Analysis of perfluorinated carboxylic acids in soils II: Optimization of chromatography and extraction

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Journal of Chromatography A, 1181 (2008) 21–32

Analysis of perfluorinated carboxylic acids in soils II: Optimization of chromatography and extraction夽 John W. Washington a,∗ , W. Matthew Henderson a , J. Jackson Ellington a , Thomas M. Jenkins b , John J. Evans b a

United States Environmental Protection Agency, National Exposure Research Laboratory, 960 College Station Road, Athens, GA 30605, USA b Senior Service America Inc., United States Environmental Protection Agency, National Exposure Research Laboratory, 960 College Station Road, Athens, GA 30605, USA Received 4 September 2007; received in revised form 28 November 2007; accepted 6 December 2007 Available online 23 December 2007

Abstract With the objective of detecting and quantitating low concentrations of perfluorinated carboxylic acids (PFCAs), including perfluorooctanoic acid (PFOA), in soils, we compared the analytical suitability of liquid chromatography columns containing three different stationary phases, two different liquid chromatography–tandem mass spectrometry (LC/MS/MS) systems, and eight combinations of sample-extract pretreatments, extractions and cleanups on three test soils. For the columns and systems we tested, we achieved the greatest analytical sensitivity for PFCAs using a column with a C18 stationary phase in a Waters LC/MS/MS. In this system we achieved an instrument detection limit for PFOA of 270 ag/␮L, equating to about 14 fg of PFOA on-column. While an elementary acetonitrile/water extraction of soils recovers PFCAs effectively, natural soil organic matter also dissolved in the extracts commonly imparts significant noise that appears as broad, multi-nodal, asymmetric peaks that coelute with several PFCAs. The intensity and elution profile of this noise is highly variable among soils and it challenges detection of low concentrations of PFCAs by decreasing the signal-to-noise contrast. In an effort to decrease this background noise, we investigated several methods of pretreatment, extraction and cleanup, in a variety of combinations, that used alkaline and unbuffered water, acetonitrile, tetrabutylammonium hydrogen sulfate, methyl-tert-butyl ether, dispersed activated carbon and solid-phase extraction. For the combined objectives of complete recovery and minimization of background noise, we have chosen: (1) alkaline pretreatment; (2) extraction with acetonitrile/water; (3) evaporation to dryness; (4) reconstitution with tetrabutylammonium–hydrogen–sulfate ion-pairing solution; (5) ion-pair extraction to methyl-tert-butyl ether; (6) evaporation to dryness; (7) reconstitution with 60/40 acetonitrile/water (v/v); and (8) analysis by LC/MS/MS. Using this method, we detected in all three of our test soils, endogenous concentrations of all of our PFCA analytes, C6 through C10 —the lowest concentrations being roughly 30 pg/g of dry soil for perfluorinated hexanoic and decanoic acids in an agricultural soil. © 2007 Elsevier B.V. All rights reserved. Keywords: Perfluorinated octanoic acid; PFOA; LC/MS/MS; Soil extracts; Extract cleanup; TBA

1. Introduction Recent research has documented the widespread distribution, on the global scale, of perfluorooctane sulfonate (PFOS; [1]) and perfluorinated carboxylic acids (PFCAs), including perflu夽 This paper has been reviewed in accordance with the U.S. Environmental Protection Agency’s peer and administrative review policies and approved for publication. Mention of trade names or commercial products does not constitute endorsement or recommendation for use. ∗ Corresponding author. Tel.: +1 706 355 8227; fax: +1 706 355 8202. E-mail address: [email protected] (J.W. Washington).

0021-9673/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2007.12.042

orinated octanoic acid (PFOA; [2–4]), in ecosystems, biota and humans. Inspired to investigation by these findings, toxicologists have identified numerous biological effects for these compounds in mammalian systems including increased liver-to-body-weight ratio, lag of weight gain of newborns, increased mortality of weaned offspring, and alteration of gene expression related to lipid metabolism, immunity and hormone regulation [5–8]. Given these and similar reports, the United States Environmental Protection Agency (EPA), Office of Pollution Prevention and Toxics (OPPT) is keenly interested in defining better the sources, distribution and effects of perfluorinated compounds (PFCs) in the environment. Toward this end, the EPA, Office of

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Research & Development (ORD) has undertaken a broad spectrum of research programs on perfluorinated compounds in its national laboratories, the National Risk Management Research Laboratory (NRMRL), the National Health & Environmental Effects Research Laboratory (NHEERL) and the National Exposure Research Laboratory (NERL). At the NERL laboratory in Athens, GA we have embarked on studies of these compounds in soils including: (1) the possibility that fluorotelomer-based polymer products might degrade to act as a source of PFCs to the environment; and (2) the distribution of these compounds in soils. Success in these research studies requires unambiguous, confident detection and quantitation of trace levels of PFCs in soils. These objectives are challenged by the complex nature of soils, which offer a suite of organic and inorganic surfaces, and organic and inorganic solutes [9] that might affect the fate of PFOA. For example, natural organic matter (NOM) in soils commonly comprises from about 0.05% to 10% of the dry mass. Whereas the net charge of NOM generally is negative, and typically ranges as high as about 500 milliequivalents per 100 g (mequiv./100 g)[10], this net charge reflects the contributions of a variety of functional groups having range of charges, both positive and negative. In the mildly acidic pH ranges commonly found in soils, PFOA usually dissociates to its monovalent anion, perfluorooctanoate (PFO− ), and, consequently, the complexly charged nature of NOM offers a rich potential for electrostatic binding of PFO− , as well as van der Waals interactions with the perfluorinated alkyl chain. As for minerals, oxy-hydroxide minerals of Fe(III) and other metals, and layer-silicate clays generally provide the dominant part of the electrostatically charged mineral surfaces in soils. The surface charges of the metal oxy-hydroxides are pH-dependent. In the case of ferric oxy-hydroxides, such as ferrihydrite, goethite and hematite, the zero-point of charge (ZPC) commonly is about 7.5 [11]; since the pH of many soils falls below this ZPC, ferric oxy-hydroxides commonly are positively charged. As such, the ferric oxy-hydroxides potentially can directly scavenge PFO− by electrostatic sorption. In contrast, the surfaces of the layer-silicate clays mostly are permanently negatively charged, and have cation-exchange capacities ranging to about 140 mequiv./100 g [12]. While these negatively charged clay surfaces are unlikely to sorb the dissociated anion, PFO− , directly to any great extent, divalent cations such as Ca2+ , Mg2+ and Fe2+ commonly are present in soil solutions at moderately

high concentrations, e.g., in the mM range. To the extent that these cations might form complexes with PFO− , the resultant monovalent complexes, e.g., (PFO-Ca)+ , potentially can sorb to the clay surfaces. In addition to the complexities that soils offer with regard to the partitioning and fate of PFOA, they also impose analytical challenges beyond those of many biological materials. Specifically, the bulk chemistry of living tissues, e.g., liver or plasma, generally is identical among individuals within a species and, consequently, the matrix interference often exhibits relatively little variability between extracts within a single tissue for a single species [13]. In contrast, the matrices of non-living environmental extracts such as soils, sediments, sludges or waters, contain varying proportions of organic and inorganic molecules from a variety of sources, each of which have been subject to unique degrees of decay and transformation. As such, the appearance of the matrices in these environmental extracts, as well as their tendency to enhance or suppress analytical signal, is unique for each sample [14]. Spurred by these factors, in this paper we report on our efforts to optimize analyses by comparison of chromatographic columns, and techniques for cleanup of soil extracts of an agricultural soil, a soil unmodified by recent direct human manipulation and a commercially distributed topsoil. 2. Experimental 2.1. Chemicals Except as noted below, all chemicals used in this study were of the highest purity offered by the suppliers, uniformly ≥97% purity. Perfluoro-n-heptanoic acid (CAS Number 375-85-9) was purchased from Oakwood Products Inc. (West Columbia, SC, USA). Perfluoro-n-hexanoic acid (CAS Number 307-24-2), perfluoro-n-octanoic acid (CAS Number 335-67-1), perfluoro-n-nonanoic acid (CAS Number 37595-1), perfluoro-n-decanoic acid (CAS Number 335-76-2), perfluoro-n-[1,2,3,4-13 C]octanoic acid, perfluoro-n-[1,2,3,4,513 C]nonanoic acid, perfluoro-n-[1,2-13 C]decanoic acid all were purchased as certified standards from Wellington Laboratories through TerraChem (Shawnee Mission, KS, USA). Formulae for these PFCAs, and the acronyms used herein for these compounds, are summarized in Table 1. Tetrabutylammonium hydrogen sulfate (TBAHS CAS Number 32503-27-8) and

