Effects of biodegradation and sorption by humic acid on the estrogenicity of 17β-estradiol

Chemosphere 85 (2011) 1383–1389

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Effects of biodegradation and sorption by humic acid on the estrogenicity of 17b-estradiol Ji Ho Lee a,b, John L. Zhou c, Sang Don Kim a,⇑ a

School of Environmental Science and Engineering, Gwangju Institute of Science and Technology, 261 Cheomdan-Gwagiro, Buk-gu, Gwangju 500-7112, Republic of Korea Hazardous Chemistry Division, National Academy of Agricultural Science, Seodundong, Gwonseongu, Suwon 441-707, Republic of Korea c Department of Applied Sciences, London South Bank University, 103 Borough Road, London SE1 0AA, United Kingdom b

a r t i c l e

i n f o

Article history: Received 16 March 2011 Received in revised form 29 July 2011 Accepted 1 August 2011 Available online 27 August 2011 Keywords: 17b-Estradiol Humic acid Activated sludge Sorption Biodegradation Estrogenic activity

a b s t r a c t The removal of 17b-estradiol (E2) by biodegradation and sorption onto humic acid (HA) was examined at various HA concentrations. Subsequently, estrogenicity associated with E2 removal was estimated using E-screen bioassay. Results showed that E2 biodegradation and its subsequent transformation to estrone (E1) were significantly reduced with increasing HA concentration. In addition, the presence of nutrients enhanced the biodegradation of E2. Overall, E2 biodegradation was the dominating contributor to its removal, which demonstrated a significantly negative correlation with E2 sorption at various HA concentrations. The sorption of E2 by HA was significantly enhanced with increasing HA concentration. Estrogenicity associated with residual E2 showed that there existed a significant difference among various HA concentrations, with the lowest value in the absence of HA. The findings suggest that the presence of HA and nutrients in natural waters should be considered in assessing estrogenicity of environmental samples due to complex sorption and biodegradation processes. Ó 2011 Elsevier Ltd. All rights reserved.

1. Introduction Endocrine disrupting chemicals (EDCs) such as estrone (E1) and 17b-estradiol (E2), released predominantly from sewage effluents and animal manure may be responsible for disrupting the reproduction of fish as well as other wildlife species (Rodger-Gray et al., 2001; Snyder et al., 2001; Anderson et al., 2003). Such chemicals are excreted from human and animal urine and feces as free estrogens or as glucuronide and/or sulfate conjugates. Conjugated estrogens, being biologically inactive, are converted into deconjugated forms during sewage treatment (e.g., activated sludge process), thus may lead to adverse reproductive effects on humans and ecosystems (Johnson and Sumpter, 2001; Anderson et al., 2003). In particular, E2, as a natural hormone, is one of the most potent estrogens among various EDCs (Khanal et al., 2006). Although the average removal efficiency of E2 can reach over 60% during activated sludge treatment (de Mes et al., 2005), E2 concentration may still be present at a few ng L1 levels in treated effluent and natural waters. This suggests that activated sludge treatment process cannot fully remove compounds such as E2. E2 at a concentration of 1 ng L1 has been shown to increase the vitellogenin (a female protein) level in the Japanese male medaka

⇑ Corresponding author. Tel.: +82 62 970 2445; fax: +82 62 970 2434. E-mail address: [email protected] (S.D. Kim). 0045-6535/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.chemosphere.2011.08.003

(Hansen et al., 1998). EA (2002) has been proposed that a predicted no effect concentration (PNEC) of E2 derived from reproductive effects for Japanese medaka during 28 d exposure is 1 ng L1. E2 was initially sorbed onto activated sludge as well as particles, and then degraded by bacteria attached to them (Joss et al., 2004). During activated sludge treatment, E2 sorption falls in the range of 23–55% (Titia de Mes et al., 2005). In addition, several studies have reported E2 being eliminated dominantly between the range 62% and 99% in activated sludge systems via bacterial activity (Johnson and Sumpter, 2001; Suzuki and Maruyama, 2006). E2 has been shown to be oxidized rapidly into estrone (E1), further degrading into estriol (E3) (Ternes et al., 1999; Layton et al., 2000; Lee and Liu, 2002). In activated sludge treatment, rapid transformation of E2 into E1 has been observed at a rate of over 90%; E1 being removed at a rate of over 50% at 24 h (Ternes et al., 1999). E2 biodegradation is affected by various environmental factors such as temperature, salinity, pH, oxygen, and inorganic nutrient availability (Shimp and Pfaender, 1985). An additional key factor is the presence of dissolved organic mattter (DOM) in natural waters. DOM possessing aromatic/aliphatic structures as well as various functional groups (e.g., hydroxyl, carboxylic acid, ketone, and amino groups) is mainly composed of humic substances such as humic acid (HA) and fulvic acids (FA). The humic substances account for approximately 50–80% of DOM and 30–40% aromatic carbon in natural waters (Aiken, 1985). The HA, having vital properties to aquatic environments (e.g., the mobilization and availability of nutrients

