Electro-Thermal Vaporization Direct Analysis in Real Time-Mass

Sep 20, 2013 - Spectrometry for Water Contaminant Analysis during Space Missions ... contaminants.1 Mass spectrometric detectors based on time- of-fli...
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Electro-Thermal Vaporization Direct Analysis in Real Time-Mass Spectrometry for Water Contaminant Analysis during Space Missions Prabha Dwivedi,† Daniel B. Gazda,‡ Joel D. Keelor,† Thomas F. Limero,‡ William T. Wallace,‡ Ariel V. Macatangay,§ and Facundo M. Fernández*,† †

School of Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332, United States Wyle Science, Technology and Engineering Group, Houston, Texas 77058, United States § NASA Johnson Space Center, Houston, Texas 77058, United States ‡

S Supporting Information *

ABSTRACT: The development of a direct analysis in real time-mass spectrometry (DART-MS) method and first prototype vaporizer for the detection of low molecular weight (∼30−100 Da) contaminants representative of those detected in water samples from the International Space Station is reported. A temperature-programmable, electro-thermal vaporizer (ETV) was designed, constructed, and evaluated as a sampling interface for DART-MS. The ETV facilitates analysis of water samples with minimum user intervention while maximizing analytical sensitivity and sample throughput. The integrated DART-ETV-MS methodology was evaluated in both positive and negative ion modes to (1) determine experimental conditions suitable for coupling DART with ETV as a sample inlet and ionization platform for time-of-flight MS, (2) to identify analyte response ions, (3) to determine the detection limit and dynamic range for target analyte measurement, and (4) to determine the reproducibility of measurements made with the method when using manual sample introduction into the vaporizer. Nitrogen was used as the DART working gas, and the target analytes chosen for the study were ethyl acetate, acetone, acetaldehyde, ethanol, ethylene glycol, dimethylsilanediol, formaldehyde, isopropanol, methanol, methylethyl ketone, methylsulfone, propylene glycol, and trimethylsilanol.

T

of analytical methods for environmental contaminants.1−3 However, many of these techniques are not suitable for space flight, often because of some combination of mass and/or volume limitations, power requirements, consumables, safety concerns, or the need for skilled operators. In terms of sample preparation, solid-phase microextraction (SPME), liquid−liquid extraction, stir bar sorptive extraction, purge and trap (PT), and headspace (HS) procedures are the most widely used techniques for low-molecular-weight organics.4,5 For example, formaldehyde and acetaldehyde are most frequently analyzed by HPLC/UV−vis after 2,4dinitrophenylhydrazine (DNPH) derivatization and solidphase extraction (SPE).6,7 Gas-phase aldehyde measurements traditionally use derivatization by o-2,3,4,5,6-(pentafluorobenzyl) hydroxylamine hydrochloride (PFBHA)8,9 followed by gas chromatographic analysis. Environmental monitoring on the International Space Station (ISS) is accomplished using data from several onboard

race contaminants in the spacecraft environment can have serious effects on the overall health of crewmembers and pose significant risks to the success of manned space missions. Environmental monitoring, along with a combination of active and passive controls, are key aspects of managing these risks. The National Aeronautics and Space Administration (NASA), in collaboration with the National Research Council (NRC), has been actively working toward identifying environmental health hazards for spaceflight crews and advancing analytical technologies for their determination. Low-molecular-weight polar organics are one class of compounds that can have a negative impact on crew health and are likely to be present in the spacecraft atmosphere. If a water recovery system is used, these compounds may also be present as contaminants in the water produced by that system. Most analytical laboratories use methods such as gas chromatography (GC) with flame ionization, electron capture, or photoionization detectors and/or high-pressure liquid chromatography (HPLC) with ultraviolet−visible (UV−vis) or electrochemical detectors for the determination of low-molecular-weight polar organic contaminants.1 Mass spectrometric detectors based on timeof-flight (TOF), quadrupole ion trap (QIT), and Fourier transform (FT) analyzers have also found effective use as part © 2013 American Chemical Society

Received: August 2, 2013 Accepted: September 20, 2013 Published: September 20, 2013 9898