Table 1 Analyzed compounds and selected reaction monitoring transitions Compound

Formula

Acronym (short formula)

Quantitation transition (m/z)

Confirmation transition (m/z)

Perfluoro-n-hexanoic acid Perfluoro-n-heptanoic acid Perfluoro-n-octanoic acid Perfluoro-n-nonanoic acid Perfluoro-n-decanoic acid Perfluoro-n-[1,2,3,4-13 C]butanoic acid Perfluoro-n-[1,2,3,4-13 C]octanoic acid Perfluoro-n-[1,2,3,4,5-13 C]nonanoic acid Perfluoro-n-[1,2-13 C]decanoic acid

CF3 (CF2 )4 COOH CF3 (CF2 )5 COOH CF3 (CF2 )6 COOH CF3 (CF2 )7 COOH CF3 (CF2 )8 COOH 13 CF (13 CF ) 13 COOH 3 2 2 13 CF (13 CF ) (CF ) COOH 3 2 3 2 3 13 CF (13 CF ) (CF ) COOH 3 2 4 2 3 13 CF (13 CF )(CF ) COOH 3 2 2 7

PFXA (C6 ) PFHA (C7 ) PFOA (C8 ) PFNA (C9 ) PFDA (C10 ) [13 C4 ]PFBA [13 C4 ]PFOA [13 C5 ]PFNA [13 C2 ]PFDA

313 → 269 363 → 319 413 → 369 463 → 419 513 → 469 217 → 172 417 → 372 468 → 423 515 → 470

313 → 119 363 → 169 413 → 169 463 → 169 513 → 169

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Table 2 Characterization properties of tested soils Soil name

P-1 Picnic Cowart a b c

Dry-mass % of <2 mm fraction Sand

Silt

Clay

69 58 58

19 20 32

11 22 10

%OMa (dry-mass basis)

CECb (mequiv./100 g)

Soil pHc

3.0 0.66 8.7

8.9 11.3 8.2

6.2 4.8 5.9

Percent organic matter, by loss of dry-mass on ignition. Cation-exchange capacity, milliequivalents per 100 g dry soil. Soil pH, 1:1 dry-soil:water on a mass basis.

sodium carbonate (CAS Number 497-19-8), were purchased from Aldrich Chemical (Milwaukee, WI, USA). Acetonitrile (ACN; CAS Number 75-05-8), glacial acetic acid (CAS Number 64-19-7), methanol (MeOH; CAS Number 67-56-1), methylene chloride (MeCl2 ; CAS Number 75-09-2) and methyl-tert-butyl ether (MTBE; CAS Number 1634-04-4) all were purchased from Fisher Chemical (Fairlawn, NJ, USA). Supelco Envicarb 120/400 activated carbon (CAS Number 1333-86-4) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Ammonium acetate (CAS Number 631-61-8) was purchased from J.T. Baker (Phillipsburg, NJ, USA). Oasis HLB solid-phase extraction (SPE) cartridges, 35-cm3 and 3-cm3 capacity were purchased from Waters (Milford, MA, USA). Formic acid (ACS grade; CAS Number 64-18-6) was purchased from Aldrich. Argon gas was ≥99.999% purity supplied by Airgas (Radnor, PA, USA). 2.2. Soils studied We performed the exploratory evaluations described herein with three soils, an agricultural sandy-loam soil (P-1), a sandyclay-loam soil from a wooded picnic area (Picnic), and a commercial sandy-loam topsoil (Cowart). Conventional characterization properties for these soils are tabulated in Table 2 and these soils are described more extensively in our earlier papers [9,14]. We did not spike any of these soils with any unlabeled PFCA. 2.3. Analytical methods Most analyses reported herein were performed on a Varian 1200L tandem mass spectrometer (MS/MS) interfaced with a Varian 430 autosampler, a Varian 230 high-performance liquid chromatograph (HPLC) and an Alltech 631 column heater. All system operations were controlled by Varian MS Workstation version 6. To conserve sample extracts while maintaining sensitivity, 25 ␮L of extract were introduced using ␮L-pick-up mode to guard/analytical columns as described in Section 2.5. Eluent flow rates were set as described in Section 2.5. Column temperature was maintained at 30 ◦ C. To minimize background for PFOA on the HPLC, we bypassed the solvent degassers which commonly are composed of fluorinated polymers that may contain trace levels of some perfluorinated carboxylic acids. Upon elution from the HPLC, sample extracts were introduced to the mass spectrometer in negative electrospray-

ionization mode (ESI(−)) with the needle potential set at −3950 V. The atmospheric-pressure-ionization (API) drying gas, from a Peak Scientific NM30LA N2 generator (99.9% purity), was maintained at 1.3 kTorr (25 psi) and 150 ◦ C. Capillary potential was set to −5 eV and the shield offset was −600 V. The collision gas was Ar and collision-cell pressure was 2.7 mTorr (5.2 × 10−5 psi). Collision energy for the quantification ions was 8 eV for the C6 acid and 10 eV for the C7 through C10 acids. Collision energy for the confirmation ions was set to 20 eV for C6 , C9 and mass-labeled C10 ; 16 eV for C7 ; 15 eV for C8 ; and 18 eV for mass-labeled C8 . The detector potential was 1800 V and it was run in selected-reaction-monitoring (SRM) mode. Quantitation and confirmation transitions were monitored for each of the PFCA homologues (Table 1). Scan time was set to 3.5 s, so that 14 monitored transitions each received about 17 scans over a typical peak-elution time of about 1 min, with a dwell time for each transition of over 200 ms. Complex extract matrices commonly impose ionization suppression or enhancement in LC/MS/MS analyses and these effects commonly are accounted for through addition of constant final concentrations of one or more matrix internal standards to the extracts immediately prior to analysis. By normalizing analyte signals to those of the matrix internal standards, fluctuations in instrument performance and matrix effects on ionization efficiency can be compensated for. In our recent, associated paper, one of our major findings was that the variably complex matrices of soil extracts, that have not been subjected to cleanup procedures, renders extracts unsuitable for quantitative analysis using matrix internal standards except when the analyte and matrix internal standard coelute [14]. Using the same three soils that we report upon herein, we found that concentrations of PFNA and PFDA that we calculated from analytical data using [13 C4 ]PFOA as a matrix internal standard differed from the values that we calculated using the method of standard additions by as much as those values calculated using external calibrations. In this effort, we took the values generated from the method of standard additions as being correct, so this observation indicated that using [13 C4 ]PFOA as a matrix internal standard did not increase accuracy of efforts at quantitation for non-coeluting compounds. Nevertheless, because our purpose in the present paper is to evaluate efficacy of a variety of matrix cleanup methods, and not to determine the quantitative PFCA concentrations of these soils, on the Varian instrument we generally calculate values herein as we have reported in the past

