Combining passive sampling and toxicity testing for evaluation of mixtures of polar organic chemicals in sewage treatment plant effluent

Abstract

Effluent from sewage treatment plants has been associated with a range of pollutant effects. Depending on the influent composition and treatment processes the effluent may contain a myriad of different chemicals which makes monitoring very complex. In this study we aimed to monitor relatively polar organic pollutant mixtures using a combination of passive sampling techniques and a set of biochemistry based assays covering acute bacterial toxicity (Microtox™), phytotoxicity (Max-I-PAM assay) and genotoxicity (umuC assay). The study showed that all of the assays were able to detect effects in the samples and allowed a comparison of the two plants as well as a comparison between the two sampling periods. Distinct improvements in water quality were observed in one of the plants as result of an upgrade to a UV disinfection system, which improved from 24× sample enrichment required to induce a 50% response in the Microtox™ assay to 84×, from 30× sample enrichment to induce a 50% reduction in photosynthetic yield to 125×, and the genotoxicity observed in the first sampling period was eliminated. Thus we propose that biochemical assay techniques in combination with time integrated passive sampling can substantially contribute to the monitoring of polar organic toxicants in STP effluents .

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Introduction

Sewage treatment plant (STP) effluent is an important source for entry of anthropogenic organic chemical pollutants and their degradation products into the aquatic environment.^1,2 Mixtures of a diverse range of potentially bioactive chemicals can be released, including pharmaceuticals, legal and illicit drugs, dietary supplements, active ingredients in personal care products, diagnostic agents, sun-screens, and numerous others.^1,3 Treatment related processes within STPs such as phase partitioning (liquid to sludge) of more hydrophobic chemicals and production of hydrophilic (‘polar’) biodegradation products within STPs should lead to a proportional dominance of polar compounds in sewage effluent when compared with influent composition. Hence estimation of risk associated with polar chemicals in sewage effluent is important.

Monitoring STP effluent is difficult due to matrix complexity and because the concentration and type of chemicals in the effluent may vary substantially within a small time frame depending on the composition and volume of incoming sewage and treatment efficiency.⁴ As such, reliance of monitoring programs on spot sampling provides limited environmentally relevant information and data require careful interpretation where the number of samples collected is inadequate.

The available methodology for assessment of organic pollutant mixture toxicity is often either inadequate or too labour intensive for routine monitoring applications. Sample chemical analysis for comparison with toxicity data for individual chemicals is a reasonably accessible method but frequently fails to account for all sample toxicity .^2,5,6In vivo analysis of whole samples can be confounded by dominant ammonium toxicity unless extensive ‘toxicity identification evalution’ (TIE) procedures are followed.⁷ The method requires toxicants to be present at environmental levels which cause acute or short term biological effects as samples are tested in dilution series. Assessment of biomarkers such as vitellogenin or ethoxyresorufin-O-deethylase (EROD) activity induction is effective for determining toxicant induced changes in biological systems, but is labour intensive due to the required use of caged fish and rarely applied for routine monitoring.

Combining passive samplers for polar pollutants with high through-put, biochemical based assays has the potential to offer a simple and cost effective means to determine potential acute and chronic impacts of polar organic chemicals in sewage effluent . While assessment of toxicity of polar passive sampler extracts is relatively novel, the more established semi-permeable membrane device (SPMD)^8,9 passive samplers for non-polar pollutants have been shown to provide a useful pre-concentration technique for bioassays .^10–13 Highly concentrated extracts can be produced which are well suited for use in high throughput biological and biochemical testing, and the method has been identified as a potentially useful pre-concentration step for effect driven analysis (EDA).⁶

