and by applying a solvent (total) extraction of water for the determination of Cws; (2) a cutoff involving a leveling-off only at log Kow of 6, as observed for approach 2. Here, BCFs were obtained by applying a short equilibration time (11 days) and SPME for the determination of aqueous concentrations; (3) a linear log BCF-log Kow relationship for approaches 3-6. In these cases, extended equilibration times VOL. 41, NO. 21, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 2. Lipid-normalized log ‘BCF’-log Kow plots resulting from the six different ‘BCF’ measurement approaches: (a) batch-shake method; (b) batch-shake method with SPME; (c) batch-shake method with SPME, substrate, and prolonged exposure time; (d) in situ exposures (solid circles, location 1; open circles, location 2); (e) dead worm-water partition coefficients (NCFs; open squares are POM-SPE-derived; solid squares are SPME-derived); (f) liposome-water partition coefficients (Klip; open triangles are POM-SPE-derived; solid triangles are SPME-derived). Dotted lines represent the 1:1 relationships. were applied (4 weeks) and aqueous concentrations were measured with SPME and/or POM-SPE. The only difference between approaches 1 and 2 is the way in which Cw was determined. Accordingly, this difference must have caused the different BCF trends described under 1 and 2, i.e., the divergence of the lower two lines in Figure 3. Obviously, solvent extractions resulted in higher Cws than SPME fiber measurements for PAHs having a log Kow >5.5, with the difference increasing with increasing PAH hydrophobicity. This dissimilarity can be explained by the fact that SPME solely quantifies the freely dissolved aqueous concentration (24), which for these chemicals must have been lower than total concentrations. This phenomenon is analogous to that often observed during sorption studies with HOCs and sediments or soils, where dissolved organic carbon (DOC) molecules originating from the sorbents can cause lower-than-expected sediment- or soil-water distribution coefficients (with the deviation increasing with sorbate hydrophobicity) (25, 26). HOCs associate with these water-soluble macromolecules, which thereby increase the overall (solvent-extractable) HOC concentration in water, leading to an overestimation of Cw. This overestimation is most pronounced for the more hydrophobic HOCs, as HOCDOC binding is mainly driven by hydrophobic interactions (27). In approaches 1 and 2, no sediment or soil was present, but the worms will have excreted DOC-like biomolecules (e.g., lipids or proteins) or small tissue fragments into the aqueous phase. Although not visible in the exposure systems, these compounds became noticeable during solvent extractions, where considerable emulsification was observed at the hexane-water interphase, which increased in size with exposure time (emulsions generally could be counteracted by addition of hydrochloric acid, though, allowing a reliable extraction). The artifact induced by the presence of these biomolecules will hereafter be referred to as ‘third phase artifact’, and it causes the typical drop in log BCF-log Kow curves at higher hydrophobicities (see Supporting Informa7366
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tion for a modeling approach demonstrating the third phase effect on log BCF-log Kow curve shape). Although in approach 2 the third phase artifact was circumvented by using SPME, the exposure time in this setup (11 days) was too short to allow full worm-water equilibration for all PAHs, as demonstrated by the fact that the log BCFlog Kow relationship for this approach levels off at log Kow ≈ 6, whereas for approaches 3-6 there is no leveling off (upper line in Figure 3). Even though there are several differences between approaches 2 and 3-6, exposure time is the only crucial one. In fact, approach 2 closely resembles the NCF approach (approach 5), where no substrate was present as well, and initial aqueous concentrations dropped due to uptake in worm tissue (see last section). For approaches 3-6, full system equilibrium can be assumed, since a linear log BCF-log Kow relationship reflects equilibrium as long as the chemicals under consideration have the same mode of interaction with the sorbent (28), as in the present study. Hence, for all chemicals tested in this study, having log Kows up to 7, four weeks (either under well-agitated or static, in situ conditions) was sufficiently long to reach equilibrium. In contrast, for approach 2 a bilinear curve is observed. Assuming full SPME fiber-water equilibration for all PAHs after 11 days in these shaken systems (21), the leveling-off at log Kow of 6 indicates that PAHs with log Kow < 6 achieved worm-water equilibrium, whereas for the more hydrophobic ones the ratio Cb/Cw was constant. This constant ratio is explicable, as initial aqueous concentrations of all PAHs were identical and uptake kinetics are similar for different PAHs (29), but it clearly reflects nonequilibrium conditions. As time goes on, the point of leveling-off will shift toward higher Kows, slowly extending the curve’s linear range. In summary, the difference between the upper two lines in Figure 3 resulted from worm-water nonequilibrium conditions (in Figure 3 referred to as nonequilibrium artifact). Analogous to the third phase artifact, this difference increases with log Kow (lines diverge), in this case, however, because