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[14] using [13 C4 ]PFOA as a matrix internal standard, except as noted. Selected extracts also were analyzed on a Waters Quattro Premier XE tandem mass spectrometer interfaced with a Waters Acquity ultra-performance liquid chromatograph (UPLC). All system operations were controlled by Waters MassLynx 4.1 and QuanLynx 4.1. To conserve sample extracts while maintaining sensitivity, 20 ␮L was withdrawn from extracts maintained in the autosampler at 4 ◦ C and introduced into a 50 ␮L loop using “partial loop with needle overfill” mode to a Waters BEH C18 trapping cartridge followed by a Waters BEH C18 analytical column, 100 × 2.1 × 2.1 (mm length × mm inside diameter × ␮m particle size), kept at 35 ◦ C. To minimize background for PFOA, we altered the UPLC plumbing by (1) substituting in polyetheretherketone (PEEK) tubing to carry the solvents; (2) by-passing the solvent degassers which are composed of fluorinated polymers and which we observed to impart significant noise to the PFOA transition; (3) inserting a C18 trap column (100 × 2.1 × 3.5) at the downgradient-most point in the water-eluent line, immediately above the solvent-mixing cell; and (4) injecting ∼1000 blanks and sample extracts, operating 24/7 to cleanse the system from its as-delivered state. In addition, we polished our doubly deionized eluent water by passing it through Waters 35-cm3 Oasis HLB extraction cartridges, then degassing manually by mild heating under vacuum and stirring with a glass stir bar, all in dedicated glassware. Regarding the C18 trap column that we installed at the down-gradient-most point in the water-eluent line, this design scavenges any PFCAs that might be in the water rather than merely delaying their elution because PFCAs do not elute from C18 columns in pure water and, instead, are displaced from the solid phase only at about 60% ACN and greater. Given this function, the trap column requires cleaning periodically which we do with 70/30, volume-to-volume (v/v), ACN/water. The UPLC was operated using ACN and water eluents, both containing 0.075% (v/v) glacial acetic acid. Pumping at a constant total flow rate of 0.5 mL/min, we started runs with an eluent of 65/35 (v/v) ACN/water, then linearly ramping to 90/10 at 5 min, holding composition constant until 11 min, linearly ramping down to 65/35 at 11.1 min, from which time we held composition constant until the end of analysis at 13 min. Upon elution from the UPLC, extracts were introduced to the mass spectrometer in ESI(−) mode with the capillary potential set at −600 V, the method-default cone potential at −17 V, the extractor potential at −2 V and the radio-frequency (rf) lens potential at 0.3 V. The cone potential was altered to −14 V for C6 and −15 V for C7 , C8 and [13 C4 ]PFOA. The source temperature was maintained at 140 ◦ C. The desolvation gas, from the Peak N2 generator, was maintained at 140 ◦ C and flowed at 800 L/h. The cone gas, also supplied by the N2 generator, was set to flow at 25 L/h. The low- and high-mass resolutions in the first quadrupole both were set to 13.0 (unitless ratio of direct to rf current voltages) and the ion energy was set to 0.7 eV. In the collision cell, the entrance was set to −3 V, the interior set to −16 V and the exit set to −1 V. The Ar collision gas was set to flow at 0.45 mL/m.

The low- and high-mass resolutions in the third quadrupole both were set to 12.0 and the ion energy was set to 1.0 eV. The detector multiplier was set to −700 V. Collision energy for the quantification ions was 13 eV for the C6 and C7 acids, and 15 eV for the C8 through C10 acids. Collision energy for the confirmation ions was set to 24 eV for C6 and 22 eV for C7 through C10 . To maximize dwell time, our Waters analytical method called for monitoring each of the homologues, C6 through C10 , as independent functions; two transitions, the quantitation-ion and the confirmation-ion, on each of the C6 , C7 and C10 functions, and three transitions, the quantitation-ion, confirmation-ion and mass-labeled analyte, on C8 and C9 . Setting dwell time to 150 ms for the two-transition channels and 100 ms for the threetransition channels, we achieved 20–30 scans per peak over typical peak-elution times of 0.2 m. Performed during methods development for the instrument, the chromatographic profile for our determination of the instrument detection limit (IDL) on the Waters instrument varied from the above conditions which we used with soil extracts to assure all matrix components were purged from the system within analytical runs. For determination of the IDL using blanks and low standards, flow rate was set to 0.5 mL/min, we started runs with an eluent of 60/40 (v/v) ACN/water, held constant for 1.5 min, then linearly ramped to 70/30 at 5 min, then linearly ramped to 90/10 at 5.1 min, holding composition constant until 6.1 min, linearly ramping down to 60/40 at 8 min. We also used a simpler monitoring scheme in that all transitions were monitored simultaneously with a 70-ms dwell time. SRM chromatograms were smoothed using a second-order Savitsky–Golay algorithm, one five-point smooth for the Varian and two five-point smooths for the Waters, which we determined accentuated the signal without imparting negative effects of peak broadening or deterioration of the separation of closely proximate peaks [15]. Signal peaks were delimited by the valleys immediately bounding a peak having an apex exceeding noise or, should they be present, contiguous peaks exceeding noise so long as linking valleys exceeded the apices defining the general amplitude of surrounding noise peaks as well. We define method detection limits (MDLs) with a statistical comparison of signal strength with noise [14] and, as such, the conservative integrity of these values requires use of the largest noise for which there is reasonable potential it might be confused with analyte signal. For the Varian, the detector sensitivity and background for our analytes was such that the blanks registered no analytical peak. Consequently, sample-extract noise on this instrument is best characterized using noise on the extracts themselves so that the complex matrices that might depress the contrast of signal-to-noise is accounted for [14]. On the other hand, the Waters instrument was sensitive enough to register an analytical peak for PFOA on blanks; despite extensive investigations and efforts including direct infusion on the MS/MS of ACN and H2 O, it remained unclear whether the PFOA was from the eluents or from the instrument. Regardless, on this instrument, noise was defined for MDL determination by quantifying the analytical peak area of the blanks.

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2.4. Calibration standards We performed an informal analytical survey of PFCAs in numerous soils to identify the range over which to calibrate (data not shown). Based on this work, we prepared a calibration for the C6 through C10 PFCAs consisting of nine concentrations spanning 18 pg/g to 11 ng/g. For those curves normalized to a matrix internal standard [16], the internal standard was [13 C4 ]PFOA, routinely spiked at ∼900 pg/g. Calibration standards were analyzed during sample-extract sequences. For the Varian instrument, each standard and extract was injected on column six times; standards were run at the beginning and end of each sample-extract sequence. Except as noted below, calibrations were comprised of linear regressions of untransformed data; regression weighting was determined on an analyte-specific basis with the objective of achieving central tendency for the calibration with respect to the standards, consistent with that described in our earlier paper [14]. For the Waters instrument, each standard was injected on column three times and each soil extract was injected six times; standards were interspersed with extracts and blanks throughout the sample-extract sequences. Calibrations were comprised of linear regressions of untransformed data; regression weighting was determined on an analyte-specific basis using residualanalysis plots supported by the software with the objective of achieving a central tendency for the calibration with respect to the standards as described in our earlier paper [14]. 2.5. Investigation of low-PFCA concentrations on each of three columns In this study, we compared the performance of three analytical-column systems with respect to detecting low concentrations of PFCAs. This effort entailed evaluation of: (1) chromatographic separation of the PFCAs; (2) peak shape, i.e., mono-nodality, symmetry, pre- or post-peak tails; and (3) IDL using PFOA as a representative PFCA. For each column system, we injected PFCA standards ranging from 12.5 to 400 fg/␮L using a variety of binary eluent mixes of ACN, MeOH, and aqueous solutions of acetic acid and ammonium acetate, both isocratically and with gradient programs. From these outputs, we chose the best eluent mix and program for each column based on: (1) peak area of low-concentration standards; (2) peak shape; and (3) chromatographic separation of PFCA peaks. Based on these reconnaissance runs, we chose to examine more closely the following systems: (1) On a Hypersil Gold C18 system consisting of a 10 × 3 × 3 guard column and a 150 × 3 × 3 separation column, we ran isocratically 60/40 volume to volume (v/v) ACN/26 ␮M glacial acetic acid at a flow rate of 0.30 mL/min. (2) On a Wako Fluofix-II 120E system consisting of 150 × 2 × 5 separation column, we ran isocratically 58/42 (v/v) ACN/26 ␮M glacial acetic acid at a flow rate of 0.20 mL/min. We used no guard column for this system while we were injecting standards.