Polar organic passive samplers (POS) have been developed more recently by Kingston et al.¹⁴ (3M Empore Disks™ samplers, Chemcatcher) and Alvarez et al.¹⁵ (polar organic chemical integrated sampler, POCIS). Both utilise solid phase sorbents usually applied for laboratory sample extraction. 3M Empore Disks™ samplers allow some flexibility in chemical selectivity and uptake kinetics through application of different sampling matrices¹⁶ and addition of specific membranes.¹⁴ Stephens et al.¹⁶ calibrated styrene divenylbenzene (SDB-RPS) and C18 based Empore Disks™ exposed in a Teflon Chemcatcher type sampler housing¹⁴ without a membrane (‘naked’) and determined aqueous boundary layer mass transfer coefficients for several polar organic herbicides. Recently in our laboratory, Stephens et al. (publication in preparation) have undertaken field- and laboratory-based calibration studies utilizing different configurations of the sampler (deployment devices, use of membranes). One configuration of the sampler using SDB-RPS Empore Disks™ with a polysulfone ethoxylate membrane, in a Chemcatcher type sampler housing¹⁴ showed particular promise as a sampler for polar organic pollutants. The work demonstrated that this specific configuration allows relatively long term (>1 month) time integrated sampling for a wide range of polar organic pollutants.

To date, few studies are available combining passive samplers for polar organic chemicals with toxicity testing. Petty et al.¹⁷ and Vermeirssen et al.¹⁸ have tested extracts from POCIS using the YES assays for estrogenicity, and Escher et al.¹⁹ have recently published the first results combining ‘naked’ 3M Empore Disk™ passive samplers with the new Max-I-PAM phytotoxicity assay.

The objective of this study was to assess the biological effects of polar organic pollutants in STP effluent and to demonstrate the value of combining polar passive sampling with multiple biochemical assays. We present the first results of testing Empore Disk™ polar passive sampler extracts for both genotoxicity and bacterial acute toxicity and have also applied the same phytotoxicity assay reported by Escher et al.¹⁷ Results are discussed with reference to method efficacy, for inclusion in a comprehensive screening system for mixture toxicity of organic pollutants that employs passive samplers for both polar and non-polar pollutants and covers a wider range of toxicological endpoints.

Experimental

Sampling methodology

Sampling sites. Polar organic passive samplers (POS) were deployed at two sewage treatment plants (STPs) for two, one month periods (March and May 2005). The treatment plants were selected as they are in close proximity (approximately 10 km apart), but differ in their influent sources and treatment systems. STP 1 treats approximately 150 000 m³ of effluent per day and is a biological nutrient removal system producing secondary treated effluent (nutrient removal, no disinfection). Approximately 40% of waste received is trade waste (industrial, medical and commercial sources) and the balance consists of domestic waste. STP 2 produces secondary quality effluent through primary settlement and a trickling filter system with chlorine disinfection (no nutrient removal). Prior to the second month of sampling STP 2 was converted to a UV disinfection system. More than 90% of the 20 000 m³ treated per day by STP 2 is of domestic origin. Samplers at this site were deployed in the chlorine contact tank during month 1 and in the effluent stream prior to release during month 2.

Passive sampling. Polar organic samplers (POS) used to sequester pollutants consisted of 3M stryrene divinylbenzene–reverse phase sorbent (SDB-RPS) Empore Extraction Disks™ (Varian Inc) with polysulfone ethoxylate membranes (0.2 µm, Pall), housed in a patented Teflon sampling device (‘Chemcatcher’).¹⁴ POS were deployed in duplicate for 31 days (for both months 1 and 2) at STP 1, and 34 and 31 days in months 1 and 2, respectively, at STP 2. Published data are not yet available regarding the range of chemicals which would be sampled integratively using this specific sampler configuration. However, in a calibration study in our laboratories, the same sampling devices remained in the linear (integrative) range for 30 days when exposed under a moderate flow regime to 70 ng L^–1 of nine herbicides varying from log₁₀K_OW 1.2–3.1 (Stephens et al., publication in preparation).

Empore Disks™ were prepared by conditioning in 100% methanol (HPLC grade, Merck). Samplers were transported in Chemcatcher sampling devices containing MilliQ water. Procedural blanks were produced for each site, consisting of unexposed disks contained within Teflon sampling devices which were stored at 4 °C for the duration of the sampling period.

