Environ. Sci. Technol. 1997, 31, 2679-2685
Detection of Poly(Ethylene Glycols) and Related Acidic Forms in Environmental Waters By Liquid Chromatography/Electrospray/Mass Spectrometry CARLO CRESCENZI,† A N T O N I O D I C O R C I A , * ,† ANTONIO MARCOMINI,‡ AND ROBERTO SAMPERI† Dipartimento di Chimica, Universita` “La Sapienza”, Piazza Aldo Moro 5, 00185 Roma, Italy, and Dipartimento di Scienze Ambientali, Universita` di Venezia, Calle Larga S. Marta 2137, I-30123, Venezia, Italy
Poly(ethylene glycols) (PEGs) and related mono- and dicarboxylated forms constitute one of the most abundant classes of contaminants in natural waters. Because of the lack of efficient analytical methods, only one study has been devoted to estimate levels of these analytes in some river waters. The present method involves analyte extraction from 50 mL of raw sewage, 100 mL of treated sewage, 500 mL of river water and seawater, and 2000 mL of groundwater by a 0.5-g Carbograph 4 cartridge. Isolation of acidic analytes from co-extracted neutral ones was accomplished by differential elution. During removal of the HCl-acidified CH2Cl2/CH3OH eluent mixture, carboxylated PEGs were purposely allowed to convert into their methyl esters. This offered the advantage of analyzing for all the species of interest using the same instrumental arrangement. Extracts were analyzed by liquid chromatography/ electrospray/mass spectrometry operated in the positive-ion mode. For species with ethoxy chain lengths g4, analyte recoveries were better than 83%, while lower homologues were poorly recovered, and thus, they were not considered when analyzing environmental aqueous samples. Under full-scan conditions, analytes in real water samples could be quantified at few tens of nanograms per liter, while data acquisition in the selected ion monitoring mode afforded limits of quantification of 0.1-0.3 ng/L. Analyses of influents and effluents of activated sludge wastewater treatment facilities showed that even high molecular mass PEGs, so far considered rather resistant to biodegradation, were very efficiently removed from sewages. The ubiquitous nature of PEG-type compounds was evidenced by the fact that they were detected at parts per trillion levels in marine waters 16 nautical miles from the Italian coast and in five groundwaters samples collected from subsurface depths ranging between 60 and 208 m.
Introduction Poly(ethylene oxides) having molecular masses ranging from less than 200 to more than 100 000 Da are very widely used * To whom correspondence should be addressed. Fax: +39-6490631; e-mail:
[email protected]. † Universita ` “La Sapienza”. ‡ Universita ` di Venezia.
S0013-936X(97)00096-5 CCC: $14.00
1997 American Chemical Society
in a number of human activities. Poly(ethylene oxides) with a chain length from 2 to 22 ethoxy units and an average ethoxy number (nEO) of 9-11 constitute the hydrophilic moiety of aliphatic ethoxylate alcohols (AEs) surfactants. In 1993, ca. 0.7 billion kg of AEs were used in both household and industry and disposed in the environment. AEs are rapidly biodegraded in water. There is abundant evidence (1) that the initial biodegradation of linear alkyl chain AEs occurs by cleavage at the ether bridge between the alkyl group and the polyoxyethylene moiety, following which both poly(ethylene glycols) (PEGs) and aliphatic alcohols are degraded independently. PEG biodegradation proceeds by successive depolymerization of the ethoxy chain via non oxidative and oxidative cleavage of C2 units leading to formation of shorter chain neutral and acidic PEGs (2-7). Generic structures of AEs and related PEG-type intermediates are shown in Figure 1. Depending on the particular aquatic system in which polyethoxylated species are found, their half-lives can vary from weeks to months (8, 9). For the reasons mentioned above, PEGs are considered one of the most abundant classes of contaminants in water. Monitoring of PEGs, monocarboxylated PEGs (MCPEGs), and dicarboxylated PEGs (DCPEGs) in various aquatic systems is of interest for