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(3) On a Zirchrom SAX Anion system consisting of a 10 × 3 × 3 guard column and a 150 × 2.1 × 3 separation column, we ran isocratically 60/40 (v/v) ACN/25 mM ammonium acetate with triethylamine buffer to pH 10.5 at a flow rate of 0.20 mL/min. We chose the Hypersil system to represent the C18 columns that are widely used for perfluorinated-compound separation, the Wako Fluofix column has been used for perfluorinated compounds at our sister laboratory in Research Triangle Park, NC with great success [17], and the Zirchrom SAX column is designed for anion separation. To perform this investigation, we chose a standard concentration for which we could identify visibly the analyte peak in most of the runs. This standard was injected 11 times. Using these data, we evaluated the IDL using the definitions and methods described in our earlier work [14]. 2.6. Sample preparation and extraction methods In all of the following pretreatment, extraction, cleanup and concentration procedures, we describe several steps in terms of nominal volumes; however, mass was recorded at every step of all procedures and all calculations are on a mass basis. All methods that we report included a recovery internal standard that we added to the soil sample immediately before we began extraction to function as an indicator of recovery. And all methods included use of a matrix internal standard that we added to the extract as a last step before loading the sample vial in the LC/MS/MS autosampler to account for instrument-performance variability, and matrix suppression and enhancement. Starting with the soils prepared according to our earlier paper [14], ∼1-g aliquots were weighed then dried under vacuum and over Drierite to constant mass in order to calculate moisture content. 2.6.1. Elementary ACN/H2 O extraction This extraction is identical to that which we described in Washington et al. [14]. Reiterating here, for each sample, extractions were performed on three 5-g aliquots in pre-weighed, MeOH-washed, 16-mL polypropylene copolymer (PPCO) centrifuge tubes with size-18 PPCO caps. Each aliquot was extracted according to: (1) add 18-M H2 O to obtain ∼50% moisture on a dry-mass basis, vortex until visually homogenized and reweigh; (2) autopipette 70 ␮L of 83 ng/g [13 C5 ]PFNA in 60:40 (v/v) ACN:H2 O as a recovery internal standard, reweigh; (3) add 4.75 mL of ACN to yield a 60:40 (v/v) solution of ACN:H2 O, vortex until visually homogenized and reweigh; (4) sonicate for 60 min; (5) centrifuge at 36.6 kG and 18–22 ◦ C for 30 min; (6) decant supernatant into a tared autosample vial and weigh; and (7) autopipette 100 ␮L of 15 ng/g [13 C4 ]PFOA in 60:40 (v/v) ACN:H2 O to achieve about 1 ng/g as a matrix internal standard and reweigh. 2.6.2. ACN/H2 O extraction with alkaline pretreatment Based on our experience [14] with the elementary ACN/H2 O extraction, i.e., the method described above in Section 2.6.1, selected reaction monitoring (SRM) chromatograms of soil

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extracts typically include a broad matrix peak that elutes contemporaneously with shorter-chained PFCAs, confounding their detection at low concentrations. Powley et al. [18] reported a procedure for “matrix effect-free” analysis of PFCAs in soils and other environmental matrices that entailed a 30-min alkaline pretreatment of the soil and a 15-min post-extraction cleanup with Envicarb. We investigated the efficacy of these pre- and post-extraction steps by following steps 1 and 2 of the elementary procedure above and then: (3) add 200 ␮L of 2.0 M NaOH, vortex, reweigh and allow to react for 30 min; (4) add 4.75 mL of ACN to yield an ∼60:40 ACN:H2 O (v/v) solution, vortex until visually homogenized and reweigh; (5) neutralize with 200 ␮L of 2.0 M HCl, vortex and reweigh; (6) sonicate for 60 min; (7) centrifuge at 36.6 kG and 18–22 ◦ C for 30 min; (8) decant 1 mL of supernatant into a tared autosample vial and weigh; (9) autopipette 70 ␮L of 15 ng/g [13 C4 ]PFOA in 60:40 ACN:H2 O (v/v) as a matrix internal standard and reweigh. 2.6.3. ACN/H2 O extraction with alkaline pretreatment and 15-min dispersed activated-C cleanup To test the effect of activated C, independently of the alkaline pretreatment described above in Section 2.6.2, we took a second split of the supernatant in step 7 and: (8) decant 1 mL of supernatant into 2-mL microcentrifuge tubes containing ∼25 mg Envicarb, vortex until visually homogenized and let rest for 15 min; (9) centrifuge at 8.6 kG for 10 min; (10) decant supernatant into a tared autosample vial and weigh; and (11) autopipette 70 ␮L of 15 ng/g [13 C4 ]PFOA in 60:40 ACN:H2 O (v/v) as a matrix internal standard and reweigh. 2.6.4. ACN/H2 O extraction with 16-h dispersed activated-C cleanup This procedure is identical to that of Section 2.6.3 above excepting, in step 8, the dispersed activated C is allowed to react with the extractant for 16 h instead of 15 min based on the premise that sorption of interfering, matrix molecules from solvents to the solid phase might be slow enough that added reaction time would benefit cleanup. 2.6.5. ACN/H2 O extraction with ion-pair cleanup Brandstrom [19] described an approach to cleanup complex extract matrices in which analytical ions are paired with aqueous counter ions so that the resulting neutral complex can be exchanged into a water-immiscible organic solvent. A salt commonly used for this purpose is TBAHS which dissociates in water to form the tetrabutylammonium cation (TBA) that pairs with analyte anions. In our first efforts with this approach, our tube blanks had significant levels of PFOA. Looking for the source of this contamination, we analyzed an aqueous solution of TBAHS and found PFOA. As a result, in all ion-pair work we describe below, we polished the TBAHS solutions by passing them through HLB SPE cartridges, to trap the PFCA contamination. For the effort with this approach that we report upon herein, we followed steps 1–5 of our elementary extraction described in

Section 2.6.1 and then (6) decant supernatant into a weighed 12-mL vial and reweigh; (7) autopipette 200 ␮L of 15 ng/g [13 C4 ]PFBA in 60:40 (v/v) ACN:H2 O as a recovery internal standard with respect to the cleanup in case the [13 C5 ]PFNA recovery internal standard with respect to extraction is determined to be lost, reweigh; (8) evaporate to dryness (using the SPE stand, with nylon filters fitted at the needle inlets, using ∼0.78 kTorr (∼15 psi) vacuum – filtering air only, these filters did not touch any liquid), reweigh; (9) add 4 mL of 0.25 M polished TBAHS and 0.125 M Na2 CO3 , vortex until visually homogenized and reweigh; (10) add 5 mL of MTBE, vortex and reweigh; (11) transfer MTBE to a tared vial, reweigh; (12) evaporate to dryness with filtered air, reweigh; (13) reconstitute with 2 mL of 60:40 ACN:H2 O, vortex and reweigh; and (14) autopipette 200 ␮L of 15 ng/g [13 C4 ]PFOA in 60:40 ACN:H2 O (v/v) as a matrix internal standard and reweigh. 2.6.6. ACN/H2 O extraction with alkaline pretreatment and ion-pair cleanup In a second effort with ion-pair cleanup, we followed steps 1–6 of our ACN/H2 O extraction with alkaline pretreatment described in Section 2.6.2 and then: (7) autopipette 200 ␮L of 15 ng/g [13 C4 ]PFBA in 60:40 (v/v) ACN:H2 O as a recovery internal standard with respect to the cleanup in case the [13 C5 ]PFNA recovery internal standard with respect to extraction is determined to be lost, reweigh; (8) evaporate to dryness with filtered air, reweigh; (9) add 4 mL of 0.25 M polished TBAHS and 0.125 M Na2 CO3 , vortex until visually homogenized and reweigh; (10) add 5 mL of MTBE, vortex and reweigh; (11) sonicate for 60 min; (12) transfer MTBE to a tared vial, reweigh; (13) evaporate to dryness with filtered air, reweigh; (14) reconstitute with 2 mL of 60:40 ACN:H2 O, vortex and reweigh; and (15) autopipette 200 ␮L of 15 ng/g [13 C4 ]PFOA in 60:40 ACN:H2 O (v/v) as a matrix internal standard and reweigh. 2.6.7. TBA 60-min extraction with ion-pair cleanup We adapted our TBA, ion-pair extraction from the method employed by Henderson et al. [7] which, in turn, was based on the method of Hansen et al. [20]. Starting with two 5g aliquots of soil of known moisture content in pre-weighed, MeOH-washed, 16-mL PPCO centrifuge tubes and caps, each aliquot was extracted according to: (1) add 0.167 M polished TBA + 0.167 M Na2 CO3 solution to the soil samples to obtain 100% moisture on a dry-mass basis including the soil moisture already present in the soil, vortex until visually homogenized and reweigh; (2) autopipette 100 ␮L of 15 ng/g [13 C5 ]PFNA in 60:40 (v/v) ACN:H2 O as a recovery internal standard, vortex and reweigh; (3) add 3.0 mL of MTBE, vortex and reweigh; (4) sonicate for 2 h on ice, dry and reweigh; (5) centrifuge at 36.6 kG and 18–22 ◦ C for 30 min; (6) transfer MTBE supernatant into a tared vial and reweigh; (7) autopipette 100 ␮L of 15 ng/g [13 C4 ]PFBA in 60:40 ACN:H2 O (v/v) as a recovery internal standard with respect to the cleanup, vortex and reweigh; (8) evaporate to dryness with filtered air; (9) add 2 mL 60:40 ACN:H2 O (v/v), vortex and reweigh; (10) autopipette 100 ␮L of 15 ng/g [13 C4 ]PFOA in

J.W. Washington et al. / J. Chromatogr. A 1181 (2008) 21–32

27

Figs. 1–3. SRM chromatograms on the Hypersil Gold C18 column (Fig. 1), the Wako Fluofix-II 120E column (Fig. 2) and the Zirchrom SAX Anion column (Fig. 3) for an ∼550 fg/g standard of C6 (a), C7 (b), C8 (c), C9 (d) and C10 (e). Note the sharp symmetric peaks and separation among peaks on the Hypersil column as opposed to the poor symmetry we had on the Wako and the contemporaneous elution of the acids on the SAX column. Also note the near-baseline separation of the branched-chain isomer from the straight-chain isomer for C8 on the Hypersil column.

60:40 ACN:H2 O (v/v) to achieve about 1 ng/g as an internal standard, vortex and reweigh. 2.6.8. Solid-phase extraction PFCAs have been analyzed with success following cleanup with SPE, for a variety of extracts having complex matrices, e.g., [21,22]. For this effort, we followed steps 1–5 of our elementary extraction described in Section 2.6.1 and then: (6) decant 2 mL of supernatant into weighed 12-mL vials and reweigh; (7) autopipette 100 ␮L of 15 ng/g [13 C4 ]PFBA in 60:40 ACN:H2 O (v/v) as a recovery internal standard with respect to the cleanup, vortex and reweigh; (8) evaporate to dryness with filtered air; (9) reweigh vials after evaporation; (10) reconstitute using 2.5 mL 18-M H2 O; (11) prepare HLB 3-cm3 cartridge per instructions on package; (12) load cartridge with 2 mL of aqueous extract, reserving 0.5 mL for ‘starting-point’ analysis and the 2 mL of loading eluate water to check for analyte ‘break through;’ (13) wash cartridge with 2 mL 0.1 M formic acid using minimum vacuum reserving eluate acid to check for ‘break through;’ (14) elute with 2 mL ACN using minimum vacuum; (15) dilute with 18-M H2 O to 60:40 ACN:H2 O and reserve for analysis; (16) post-elute cartridge with 2 mL MeCl2 , again using minimum vacuum; (17) evaporate MeCl2 to dryness with filtered air; (18) reconstitute to 1 mL of 60:40 ACN:H2 O and reserve for analysis. 3. Results and discussion 3.1. Analytical column/system comparisons For the Hypersil Gold C18 column on the Varian system, all monitored acids, C6 through C10 , eluted with baseline separation in all cases (Fig. 1a–e). Peaks were mono-nodal in all cases excepting intermittently at the lowest concentration. Symmetry

generally was good, excepting a little post-peak tailing (e.g., Fig. 1e). The IDL, as defined in Washington et al. [14], for PFOA on the Hypersil Gold column was 9.1 fg/␮L, or about 180 fg on column. For the Wako Fluofix-II 120E column on the Varian system, the peaks for C6 through C8 were not separated to baseline, but C9 and C10 did separate from the other homologues (Fig. 2a–e). The C6 and C7 peaks were sharp, albeit with post-peak tailing, but the longer-chained homologues had broad, multi-nodal, asymmetric peaks (Fig. 2a–e). The IDL for PFOA on the Wako column was 36 fg/␮L, or about 720 fg on column. At least part of the contrast in performance achieved with the Wako column between Triangle Park/EPA group who achieved good chromatography [17] and us likely is related to choice of solvents. Whereas the Triangle Park/EPA group used MeOH/H2 O, we used ACN/H2 O. We chose to stay with ACN/H2 O because we had better sensitivity with this solvent matrix than we did with MeOH/H2 O. For the Zirchrom SAX Anion column on the Varian system, the peak apices for C6 through C10 eluted over approximately 0.6 min and the peaks were not separated to baseline (Fig. 3a–e). The short retention time of these co-eluting peaks is approximately equal to that expected for conservative flow through the stationary phase, perhaps related to the perfluorinated composition of the analytes. Whereas co-elution of analytical peaks generally is undesirable, this is particularly true for the PFCAs because several of the homologues share a common confirmation ion (Table 1). For this column, the peaks were broader and less symmetric than with the Hypersil Gold, and not always mono-nodal (e.g., Fig. 3e). The IDL for PFOA on the SAX column was 10 fg/␮L, or about 200 fg on column. Based on these observations, the C18 columns gave the best combination of sensitivity, peak shape and chromatographic sep-

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J.W. Washington et al. / J. Chromatogr. A 1181 (2008) 21–32

Fig. 4. Detection of the 413 → 369 transition for PFOA at 313 ag/␮L (ppq). The upper mass chromatogram is a blank consisting of 60% Optima-Grade ACN, 40% 18 M H2 O polished with an HLB cartridge, as described in the text. The lower mass chromatogram is 313 ag/␮L PFOA in the same matrix as the blank. The peak-to-peak signal to noise for the standard is S/N = 3.3.

aration of analytical peaks. For the Varian, we chose to continue our investigations using the Hypersil Gold C18 system. Based on these results, for the Waters system we chose to stay with the C18 stationary phase, installing a Waters BEH C18 column in the system. For the Waters system using the Waters BEH C18 analytical column and with the UPLC plumbing altered as described in Section 2.3, we explored the IDL capability with 50 ␮L injections in “partial loop with needle overfill” mode, seven injections of a 313 ag/␮L standard and seven injections of solvent blanks. Using these data and the IDL definition we derived in our earlier work [14], which accounts for the difference in peak area between a standard and a blank, our IDL was calculated to be 270 ag/␮L or about 14 fg on column (Fig. 4). To our knowledge, nobody has reported in peer-reviewed literature a PFOA IDL as low as this before. 3.2. Extraction method comparisons We summarize our analytical results for the three test soils by extraction method in Table 3. These and other soil data reported herein represent the mean ± one standard deviation of three analytical runs on each of three replicate vials for each soil extraction, so each concentration is the mean of nine analyses except when rare outliers were omitted. Except as noted in Table 3, the values were calculated using [13 C4 ]PFOA as a matrix internal standard. Whereas [13 C4 ]PFOA can be used as a matrix internal standard to quantitate unlabelled PFOA accurately, values generated by this method for PFCAs that elute at times other than that of [13 C4 ]PFOA are only rough approximations, generally ranging from about half to double the correct concentration with these analytes in these soils in our experience [14]. Given the central objective of this work, to evaluate the efficacy soil-extract clean-up methods, the approximate nature of PFCA data other than PFOA is not an impediment. However, for efforts in which quantitation of all the PFCAs is desired, additional efforts are necessary. Commercially produced, mass-labeled PFCAs exist for most of the homologues from C4 to C12 , including numerous mass-labelings of PFOA,

e.g., [13 C2 ]PFOA, [13 C4 ]PFOA and [13 C8 ]PFOA. As such, when quantitation of multiple PFCAs is an objective of analyses, most PFCAs can be quantitated by normalizing to their isotopically enriched molecule, effectively using isotopic dilution to quantitate, and recovery could be calculated using a second isotopically enriched PFOA molecule chosen from the variety available. Alternatively, the method of standard additions could be performed for all analytes. For reference, the first extraction listed in Table 3 for each soil is our elementary extraction with no pretreatment or cleanup step. In earlier research, we established that this method achieves good recovery of recovery internal standards as well as PFCAs spiked and aged in the soil, under laboratory conditions, for up to 70 days [23]. Concerned with recovery of endogenous PFCAs exposed to the soils for still longer times under environmental conditions, we also inferred good recovery by extracting and analyzing PFCA levels in highly contaminated soils, for which we did not know the concentrations a priori, finding levels closely comparable with those found by our colleagues at the Triangle Park/EPA laboratory using different extraction and analysis methods [23]. Evaluating the PFOA results in Table 3, for which we have established that [13 C4 ]PFOA is a suitable matrix internal standard to quantitate upon [14], values generally are reasonably comparable between extractions within soils; with few exceptions, PFOA values vary among the extraction methods, within soils, by less than twofold. The most prominent outlier among the extraction methods, for both the quantitated PFOA as well as the other, approximated PFCAs, is the SPE cleanup. For this method, C8 in the Cowart soil was the only detect (Table 3). The results we report in Table 3 for the SPE method are from our second attempt at this method. In our first attempt, we achieved very poor recovery as well. Reflecting on this first effort, we tentatively ascribed our low recovery to application of too-high a vacuum during elution of the PFCAs trapped on the SPE solid phase resulting in a ‘shortcircuit’ flow of the eluent through the SPE. Resultingly, as we describe above for the cleanup leading to these data, we applied the minimum vacuum necessary to draw the eluents through the

Table 3 Soil [PFCA] analytical results on Varian LC/MS/MS for each pre-extraction, extraction and post-extraction Experimental system Soil

Picnic

Cowart (6)

Alkaline treatment

Extractant

0.5 M NaOH 0.5 M NaOH 0.5 M NaOH (1) 0.5 M NaOH(1) 0.2 M Na2 CO3 (2) (1)

ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O TBA ACN/H2 O (3)

0.5 M NaOH 0.5 M NaOH 0.5 M NaOH (1) 0.5 M NaOH(1) 0.2 M Na2 CO3 (2) (1)

ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O TBA ACN/H2 O (3)

0.5 M NaOH 0.5 M NaOH 0.5 M NaOH (1) 0.5 M NaOH(1) 0.2 M Na2 CO3 (2) (1)

ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O TBA ACN/H2 O (3)

Cleanup

PFXA (C6 )

PFHA (C7 )

PFOA (C8 )

PFNA (C9 )

PFDA (C10 )

C (15 m) C (16 h) TBA/MTBE TBA/MTBE MTBE (2 h) SPE

ND 0.062 ± 0.094 0.038 ± 0.057 ND 0.701 ± 0.471 ND 0.096 ± 0.014 ND

ND ND ND ND ND 0.372 ± 0.156 0.087 ± 0.037 ND

0.084 ± 0.026 0.147 ± 0.047 0.092 ± 0.092 0.120 ± 0.033 0.111 ± 0.222 0.089 ± 0.083 0.315 ± 0.063 ND (4)

0.074 ± 0.013 0.109 ± 0.027 0.079 ± 0.016 0.062 ± 0.019 0.174 ± 0.236 0.514 ± 0.175 0.111 ± 0.027 ND

0.019 ± 0.016 0.034 ± 0.038 0.050 ± 0.013 0.040 ± 0.019 ND 0.145 ± 0.075 0.065 ± 0.024 ND

137 ± 13 152 ± 20 161 ± 14 124 ± 17 87 ± 22 139 ± 21 82 ± 11 16 ± 6

C (15 m) C (16 h) TBA/MTBE TBA/MTBE MTBE (2 h) SPE

0.153 ± 0.059 0.028 ± 0.044 ND ND ND 1.584 ± 0.767 0.320 ± 0.032 ND

0.076 ± 0.030 ND ND ND ND 0.592 ± 0.269 0.142 ± 0.035 ND

0.539 ± 0.102 0.428 ± 0.142 0.423 ± 0.078 0.487 ± 0.101 0.503 ± 0.138 1.174 ± 0.157 0.779 ± 0.129 ND

0.411 ± 0.701 0.237 ± 0.083 0.225 ± 0.085 0.159 ± 0.053 0.143 ± 0.063 0.377 ± 0.219 0.149 ± 0.050 ND(5)

0.231 ± 0.035 0.258 ± 0.022 0.407 ± 0.649 0.197 ± 0.063 0.226 ± 0.211 0.323 ± 0.111 0.130 ± 0.029 ND

103 ± 7 155 ± 24 134 ± 17 167 ± 33 75 ± 12 97 ± 38 58 ± 8 23 ± 33

C (15 m) C (16 h) TBA/MTBE TBA/MTBE MTBE (2 h) SPE

0.198 ± 0.061 ND 0.214 ± 0.326 0.540 ± 0.191 ND ND 0.296 ± 0.051 ND

0.184 ± 0.113 ND ND ND 0.149 ± 0.063 0.544 ± 0.295 0.210 ± 0.086 ND

0.732 ± 0.114 0.452 ± 0.104 0.459 ± 0.159 0.539 ± 0.144 0.201 ± 0.054 0.811 ± 0.180 0.740 ± 0.156 0.031 ± 0.013 (7)

0.140 ± 0.049 ND ND ND 0.076 ± 0.037 0.360 ± 0.127 0.120 ± 0.044 ND

0.168 ± 0.023 ND 0.036 ± 0.058 ND ND 0.169 ± 0.055 0.080 ± 0.035 ND

107 ± 21 109 ± 16 98 ± 16 148 ± 29 28 ± 12 88 ± 18 64 ± 10 0

J.W. Washington et al. / J. Chromatogr. A 1181 (2008) 21–32

P1

[13 C5 ]PFNA Recovery (%)

[PFCA] (ng/g dry soil)

Notes: ND indicates none detected due to absence of an analytical peak or uncertainty imparted by matrix. (1) These quantifications based on external calibrations due to instrumental instability for this matrix, see text for details. (2) These experimental results obtained using TBAHS contaminated with unlabeled PFCAs. (3) Recovery from two elution steps, the first with 2 ml ACN and the second with 1 mL MeCl2 . (4) For the third soil sample of this series, 0.166 ± 0.058 ng of PFOA passed through the SPE cartridge with the loading water, effectively short-circuiting past the intended loading of the cartridge—roughly 1/3 of the PFOA present in this 5-g soil sample. (5) C9 was detected in a minority of the SPE extracts for the Picnic soil, 3 of 12 injections, and we report here the majority result of ND. (6) Matrix interference was observed during the C8 elution time for the Cowart extracts. (7) Detected in 2 of 3 soil extracts; average does not include non-detects.

29

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J.W. Washington et al. / J. Chromatogr. A 1181 (2008) 21–32

Fig. 5. 413 → 369 SRM chromatograms for the P-1 soil on the Varian 1200L system using the Hypersil Gold C18 column for all pretreatment-extraction-cleanup combinations we tested. Despite selective analytical steps including chromatographic separation, electrospray ionization, precursor-ion mass selection and transition-ion mass selection, without cleanup, the soil matrix can impart sufficient complexity to dampen the contrast of the PFOA signal with noise (a). As the PFOA quantitation signal approaches the MDL, the PFOA confirmation signal can become indecipherable from noise stymieing confirmation of low PFOA concentrations. The most prominent noise usually precedes PFOA in most soils, eluting contemporaneously with shorter-chain PFCAs. Of the cleanups we tried, the ion-pairing cleanups (e and f) tended to be most effective in minimizing noise while retaining full analytical signal.

SPE column. The results we report for our second SPE effort (Table 3) were not an improvement over our first effort and argue that our shortcoming did not lay dominantly in rate of solvent elution. Compounding these poor recovery results, we also analyzed the loading eluate water after percolating it through the SPE. In one of six replicates, the loading eluate water retained

about 1/3 of the PFOA (Table 3) after percolating through the SPE indicating that the SPE did not properly trap the PFCAs. Judging our experience with SPE in light of the literature, van Leeuwen et al. [24] compared the analytical results of a ‘roundrobin’ survey of 38 laboratories for PFCAs in several types of samples including a simple water sample containing 19.4 ng/L of PFOA. For these water samples, 15 labs used SPE to isolate the PFCA analytes and, of these, seven labs appear to have used HLB media as we have in our efforts. The PFOA data reported by these labs using SPE ranged from 3.4 to 190 ng/L, and the low and high limiting values were from laboratories using HLB. The mean PFOA value for 15 SPE-using labs was 46 ± 54 ng/L (mean ± 1 standard deviation), an average of 240% the actual value. On the other hand, judging by good results for recovery internal standards, several groups have reported accurate and precise results using SPE (e.g., [22,25,26]); one common factor among these successful studies is that they eluted the trapped analytes from the SPE using large volumes of MeOH, at least 10 mL of MeOH for these studies, verses 2 mL of ACN and 2 mL of MeCl2 for our study. Regardless of the cause, our cleanups of soil extracts with SPE for analysis of PFCAs did not fare well. The efficacy of various cleanup procedures can be observed qualitatively in Fig. 5a–h which depicts typical SRM PFOA (413 → 369) chromatograms for each procedure on the P-1 soil. Fig. 5a illustrates typical matrix noise for the elementary ACN/H2 O extraction for which there was no cleanup. In this soil, a broad, multi-nodal, asymmetric noise peak elutes immediately prior to the analytical PFOA peak. For soils having smaller analytical PFOA peaks, or longer noisy tailing of the matrix, than for this P-1 soil, the presence of the analyte becomes ambiguous regardless of reference to multiple detection criteria including elution time, precursor-ion m/z, quantitation product-ion m/z and confirmation product-ion m/z. Referring to the SRM chromatograms of the extracts cleaned with the dispersed activated C, the Envicarb (Fig. 5c and d), the matrix noise does not appear to be appreciably less than the extract with no cleanup (Fig. 5a and b). The efficacy of cleanup in maximizing the contrast of signal to noise can be quantitatively evaluated simply by taking the ratio of the areas of the PFOA peak to the maximum noise peak eluting between 2.5 min and the PFOA signal; this ratio is summarized for all the extraction methods and tested soils in Table 4. The limited efficacy of Envicarb (Fig. 5c and d) is confirmed by the ratio in Table 4 in that the Envicarb cleanups fall in the same general range (e.g.,

Table 4 Soil (PFOA/maximum noise) peak area ratios for tested extractions Extraction procedure

(PFOA/maximum noise) peak area ratio

Alkaline treatment

Extractant

Fig. 5a 0.5 M NaOH (Fig. 5b) 0.5 M NaOH (Fig. 5c) 0.5 M NaOH (Fig. 5d) Fig. 5e 0.5 M NaOH (Fig. 5f) 0.2 M Na2 CO3 (Fig. 5g) Fig. 5h

ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O ACN/H2 O TBA ACN/H2 O

Cleanup

P-1

Picnic

Cowart

C (15 m) C (16 h) TBA/MTBE TBA/MTBE MTBE (2 h) SPE

1.04 ± 0.64 1.23 ± 0.58 1.97 ± 1.17 2.25 ± 1.70 1.65 ± 0.48 2.09 ± 1.03 8.62 ± 2.90 <1.0

5.96 ± 2.01 4.14 ± 1.22 3.37 ± 1.19 5.57 ± 2.21 5.71 ± 1.28 16.4 ± 5.77 24.1 ± 7.51 <1.0

10.5 2.48 2.07 2.41 2.75 13.6 26.6 0.72

± ± ± ± ± ± ± ±

2.34 1.12 0.65 0.78 1.19 6.79 6.41 0.52

88 ± 2 91 ± 6 77 ± 8 75 ± 7 0.106 ± 0.015 0.102 ± 0.010 0.647 ± 0.048 0.678 ± 0.057 NA indicates not analyzed.

Cw

0.5 M NaOH

ACN/H2 O ACN/H2 O

TBA/MTBE TBA/MTBE

NA 0.432 ± 0.051

0.160 ± 0.010 0.236 ± 0.058

0.120 ± 0.010 0.259 ± 0.118

87 ± 11 103 ± 4 87 ± 12 81 ± 11 0.154 ± 0.032 0.191 ± 0.023 0.689 ± 0.062 0.609 ± 0.047 0.5 M NaOH Pc

P1

0.5 M NaOH

ACN/H2 O ACN/H2 O

TBA/MTBE TBA/MTBE

NA 0.526 ± 0.112

0.122 ± 0.012 0.153 ± 0.042

0.174 ± 0.017 0.154 ± 0.010

103 ± 13 120 ± 8 75 ± 17 91 ± 11 0.038 ± 0.004 0.044 ± 0.006 0.084 ± 0.005 0.081 ± 0.015 ACN/H2 O ACN/H2 O

TBA/MTBE TBA/MTBE

NA 0.064 ± 0.030

0.030 ± 0.009 0.029 ± 0.006

0.057 ± 0.009 0.062 ± 0.015

External Calib. PFNA (C9 ) Alkaline treatment

Extractant

Cleanup

PFOA (C8 ) PFXA (C6 ) Soil

PFHA (C7 )

[PFCA] (ng/g dry soil using [13 C4 ]PFOA internal standard) Experimental system

Table 5 Soil [PFCA] analytical results on Waters LC/MS/MS for each pre-extraction, extraction and post-extraction

3.37–5.57 for the Picnic soil) as the elementary extractions (e.g., 4.14–5.96 for the Picnic soil). In contrast, the SRM chromatograms for the various ionpairing procedures all have much less noise than the elementary extraction (Fig. 5e–g vs. Fig. 5a). Among the ion-pairing procedures, the Na2 CO3 /TBA/MTBE (Fig. 5g) exhibited the strongest contrast in terms of the signal-to-early-noise peak-area ratios (Table 4). However, background for the Na2 CO3 /TBA/MTBE extraction shows a slight elevation starting at the PFOA peak and lasting until the end of the run compared to the other ion-pair extractions (Fig. 5e and f vs. Fig. 5g), a period during which longer-chain homologues elute. Also, since the ion-pair extractions (Fig. 5e–g) all include evaporation of the extract-cleanup MTBE and reconstitution in a common solvent mix, 60/40 ACN/H2 O, matrix effects for these three clean extracts likely is similar among the three extraction methods. Despite this similar matrix, the Na2 CO3 /TBA/MTBE (Fig. 5g) shows a markedly lower recovery (Table 3) than the other two ion-pair procedures (Fig. 5e and f). Taking the early noise, signal-to-noise ratios, late noise and recovery all together, the NaOH/ACN/H2 O/TBA/MTBE (Fig. 5f) appears to provide the best results for extraction of PFCAs from soil of the methods we tested. A major drawback to the ion-pairing extractions (Fig. 5e–g), however, is that, when we ran these extracts on the Varian 1200L LC/MS/MS, the instrument’s sensitivity decreased steadily during the sequence, dropping about sixfold over about 100 analytical runs (Fig. 6). When we stopped a sequence and cleaned the MS, the instrument’s sensitivity returned, only to decrease once again when the sequence was continued (Fig. 6). This effect was so pronounced that we could not establish a multi-point standard curve with the ion-pair extracts on the Varian. Instead, the values reported for the ion-pair extracts in Table 3 are from our best-estimate, single-point calibrations, forced through zero, using the standard that was closest in run sequence to each individual extract. Such instrumental instability obviously is a fatal shortcoming for the purpose of quantitation.