The Empore Disk™ extraction procedure followed a method similar to that of Shaw and Müller.²⁰ Disks were extracted twice in an ultrasonic bath for 5 min with 5 mL methanol. The two 5 mL volumes were combined and reduced in volume to approximately 1 mL before being filtered through a Millex 0.45 µM syringe driven filter unit. Extracts were transferred to glass vials and evaporated under a gentle stream of nitrogen to 250 µL, and then made up to exactly 1 mL with MilliQ water. Extracts from 2 samplers were combined prior to bioassay analysis.

Sampling rates. Sampling rates (litres per day) represent the volume of water from which chemicals are accumulated throughout the exposure period based on uptake kinetics determined in calibration studies.⁹ We have undertaken both field and laboratory experiments on the calibration of Empore Disk™ passive samplers both with¹⁶ and without (Stephens et al., publication in preparation) inclusion of a polysulfone ethoxylate membrane, and found that for SDB-RPS Empore Disks™ POS deployed with membranes in a Chemcatcher housing, in a turbulent system, an average sampling rate of 0.12 L day^–1 can be applied for exposure periods in excess of 1 month. This value represents the sampling rate of the herbicide diuron, which fell close to the mean rate of those chemicals analysed (variation up to a factor 2). Here the diuron sampling rate has been applied to approximate the uptake rate for the entire mixture.

Sampling rates may also vary depending on environmental conditions such as flow or turbulences affecting the POS sampler : water boundary layer, temperature and biofouling. These have not been accounted for in the present study as limited methods are presently available for in situ calibration of these devices. Variation due to flow is likely to be the greatest source of sampling rate variability and this has been minimized through the design of the sampling device which promotes an increased boundary layer.

Dose response assessments

Expression of results. Toxicity testing results are expressed using a diuron based relative enrichment factor (REF_D).^11,19 This value represents the enrichment of toxicants in the bioassay test solution when compared with source water concentrations. Extracts are enriched from a certain volume of source water and a proportion of the sample applied to bioassays , where it is diluted by test media. Hence the REF_D value is calculated from the volume of water enriched by the POS determined using the diuron sampling rate (V_W in mL), the final extract volume (V_E in mL), the volume of extract introduced to the bioassay (V_EA in mL) and the volume of bioassay media (V_A) (eqn (1)).EC_X (X% effect concentration, calculated by regression analysis) or LOEC (lowest observed effect concentration) values are calculated for each assay using the REF_D values to derive sample concentration (nomenclature: EC(REF_D)₅₀ or LOEC(REF_D)).

Bacterial cytotoxicity. Bacterial (Vibrio fischeri) acute toxicity tests were conducted using a Microtox 500 Analyzer™ according to test procedures slightly modified from the standard 15 min EC₅₀ (50% inhibition of luminescence ).²¹ Samples and procedural blanks were tested at eight concentrations in duplicates up to a maximum REF_D of approximately 370. Concentration–effect curves were fitted to a log-logistic function (eqn (2)) using Prism 4.0 Software (GraphPad, USA), which used the least squares method to compute the fit for experimental data. Minimum (0%) and maximum (100%) effect values and slope (s = 1) were fixed to allow comparison of EC(REF_D)₅₀ values between samples.

Phytotoxicity. A fluorescence based photosynthetic yield analysis technique was applied for sample phytotoxicity assessment using a prototype of a new chlorophyll fluorescence imaging system for saturation pulse quenching analysis of algae suspensions on multi-well plates: Maxi-Imaging-PAM chlorophyll fluorometer (Max-I-PAM) (first prototype manufactured by J. Kolbowski and U. Schreiber, Würzburg, Germany; series production by Heinz Walz GmbH, Germany). The methodology of the Max-I-PAM bioassay is similar to that of the previously introduced ToxY-PAM bioassay ²² that has been applied for assessment of phytotoxicity.^23–25 While with the ToxY-PAM only one control and one sample can be compared, the Max-I-PAM allows simultaneous assessment of photosynthetic activity in a large number of samples (using 96 well plates). PAM fluorometry in conjunction with the saturation pulse method provides highly accurate information on the quantum yield of energy conversion at photosystem II (PSII) reaction centres (for a recent review see ref. 26). Inhibition of photosynthesis is calculated by comparison of the quantum yield of a test well with that of a control well. The Max-I-PAM assay is particularly sensitive for the presence of PSII herbicides, which bind to the quinone binding site of PSII in photosynthesis and inhibit energy conversion.²²