assessing their impact on aquatic life. Moreover, they could be effective markers of potable water contamination from anthropogenic sources. Investigation on the levels of PEG-type compounds in the environment is made very difficult by the lack of simple and effective methods for detecting them at trace levels. Less than 10% of 6 ethoxy unit-containing PEG (PEG 6) can be recovered from water by solvent extraction (10). The highly polar nature of neutral and acidic PEGs as well as the absence of any chromophore group in the structure of these analytes make them not amenable to direct analysis by both gas chromatography and liquid chromatography (LC). Therefore, it is not surprising that only one environmental survey aimed at detecting PEG residues in the Mississippi River and its tributaries by nuclear magnetic resonance (NMR) spectrometry has been conducted (11). Although notable results were obtained, the authors recognized that the large sample size and extensive processing requirements of the quantitative determination obviates use of this method for environmental monitoring of PEG residues in water. Carbograph 4, which is a recently introduced form of graphitized carbon black (GCB), has been successfully applied to solid-phase extraction (SPE) of very polar pesticides from large water volumes (12, 13). Although GCBs are essentially reversed-phase sorbents, the presence of positively charged active centers on their surfaces enable them to behave also as anion exchangers. This feature has been exploited in the past for neutral/acid fractionation of analytes by stepwise desorption (14-17). Polyethoxylated compounds are particularly amenable to analysis by LC/MS equipped with thermospray (TS) or electrospray (ES) ion sources, as they have a tendency to form complexes with protons and inorganic ions. LC/TS/ MS has been used to detect neutral PEGs in wastewaters (18) and AEs (19) in spiked environmental waters. A method based on SPE and LC/ES/MS has been developed by us to detect AEs in various types of aqueous samples (20). The purpose of this work has been twofold. One has been to develop a simple, specific, and very sensitive method for monitoring PEGs and related acidic forms in aqueous environmental samples from various sources. Extraction of analytes and their neutral/acid fractionation was performed with a Carbograph 4 extraction cartridge. Analyte subfractionation and quantitation was made by a benchtop LC/ES/
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FIGURE 1. Structures and acronyms of linear alcohol polyethoxylates (AEO), polyethylene glycols (PEGs), monocarboxylated PEGs (MCPEGs), and dicarboxylated PEGs (DCPEGs). MS instrumentation. The second objective has been to assess levels of the analytes in various aqueous environmental compartments. In this work, PEGs with nEO less than 4 and related acidic forms were not considered, as they were poorly recovered by the extraction apparatus.
Experimental Section Reagents and Chemicals. Individual PEGs 4-6 (purities higher than 95%) and DCPEG 3 (ca. 90% purity) were obtained from Aldrich Chemical Co., Milwaukee, WI, while DCPEG 4 (ca. 90% purity) was obtained from Fluka Buchs, Switzerland. Numbers in acronyms indicating DCPEGs do not refer to the number of ethoxy units effectively present in the molecules (as in the case of acronyms relative to PEGs). Rather they are used to indicate species that can be formed by oxidation of PEG 3 and PEG 4. Such expressions will be used throughout this paper. PEG 7, 8, and 11 (purities higher than 90%) were kindly supplied by B. Masci, who synthesized them as previously reported (21). PEG 400 and DCPEG 600 (numbers indicate average molecular weights) were obtained from Aldrich. Both mixtures were of unknown purities. After their standardization (see below), stock solutions of the PEG 400 and DCPEG 600 mixtures were separately prepared by dissolving 1 g of each mixture in 100 mL of acetonitrile. Working standard solutions of both mixtures were prepared by further diluting stock solutions with acetonitrile. These solutions were used for recovery studies and to quantify analytes in environmental samples. For LC analysis, distilled water was further purified by passing it through a Milli-Q Plus apparatus (Millipore, Bedford, MA). Methanol “Plus” of gradient grade, was obtained from Carlo Erba, Milano, Italy. To eliminate traces of inorganic cations, the organic solvent was distilled in a glass apparatus. Other solvents were of analytical grade (Carlo Erba), and they were used as supplied. Apparatus. Extraction cartridges were prepared by gently packing 0.5 g of Carbograph 4 (surface area, 210 m2/g; 120400 mesh size, Carbochimica Romana, Rome, Italy) in syringelike glass tubes (Baker, Milan, Italy). Frits above and below the sorbent bed were made of Teflon (Baker). Conventional polypropylene tubes and polyethylene frits could not be used as they released small variable amounts of low molecular weight PEG impurities that affected analysis at nanogram per liter levels of these analytes. These artifacts could not be eliminated even after repeated washing with organic solvents. The Carbograph 4 cartridge was attached to a side-arm filtration flask, and liquids were forced to pass through the cartridge by vacuum (water pump). Before processing water samples, the cartridge was washed with 10 mL of the eluent phase for acidic analytes (see below), followed by 2 mL of methanol and 10 mL of HCl-acidified water (pH 2). Samples. The 24-h composite samples of raw and treated sewages were obtained by using flow proportional samplers. These samples were from various mechanical-biological
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sewage treatment plants (STPs) that receive mainly domestic wastes and are located in the area of Rome. Samples of surface water were collected from the river Tiber. Groundwater samples were obtained from wells near Rome. These aquifers are used for drinking water and are situated in areas where no sewage treatment facilities are operative and domestic wastewaters are discharged through cesspools. Representative samples were obtained after purging 5-10 well volumes to eliminate stagnant water. Seawater samples were collected at 2 m depth, following the mixing zone between seawater and Tiber water. All samples were collected in glass bottles and preserved by addition of HgCl2 (20 mg/L). Formaldehyde could not be used as preservative because it can react with glycols to form cyclic acetals. All samples were stored at 4 °C until analysis. Sample Preparation. Analytes were extracted from 50 mL of raw sewages, 100 mL of treated sewages, 500 mL of river waters and seawaters, and 2000 mL of groundwaters. Before processing samples rich in suspended particulate matter, such as river waters and raw and treated sewages, particles were removed by filtration with a 1.5-µm pore size Whatman GF/C glass fiber pad (Maidstone, U.K.). Besides plugging the extraction cartridge, the presence of relevant amounts of particulate matter in the aqueous sample affected recovery of PEGs and MCPEGs. This effect is hard to explain considering that no appreciable amounts of PEGs and MCPEGs were detected in methanolic extracts of particles isolated from the aqueous matrix by filtration. The extraction step was carried out as previously reported (12). Neutral PEGs were eluted by passing 1.5 mL of methanol followed by 8 mL of CH2Cl2/CH3OH (80:20, v/v) through the cartridge, at a flow rate of about 5 mL/min. The eluate was collected in a 1.4 cm i.d. glass vial with a conical bottom. The last drops of this solvent mixture were collected by further decreasing the pressure inside the vacuum flask. Carboxylated PEGs were removed from the sorbent bed and collected in a second vial by elution with 8 mL of CH2Cl2/CH3OH (80:20, v/v) acidified with HCl, 20 mmol/L. During the solvent removal step, these compounds were purposely allowed to convert into their methyl esters. With seawater samples, the extraction procedure described above was modified so that water was acidified to pH 3 in advance, and following the passage of the