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and the oxidative and reductive transformation of trace metals and organic chemicals), is widely used in studying the effect of DOM on the sorption of organic pollutants (Cheng et al., 2007). Recent research suggests that DOM dominantly contributes to determining the sorption of EDCs (Zhou et al., 2007). About 15–30% of E2 was bound with river-derived DOM (Yamamoto and Liljestrand, 2003; Zhou et al., 2007). The present study focuses on E2 interaction with HA which plays an important role in determining the fate of E2. Such interactions result in the formation of sorbed E2 and hence influence the bioavailability and biodegradability of E2 (Weber et al., 2005). Previous studies on E2 biodegradation has been performed extensively within activated sludge treatment plant (Ternes et al., 1999; Shi et al., 2004; Li et al., 2008). Little research into the effects of E2 biodegradation and sorption by HA on its estrogenicity has been documented. As a consequence, the objective of the study was to investigate E2 biodegradation behavior at various HA concentrations. The contributions of sorption and biodegradation on the fate of E2 and its estrogenicity were also evaluated by means of E-screen bioassay. 2. Materials and methods 2.1. Materials All chemicals used in the study were purchased from Sigma Aldrich (St. Louis, MO, USA) with a high purity (>98%). Aldrich HA, having a molecular weight of 3000–4000 Da was used to evaluate reliable E2 sorption via clear separation using 1-kDa ultrafiltration membrane (Chin et al., 1994). E2 has the following physical and chemical properties: molecular formula (C18H24O2), molecular weight (272.4 g mol1), water solubility (3.1–12.96 mg L1) (Shareef et al., 2006), log Kow (3.1–4.01), and pKa (10.2–10.7). The stock solution was prepared with HPLC grade methanol (Fisher Scientific, Pittsburgh, PA, USA) with a final concentration of 400 mg L1. To inhibit bacterial activity, sodium azide was added. For E-screen bioassay, Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with and without phenol red, fetal bovine serum (FBS), trypsin, and 0.4% sulforhodamine B (SRB) dying solution were purchased from Gibco-BRL life technologies (Invitrogen™, Paisley, UK). DMEM, supplemented with 10% FBS, and 10% FBS treated with 5% charcoal and 0.5% dextran (Sigma, St. Louis, Mo, USA) were used for normal cell culture and the bioassay. Stirred ultrafiltration cells (Model 8200, Amicon and Millipore, Bellerica, MA, USA) and regenerated cellulose membranes (1- kDa, membrane area 28.7 cm2, Ultracel, Millipore, Bellerica, MA, USA) were used for E2 sorption experiments. 2.2. Activated sludge To use as bacterial inoculum, 4 L of activated sludge was obtained from the Gwangju waste water treatment plant (WWTP) discharged through the Yeongsan River, Korea, in December 2008. The WWTP, discharging 600,000 tons d1, treated domestic sewage and landfill leachate via nutrient removal processes combined with a conventional activated sludge system. The plant operated with a hydraulic retention time (HRT) of 10 h and a sludge retention time (SRT) of 10–15 d. The activated sludge was collected in glass bottles rinsed with methanol and ultrapure Milli-QÒ water (Millipore Co., Billerica, MA, USA). The sludge was delivered to the laboratory at 4 °C to maintain original state. To obtain stable and reproducible bacterial activity, the sludge was then preconditioned with an air diffuser for 7 d by following the 301 protocol of the OECD chemical testing guidelines (OECD, 1992). During continuous aeration, 100 mL of supernatant was withdrawn after 1 h of