The marine diatom Phaeodactylum tricornutum was used for phytotoxicity assessment. Algae were grown in a culture chamber at 23 °C with a 12 h natural dark/light cycle receiving 50 µE m^–2 s^–1. Log phase cultures were used for assays, by subculturing algae 3 days prior to analysis. For a detailed description of growth conditions and media see Bengtson Nash et al.²⁴ Testing was undertaken using a measuring light intensity of 10 and measuring frequency of 8 (3 µmol quanta m^–2 s^–1 photosynthetically active radiation (PAR), no actinic light). Five readings were averaged after 1 h incubation. A full description of the methodology is published elsewhere.²⁷ Samples and procedural blanks were tested at twelve concentrations (maximum REF_D 530) in duplicate. A diuron positive control was also tested at 16 concentrations (0.07–34 µg L^–1). Concentration–effect curves were fitted using eqn (2), minimum (0%) and maximum (100%) effect values and slope (s = 1) were fixed to allow comparison of EC(REF_D)₅₀ values between samples and positive control.

Diuron equivalent concentrations (DEC) represent the concentration of diuron (µg L^–1) which would be required to produce an effect equal to that observed in samples.^19,23 These were calculated from EC₅₀ values as per eqn (3).

Genotoxicity. The umuC assay was carried out according to the procedure described by ISO (2000). The bacteria Salmonella typhimurium TA1535/pSK1002, a gift from Prof. Reifferscheid, University of Mainz, Germany, was used. To enable detection of progenotoxins that require metabolic activation to become genotoxic, samples were tested both with and without inclusion of liver 9000 g supernatant fraction (S9), prepared form rats treated with Aroclor 1254 (Moltox Inc, USA). In brief, the bacteria were grown in Luria–Bertani medium containing 50 µg mL^–1ampicillin (Sigma, USA) at 37 °C for ca. 16 h with vigorous shaking. The overnight culture was diluted 20-fold in TGA medium (1% Bactotryptone, 0.5% NaCl, 0.2% glucose, 50 µg mL^–1ampicillin) and the culture was continued until the optical density at 600 nm reached about 0.25. 96 well plates were prepared with half of the treatments containing 3% S9 mix. Testing extracts were then added, followed by the bacterial culture. The mixture was incubated at 37 °C for 2 h with shaking. 300 µL of the mixture was transferred to a new plate with pre-warmed fresh TGA medium for a further incubation of 2 h with shaking. The bacterial density at 600 nm was measured and ß-galactosidase activity was assayed using 2-nitrophenyl-ß-D-galactopyranoside (OPNG, 4 mg mL^–1) as a substrate. The activity was determined by the method of Miller²⁸ with a slight modification by Oda et al.²⁹ The results are presented as means of triplicate determinations. Positive controls with 4-nitroquinoline-N-oxide (4-NQO) and 2-aminoanthracene (2-AA), negative control and DMSO and procedural control were also tested.

Results

Bacterial acute toxicity

All samples showed toxicity in the Microtox™ bioassay (EC(REF_D)₅₀ values are provided in Table 1), though results indicated only minimal acute toxicity due to organic chemical contaminants would occur at environmental levels (REF_D = 1) (Fig. 1). STP 2 produced the most toxic sample during the first month of sampling; following the STP 2 plant upgrade this toxicity was reduced below levels observed in samples collected from STP 1. Responses observed at STP 1 were highly consistent between sampling periods. Some toxicity was observed in procedural blanks, up to about 30% toxicity at the highest enrichment level.