sample through the cartridge, the cartridge was washed with 40 mL of distilled water. The reason for these modifications will be discussed later. Solvent evaporation was carried out in a water bath at 40 °C, under a gentle stream of nitrogen. When both neutral and acidic extracts reached volumes of about 200 µL, the walls of the vials were washed respectively with 100 µL of methanol and 100 µL of HCl-acidified methanol (20 mmol/ L). The extract containing neutral PEGs was concentrated to ca. 50 µL, while that containing acidic PEGs was taken to dryness. After adding 50 µL of a water/methanol solution (80:20, v/v) and measuring the final volume of the neutral extract, 20% of the extract was injected into the LC column. The residue containing methyl esters of the acidic PEGs was reconstituted with 100 µL of a water/methanol (80/20, v/v) mixture, and 20 µL was injected into the LC column. When extracts were found to contain analyte concentrations sufficient to saturate the ES/MS detector, smaller volumes of the extract were re-injected. LC/ES/MS Analysis. Liquid chromatography was carried out with a Varian (Walnut Creek, CA) Model 9010 equipped with a Rheodyne Model 7125 injector having a 25-µL loop. The analytes were chromatographed on an Alltima 25 cm × 4.6 mm i.d. column filled with 5-µm C-18 reversed-phase packing (Alltech). For fractionating both PEGs and methyl esters of acidic PEGs, phase A was glass-distilled methanol, and phase B was water. Both solvents contained recrystallized NaCl at 10 µmol/L. The mobile-phase composition was
initially 20% A that was increased linearly to 60% A in 25 min. Gases in both solvents were removed by sparging with helium. The flow rate of the mobile phase was 1 mL/min, and 40 µL/min of the column effluent was diverted to the ES source. A Fisons VG Platform benchtop mass spectrometer (Fisons Instruments/VG BioTech, Milan, Italy) consisting of a pneumatically-assisted ES interface and a single quadrupole was used for detecting and quantifying target compounds in the LC column effluent. This was introduced into the ES interface through a 40 cm length of 75 µm diameter PEEK capillary tube. The MS was operated in the positive-ion mode by applying to the capillary a voltage of 4 kV. The source temperature was maintained at 70 °C. Full-scan LC/MS chromatograms were obtained by scanning the quadrupole from 200 to 900 m/z with a 4-s scan, after setting the skimmer cone voltage at 20 V. To detect PEGs 4-21, MCPEGs 4-13, and DCPEGs 4-13 in groundwater extracts, the ES/MS system was operated in the time-scheduled selected-ion monitoring (SIM) acquisition mode with six retention time windows. Each window contained 3-4 acquisition channels relative to MNa+ adduct ions of the selected analytes. Quantitation. To quantify both neutral and acidic PEGs in spiked and unspiked aqueous samples, the external standard quantification procedure was followed. Standard solutions were prepared at 10 levels using each of two working standard solutions, one containing neutral PEGs and the other containing acidic ones, in the respective solvent mixtures used to elute the two classes of analytes from the extraction cartridge. Thereafter, the rest of the procedure described above was followed. For each analyte, calibration curves were constructed by plotting peak areas from selected extractedion-current profiles against the amounts injected from the standard solutions into the LC column. The response of the ES/MS detector was linearly related to injected amounts of each neutral and acidic PEG homolog up to 60-80 ng. Saturation of the ES/MS detector occurred for amounts larger than about 600-800 ng. For any analyte, recovery was assessed by measuring the peak area produced on analyzing spiked samples and subtracting the matrix blanks. A neutral PEG-containing standard solution was prepared and analyzed each working day to check the sensitivity of the ES/MS detector. When following cleaning procedures reported elsewhere (12), day-to-day variations of the mass detector response were no larger than 10%.
Results and Discussion Optimization of Instrumental Conditions. Although 10 µmol/L trifluoroacetic acid (TFA) was added to the LC mobile phase consisting of deionized water and distilled methanol, resulting spectra of both neutral and acidic PEGs displayed, besides protonated adduct ions, intense signals for adduct ions with Na+, NH4+, and K+ ions. This effect can be attributed to the flexible structure of polyethoxylated species, making them particularly able to form stable complexes with inorganic cations (20). To simplify spectra interpretation, we replaced TFA with 10 µmol/L NaCl. This offered the additional advantage of enhancing the ion signal intensities of analytes by about 60%. Although the upper limit of the mass scan range was set at 900 m/z, high molecular weight PEGs could be analyzed with the ES/MS system. This is due to the known ability of the ES process to generate multiply charged adduct ions from large molecules. In particular, we observed that PEG homologs were able to pick up an additional sodium ion for every increment of 12 ethoxy units, starting from PEG 12. As an example, the spectrum for PEG 36 (1646 Da) displayed a base peak at m/z 572, resulting from formation of the [M + 3Na]3+ adduct ion. The decision to add HCl to the CH2Cl2/CH3OH solution to displace acidic PEGs from the anion-exchange sites on the Carbograph surface instead of other previously proposed
FIGURE 2. Time course of concentrations of PEG 4, PEG 11, and related intermediates from a biodegradation test. displacers (16, 17) was not casual. This condition led to conversion of mono- and dicarboxylic acids to the corresponding methyl esters during the solvent removal step. Esterification yields of MCPEGs and DCPEGs were 92 and 86%, respectively, as compared to those obtained by following the classical procedure involving diazomethane as derivatizing agent (22). LC/ES/MS analysis of carboxylated PEGs as methyl esters offered two important advantages. One is that, with the ES/MS system set in the positive-ion mode, esterified acidic PEGs exhibited much higher molar responses than those of intact acidic PEGs. This contrasts with analyses conducted under chromatographic conditions reported elsewhere (23) with the ES/MS system operated in the negativeion mode. The second advantage is that analysis of the acidic fraction could be performed by using the same instrumental conditions as those used for analyzing neutral PEGs. PEG 400 and DCPEG 600 Calibration. The procedure of calibration of the two mixtures was made difficult because the molar response of the ES/MS detector is strictly dependent on the structure of the analyte. The percentage weight distribution of the various oligomers in the PEG 400 commercial mixture was determined by preparing a water/ methanol (90:10, v/v) solution containing 5 µmol/L of each pure individual PEG. This solution was injected three times into the LC column. The molar responses of each pure PEG were calculated by measuring its peak area and dividing by the number of moles injected. These response factors were then plotted against nEO. As in the case of AEs (20), the resulting graph showed that molar responses of PEGs increased steadily from nEO 4 to nEO 8, whereas no significant change was observed passing from PEG 8 to PEG 11. This behavior results from the fact that the flexibility of the molecular structure of short-chain polyethoxylates increase as nEO increases. This enables them to form increasingly stable complexes with cations. By assigning to PEGs with nEC > 8 the same molar response factor as that of PEG 8, the percentage weights of any individual PEGs in the PEG 400
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TABLE 1. Concentrations of Neutral and Carboxylated PEGs in Influents and Effluents of Four Sewage Treatment Plants (STPs) concentration, µg/L STP 1
PEG 4 PEG 5 PEG 6 PEG > 6 MCPEG 4 MCPEG 5 MCPEG 6 MCPEG > 6 DCPEG 4 DCPEG 5 DCPEG 6 DCPEG > 6 a
STP 2
STP 3
STP 4
influent
effluent
influent
effluent
influent
effluent
influent
effluent
4.4 4.2 3.7 140 2.7 1.8 1.8 16 0.72 1.5 1.5 6
2.1 (0.48)a 1.8 (0.42) 1.0 (0.27) 6 (0.04) 0.6 (0.22) 0.5 (0.27) 0.6 (0.33) 6 (0.38) 0.32 (0.44) 0.82 (0.54) 0.69 (0.46) 4 (0.7)
1.8 1.7 4.7 210 7.8 7.0 3.2 42 7.3 11 10 72
0.90 (0.50) 0.73 (0.43) 0.65 (0.14) 4 (0.02) 0.4 (0.05) 0.3 (0.04) 0.2 (0.06) 2 (0.05) 0.15 (0.02) 0.15 (0.01) 0.18 (0.02) 0.3 (0.01)
14 13 11 230 4.2 2.9 2.5 20 0.9 1.8 1.8 13
1.1 (0.01) 0.6 (0.05) 0.4 (0.04) 4 (0.02) 0.24 (0.06) 0.13 (0.03) 0.07 (0.03) 1 (0.05) 0.1 (0.11) 0.2 (0.11) 0.3 (0.17) 0.3 (0.02)
7.3 8.7 7.9 62 9.1 8.3 7.7 60 5.5 9.2 8.8 67
0.31 (0.04) 0.22 (0.03) 0.14 (0.02) 3 (0.05) 0.1 (0.01) 0.06 (0.01) 0.05 (0.01) 0.3 (0.01) 0.03 (0.01) 0.03 (