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analytical platforms, augmented by ground analysis of archive samples.10−13 Although environmental monitors on ISS provide very limited information, the state of the environment can be adequately assessed from these key parameters. The Total Organic Carbon Analyzer (TOCA)14 and the Colorimetric Water Quality Monitoring Kit (CWQMK)15 are used to monitor total organic carbon and biocide concentrations, respectively. Culture-based microbial assay kits are available to monitor total heterotrophic plate counts for surface, air, and water samples, and a colorimetric assay is provided to check for coliform bacteria in water. The air monitoring capabilities on the ISS are more advanced; instruments based on GCdifferential ion mobility spectrometry (DMS) are capable of monitoring at least 20 target compounds. 16 Common atmospheric gases, such as oxygen, nitrogen, carbon dioxide, and water vapor, are measured on separate instruments such as the Major Constituents Analyzer (MCA),17 the ISS Portable Oxygen Monitor (iPOM), and the Carbon Dioxide Monitoring Kit (CDMK).16 Detailed chemical analyses of ISS archival air and water samples are performed at the Johnson Space Center (JSC) in the Toxicology Laboratory and the Water and Food Analytical Laboratory (WAFAL), respectively. These laboratories use suites of analytical instrumentation to monitor hundreds of analytes in the archive samples. Unfortunately, most of the instrumentation used to perform these analyses is not adaptable for in-flight monitoring for the reasons listed previously. The use of multiple, highly specialized monitors and reliance on ground analyses is a weakness in the current approach to environmental monitoring on manned spacecraft. This limitation has the potential to affect operations on future exploration missions beyond low Earth orbit, from which return of archive samples will not be possible. Having the ability to collect and interpret data in near-real time will help manage nominal and off-nominal scenarios, reduce crew time associated with archive sampling, and help ensure crew health. Moreover, because spacecraft are closed systems, many of the target compounds for air and water samples are the same. The ability to monitor air and water trace contaminants using a single instrument would provide savings in both crew time and mass/ volume. Developments in the last 8−9 years in the field of open air ″ambient″ ionization technology, including plasma-based ionization sources such as direct analysis in real time (DART) and spray-based sources such as desorption electrospray ionization (DESI), have provided new avenues for developing high-throughput analytical applications that were not previously possible.18−20 Plasma-based ambient ionization techniques involve direct current or radiofrequency electrical discharges sustained between a pair of electrodes in direct or indirect contact with a flowing gas such as nitrogen or helium. Ionized molecules, radicals, excited-state neutrals, and electrons generated in the plasma are carried by the discharge gas and used for ionization of samples. Coupling of plasma-based ionization techniques to mass spectrometry (MS) allows the analysis of samples in solid, liquid, or gas phase with high sensitivity and throughput.19 DART is one of the most popular plasma-based ambient ionization techniques because of its simplicity, robustness, and commercial availability.21,22 DART applications ranging from fast screening of sample composition to applications in metabolomics have been reported in recent years.23−26

Here, we investigate the analytical performance of a proof-ofconcept system comprising a DART plasma ion source coupled to an electro-thermal vaporization (ETV) unit followed by MS detection for the purpose of direct, rapid water quality assessment with minimal sample preparation. The choice of ETV instead of the more conventional neutral desorption approach using a heated plasma discharge gas has the advantage of allowing controlled introduction of the sample before it is desorbed with minimal analyte losses, followed by a rapid desorption event driven by the ETV element. This is critical for quantification purposes, as maximum signal reproducibility is required to achieve acceptable linearity of the calibration curve, even with manual sample introduction. We envision that the DART-ETV-MS concept system presented here could also have applications beyond space mission water analysis, including routine water quality assessment on Earth.

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EXPERIMENTAL SECTION Chemicals. Selection of the target analytes used in the study was based on comparison of the target analyte lists for air and water quality monitoring on the ISS. The target compounds represent low-molecular-weight polar organics that are routinely detected in the ISS atmosphere and have the potential to be present in water produced by the Water Processing Assembly (WPA). All of these compounds are either contaminants that have been previously detected in the WPA product water or species that could be produced by incomplete oxidation of larger organic molecules. They include ethyl acetate (EtOAc), acetone (ACTN), ethanol (EtOH), ethylene glycol (EG), dimethylsilanediol (DMSD), formaldehyde (FM), acetaldehyde (AcA), isopropanol (IPA), methanol (MeOH), methylethyl ketone (MEK), methylsulfone (DMSO2), propylene glycol (PG), and trimethylsilanol (TMS). MeOH, ACTN, and IPA were HPLC grade (EMD Chemicals, Gibbstown, NJ). Ultrapure water (18.2 MΩ cm−1) used for dilution was obtained from a Nanopure purification unit (Barnstead, San Jose, CA). DMSD was provided by NASA at 100 ppm concentration in water. The rest of the target analytes were purchased from Fisher Scientific Inc. (≥98% purity). DART Instrumentation and Operation. A commercial DART-100 ion source (IonSense Inc., Saugus, MA) was used in this prototype proof-of-principle configuration with the goal of later using a miniaturized plasma ion source if initial results were promising in terms of sensitivity and reproducibility. Detailed description of the DART-100 ion source is available elsewhere.21,27 In short, the ion source consists of a needle electrode and a grounded perforated electrode, between which a discharge is initiated through a potential difference of several kilovolts. This discharge results in a plasma plume that is driven toward the source nozzle and is stripped of most ions by two perforated (grid 1 and grid 2) electrodes. The ceramic nozzle of the DART ion source was positioned in line with the entrance of the ETV unit 1 cm away from the inlet (described below). The DART gas (high-purity N2, 99.995%, Airgas, Atlanta, GA) was not heated. Table S1 (Supporting Information) lists typical DART operation parameters that provided optimum sensitivity for all species investigated. TOF MS Detection. The DART-100 ion source was coupled to a Bruker micrOTOF-Q I mass spectrometer (Bremen, Germany) fitted with the ETV unit. To prevent gas overload on the mass spectrometer vacuum system, a VAPUR gas-ion separator tube (GIST) interface (IonSense, Saugus, MA) was used. This VAPUR interface was connected to a 9899

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Figure 1. Schematic diagram (A) and photograph (B) of the DART-ETV-MS setup. A close-up picture of the ETV is shown in part C. Part D (1−4) illustrates the mass spectra of background and acetaminophen ions (used as test compound) in positive mode (D1 and D2, respectively) and of background and acetaminophen ions in negative mode (D3 and D4, respectively). Spectra were acquired with an ETV current of 5 A.

Vacuubrand 2C diaphragm pump (Vacuubrand, