Electrochemical oxidation of quinoline aqueous solution on β-PbO₂ anode and the evolution of phytotoxicity on duckweed

Nitrogen-heterocyclic compounds (NHC_s) like pyridine and its derivatives, indole, quinoline, etc. have received increasing attention due to their toxic, carcinogenic, mutagenic, and severe odour potential (Padoley et al. 2008), and their azardous effects on ecosystem and human health (Stapleton et al. 2010). Pyridines have been rated as priority pollutants by the United States Environmental Protection Agency (Mathur et al. 2008) and extensively used for the production of pesticides, herbicides, pharmaceuticals, industrial solvents, dyes, rubber chemicals and so on (Acheson 1960), thereby frequently existing in the effluent of the aforementioned industries. Once NHC_s enter into the environment, they can persist for long periods (Rogers et al. 1985), get easily transported through the soil and then contaminate ground water (Kuhn & Suflita 1989) due to the fact that their heterocyclic structure makes them more soluble than their homocyclic analogues (Padoley et al. 2008). In addition, the mutagenic potential of NHC_s increases with the increasing number of rings: for example, quinoline is confirmed to be more mutagenic than pyridine (Mohammad et al. 1983). As a cost-effective process, biological oxidation using specific degradation bacteria is definitely becoming the primary choice to treat industrial wastewater containing quinoline. Various quinoline-degrading bacteria have been obtained, such as Pseudomonas sp. (Griese et al. 2006; Sun et al. 2009; Qiao & Wang 2010), Burkholderia sp. (Wang et al. 2001), Rhodococcus sp. (Zhu et al. 2008), Comamonas sp. (Cui et al. 2004), Bacillus sp. (Tuo et al. 2012), Brevundimonas sp. (Wang et al. 2015b) and so on. However, conventional biological processes are still restricted in practical application because of some inherent shortcomings, such as the time-consuming acclimation of special microorganisms and high selectivity (Xing et al. 2012), long hydraulic retention time and poor quinoline concentration tolerance (Padoley et al. 2008). Advanced oxidation processes (AOP_s) need to be eventually employed, which are capable of either the complete removal of the pollutant from wastewater or at the very least the destruction of the inhibition on biological activity due to the hydroxyl radicals (·OH) reaction (Yu et al. 2013). 91.5% of quinoline could be removed through photo-catalytic oxidation using TiO₂ nano-particles as a catalyst within 240 min under its optimized conditions (Jing et al. 2012). However, TiO₂ is difficult to recover or separate from treated effluent, and may bring secondary environmental contamination (Ma et al. 2012). Electrochemical oxidation could overcome the aforementioned issues and is easy to regulate under a wide range of conditions by adjusting the current. ·OH electro-generated on the surface of a β-PbO₂ anode (Cong et al. 2008) could effectively convert an aromatic pollutant to non-toxic and biodegradable organic acids (Wu et al. 2005). Nevertheless, it is worthwhile stressing that some intermediates formed during AOP_s were more toxic than the parent pollutant. For instance, benzoquinone is the most toxic intermediate during electrochemical oxidation of phenol (Wu & Zhou 2001). 3,4-Dichloroaniline (3,4-DCA), one of the intermediates formed during electro-Fenton of diuron, was found to present a specific toxicity 90 times higher than diuron (Oturan et al. 2008). Therefore, not only the removal of parent compound but also the toxicity of degradation intermediates should be equally considered. However, separation and identification of all intermediates using chemical analysis are usually insufficient because of the enormous number of intermediates, which may be at a concentration too low to allow analytical determination. Consequently, it is difficult to predict the toxic property of the effluent using a physicochemical approach. Hence, utilizing a biological test system with living cells or organisms may be the promising option, which can give a complete response to the complex mixture of intermediates without any prior knowledge of its composition or chemical properties (Radić et al. 2011).

Currently, the bioluminescence Microtox^® standard method (Oturan et al. 2008; Dirany et al. 2010; Mousset et al. 2014), green alga (Oturan et al. 2008), specific oxygen uptake rate (Wang et al. 2015a) and duckweed test (Abdul et al. 2012) have been used to assess the comprehensive toxicity of treated effluent during AOP_s. Among them, duckweed possesses physiological properties (small size, high multiplication rate and vegetative propagation) and may selectively accumulate certain chemicals, thus it could serve as an ideal biological monitor to assess the toxicity of surface waters and wastewaters (Radić et al. 2011) that is more economical, intuitional and convenient. It is not necessary to conduct the test procedure axenically, and it is feasible to keep contamination by other organisms to a minimum. When electrochemical oxidation is employed to degrade the refractory pollutant, the electro-generated ·OH has the ability to disinfect the aqueous solution, implying that duckweed could be exposed directly to the effluent after oxidation.

In this study, electrochemical oxidation of quinoline using a β-PbO₂ electrode modified with fluoride resin (Wu & Zhou 2001; Zhou et al. 2005) as anode was firstly carried out, and the effects of current density, initial pH and Na₂SO₄ dosage on the removal of quinoline were investigated. High performance liquid chromatography (HPLC), ion chromatography (IC) and gas chromatography-mass spectrometry (GC-MS) analysis were applied to determine the formation of intermediates, based on which the electrochemical oxidation mechanism of quinoline on the β-PbO₂ anode was postulated. Then the evolution of the toxicity of the quinoline solution with electrochemical oxidation time was determined through the exposure of duckweed to the oxidized solution over a period of 3–7 d. Chlorophyll a content (C_a) and the frond number (FN) of duckweed measured with and without pH adjustment and the addition of nutrients to the oxidized quinoline solution prior to culturing the duckweed were chosen as the growth parameters.

Quinoline was purchased from Sigma-Aldrich. Then 150 mg/L of quinoline aqueous solution was prepared by diluting quinoline in double distilled water, and the initial pH was 6.2. Methanol for HPLC analysis was chromatographic grade. Other chemicals used were analytical grade and purchased from Hangzhou Huipu Chemical Reagent Co. Ltd (Hangzhou, China).

Electrochemical oxidation of quinoline was performed in an open, cylindrical, one-compartment and double layer Pyrex reactor (inner layer size: Ф 8.0 × 9.0 cm). Cooling water was passed through the hollow layer of the reactor to keep the solution at room temperature. For each set of experiments, 200 mL of quinoline solution was added into the reactor. The pH was adjusted to the desired value using 0.1 mol/L of H₂SO₄ and 0.1 mol/L of NaOH solutions and measured with a pH meter (pH3110 SET2, Germany). A modified β-PbO₂ anode (Ф 4.5 × 6.4 cm, effective area 90 cm²) was centered in the electrochemical reactor and surrounded by a stainless steel net cathode that covered the inner wall of the reactor. Constant current was maintained at the desired level with a digital DC power supply (WYL3010, Hangzhou, China). Samples were periodically taken for HPLC, GC-MS and IC analysis to examine the variation of quinoline concentration, the formation of main organic components and the transformation of quinoline-N. The removal of quinoline was calculated according to the concentration of quinoline before and after oxidation.

To ensure an adequate solution to cultivate the duckweed for the toxicity test, electrochemical oxidation of quinoline was performed in a bigger reactor (inner layer size: Ф10 × 20 cm) using a β-PbO₂ anode (Ф 4.5 × 20 cm, effective area 200 cm²) under 30 mA/cm² of current density, 2.0 g/L of Na₂SO₄ and pH 6.2. 900 mL of quinoline solution was added into the reactor. Samples were taken after variable periods of oxidation time (up to 4 h) for the duckweed growth test to identify the phytotoxicity of the quinoline solution during electrochemical oxidation. The concentration of quinoline was also examined by HPLC. Chemical oxygen demand (COD) was determined in accordance with method 508C (closed reflux, colorimetric method) in Standard Methods (APHA 1998).

Duckweed was originally collected from a small pond at Xiaoshan City (China), which was free of obvious pollution sources, and maintained in nutrient solution for three months prior to toxicity testing. Nutrient solution for duckweed culture was prepared according to the OECD protocol (OECD 2006) and was adjusted to pH 7.5 ± 0.1. The duckweed was pre-cultivated in nutrient solution for one week under continuous cool fluorescent light at an intensity of 8,000 lux and at 24 ± 2 °C. Following the adaptation period, 20 healthy colonies with 2 fronds per colony were transferred to each 100 mL beaker containing 50 mL of oxidized quinoline solution. Duckweed grown on 2.0 g/L of Na₂SO₄ solution served as a control. Then the beakers were covered with cling film that had 20 needle holes punched through the film to reduce evaporation and ensure that air could circulate. Finally, the beakers were placed into a biochemical incubator (GZX-300BSH-III, Shanghai, China) and taken out to be shaken by hand once every day. The temperature and intensity of light in the biochemical incubator were controlled at 24 ± 2 °C and 8,000 lux, respectively. Three replicates were prepared from each treatment. The FN in each beaker was recorded from 0 d to 7 d.