Table 1 Results of bioassay analysis^a

	Microtox		Max-I-PAM		umuC
	EC(REFD)50	95% CI	EC(REFD)50	95% CI	LOEC(REFD) (–S9)	LOEC(REFD) (+S9)
a NR: no response in bioassay . b No dose response analysis was undertaken, 4-NQO tested at 50 ng mL^–1 and 2-AA at 200 ng mL^–1.
STP 1
Month 1	64	(60–69)	24	(20–29)	NR	NR
Month 2	68	(63–74)	35	(31–42)	119	119
STP 2
Month 1	26	(23–29)	30	(117–135)	33	NR
Month 2	84	(74–95)	125	(117–135)	NR	NR
Positive control	17 mg l^–1 (phenol)	(13–22) mg l^–1	6.7 µg L^–1 (diuron)	(6.1–7.4) µg L^–1	Positive^b (4-NQO)	Positive^b (2-AA)
Blank	1500	(920–2500)	NR	—	NR	—

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Fig. 1 Concentration–effect curves for Microtox™ analysis at (a) STP 1 and (b) STP 2 during months 1 and 2 sampling (M1 and M2, respectively). Duplicate results plotted. R₂ values for concentration–effect curves > 0.9870. EC(REF_D)₅₀ values are listed in Table 1.

Fig. 2 Concentration–effect curves for Max-I-PAM analysis, inhibition of PSII photosynthetic yield at (a) STP 1 and (b) STP 2 during months 1 and 2 sampling (M1 and M2, respectively). Duplicate results plotted. R₂ values for concentration–effect curves > 0.9577. EC(REF_D)₅₀ values are listed in Table 1.

Phytotoxicity

Phytotoxicity was relatively consistent between months 1 and 2 of sampling at STP 1 and month 1 of sampling at STP 2, where diuron equivalent concentrations (DEC) of 0.28, 0.19 and 0.22 µg L^–1, respectively, could be calculated from bioassay results. Following the upgrade of STP 2, a DEC of 0.05 µg L^–1 was recorded. No response was observed in procedural blanks tested.

Genotoxicity

Genotoxicity was observed in samples from both treatment plant in the absence of metabolic activation (S9 mix). At STP 1, 100× sample enrichment was required to produce a positive response. Where chlorination treatment was employed at STP 2, a genotoxic response was exhibited in samples enriched up to 30×, this effect was not present in samples collected following the plant upgrade. Table 1 displays LOEC(REF_D) values for samples tested without metabolic activation. Genotoxicity figures are provided in the ESI.†

Testing of samples in the presence of an S9 mix is expected to result in an increase in genotoxicity due to metabolic activation of progenotoxins. Our results indicated the opposite response and genotoxicity was reduced in all samples (Table 1). Similar results have been observed by others and it has been suggested that sorption of genotoxins to proteins in the S9 mix results in the non-expression of the mutagenicity of some direct and indirect mutagens .^23,24 The observed reduction in genotoxicity in the presence of S9 in samples tested in this study cannot therefore be considered to have implications for sample genotoxicity.

Discussion

The results from the bioassays suggest that different biological effects may be expected as a result of organic chemical pollution in the receiving zones of the two treatment plants. Fig. 3 suggests phytotoxicity is the most toxicologically relevant endpoint (of those tested) at STP 1. At STP 2, bacterial cytotoxicity and genotoxicity were potential endpoints of concern whilst chlorination disinfection was employed. Following conversion of STP 2 to UV treatment, all assays exhibited lowered toxicity .

Fig. 3 Comparison of sample enrichment required to produce a defined effect in each assay (Microtox: EC(REF_D)₅₀; Max-I-PAM: EC(REF_D)₅₀; umuC: LOEC(REF_D). Values available in Table 1.

The effluent undergoing chlorination disinfection exhibited higher levels of both bacterial cytotoxicity and genotoxicity when compared with both UV treated effluent from the same plant and effluent from STP 1 (no disinfection). As chlorine is not expected to accumulate in POS, elevated cytotoxicity and genotoxicity responses in chlorine treated effluent can be tentatively attributed to chlorination byproduct toxicity , as has been observed previously in chlorine treated effluent ,^30,31 however further data are required to support this hypothesis.

The results of the three bioanalytical endpoints employed in this study indicate that a substantial improvement in effluent quality was achieved through the upgrade of STP 2. Such data provides useful information to guide both ecological and, where wastewater is intended for agricultural or potable water reuse, human health risk assessment and can be used to guide management initiatives.