settlement, a mineral medium of an identical volume being subsequently added to assist bacterial communities to adapt on a daily basis. In preconditioned sludge, pH, dissolved oxygen (DO), dissolved organic carbon (DOC), and mixed liquor suspended solid (MLSS) concentration were measured with five replicates. DO and pH were measured by using a pH meter (250A, Thermo Orion, Beverly, MA, USA) and DO meter (WTW multi 350i, Weilhein, Germany). For DOC and MLSS measurements, the preconditioned activated sludge was filtered with 0.45 lm mixed cellulose ester filter (WhatmanÒ, Brentford, UK). DOC concentration was immediately measured by the combustion catalytic oxidation method using TOC analyzer (TOC-vCSH, Shimadzu Co., Kyoto, Japan). The pH, temperature, conductivity, DO, DOC, and MLSS concentrations of preconditioning activated sludge were shown in Table 1. E1 and E2 concentrations were not detected in preconditioned sludge. 2.3. E2 degradation experiments Microcosm experiments were conducted to assess HA effects on E2 degradation in a batch mode. One group of samples contained the fresh mineral medium consisted of 1.0 g L1 of (NH4)2SO4, 0.8 g L1 of Na2HPO4, 0.2 g L1 of MgSO47H2O, 0.2 g L1 of KH2PO4, 0.005 g L1 of FeCl3, 0.001 g L1 of (NH4)6Mo7O24, 0.1 g L1 of CaCl2, 0.937 g L1 of NaHCO3 and 0.005 g L1 of KCl dissolved in autoclaved ultrapure Milli-QÒ water and adjusted to a pH of 7.0–7.5. The other group of samples includes only autoclaved ultrapure Milli-QÒ water under same condition. Then E2 was added to each flask at a nominal concentration of 1 mg L1. After methanol evaporation by means of purging the air, a mineral medium of 100 mL was added to half of the samples. The HA (0.1 g) was dissolved in 10 mL of 0.1 M NaOH solution and then diluted to 100 mL with ultrapure Milli-QÒ water. After adjusting to pH 7.0, stock solution of HA was filtered through a 0.2 lm mixed cellulose ester filter (WhatmanÒ, Brentford, UK) and then diluted to make final HA concentrations of 10, 30, and 50 mg C L1, respectively. HA solutions of 100 mL were likewise transferred to each 500 mL flask to ensure concentration ranging between 0 and 50 mg C L1. In addition, a 0.5 mL aliquot of preconditioned activated sludge was inoculated. All samples were incubated in duplicate at 25 °C on a rotary shaker at 180 rpm for 3 d under aerobic conditions. An aliquot of 40 mL was taken at intervals. Abiotic control tests were performed with 40 mM sodium azide (NaN3) inoculations to prevent bacterial activity while no inoculation of E2 was necessary for biotic control tests. In abiotic control, the average recovery (%) of E2 ranged from 102.4 ± 16.6 to 116.5 ± 6.2 for all HA concentrations during incubation time. The relative standard deviation (RSD) was less than 16%, with majority of samples with RSD < 10%. There was little degradation of E2 as well as transformation to E1 in abiotic control. The pH and DO values of all samples were 6.4–8.4 and 6.5–8.8 mg L1, respectively. 2.4. Sorption experimentation Ultrafiltration tests were conducted with a static mode to evaluate the extent of E2 sorption by HA (10–50 mgC L1), which were slightly higher than observed in typical natural water. The samples were filtered through 0.45 lm mixed cellulose ester filter (WhatmanÒ, Brentford, UK) and then adjusted to pH 7.0 with 0.1 M HCl and NaOH. In addition, the samples were treated with sodium azide and wrapped an aluminum foil to avoid E2 losses (i.e. sorption to wall, biodegradation, and photodegradation). E2 (1 mg L1) used in E2 biodegradation tests was equally added to each HA solution of 100 mL to assess the fraction of E2 sorbed by HA during E2 biodegradation. The highest E2 sorbed fraction at various HA concentrations was observed at 24 h. All samples were thoroughly shaken at 180 rpm for 24 h at 25 °C to maintain a state

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J.H. Lee et al. / Chemosphere 85 (2011) 1383–1389 Table 1 The characteristic of preconditioned activated sludge. pH

Temp. (°C)

6.68–6.72 a b c d

22–23

Conductivity (ls cm1) 1596 ± 12

DOa (mg L1) 2.1–2.9

DOCb (mg L1) 7.8

MLSSc (mg L1) 4840 ± 15

E2 (ng L1) d


E1 (ng L1)
Dissolved oxygen. Dissolved organic carbon. Mixed liquor suspended solid. LOD = limit of detection determined at 1.0 ng L1.

of equilibrium. Those were continuously operated to separate the sorbed and free E2 by DOM using a stirred ultrafiltration cell which held the 1 kDa regenerated cellulose ultrafiltration membrane (Millipore, Bellerica, MA, USA). Such a membrane has a contact angle of 36 ± 2.6° indicating a hydrophilic surface. Prior to running the system, membranes were initially soaked with ultrapure Milli-QÒ water and washed three times at intervals of 12 h to remove glycerin preservatives. The membrane was subsequently flushed with ultrapure Milli-QÒ water during all experiments and preconditioned until a stable flow rate was observed. The 100 mL samples were poured into ultrafiltration cell system and then passed through the membrane under a N2 gas pressure of 3.5 kfg to ensure a low flow rate of 0.8 mL min1 in permeate. A total of 90 mL permeate was separately collected in borosilicate bottles covered with aluminum foil, after discarding the 10 mL permeate to avoid permeate dilution by interaction with remaining MilliQÒ water. E2 concentration in permeate taken from all samples was determined by LC–MS analysis without pretreatment. E2 concentration in retentate was obtained by subtracting its concentration in permeate from initial E2 concentration. Sorbed E2 fractions to various HA concentrations were calculated using the following equations (Zhou et al., 2007):

kE2HA ¼

C E2HA ; C E2w

ð1Þ

where CE2HA (mg L1) and CE2w (mg L1) are E2 concentrations in HA solution (retentate) and in water (permeate), respectively. 1

koc ðL kgoc Þ ¼

kE2HA ð½DOC : DOC conc:; kg C L1 Þ; ½DOC

dichloromethane/hexane (1:1, v/v). The dry residual was reconstituted into 1 mL with methanol, and then divided into 300 lL aliquot for chemical analysis as well as for E-screen bioassay. Chemical analysis was performed by a liquid chromatography mass spectrophotometer (LC-MS-2010 EV, Kyoto, Japan) equipped with an ACE 5 C18-HL column (i.e. 5 lm particle size, 2.1 mm internal diameter, and 1.5 m length, ACEÒ, Kyoto, Japan) combined with a guard cartridge (2.1 mm internal diameter, ACEÒ, Kyoto, Japan). It was analyzed with selective ion monitoring (SIM) mode using electrospray interface (ESI) being operated in the negative mode. The mobile phase is composed of HPLC grade methanol and water, with a flow rate of 0.3 mL min1. The LC–MS binary mobile phase representing E2 and E1 analysis was programmed as follows: methanol was increased from 30% to 100% for 18 min and then decreased to 20% for 5 min. Nebulizing N2 gas was continuously introduced with a flow rate of 1.5 mL min1. The temperature of the curved desolvation line (CDL) and heat block stood at 250 °C. A detector voltage of 1.5 kv, a probe voltage of 4.5 v, and a CDL voltage of 25 V were set up and then analyzed. For quality control, the methanol as blank and E2 standard solution of 100 lg L1 were checked every ten sample. The other interfering peaks or matrix suppression was not significantly observed in all samples. The E1 and E2 peaks were identified based on an m/z value of 269/287 and 271/227, respectively. The limit of detection (LOD) calculated as three times signal to noise level was determined at 1.0 ng L1. The recovery for sample spiked with 500 ng L1of E2 and E1 in various HA concentrations was ranged from 95.1 ± 4.8 to 108.6 ± 4.5.