Samples of quinoline solution taken after several different reaction times were directly used to culture duckweed as soon as possible, this is because duckweed can grow easily in a wide range of pH values (3.5–10.5) (Khellaf & Zerdaoui 2010). C_a and FN were used to evaluate the growth of duckweed. Additionally, in order to reduce the effect of pH variation on the growth of duckweed, the pH of the treated quinoline samples at different oxidation times, including the control, was adjusted to 7.5 and then the duckweed was cultured again. Similarly, in order to consider the effect of additional nutrients, the distilled water was replaced with partially-degraded quinoline solution for dissolving the nutrient salts to prepare the nutrient samples that were used to culture duckweed.

The concentration of quinoline was analyzed using an HPLC equipped with a UV detector (Agilent 1200, USA) and the wavelength was set at 254 nm for quinoline and 210 nm for carboxylic acids (Bielicka-Daszkiewicz et al. 2012). The samples were analyzed using a TC-C18 column (Agilent, 4.6 mm × 250 mm × 5 μm). The mobile phase consisted of methanol:water (60:40, v/v) with a flow rate of 1.0 mL/min. Water buffered with phosphoric acid to pH 2.5 at a flow rate of 0.6 mL/min was used as the mobile phase during the separation of carboxylic acids. The column temperature was 30 °C and the injection volume was 20 μL.

The main organic intermediates formed during quinoline degradation were analyzed by GC-MS (Agilent 7890A - 5973C, USA) with an HP-5MS column (30 m × 0.25 mm × 0.25 μm). The samples were extracted by dichloromethane twice at a total volume ratio of 1 and nearly dried by a rotary evaporator (RE3000, China) at 40 °C, then filtered over anhydrous Na₂SO₄, and subsequently concentrated to 1.0 mL by pressure blowing concentrator (MTN-28000, China) with high pure nitrogen. The injection volume was 2 μL. The column temperature program for GC-MS was 40 °C isothermal for 2 min, and ramped to 300 °C with an increment of 7 °C/min, then holding for 15 min. MS analysis was performed at MS ion source of 230 °C and electron energy of 70 eV (Tuo et al. 2012). High pure helium was used as the carrier gas (1.0 mL/min). The injector temperature was 250 °C.

Evolution of inorganic ions during quinoline degradation was determined by IC (Dionex, ICS-90) equipped with a DS5 conductivity detector and an IonPac AS14 (250 mm × 4 mm (id)) anion-exchange column, linked to an IonPac AG14 (50 mm × 4 mm (id)) column guard. The mobile phase was a mixture of 3.5 mmol/L Na₂CO₃ and 1.0 mmol/L NaHCO₃ with a flow rate of 1.2 mL/min. The injection volume was 10 μL. All samples were filtered through a 0.22 μm membrane before IC analysis.

The live FN (green part/total part ≥60%) was counted and recorded every day during the exposure period (7 d) and the protruding, distinctly visible fronds counted.

Many literatures have reported the influence of solution pH during anodic oxidation of organics on a PbO₂ anode, but the results are diverse and even contrary (Samet et al. 2006; Panizza et al. 2008), hence the effect of pH on the degradation performance of quinoline needs to be investigated. Considering that the β-PbO₂ anode is more favorable in acidic medium, the experiments were carried out within the pH range of 4.0–6.2 at 150 mg/L of quinoline, 2.0 g/L of Na₂SO₄ and 50 mA/cm² of current density.

The aforementioned results demonstrate that quinoline could be effectively removed by electro-generated ·OH, and some intermediates at low overall yields were formed during oxidation. It is crucial to assess the comprehensive toxicity of the effluent at different reaction times. In this part of the study, duckweed was directly or indirectly cultured in the effluent after electrochemical oxidation of quinoline under the following conditions: 30 mA/cm² of current density, 2.0 g/L of Na₂SO₄, and 150 mg/L of quinoline solution with an initial pH of 6.2. The phytotoxicity potential of the treated solutions was assessed according to the growth inhibition of duckweed.