Method efficacy for aquatic pollutant risk assessment

The limitations of conventional monitoring approaches have been widely recognized. For example, Eggen et al.³² have listed among the current challenges in ecotoxicology: low concentrations of pollutants and long exposure times (chronic effects), multiple effects by single pollutants, and complex mixtures of pollutants. Daughton and Ternes¹ have also highlighted potential long term (multigenerational) impacts which may go undetected by the use of conventional risk assessment techniques. In the present study we have shown that combining passive sampling for polar organic pollutants with bioassays can provide a sensitive and relatively simple method for detecting effects of organic chemical pollutants across acute and chronic endpoints.

Passive sampling for polar pollutants is an attractive option in sewage effluent monitoring, allowing a time weighted average concentration of pollutants across the sampling period to be obtained and minimizing the need to handle effluent samples. The method enables extraction of larger volumes of water (up to several litres), required for detection of hormones and other pharmaceuticals which are potentially bioactive at trace levels (nanogram per litre). While use of passive samplers for non-polar pollutants is well established, polar sampling devices have been developed more recently. Further developmental and calibration work with these types of samplers is required to characterize uptake rates for a greater range of environmental pollutants and difficulties remain with regards to in situ calibration of the devices (correction for changes in uptake rates relating to flow, temperature, biofouling etc. As calibration data become available for a wider range of chemicals, sampling rates for relevant chemicals may be applied to different bioassays .

Consistency between results obtained in months 1 and 2 of sampling at STP 1 (Fig. 1 and 2), supports the advantages of implementing a time integrated sampling approach. Chemical concentrations in sewage effluent can be expected to vary substantially depending on the time of day and week, and the ability to collect time integrated samples improves the environmental relevance of results by more closely representing concentrations to which organisms are exposed in the effluent receiving zone.

To assess potential biological impacts of polar organic chemicals in sewage effluent , we have applied low volume, biochemical based assays. This approach precludes environmental realism as whole organisms and ecosystems can be expected to react differently than might be predicted from a ‘test tube’ based approach. Thus the method does not replace traditional ecotoxicological risk assessment but should instead be applied to complement available techniques and can be considered a bio-recognition system for detection of pollutants through toxic effects. In this capacity, the use of passive samplers for in situpreconcnetration shows promise for inclusion in effects driven analysis (EDA) studies. Use of passive samplers for both polar and non-polar organic pollutants samplers will enable in situ sample fractionation , and use of an increased battery of assays (including additional endpoints such as estrogenicity, cellular detoxification mechanisms, or neurotoxicological endpoints such as acetylcholine esteraseinhibition ) will provide a powerful method for assessing effects of site specific mixtures of organic pollutants.

Use of enriched extracts allows assessment of effects even where chemicals are present in source water at concentrations well below those that would cause environmental harm. Such sensitivity will allow preventative management initiatives and may prove vital for assessment of effects of trace contaminants in re-use water.

Conclusions

Combining time integrated passive sampling with a battery of high through-put bioassays allows us to undertake meaningful assessment of organic chemical pollutants in complex exposure environments. The method shows promise as an effective means to characterize sample toxicity in STP effluent and in this study important endpoints of concern have been identified. Further field and laboratory testing is required to determine the extent of method sensitivity in more pristine environments and at sites subjected to other sources of contamination.

Acknowledgements

The contributions of Dr Ulrich Schreiber in development of the Max-I-PAM instrument and methodology as well as comments on the manuscript are gratefully acknowledged. The laboratory assistance of Pamela Quayle and Kristie Lee Chue is appreciated. Thanks also to Dr Frederick Leusch for comments on the manuscript. This project is funded by an Australian Research Council Linkage Grant, with industry partners including: Queensland EPA, New South Wales DEH, Victoria EPA, SA Water, Brisbane Water, SEQ Water, Queensland Health, Australian Plague Locust Commission, Derwent Estuary Program and Environment Agency UK. Queensland Health provides funding for the National Research Centre for Environmental Toxicology.

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Footnote

† Electronic supplementary information (ESI) available: Figures displaying genotoxicity response and cytotoxicity in the umuC assay. See DOI: 10.1039/b612430e

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