PFDA (C10 )

Fig. 6. PFOA peak area of a standard injected alternately with an ion-pair extract on the Varian LC/MS/MS () and the Waters Acquity/Quattro () LC/MS/MS systems as a function of sample injection number. Because of decreasing sensitivity on the Varian, the run was suspended at about 40 injections, the MS baked out overnight, the capillary and inner needle flushed with MeOH, the ESI chamber cleaned, the MS vented and pumped down anew. After this cleanup, the sensitivity rebounded, but dropped once again as more ion-pair extracts were injected. In contrast, the Waters system retained constant sensitivity during the entire run.

31

Internal calib.

13 C -PFNA 5

Recovery (%)

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Presented with this unacceptable instrument instability, we tested the Waters Acquity/Quattro system for stability with the ion-pair extracts alternating same ion-pair extracts used on the Varian with standards for a total of 100 analytical runs. After a small initial drop-off in sensitivity, the Waters system remained steady for the remainder of the sequence of 100 runs (Fig. 6). Given this steady instrumental response on the Waters, we reanalyzed the ACN/H2 O/TBA/MTBE and NaOH/ACN/H2 O/TBA/MTBE extracts on the Waters system. Because of limited extract volume remaining after all the previous analyses, for these analyses we had to dilute them about two times. In turn, for the internal standard calibration, we had to compensate the analytical results algebraically for the dilution of the [13 C4 ]PFOA contained in the extracts. This compensation effectively was a one-point calibration forced through zero, so the precision of the results reported for these values (Table 5) are limited. Despite this, the values for PFOA and % recovery all compare well with the elementary and Envicarb values listed in Table 3, indicating satisfactory analysis for the ion-pair extracts on the Waters instrument. Table 5 also includes, however, values for PFXA and PFHA which we could not detect with the elementary and Envicarb extractions, largely due to obfuscation of signal by the complex matrices of these latter extractions. It also is noteworthy that, in every analysis performed on the Waters system with ion-pair extracts, recovery falls within 30% of perfect recovery (Table 5). 4. Conclusions While an elementary acetonitrile/water extraction of soils recovers PFCAs effectively, natural soil organic matter also dissolved in the extracts commonly imparts significant noise that appears as broad, multi-nodal, asymmetric peaks that co-elute with several PFCAs. This noise is highly variable among soils and it challenges detection of low concentrations of PFCAs by lowering the signal-to-noise ratio. In an effort to decrease this background noise, we investigated several methods of cleanup. For the combined objectives of complete recovery and minimization of background noise, we have settled on: (1) alkaline pretreatment; (2) extraction with acetonitrile/water by sonication; (3) evaporation to dryness; (4) reconstitution with tetrabutylammonium hydrogen sulfate ion pairing; (5) ion-pair extraction to methyl-tert-butyl ether; (6) evaporation to dryness; (7) reconstitution with 60/40 acetonitrile/water (v/v); and (8) LC separation using a C18 column, MS/MS analysis and quantitation with mass-labeled matrix internal standards corresponding to each analyte or by use of the method of standard additions. Acknowledgements The authors thank USEPA/OPPT for contributing financially to this research. We thank Cathy Fehrenbacher, Greg Fritz, Lau-

rence Libelo, David Lynch, Carolyn Acheson, Bryan Boulanger, Mark Strynar, Shoji Nakayama and Tim Collette for stimulating conversations regarding PFOA and/or reviews of this manuscript, and we thank anonymous reviewers for helpful reviews. References [1] J.P. Giesy, K. Kannan, Environ. Sci. Technol. 35 (2001) 1339. [2] A.M. Calafat, L.L. Needham, Z. Kuklenyik, J.A. Reidy, J.S. Tully, M. Aguilar-Villalobos, L.P. Naeher, Chemosphere 63 (2006) 490. [3] J.W. Martin, M.M. Smithwick, B.M. Braune, P.F. Hoekstra, D.C. Muir, S.A. Mabury, Environ. Sci. Technol. 38 (2004) 373. [4] J. Martin, D. Whittle, D. Muir, S. Mabury, Environ. Sci. Technol. 38 (2004) 5379. [5] K.S. Guruge, L.W. Yeung, N. Yamanaka, S. Miyazaki, P.K. Lam, J.P. Giesy, P.D. Jones, N. Yamashita, Toxicol. Sci. 89 (2006) 93. [6] W. Henderson, M.M.A. Smith, Toxicol. Sci. 95 (2007) 452. [7] W. Henderson, E. Weber, S. Duirk, J. Washington, M. Smith, J. Chromatogr. B 846 (2006) 155. [8] C. Lau, J.L. Butenhoff, J.M. Rogers, Toxicol. Appl. Pharmacol. 198 (2004) 231. [9] J. Washington, D. Endale, L. Samarkina, K. Chappell, Geochim. Cosmochim. Acta 68 (2004) 4831. [10] J. Oades, in: J. Dixon, S. Weed (Eds.), Minerals in Soil Environments, Soil Science Society of America, Madison, WI, 1989, p. 89. [11] U. Schwertmann, R. Taylor, in: J. Dixon, S. Weed (Eds.), Minerals in Soil Environments, Soil Science Society of America, Madison, WI, 1989, p. 379. [12] G. Borchardt, in: J. Dixon, S. Weed (Eds.), Minerals in Soil Environments, Soil Science Society of America, Madison, WI, 1989, p. 675. [13] U. Berger, M. Haukas, J. Chromatogr. A 1081 (2005) 210. [14] J. Washington, J. Ellington, T. Jenkins, J. Evans, J. Chromatogr. A 1154 (2007) 111. [15] M. Oehme, Practical Introduction to GC–MS Analysis with Quadrupoles, Huthig Verlag, Heidelberg, 1998. [16] D. Coleman, L. Vanatta, Am. Lab. 37 (2005) 23. [17] S. Nakayama, M. Strynar, L. Helfant, P. Egeghy, X. Ye, A. Lindstrom, Environ. Sci. Technol. 41 (2007) 5271. [18] C.R. Powley, S.W. George, T.W. Ryan, R.C. Buck, Anal. Chem. 77 (2005) 6353. [19] A. Brandstrom, Pure Appl. Chem. 54 (1982) 1769. [20] K. Hansen, L. Clemen, M. Ellefson, H. Johnson, Environ. Sci. Technol. 35 (2001) 766. [21] M.K. So, S. Taniyasu, P.K. Lam, G.J. Zheng, J.P. Giesy, N. Yamashita, Arch. Environ. Contam. Toxicol. 50 (2006) 240. [22] M.K. So, N. Yamashita, S. Taniyasu, Q. Jiang, J.P. Giesy, K. Chen, P.K. Lam, Environ. Sci. Technol. 40 (2006) 2924. [23] J. Ellington, J. Washington, M. Strynar, J. Evans, T. Jenkins, W. Henderson, Society of Ecological Toxicology and Chemistry National Meeting, Society of Ecological Toxicology and Chemistry, Baltimore, MD, 2005. [24] S. van Leeuwen, A. Karrman, B. van Bavel, G. Lindstrom, Environ. Sci. Technol. 40 (2006) 7854. [25] B. Scott, C. Moody, J. Small, D. Muir, S. Mabury, Environ. Sci. Technol. 40 (2006) 6405. [26] E. Sinclair, S. Kim, H. Akinleye, K. Kannan, Environ. Sci. Technol. 41 (2007) 1180.