When the duckweed was directly exposed to the effluent after 0 min and 60 min of oxidation time, the C_a of the duckweed decreased from 0.82 mg/L to 0.21 mg/L, indicating that the toxicity of the quinoline solution increased with the increase in oxidation time. When the pH of the treated solution was adjusted to 7.5, the C_a content of the duckweed correspondingly decreased from 1.16 mg/L to 0.43 mg/L. These observations indicate that the toxicity evolution of the effluent during the first 60 min of electrochemical oxidation is related not only to the variation in pH value but also to the toxicity of the intermediates formed. When the oxidation time was 60 min, the pH of the solution decreased sharply from 6.2 to 4.3, which significantly inhibited the growth of the duckweed. On the other hand, the formation of some cyclic intermediates might possess a strong inhibition to duckweed growth. Although 50% of the quinoline was removed at this time, the COD removal was only 36.8%, implying that the quinoline was not mineralized. Consequently, with the pH adjustment to 7.5, the growth inhibition of duckweed was not obviously alleviated, indicating that the toxicity of the effluent at the first 60 min might mainly come from some intermediates that may be more toxic to duckweed than quinoline. It was reported that the addition of one hydroxyl group at the C-8 position on quinoline caused a 40-fold increase in the inhibitory effect on the growth of Toxoplasma (Kadri et al. 2014) and 8-hydroxyquinoline had significant acute toxicity in adult Cryprinus carpio (Yan et al. 2015). With the prolonging of oxidation time to above 100 min, the intermediates were gradually converted to organic acids under the attack of ·OH, and the growth inhibition of duckweed was primarily caused by the decrease in the pH of the solution (from 4.3 to 3.8–4.0), and this could be elucidated from the reduction of the growth inhibition of duckweed. For example, the C_a contents of duckweed exposed to the solution after 160 min of oxidation with and without pH adjustment were 1.06 mg/L and 0.17 mg/L, respectively.

A similar trend of growth inhibition of duckweed was noticed with FN (Figure 6(b)). When the duckweed was exposed to the quinoline solutions after 60–160 min of oxidation, duckweed leaves started to look brown and gradually became black, and even died. However, once the pH was adjusted to 7.5, the FN significantly increased, especially when the duckweed was exposed to quinoline solutions after 120 and 160 min of oxidation, when the FN increased from 0 to 38 and 0.3 to 42, respectively. This also showed that the toxicity evolution of the quinoline solution after 100 min of treatment was mainly affected by the pH variation due to the formation of organic acids. Although pH adjustment could alleviate the growth inhibition of duckweed, the increasing amount of FN at 20 and 60 min was far less than that at 100–200 min. Therefore, the toxicity of the quinoline solution during electrochemical oxidation was mainly governed by the formation of toxic intermediates during the initial reaction period. The acidic medium played a dominant role in the phytotoxicity of duckweed due to the abundant accumulation of organic acids in the subsequent reaction period. On this account, ensuring a reaction time above 120 min and pH adjusted to 7.5, the toxicity of the effluent on duckweed could be disregarded.

Quinoline could be effectively removed through electrochemical oxidation using a β-PbO₂ anode doped with fluoride resin, and was eventually converted to some biodegradable organic acids (such as oxalic acid, formic acid, acetic acid, propionic acid and fumaric acid); the quinoline-N was finally transformed to ⁠. The test of duckweed growth demonstrated that the formation of some intermediates indeed increased the comprehensive toxicity of the quinoline solution during electrochemical oxidation, especially after 60 min of oxidation. The inhibition of duckweed growth caused by the decrease in pH value due to the formation of organic acids could be alleviated through adjusting the pH of the treated quinoline solution to 7.5. Furthermore, the addition of nutrients to the treated quinoline solution was beneficial to the growth of the duckweed. Hence, quinoline solution under this study after 120 min of oxidation could be well used as a follow-up biological treatment.

The authors gratefully acknowledge the financial support of Zhejiang Provincial Natural Science Foundation of China (Grant No. LY12E08014).