ð2Þ 2.6. E-screen bioassay

koc  ½DOC Sorbed fraction of E2ðfE2HA Þ ¼ 1 þ koc ½DOC

ð3Þ

In order to assess the mass balance of E2, the solutions spiked with 1 mg L1 of E2 in distilled water were passed through the same system under identical condition. When E2 concentration before and after running stirred cell was compared, the loss of E2 was negligible with a good recovery of 99.2 ± 1.35%. 2.5. Sample preparation and chemical analysis Samples obtained from biodegradation experiments were filtered through 0.45 lm mixed cellulose ester filter (WhatmanÒ, Brentford, UK), and then processed by the solid phase extraction (SPE), modifying procedure described in Hashimoto and Murakami (2009). Prior to sample pretreatment, Oasis HLB cartridges (Waters, Milford, MA, USA) were repeatedly preconditioned with 15 mL methanol, followed by 15 mL ultrapure water at a flow rate of 1–2 mL min1. Then, all samples were loaded onto conditioned cartridge at a same flow rate. Cartridge was completely dried under vacuum, and then analytes attached onto cartridge were eluted with 15 mL methanol at a flow rate of 1 mL min1. The methanol was completely blown down under a gentle flow of nitrogen. The concentrated samples were reconstituted to 1 mL with dichloromethane/hexane (1:1, v/v), cleaned up with a Sep-Pak Florisil cartridge (Waters, Milford, MA, USA), and then eluted with 15 mL

E-screen bioassay using a MCF-7 BUS cell was conducted to confirm E2 degradation behavior through a modified procedure established by Soto et al. (1995). In order to assess the effect of only E2 sorption, the estrogenicity of solutions containing HA only as a background was checked. The net estrogenicity obtained by subtracting the estrogenicity for HA only as a background from those samples with both HA solutions and E2 was estimated. A 300 lL aliquot extracted from SPE for an E-screen bioassay was diluted by 4.3  105 times with DMEM culture media in the course of preparation of final 8.6  1012 M E2 concentration. The prepared E2 concentration corresponds to EC50 value based on a concentration–response curve in MCF-7 BUS cell. The MCF-7 BUS cell was inoculated with a cell density of 104 well1. A sample of 400 lL diluted with a DMEM culture media was added to 24 well plates and then cultured for 6 d in a CO2 incubator under a temperature of 37.5C with 5% CO2. After suction of samples, the MCF-7 BUS cell was fixed by adding 10% trichloroacetic acid (Merck, Frankfurter, Darmstadt, Germany) at 4C for 40 min, washed with Milli-Q water, and completely dried at room temperature. The cell was dyed with 0.4% sulforhodamine B (SRB) solution and washed with 1% acetic acid to eliminate excess SRB solution. After well plates were dried completely, 400 lL of a 10 mM tris buffer (AMRESCO, Solon, Ohio, USA) at pH 10.5 was added to each well plate. Cell proliferation was quantified by measuring the absorbance value at 560 nm with an ELISA reader (BIO-TEK Instruments Inc., Winooski, Vermont,

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USA). The estrogenic activity was expressed as an estradiol equivalent concentration (EEQ) value. 2.7. Enumeration of total bacterial cell The total number of bacteria cells was determined using a DAPI (40 ,6-diamidino-2-phenylindole) staining method to examine bacterial growth during utilization of organic substrates. The bacteria cell stained by DAPI forms a fluorescent complex by combining with a specific DNA. Thus, the live bacteria community is mainly observed with 0.1–1% bacterial aggregates (Stevik et al., 1998). The bacterial cell number in inoculums was quantified by 3.7 ± 1.9  104 cells mL1 with different conditions. Aliquots (5 mL) were taken at 72 h and filtered via 5 lm mixed cellulose ester filters (Advantec, Toyo Roshi Kaisha, Japan), stored at 4 °C refrigerator and taken. A light-sensitive DAPI with 100 lg mL1 of stock concentration was used as a staining solution. Under dark conditions, a DAPI staining solution of 10 lL was added to filtered samples and stained for 5 min. In addition, the solutions were filtered with 0.2 lm black polycarbonate Isopore™ membrane filters (Millipore, Ireland) without vacuum, rinsed with ultrapure MilliQÒ water, and then dried. Each filter was mounted on a glass microscopy slide sealed with a coverslip on top. The stained bacterial cells within mounted filters were observed at 1000 magnification using a confocal laser scanning microscope (LSM5, Zeiss, Germany). The bacterial cell was calculated using an I-solution program after randomly taking microscopic views of 10 fields. By considering the sample volume and size of the microscopic views, a cell average was thus calculated.

2000) or an inhibition of bacterial activity (Marscher and Kalbitz, 2003). A specific UV absorbance of HA at 254 nm (SUVA254) of 0.07 L mg1 cm1 indicated a higher hydrophobicity than that in natural waters (Steinberg et al., 2008). 3.2. E2 biodegradation behavior The E2 biodegradation behavior was investigated at various HA concentrations during incubation time. The degradation rate of E2 at each HA concentration was determined by plotting the E2 concentration against incubation time of 0–72 h. (Fig. 1a). When HA was absent, E2 concentration rapidly decreased by up to 60% at 6 h and then disappeared by over 90% at 24 h. In the presence of HA, the extent of E2 degradation was approximately 20–30% of initial E2 concentration at 6 h, which was about two or three times slower than that in the absence of HA. The E2 concentrations for tested HA concentrations were reduced by 55–65% at 24 h and then not detected at 48 h. The decrease in E2 biodegradability became less obvious with further increase in HA concentration. E2 biodegradation was significantly affected by HA concentration and incubation time using two-way ANOVA tests (⁄⁄⁄p < 0.001). The regression effect among dependent variables in the MLR model was not observed in terms of VIF of 1.0. In order to compare the E2 degradability among HA concentrations, biodegradation rate constant of E2 (kbioE2) was calculated by the first order degradation kinetic equation.

(a)

2.8. Statistical analysis

3.1. Characterization of HA HA degradability was assessed to confirm the recalcitrant property of HA. The DOC concentrations with incubation time were measured at various HA concentrations. The degradation rate constant of DOC (kDOC) was calculated by assuming the first order degradation kinetics as given in following equation:

ð4Þ

where C0 is the initial DOC concentration (lg L1), C is the DOC concentration (lg L1) at time t (h),and kDOC is the first order degradation rate constant of DOC (h1). The kDOC values were decreased from 0.003 ± 0.001 h1 to 0.001 ± 0.001 h1, with increasing HA concentration of 10– 50 mgC L1. However, there was no significant difference (p > 0.05). Those are due to either its hydrophobicity (Li et al.,

E2 conc. (C/C0)

0.8 0.6 0.4 0.2 0.0 0

10

20

30

40

50

60

70

Time (hr)

(b) E1 conc. (C/C0) relative to E2 conc.

3. Results and discussion

C ¼ kDOC  t C0

HA 0 mg C/L 10 mg C/L 30 mg C/L 50 mg C/L Abiotic control

1.0

One and two way analysis of variance (ANOVA) tests were used to evaluate the significant difference in the E2 removal (e.g., sorption and biodegradation), transformation to E1, and estrogenicity among HA concentrations and nutrients. Prior to ANOVA tests, multiple linear regression (MLR) analysis was performed using E2 biodegradation and transformation to E1 as dependent variables, as well as incubation time and various HA concentrations as explanatory variables. As a result, if the largest variance inflation factor (VIF) is greater than 10 or average VIF value is greater than 1, the existence of regression effect is confirmed (Bowerman and O’Connell, 1990). Statistical analysis used was performed using SPSS Inc. ver. 18.0 (Chicago, Illinois, USA), with a probability value of 0.05 or less being considered statistically significant.

 ln

1.2

1.0

HA 0 mg C/L 10 mg C/L 30 mg C/L 50 mg C/L

0.8

0.6

0.4

0.2

0.0 0

10

20

30

40

50

60

70

Time (hr) Fig. 1. The E2 biodegradation behavior (a) and subsequent transformation to E1 (b) at various HA concentrations during incubation time. Statistically significant difference was observed in E2 biodegradation as well as transformation to E1 between various HA concentrations using two-way ANOVA tests (⁄⁄⁄p < 0.001).

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J.H. Lee et al. / Chemosphere 85 (2011) 1383–1389

ln

C ¼ kbioE2  t C0

the HA effect on E2 biodegradation. The E2 transformation to E1 was significantly influenced by HA concentrations and incubation time using two-way ANOVA tests (⁄⁄⁄p < 0.001). There was little regression effect between two dependent variables based on VIF of 1.0 obtained from MLR analysis. In the absence of HA, E2 transformation to E1 was the greatest, reaching up to 50% at 6 h, which was 1.2–3.0 times higher than that in the presence of HA. Initially the extent of E2 transformation to E1 at all HA concentrations showed gradually increased pattern until 24 h and reached 20– 45% relatively to initial E2 concentration. The degree of transformation to E1 declined as HA concentration was increased. Such transformation reached the highest levels at around 24 h, before declining, finally reaching below 5% at 48 h. Maximum transformation to E1 (%) was calculated by use of the equation ðC E1 max =C E2init Þ  100; indicating that maximum E1 concentration ðC E1 max Þ was divided into initial E2 concentrations ðC E2init Þ (Li et al., 2008). In the absence of HA, the highest maximum E1 transformation (%) at 24 h was found to be 62%. However, it was sequentially reduced from 42% to 25% with the addition of HA. The maximum transformation to E1 (%) was significantly different at various HA concentration using one-way ANOVA tests (⁄⁄p < 0.01). Particularly, post hoc test for one-way variance analysis revealed significant decrease for maximum transformation to E1 at 50 mg C L1 of HA compared to 0 mg C L1 of HA (⁄⁄p < 0.01).

ð5Þ

where C0 is the initial E2 concentration (lg L1), C is the E2 concentration (lg L1) in the water phase at time t (h), and kbioE2 (first order degradation rate constant of E2; h1) is obtained from plotting the (ln C/C0) versus t. The kbioE2 values and bacterial cell numbers were examined at various HA concentrations in the presence and absence of nutrients (Table 2). Those values were strongly influenced by HA concentrations as well as nutrients using two-way ANOVA tests (⁄p < 0.05). The kbioE2 value was the highest at 0.13 ± 0.01 h1 in the absence of HA, and decreased to 0.04 ± 0.01 h1 as HA levels were increased. Significant difference in kbioE2 value was observed among HA concentrations in the presence of nutrients (F = 14.9; p = 0.01). When the kbioE2 values between each pair of HA concentration groups were compared, the absence of HA revealed a more significant increase than the presence of HA concentrations using oneway ANOVA test (⁄p < 0.05). Several causes which may inhibit E2 degradability at increasing HA levels have been reported. E2 sorbed by HA appeared to be not available to bacterial communities which are responsible for E2 biodegradation (Shimp and Pfaender, 1985; Weber et al., 2005; Albers et al., 2008). However, some studies have reported that E2 biodegradation has not been greatly influenced by E2 sorption to HA due to rapid desorption of sorbed E2 in the liquid phase (Das et al., 2004; Andersen et al., 2005; Stumpe and Marschner, 2009). Another reason may be that HA interferes with a bacterial activity through irreversible enzyme binding (Shimp and Pfaender, 1985). Such mechanism limits the E2 biodegradation and its metabolite formation. Although it was not fully identified, the addition of HA translated into negative influences of bacterial cell number, which showed a significant reduction at increasing HA level in the presence of nutrients (F = 13.8; p = 0.001). The bacterial cell number in the presence of nutrients decreased from 23.4 ± 8.5  105 to 9.7 ± 1.9  105 cells mL1. Such evidence possibly supports the premise that increased HA levels cause detrimental effects on the bacterial activity responsible for E2 degradation. Besides, the bacterial cell number in the presence of nutrients was 5–10 times higher than that in the absence of nutrients under same HA concentration (Table 2).

3.4. Removal of E2 and estrogenicity The contribution of E2 sorption and biodegradation to its removal at various HA concentrations was examined at 24 h in the absence of nutrients (Fig. 2). Increasing HA concentrations led to the higher E2 sorption, ranging from 6.0% ± 0.01 at 10 mg C L1, 12.3% ± 2.1 at 30 mg C L1, to 21.9% ± 0.7 at 50 mg C L1 HA, respectively. Besides, biodegradation was shown to be the dominant mechanism in E2 removal process. In the absence of HA, the highest E2 removal (89%) was found to be due entirely to biodegradation. In accordance with increased HA concentrations, the fraction of E2 sorption was increased from 6% to 22%, meanwhile the E2 biodegradation ability was reduced from 67% to 56%. These results demonstrated that suppressive E2 biodegradation was possibly attributable to a higher E2 sorption by HA. The negative relationship between E2 biodegradation rate and the E2 sorption capability was apparent. The aquatic DOC concentration of 5 mg L1 reduces the E2 biodegradation up to 20–30% (Andersen et al., 2005). The sorption of E2 by HA was less significant in its removal than biodegradation. However, E2 removal fractions were slightly increased from 72% to 77%, when E2 sorption was increased. There were significant differences in the sorption, biodegradation, and removal fraction of E2 at various HA concentrations using one-way ANOVA tests (⁄⁄p < 0.01). In particular, a significant increase for E2 removal and biodegradation, as well as a significant decrease for E2 sorption, were observed at 0 mg C L1 HA compared to other HA concentrations (⁄p < 0.05).

3.3. E2 transformation into E1 From E2 biodegradation, subsequent transformation into E1 was investigated at various HA concentrations (Fig. 1b). Despite the lack of information about bacterial identification responsible for E2 biodegradation in the present study, it has been recognized that various microbes (e.g., Rhodococcus zopfii, Rhodococcus equi, Pseudomonas fluorescens, Bacillus thuringiensis strain) in wastewater activated sludge play a major role in E2 transformation and the extent of E2 biodegradation is affected by bacterial strain and its activity (Khanal et al., 2006). By considering these effects, mixed bacteria present in activated sludge system were used to assess

Table 2 E2 biodegradation rate constant (kbioE2) and bacterial cell number at various HA concentrations in the presence and absence of nutrients. Parameters

With nutrients

Without nutrients 1

HA concentration (mg C L

kbioE2 (h1) (r2) Bacterial cell no. (105 cells mL1)

HA concentration (mg C L1)

)

0

10

30

50

F

pvalue

0.13 ± 0.01 (0.97) 23.4 ± 8.5

0.08 ± 0.01 (0.90) 12.8 ± 1.3

0.06 ± 0.01 (0.88) 10.4 ± 1.2

0.04 ± 0.01 (0.86) 9.7 ± 1.9

14.9 0.012⁄ 13.8 0.001⁄⁄

0

10

30

50

F

0.05 ± 0.01 (0.92) 2.9 ± 0.6

0.04 ± 0.01 (0.96) 2.0 ± 0.2

0.03 ± 0.01 (0.95) 1.2 ± 0.02

0.02 ± 0.01 (0.92) 1.2 ± 0.02

1.7 0.31

The kbioE2 and bacterial cell No. showed significant difference among HA concentrations with nutrients using one-way ANOVA test (⁄p < 0.05, Each value was also expressed as average ± SE of five replicates.

