Characterization of Biodegradation Intermediates of Nonionic

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Biomacromolecules 2003, 4, 46-51

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Characterization of Biodegradation Intermediates of Nonionic Surfactants by MALDI-MS. 2. Oxidative Biodegradation Profiles of Uniform Octylphenol Polyethoxylate in 18O-Labeled Water Hiroaki Sato,*,† Atsushi Shibata,‡ Yang Wang,‡ Hiromichi Yoshikawa,§ and Hiroto Tamura‡ Measurement Technology Group, Institute for Environmental Management Technology, National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba 305-8569, Japan, Agricultural High-Tech Research Center, Meijo University, Nagoya 468-8502, Japan, and Department of Environmental Chemistry, Faculty of Engineering, Kyushu Kyoritsu University, Kitakyusyu 807-8585, Japan Received July 15, 2002; Revised Manuscript Received September 30, 2002

This paper reports the characterization of the biodegradation intermediates of octylphenol octaethoxylate (OP8EO) by means of matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). The biodegradation test study was carried out in a pure culture (Pseudomonas putida S-5) under aerobic conditions using OP8EO as the sole carbon source and 18O-labeled water as an incubation medium. In the MALDI-MS spectra of biodegraded samples, a series of OPnEO molecules with n ) 2-8 EO units and their corresponding carboxylic acid products (OPnEC) were observed. The use of purified OP8EO enabled one to distinguish the shortened OPEO molecules as biodegradation intermediates. Furthermore, the formation of OP8EC (the oxidized product of OP8EO) supported the notion that terminal oxidation is a step in the biodegradation process. When biodegradation study was carried out in 18O-labeled water, incorporation of 18O atoms into the carboxyl group was observed for OPEC, while no incorporation was observed for the shortened OPEO products. These results could provide some rationale to the biodegradation mechanism of alkylphenol polyethoxylates. Introduction Alkylphenol polyethoxylates (APEOs), such as octylphenol polyethoxylate (OPEO) and nonylphenol polyethoxylate (NPEO), are a widely used class of nonionic surfactants with domestic, industrial, and agricultural applications as detergents, emulsifiers, adjuvants in agricultural products, and so on. After their application, these surfactants might be discharged with wastewater into the environment, where the hydrophilic ethylene oxide (EO) chains of APEOs are easily biodegraded (primary biodegradation) by most bacteria to form stable metabolic products, for example, alkylphenols (APs), APEOs with short EO chains, and their carboxylate products oxidized at the EO chain terminus (APECs).1-6 In recent years, several reports have indicated that the primary biodegradation products of APEOs exhibit estrogenic activity.7-11 Therefore, elucidation of the primary mechanisms of the biodegradation of APEOs is important to understand the fate of APEOs in the natural environment.12-16 It is almost certain that the gradual shortening of the EO chain from the hydroxyl terminal side is a common pathway of the primary biodegradation and that terminal oxidation of EO chain also occurs. However, the biodegradation mechanisms of APEOs are still not fully understood. * To whom corresponding should be addressed. Phone: +81-298-618302. Fax: +81-298-61-8308. E-mail: [email protected]. † AIST. ‡ Meijo University. § Kyushu Kyoritsu University.

Ahel et al. reported that the terminal oxidation of the EO chain was the principal mechanism in the initial biodegradation of APEOs.15 Regarding the biodegradation of poly(ethylene glycol) (PEG), the structure of which corresponds to the EO chain of APEOs, Kawai has already proposed that terminal oxidation and exo-type ether cleavage of the EO chain proceed alternately.17 Because APECs with short EO chains (mainly n < 4) have been detected in many environmental samples, it has generally believed in the field of environmental science that a terminal oxidation process of the EO chain might follow the EO shortening process. Whether an ether cleavage (exo-type) takes place with APEC has to be confirmed. Jonkers et al.18 have studied the NPEO biodegradation in river water under aerobic conditions using liquid chromatography-electrospray ionization mass spectrometry-tandem mass spectrometry (LC-ESI-MS/MS). Because NPECs (carboxylates of NPEO oxidized at EO terminal) with long EO chains at the initial stage were detected, they concluded that terminal oxidation was the initiating step of NPEO biodegradation. Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) would also be an effective analytical tool for the characterization of APEOs.19-25 In our previous study (part 1),26 the intermediates in the bacterial biodegradation of OPEO under aerobic conditions have been characterized by means of MALDI-MS. Because the formation of OPEC and the changes in molecular weight distribution during biodegradation test could be successfully monitored, we modified the

10.1021/bm025616q CCC: $25.00 © 2003 American Chemical Society Published on Web 10/31/2002

Characterization of Biodegradation Intermediates

Figure 1. Possible primary biodegradation mechanisms of OPEO under aerobic conditions.26 In this model, OPEO molecules are oxidized to form carboxylic acid products (OPEC) at the EO chain terminus (step I), followed by scission of the ether bond to form OPEO molecules shortened by one EO unit with the liberation of glyoxylic acid (step II). This cycle proceeds until n - i ) 2 or 3.

PEG biodegradation model by Kawai17 for the biodegradation of OPEO as shown in Figure 1.26 Although reports by both Jonkers et al.18 and us26 supported the terminal oxidation model, unfortunately these reports could not show clear evidence that the first step is terminal oxidation because commercial OPEOs are a mixture of molecules with different EO chain lengths and the first reacting products could not be distinguished. In this report, to clarify the mechanisms underlying the oxidative biodegradation of APEOs, we propose a novel approach for the characterization of the biodegradation intermediates of OPEO by MALDI-MS using uniform OPEO with n ) 8 EO units (octylphenol octaethoxylate, OP8EO) as a sample and using 18O-labeled water as a incubation medium. The use of uniform sample with no molecular weight distribution would be an effective approach to distinguish between partly degraded OPEO molecules and the original undegraded one. Furthermore, stable isotopelabeling studies using 18O isotope would be expected to provide an important clue as to the nature of biodegradation pathways. Experimental Section Surfactant Sample Purification. Uniform OP8EO molecules were isolated from Triton X-100 (Aldrich Chemical Co; average EO units n ) ca. 9.5) by column chromatography. About 2 mg of Triton X-100 dissolved in 5 mL of an eluting solvent (ethyl acetate/acetone/water ) 55/35/10 vol %) was applied to an open column (44 mm i.d. × 730 mm long) packed with 300 g of silica gel (Merck Japan Ltd., particle size ) 0.063-0.200 mm). Fractions of about 4 mL were collected, and the components of each fraction were checked by MALDI-MS and gas chromatography. The fractions containing OP8EO were further purified 2 times until no significant amount of impurities such as OP7EO and OP9EO was detected.