FIGURE 3. Combined log ‘BCF’-log Kow plot, showing data resulting from all six approaches used to measure ‘BCFs’. Lines are drawn by hand to indicate and distinguish between two major artifacts that can bias BCF measurements. Note that the braces are indicative only and do not point out that, for example, any solid markers under the upper line reflect nonequilibrium per se (i.e., variation around the lines is not taken into account by the braces). the time needed to reach equilibrium increases with compound hydrophobicity. It should be stressed that the precise shape and absolute height of the lower two curves in Figure 3 are strictly conditional: they apply to the present experiments only and cannot be generalized. Shape and height may vary due to differences in the extent of the artifacts or due to the occurrence of combined effects. Also, although the middle log BCF-log Kow relationship in Figure 3 reflects nonequilibrium conditions in the present study, a similarly shaped curve may represent third phase effects in experiments that are being performed under full equilibrium conditions (see modeling approach in Supporting Information). Finally, it should be mentioned that although the above discussion focuses on ‘static’ approaches, the artifacts discussed also apply to ‘flow-through’ setups (mostly used for exposure of fish). In such setups, nonequilibrium conditions are the most probable artifacts; the occurrence of third phase effects will depend on both water refreshment rate and water sampling point. Implications for Risk Assessment of Very Hydrophobic Chemicals. The present results once more demonstrate that accumulation of very hydrophobic chemicals in organisms does occur and that uptake is controlled by hydrophobicity (11-13, 15). If properly and accurately measured, no cutoff is observed around log Kow of 5.5-6, and the obtained log BCF-log Kow relationship will be linear up to at least log Kow of 7.5-8. Although Figures 2 and 3 show data up to log Kow of 7 for PAHs, NCFs were also measured with POM-SPE for PCBs with log Kows of up to 7.5 (see Supporting Information, Figure S3). The log NCF-log Kow relationship in this case was fully linear as well. The above has several implications. First of all, the present data strongly suggest that previously observed cutoff phenomena for nonmetabolizable HOCs can be ascribed to artifacts, being either third phase effects, kinetic restrictions, or both. This not only relates to organisms (e.g., worms and fish) but also to artificial phases, such as liposomes (7) and SPME fibers (30). Of course, other artifacts may occur during ‘BCF’ measurements as well (e.g., sorption of HOCs to filters when separating liposomes and water (14), which may actually force log Klip-log Kow curves upward), but a detailed discussion on these confounding factors is beyond the scope of the present paper. Second, if very hydrophobic HOCs do bioconcentrate, they will be able to express toxicity, if not via a specific then at least via a nonspecific mode of action, i.e., the chemicals
will at least act as baseline, narcotic toxicants. The fact that several studies have labeled many very hydrophobic HOCs as nontoxic might for the greater part be explained by nonequilibrium conditions, as recently suggested by Mayer and Reichenberg (31). Third, the absence of a cutoff implies that bioaccumulation models in current HOC risk assessment protocols, being based on nonlinear log BCF-log Kow relationships (8, 9), are incorrect. Such models will underpredict actual risks for chemicals with log Kow > 6, with the error sharply increasing with chemical hydrophobicity. Also note from Figures 2c-f that log BCFs might be (much) higher than log Kows (up to 1.2 log unit in this study), thereby also annulling models assuming BCFs to be equal to or lower than Kows. All in all, it is advisable to reconsider current BCF models. If the environmental conditions and the organism’s lifespan permit, very hydrophobic chemicals most likely are extremely bioaccumulative. Although the present data indicate there is no BCF cutoff up to log Kow of about 7.5-8, it cannot be concluded there is no cutoff at all. Obviously, there will be a point above which the chemical’s molecular dimension does not permit permeation of membranes anymore, but the question is whether such a point exists within the hydrophobicity/size range applicable