⁄⁄

p < 0.01).

pvalue

1.4 0.14

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1.0 HA_biodegradation HA_sorption

E2 removal fraction

0.8

0.6

0.4

0.2

0.0

0

10

50

30

DOC concentration (mg C/L) Fig. 2. The contribution of sorption and biodegradation at various HA concentrations to E2 removal at 24 h in the presence of nutrients. The sorption, biodegradation, and removal fraction of E2 were significantly different between different HA concentrations using one-way ANOVA tests (⁄⁄p < 0.01).

2.5

Estrogenicity (ng-EEQ/L)

In the presence of nutrients In the absence of nutrients

2.0

1.5

1.0

0.5

0.0

Positive control

0

30

10

50

DOC concentration (mgC/L) Fig. 3. The estrogenicity of E2 solutions at the end of 24-h interactions with different HA concentrations in the presence and absence of nutrients. The estrogenicity of HA only as a control ranged from 0.25 ± 0.01 to 0.30 ± 0.23 ngEEQ L1. Each value was also expressed as average ± SE of duplicate determinations in independent experiment. There was a significant difference in the estrogenicity among different HA concentrations using one-way ANOVA test (⁄⁄p < 0.01).

The E2 removal by biodegradation and sorption process influences its estrogenicity as a result of reduced impact on estrogen receptors. The E2 sorption with colloid organic materials causes a reduction in its estrogenic activity (Holbrook et al., 2005). In connection with E2 removal fraction by sorption and biodegradation, estrogenicities in HA solutions containing E2 were estimated at 24 h by means of E-screen bioassay (Fig. 3). The HA presence showed slightly higher estrogenicity than that with no HA, which represented the highest E2 removal. There was a significant

difference in the estrogenicity among different HA concentrations (⁄⁄p < 0.01). Moreover, estrogenicities were significantly different between the group of different HA concentrations and the positive control (⁄⁄⁄p < 0.001). When the estrogenicity for only HA as a control ranging from 0.25 ± 0.01 to 0.30 ± 0.23 ng-EEQ L1 was considered, it was estimated to be relatively low in the presence of HA even at low E2 removal. This phenomenon was directly related to the influence of anti-estrogenic activity on HA itself toward MCF-7 BUS cell. Due to exposure to 20 mg L1 of HA, the b-galactosidase activity was inhibited by 40% (Wu et al., 2009). Anti-estrogenic effects of 60% for luciferase activity of human breast carcinoma MVLN cell were observed at 50 mg L1 of HA concentration (Janošek et al., 2007). As HA level was increased, estrogenicity declined from 0.95 to 0.61 ng-EEQ L1 in the presence of nutrients and from 1.05 to 0.80 ng-EEQ L1 in the absence of nutrients, respectively. Such extent of estrogenicity did not differ significantly among various HA concentrations. 3.5. Nutrients effects The availability of nutrients, especially nitrogen and phosphorus are important factors influencing the biodegradation of organic compounds (Swindoll et al., 1988). In natural waters, NO 3 –N and 1 PO3 and 4 –P concentrations typically exists at below 1 mg L 0.1 mg L1, respectively, which are two orders of magnitude lower than those used in present study. Such insufficient nutrients will not be able to support biological activity, hence limiting the extent of EDCs biodegradation (Das et al., 2004; Ren et al., 2007; Ying et al., 2008). The E2 biodegradation, transformation to E1, E2 sorption, and E2 removal as well as relative estrogenicity among HA concentrations in the absence of nutrients were assessed by considering the limiting nutrients in natural waters (Table 3). As HA levels were increased, the kbioE2 values were decreased at a much slow rate ranging from 0.05 ± 0.01 to 0.02 ± 0.01 (h1), however, those values were not significantly different from various HA concentrations (F = 1.7; p = 0.31). Besides, E2 biodegradation was significantly reduced from 59% to 29% (p = 0.04), meanwhile E2 sorption was significantly increased from 0 to 35% (p = 0.001). A strongly negative correlation was found between biodegradation and sorption of E2 at various HA concentrations (r = 0.96; ⁄⁄ p < 0.01). In addition, sequential reduction of E2 biodegradation resulted in a significant decrease in transformation to E1 (⁄⁄p < 0.01). However, statistically significant difference in the overall E2 removal was not observed among HA concentrations (p > 0.05). The 59–65% of E2 were removed at 0–50 mg C L1 of HA in the absence of nutrients, which was 1.2–1.5 times lower than that in the presence of nutrients. Such low E2 removal resulted in higher estrogenicity than that in the presence of nutrients. However, estrogenicities among various HA concentrations were not significantly different using one-way ANOVA tests (p > 0.05). When HA was not added, the bacterial activities in the absence of nutrients were much more reduced, resulting more decreased biodegradation rate of E2 than in the presence of nutrients. As the HA concentration increased, the biodegradation rate of E2 gradually decreased. This revealed that the recalcitrant HA was