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Bacterial Biodegradation Test. The nature of biodegradation pathways of APEOs could depend on the degradation conditions, especially bacteria species (or strains). Therefore, we used the pure strain of bacteria isolated from a paddy field soil, which was the same as that used in our previous study.26 In this report, the bacteria was tentatively identified as Pseudomonas sp.26 Afterward, it was further identified as Pseudomonas putida and designated strain S-5.27 The bacterial biodegradation procedure was similar to that described in the previous study,26 except for the use of 18Olabeled water and a reduction in the volume of the culture medium. First, 200 µL of a basal salt medium containing ca. 0.1% (w/v) OP8EO as the sole carbon source prepared with normal water was taken in each of several vials with an inner diameter of 18 mm. After the freeze-drying of the basal salt medium, 200 µL of 18O-labeled water (Cambridge Isotope Laboratories Inc., ca. 95-98% purity) was added into each vial. Then, 4 µL of liquid medium containing bacteria, the optical density at 660 nm of which was adjusted to 0.5, was inoculated into the medium. Because the inoculum was prepared with normal water, the final concentration of 18O-labeled water in the culture medium was slightly diluted to be ca. 90-95%. As control tests, a medium with normal water in the presence of bacteria and a medium with 18O-labeled water without bacteria were also examined under the same conditions. The vials were incubated with constant shaking at 30 °C for 12, 24, 36, 48, and 72 h. After the designated time, 20 µL of the culture medium was collected from the vial into glass tubes with an inner diameter of 4 mm and then immediately vacuum-dried. The tubes were then sealed and stored in a refrigerator at 4 °C until MALDIMS measurement. MALDI-MS Measurement. As the matrix for sample ionization, 5,10,15,20-tetrakis(pentafluorophenyl)porphyrin (F20TPP, Sigma Chemical Co., MW ) 974.6) was used, which provides clear MALDI spectra of low molecular weight samples (< ca. m/z 750) with minimum matrix interference from matrix signals.24 About 2 mg of F20TPP was dissolved 1 mL of ethyl acetate to make the matrix solution. To extract the biodegradation intermediates from the dried culture medium, 20 µL of ethyl acetate was added to the stored glass tubes. The sample extract (1 µL) was mixed with the matrix solution (3 µL) in a glass tube with an inner diameter of 4 mm. Prior to the sample/matrix cocrystal deposition, 1 µL of a 1 mM sodium iodide solution in acetone was spotted onto a flat stainless steel sample plate, and dried in air to deposit fine NaI crystals as a cationization salt. Then, 1 µL of the sample/matrix solution was pipeted onto the thin NaI crystal layer and dried in air. The MALDI-MS measurements were performed using a Voyager DE-PRO time-of-flight mass spectrometer (Applied Biosystems, Japan) equipped with a pulsed nitrogen laser (λ ) 337 nm, 3 ns pulse width, and 3 Hz frequency) and a delayed extraction ion source. Laser beam intensity was experimentally attenuated to just above the threshold for analyte ionization. Ions generated by the laser desorption were introduced into the flight tube with an acceleration voltage of 20 kV in the high-resolution reflector (2.0 m flight path) positive ion mode. The delay time setting was typically

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Figure 2. Typical MALDI mass spectra of (a) the original OP8EO, together with its biodegradation intermediates recovered after incubation for (b) 12 and (c) 48 h. The / corresponds to matrix fragments or impurity peaks. R means p-octylphenyl group.

50 ns. All mass spectra were collected by averaging 500 individual laser shots. Results and Discussion Biodegradation Profiles of Uniform OP8EO. Figure 2 shows typical MALDI mass spectra of the original OP8EO (a) and its biodegradation intermediates recovered after incubation for 12 h (b) and 48 h (c) with normal water. In the MALDI mass spectrum of the original OP8EO (Figure 2a), an apparent single peak is clearly observed at around m/z 581, which corresponds to sodium-cationized OP8EO [R-(CH2CH2O)8-H + Na+], where R represents the poctylphenyl group. Here, it should be noted that considerably fewer matrix fragments or impurity peaks or both are observed in the mass spectrum. This unique feature of F20TPP matrix24 is convenient for a brief survey of

biodegradation intermediates as shown in Figure 2b,c. In both MALDI mass spectra of biodegraded samples, various peaks corresponding to biodegradation intermediates are mainly observed in the range ca. m/z 300-620. According to our previous report,26 the series of ion peaks labeled with circles (O) can be attributed to OPnEO molecules with a sodium cation attached [R-(CH2CH2O)n-H + Na+; m/z ) 44n + 206 + 23]. The other series (1 and 3) can be assigned to the ions of carboxylated OPnEO molecules (OPnEC) with a sodium cation attached [R-(CH2CH2O)n-1-CH2COOH + Na+; m/z ) 44(n - 1) + 264 + 23] and their sodium salts (OPnECNa) [R-(CH2CH2O)n-1-CH2COONa + Na+; m/z ) 44(n - 1) + 286 + 23], respectively. The formation of carboxylate salt ions such as OPnECNa, which might be mainly generated during sample preparation or ionization or both, is a common phenomenon in MALDI-MS measurements using sodium cationization salts.28 Here, the formation

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Figure 4. Enlarged MALDI mass spectra (trimer region) of the biodegraded OPEO recovered after 48 h incubation using (a) normal water and (b) 18O-labeled water.

Figure 3. Changes in peak distribution during OP8EO biodegradation for (a) 12, (b) 24, (c) 48, and (d) 72 h observed by MALDI-MS. Each bar consists of OPEO and oxidized products (OPEC and OPECNa). Error bars are given as standard deviation for three measurements for each of three individual tests (i.e., n ) 9).

of OP8EC and their sodium salts (OP8ECNa) strongly supports that the first step in the biodegradation reaction is the terminal oxidation. On the other hand, no formation of the products oxidized at the octyl-chain moiety was observed, which were frequently observed in the environmental samples. The use of purified OP8EO enables one to distinguish the shortened OPEO molecules as biodegradation intermediates from the original undegraded forms. In Figure 2b, the most intense peak observed at m/z 581 corresponds to “undegraded” OP8EO, while the other OPnEO molecules (n ) 2-7) are surely biodegradation intermediates. For further biodegraded samples recovered after 48 h incubation (Figure 2c), the original OP8EO peak disappears almost completely,

whereas the molecular weight distribution of the biodegradation intermediates shifts to a lower mass region with a maximum at m/z 397 corresponding to OP3ECNa molecules. The changes in the peak distribution of biodegradation intermediates up to 72 h are summarized in Figure 3. Although peak intensities might be affected by low-massdiscrimination effect19 to some extent, the profiles of gradual degradation from n ) 8 down to 2-3 with terminal oxidation could be observed semiquantitatively. In addition, we confirmed the formation of glyoxylic acid assayed by the method of Gamer and Gaunt.29 These results support the model previously proposed,26 in which terminal oxidation and exo-type ether cleavage of the EO chain proceed alternately as shown in Figure 1. Incorporation Behavior of Oxygen Atoms during Biodegradation Process. To elucidate the mechanisms behind the biodegradation of OPEO, biodegradation tests using 18O-labeled water were then carried out. To compare the MALDI mass spectra of the biodegraded OPEO using normal water and 18O-labeled water, Figure 4 shows the trimer region (OP3EO, OP3EC, and OP3ECNa) between m/z 350 and 410 of the products recovered after 48 h as an example. In both mass spectra, isotope peaks are completely separated. Interestingly, the peaks corresponding to OPEC molecules and their sodium salts shifted to higher masses by 2 and 4 Da when 18O-labeled water was used, while no peak shift was observed for the OPEO intermediates. Basically the same tendencies were observed for all other peaks. The mass increments of the carboxylic acid products clearly suggest that the origin of the oxygen atoms was water and that dehydrogenase was involved in this reaction. The incorporation of one, as well as two, 18O atoms could provide a clue as to the mechanisms of the biodegradation of OPEO, because similar phenomena have been reported in the field of proteomics using mass spectrometry, in which, on enzymatic digestion of proteins by serine proteases in highly enriched 18O-labeled water, the resulting peptides

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Figure 5. Possible formation pathways of OPEC molecules. X represents heteroatoms such as O, N, and S.

contained two 18O atoms at the carboxyl terminal group.29,30 In these cases, the incorporation of two 18O atoms could be explained by an oxygen-exchange reaction of ester compounds. While an oxygen-exchange reaction hardly takes place between free carboxyl groups and water molecules (except under extremely acidic or basic conditions), the carbonyl oxygen atoms in a functional group of an ester and amide are exchangeable for those of water molecules.31,32 Considering the oxygen-exchange reaction, Figure 5 illustrates a possible pathway for the formation of OPEC molecules. In the first stage, dehydrogenation of OPEO molecules might proceed to form aldehyde intermediates,33 which would soon be followed by the next dehydrogenation stage. At this stage, an enzyme (or coenzyme)-substrate complex linked via a covalent bond might be formed as a reaction intermediate. Here, the carbonyl oxygen (16O) of the originally formed complex (I) could be exchanged for 18 O through the formation of tetrahedral intermediates, so