to (anthropogenic) HOCs for which risk assessment is necessary (e.g., brominated flame retardants, dioxins, petrochemicals). Future research will have to reveal the possible existence and relevance of such a point. Alternative Methods for Measuring BCFs. On the basis of the results obtained in this study, it is recommended not to use traditional (short-term) approaches for the determination of BCFs of HOCs with log Kow > 5, as both third phase and nonequilibrium artifacts may be expected. Reliable BCFs can only be obtained by applying alternative approaches that take these artifacts into account. As discussed above, third phase artifacts can simply be circumvented by using a “passive sampler”, such as an SPME fiber or POM strip for the determination of freely dissolved aqueous concentrations. Also for less hydrophobic chemicals, passive sampler extractions are useful, as they are less laborious than solvent extractions. Note that solvent extractions and SPME yielded indistinguishable BCFs for the four least hydrophobic PAHs studied in this work (phenanthrene, anthracene, fluoranthene, pyrene; see Figures 2a and 2b). In addition, an important advantage of SPME is that it allows for the determination of BCFs in situ (approach 4), yielding truly environmentally relevant BCFs. VOL. 41, NO. 21, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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The solution for the second, nonequilibrium artifact seems even more simple, just increasing equilibration time, but because of biological factors this might be problematic in many cases. Exposing organisms in water-only systems for a prolonged time might result in premature death due to stress (caused by a combination of, e.g., the absence of substrate and food, and the presence of chemicals), but also the lifespan of test organisms might simply be too short (e.g., in case of Daphnids or algae). In the latter case, NCFs might offer an alternative (see below); however, the environmental relevance of the results will be questionable, as steady state might not be reached for very hydrophobic HOCs under field conditions. In the first case, survival of benthic organisms can be increased by adding substrate (cellulose or sediment), as done in approach 3 (and 4). By definition, this will yield bioaccumulation factors (BAFs) instead of BCFs, as the test chemicals will also be taken up by ingestion of contaminated substrate. However, note that for benthic organisms, BAFs in fact are the only factors of importance. Although there is no consensus yet on whether uptake via contaminated substrate would increase final Cbs (32), the present data suggest that this is not the case for L. variegatus. After all, a comparison between passive partitioning approaches (i.e., approaches 5 and 6, where uptake is via the water phase only) and approaches 3 and 4 (where PAHs are taken up via substrate as well) does not reveal higher values for the latter (cf. Figures 2c and 2d vs Figures 2e and 2f). In contrast to equilibrium conditions, kinetics will be affected by consumption of contaminated substrate (uptake rates will increase) (32), but this will only result in reduced equilibration times, which should be considered an advantage. Another benefit of using substrate is that it might serve as a buffering source of chemicals, maintaining a constant aqueous concentration. Obviously, this only holds when (1) the sorption capacity of the substrate dominates over that of the organisms, and (2) desorption rates from the substrate are not rate-limiting in the overall equilibration process, as might be the case for field-contaminated sediments from which desorption is often very slow (33). In fact, the latter might explain the somewhat lower values of the last two data points in Figure 2d (in situ exposures). With cellulose, such slow desorption is not plausible, but in the present study (approach 3), this material’s sorption capacity (34) was low as compared to that of the worms. Therefore, aqueous concentrations in approach 3 were not stable (i.e., a significant fraction of the (in particular high-molecular-weight) chemicals partitioned into the worms). Stable concentrations are often defined as requisite for the determination of BCFs for HOCs; however, this condition is questionable, as it does not affect final equilibrium distribution (32). In case initial aqueous concentrations drop due to significant partitioning into organisms, kinetics might be increased (32), but final equilibrium distribution will be identical as compared to situations with stable Cws. The present data support this view by showing identical BCFs for less hydrophobic PAHs in case of constant Cw (approach 4; Figure 2d) and declining Cw (approaches 1 and 2; Figures 2a,b), as well as for the more hydrophobic PAHs (cf. approach 4 with stable Cws versus approaches 5 and 6 with strongly decreasing Cws).