Table 3 The sorption, biodegradation, and removal (%) of E2 at various HA concentrations in the absence of nutrients. Parameters (%)

Maximum transformation to E1 Biodegradation Sorption Removal a

HA concentration (mg C L1) 0

10

30

50

F

p-valuea

33 ± 0.1 59 ± 4 – 59 ± 4

24 ± 1.7 56 ± 4 8±1 64 ± 4

24 ± 0.2 45 ± 9 19 ± 3 64 ± 6

22 ± 1.1 29 ± 9 35 ± 4 65 ± 4

48.3 7.6 62.1 0.6

0.001⁄⁄ 0.040⁄ 0.001⁄⁄ 0.628

Significant differences were found at various HA concentrations using one-way ANOVA test (⁄p < 0.05,

⁄⁄

p < 0.01).

J.H. Lee et al. / Chemosphere 85 (2011) 1383–1389

not utilized by bacterial community as a carbon source, and gives the inhibitory effect on bacterial activity (Steinberg et al., 2008). When nutrients concentrations are limited in natural waters, the biodegradation rates of organic chemicals are negatively affected (Zaidi and Imam, 1999). This study suggests poor nutrition condition in aquatic environments must be considered in estimating E2 biodegradation and its estrogenicity. 4. Conclusions The E2 biodegradation and transformation to E1 were significantly decreased with increasing HA concentration, with E2 biodegradation rate constant and maximum transformation to E1 being the highest in the absence of HA. E2 biodegradation was the dominant contributor to its removal, which showed a significantly negative relationship with E2 sorption. As HA concentration was increased, E2 removal was significantly enhanced, with a significantly elevated E2 sorption from 0% to 22%, and E2 biodegradation was decreased from 89% to 56%. Therefore, estrogenicity was estimated to be the lowest in the absence of HA, with the highest E2 removal of 89%. Besides, nutrients were also shown to be a key factor to determine the E2 biodegradation and resulting estrogenicity. As a consequence, it was demonstrated that the HA and nutrients present in natural water, which has not been focused in previous studies, should be considered in estimating the E2 removal and estrogenicity. Acknowledgments This research was supported by ‘‘Basic Science Research Program’’ through the National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (2009-0074074), and ‘‘Innovative Technology of Ecological Restoration’’ Project at GIST. References Aiken, G.R., 1985. Isolation and concentration techniques for aquatic humic substances-humic substances in soil, sediment, and water: geochemistry, isolation, and characterization. John Wiley and Sons, New York. pp. 363–385. Albers, C.N., Banta, G.T., Hansen, P.E., Jacobsen, O.S., 2008. Effect of different humic substances on the fate of diuron and its main metabolite 3,4-dichloroaniline in soil. Environ. Sci. Technol. 42, 8687–8691. Andersen, H.R., Hansen, M., Kjolholt, J., Lauridsen, F.S., Ternes, T., Sorensen, B.H., 2005. Assessment of the importance of sorption for steroid estrogens removal during activated sludge treatment. Chemosphere 61, 139–146. Anderson, H., Siegrist, H., Halling-SØrensen, B., Ternes, T.A., 2003. Fate of estrogens in municipal sewage treatment plant. Environ. Sci. Technol. 37, 4021–4026. Bowerman, B.L., O’Connell, R.T., 1990. Linear statistical models: an applied approach, second ed. Belmont, CA, Duxbury Press. Cheng, X., Zhao, L., Wang, X., Lin, J.M., 2007. Sensitive monitoring of humic acid in various aquatic environments with acidic cerium chemiluminescence detection. Anal. Sci. 23, 1189–1193. Chin, Y.P., Aiken, G., O’Loughlin, E., 1994. Molecular weight, polydispersity, and spectroscopic properties of aquatic humic substances. Environ. Sci. Technol. 28, 1853–1858. Das, B.S., Lee, L.S., Rao, P.S.C., Hultgren, R.P., 2004. Sorption and degradation of steroid hormones in soils during transport: column studies and model evaluation. Environ. Sci. Technol. 38, 1460–1470. de Mes, T., Zeeman, G., Lettinga, G., 2005. Occurrence and fate of estrone, 17bestradiol and 17a-ethynylestradiol in STPs for domestic wastewater. Rev. Environ. Sci. Biotechnol. 4, 275–311. Environment Agency (EA), 2002. Proposed Predicted-No-Effect-Concentrations (PNECs) for Natural and Synthetic Steroid Oestrogens in Surface Waters, R&D Technical Report P2-T04/1, Environmental Agency, Bristol, UK. Hansen, P.D., Dizer, H., Hock, B., Marx, A., Sherry, J., McMaster, M., Blaise, C., 1998. Vitellogenin-a biomarker for endocrine disruptors. TrAC. 17, 448–451. Hashimoto, T., Murakami, T., 2009. Removal and degradation characteristics of natural and synthetic estrogens by activated sludge in batch experiments. Water Res. 43, 573–582. Holbrook, R.D., Novak, J.T., Love, N.G., 2005. Impact of activated sludge derived colloidal organic carbon on behavior of estrogenic agonist recombinant yeast bioassay. Environ. Toxicol. Chem. 24, 2717–2724.

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