an 18O-containing complex (II) would also be formed to some extent. As results of two successive dehydrogenation reactions and an oxygen-exchange reaction, a mixture of OPEC molecules with one and two 18O atoms might be formed. One possible candidate involved in this reaction is nicotinamide adenine dinucleotide (phosphate) [NAD(P)+]dependent aldehyde dehydrogenase, of which the cysteine residue as the active site could bond to aldehyde substrates to form a thiol-ester intermediate.34-37 The further study to identify this enzyme is currently in progress. On the other hand, the absence of a shift in molecular mass for the shortened OPEO molecules despite the use of 18 O-labeled water also has meaning (Figure 4). This fact suggests that at the ether-cleavage stage when shortened OPEO molecules are formed as shown in Figure 1, the oxygen atom (16O) at the ether bond would be left in the newly formed hydroxyl terminal of the OPEO molecule. On the basis of this observation, Figure 6 shows possible

Characterization of Biodegradation Intermediates

Figure 6. Possible formation pathways of shortened OPEO molecules.

mechanisms for the formation of shortened OPEO molecules, in which the incorporation of hemiacetal intermediates with an 18O-labeled hydroxyl group might be followed by scission of the neighboring ether bond to form a new hydroxyl terminal with liberation of glyoxylic acid as already proposed by Kawai.38 Conclusions This study has demonstrated the characterization using MALDI-MS of biodegradation intermediates of uniform OPEO employing the stable isotope-labeling technique. The possible biodegradation pathways for OPEO molecules could be proposed by using uniform sample. Furthermore, the incorporation of 18O atoms into biodegradation intermediates was successfully observed by MALDI-MS. Thus, the characterization method presented here would be useful for studying the biodegradation of various nonionic surfactants. Acknowledgment. This research was supported financially by Agricultural High-Tech Research Center, Meijo University, under the “Environmental Control through the Function of Microorganisms” project and a Grant-in-Aid (Endocrine Disruptors) from the Ministry of Agriculture, Forestry, and Fisheries of Japan (ED-02-V-1-3). References and Notes (1) Reinhard, M.; Goodman, N.; Mortelmans, K. E. EnViron. Sci. Technol. 1982, 16, 351-362. (2) Giger, W.; Brunner, P. H.; Schaffner, C. Science 1984, 225, 623625.