indistinguishable (t-test) from BCFs obtained by approach 3 (substrate + SPME) and 4 (in situ exposures). Therefore, using a dead organism phase might be a promising alternative method for measuring ‘BCFs’. However, NCFs for anthracene and benzo[a]pyrene unfortunately could not be determined, as these chemicals appeared to have largely been degraded during equilibration. Interestingly, these PAHs are the most photosensitive ones, but photodegradation was ruled out (recovery determinations showed no degradation). Therefore, they were probably oxidized otherwise, possibly through tissue-generated radicals. Attempts to counteract formation of these presumable radicals (by replacing acetone with methanol as spiking solvent, and sodium azide with mercury chloride as biocide) and to neutralize them (by adding ascorbic acid or R-tocopherol acetate on a PAH-equal molar basis to the systems) did not, however, yield different results (not shown). Future experiments may unravel this degradation mechanism. The liposome approach (approach 6) did not show this problem and therefore might be preferred over the NCF approach, as it does have the same advantages. Moreover, resulting Klip values are statistically indistinguishable (t-tests) from NCFs (both POM-SPE and SPMEdetermined). The same applies to Klip values and BCF values obtained by approach 3. It should also be noted that identical Klip values were determined with POM-SPE and SPME (overlapping open and solid markers in Figure 2f) and that values determined at different liposome:water ratios were the same. This suggests a true partitioning process and clearly indicates the validity of the mass-balance approach applied, as vastly different phase distribution ratios were covered (e.g., percent of chemical absorbed by SPME fiber or POM ranged from 0.04-90). Although different sorbent-water ratios resulted in identical NCFs as well, POM-SPE and SPME applications resulted in significantly different NCF values (paired t-test; see Figure 2e). It is beyond the scope of this paper to discuss this difference in detail; however, fouling, which might occur for SPME (24) with this fine-particulate, sticky dead organism phase, but which is not expected for POM-SPE (17), might be the explanation. Finally, the observation that Klip and NCF values for the two least hydrophobic PAHs tested (phenanthrene and anthracene) are higher than observed for approaches 1-4 using living worms might either be caused by experimental artifacts (loss of these relatively soluble/volatile compounds during gutpurging and/or freeze-drying of worms in approaches 1-4) or by metabolic losses. In the first case, surrogate phase approaches clearly are the superior methods; if the second is true, the use of such phases will overpredict BCF values of metabolizable chemicals.
Another way in which exposure times might be extended is to work with a surrogate organism phase, i.e., dead organisms or liposomes (approaches 5 and 6). When using such a surrogate approach, it is implicitly assumed that HOCs are taken up in organisms by passive hydrophobic partitioning from the water phase only (which seems to be the case here; see above). The major advantage of this approach is that a biocide can be added, allowing exposure times to be extended, so that even for very hydrophobic HOCs full equilibrium can be achieved, without fearing reduced animal survival. Note that NCFs presented in Figure 2e are statistically
Table with all BCF, NCF, Klip, and Kf values; modeling approach demonstrating the third phase effect on the logBCF - logKow relationship; logNCF - logKow plot for PCBs.
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Acknowledgments We are indebted to Beate Escher and Nadine Bramaz (EAWAG, Switzerland) for providing the liposome solution. The work described in this paper was supported by the European Commission Framework 6 Integrated Project ALARM (Assessing LArge scale environmental Risks for biodiversity with tested Methods; GOCE-CT-2003-506675).
Supporting Information Available
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Received for review April 27, 2007. Revised manuscript received August 3, 2007. Accepted August 10, 2007. ES0709977
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