Biomacromolecules, Vol. 4, No. 1, 2003 51 (3) Ahel, M.; Conrad, T.; Giger, W. EnViron. Sci. Technol. 1987, 21, 697-703. (4) Di Corcia, A.; Samperi, R.; Marcomini, A. EnViron. Sci. Technol. 1994, 28, 850-858. (5) Lee, H.-B.; Peart, T. E.; Bennie, D. T.; Maguire, R. J. J. Chromatogr., A 1997, 785, 385-394. (6) Ding, W.-H.; Chen, C.-T. J. Chromatogr., A 1999, 862, 113-120. (7) Soto, A. M.; Justica, H.; Wray, J. W.; Sonnenschein, C. EnViron. Health Perspect. 1991, 92, 167-173. (8) Jobling, S.; Sumpter, J. P. Aquat. Toxicol. 1993, 27, 361-372. (9) White, R.; Jobling, S.; Hoare, S. A.; Sumpter, J. P.; Parker, M. G. Endocrinology 1994, 135, 175-182. (10) Jobling, S.; Sheahan, D.; Osborne, J. A.; Matthiessen, P.; Sumpter, J. P. EnViron. Toxicol. Chem. 1996, 15, 194-202. (11) Sonnenschein, C.; Soto, A. M. J. Steroid Biochem. Mol. Biol. 1998, 65, 143-150. (12) Swisher, R. D. Surfactant Biodegradation, 2nd ed.; Surfactant Science Series, Vol. 18; Marcel Dekker: New York, 1987. (13) Kravetz, L. J. Am. Oil Chem. Soc. 1981, 20, 58A-65A. (14) White, G. F. Pestic. Sci. 1993, 37, 159-166. (15) Ahel, M.; Hrsˇak, D.; Giger, W. Arch. EnViron. Contam. Toxicol. 1994, 26, 540-548. (16) Di Corcia, A.; Costantino, A.; Crescenzi, C.; Marinoni, E.; Samperi, R. EnViron. Sci. Technol. 1998, 32, 2401-2409. (17) Kawai, F. Spec. Publ.-R. Soc. Chem. 1992, 109, 20-29. (18) Jonkers, N.; Knepper, T. P.; De Voogt, P. EnViron. Sci. Technol. 2001, 35, 335-340. (19) Pasch, H.; Zammert, I.; Just, U. Int. J. Polym. Anal. Charact. 1995, 1, 329-338. (20) Barry, J. P.; Radtke, D. R.; Carton, W. J.; Anselmo, R. T.; Evans, J. V. J. Chromatogr., A 1998, 800, 13-19. (21) Cumme, G. A.; Blume, E.; Bublitz, R.; Hoppe, H.; Horn, A. J. Chromatogr. A 1997, 791, 245-253. (22) Willetts, M.; Clench, M. R.; Greenwood, R.; Mills, G.; Carolan, V. Rapid Commun. Mass Spectrom. 1999, 13, 251-255. (23) Ayorinde, F. O.; Eribo, B. E.; Johnson, J. H., Jr.; Elhilo, E. Rapid Commun. Mass Spectrom. 1999, 13, 1124-1128. (24) Ayorinde, F. O.; Elhilo, E. Rapid Commun. Mass Spectrom. 1999, 13, 2166-2173. (25) Ayorinde, F. O.; Hambright, P.; Porter, T. N.; Keith, Q. L., Jr. Rapid Commun. Mass Spectrom. 1999, 13, 2474-2479. (26) Sato, H.; Shibata, A.; Wang, Y.; Yoshikawa, H.; Tamura, H. Polym. Degrad. Stab. 2001, 74, 69-75. (27) Nishio, E.; Ichiki, Y.; Tamura, H.; Morita, S.; Watanabe, K.; Yoshikawa, H. Biosci., Biotechnol., Biochem., in press. (28) Laine, O.; O ¨ sterholm, H.; Ja¨rvinen, H.; Wickstro¨m, K.; Vainiotalo, P. Rapid Commun. Mass Spectrom. 2000, 14, 482-495. (29) Schnolzer, M.; Jedrzejewski, P.; Lehmann, W. D. Electrophoresis 1996, 17, 945-953. (30) Yao, X.; Freas, A.; Ramirez, J.; Demirev, P. A.; Fenselau, C. Anal. Chem. 2001, 73, 2836-2842. (31) Bender, M. L. Chem. ReV. 1960, 60, 53. (32) Hori, K.; Hashitani, Y.; Kaku, Y.; Ohkubo, K. J. Mol. Struct. 1999, 461-462, 589-596. (33) Kawai, F.; Kimura, T.; Tani, Y.; Yamanaka, H.; Ueno, T.; Fukami, H. Agric. Biol. Chem. 1983, 47, 1669-1671. (34) Wang, X.; Weiner, H. Biochemistry 1995, 34, 237-243. (35) Farres, J.; Wang, T. T. Y.; Cunningham, S. J.; Weiner, H. Biochemistry 1995, 34, 2592-2598. (36) Vedadi, M.; Szittner, R.; Smillie, L.; Meighen, E. Biochemistry 1995, 34, 16725-16732. (37) Ahvazi, B.; Coulombe, R.; Delarge, M.; Vedadi, M.; Zhang, L.; Meighen, E.; Vrielink, A. Biochem. J. 2000, 349, 853-861. (38) Kawai, F. FEMS Microbiol. Lett. 1985, 30, 273-276.

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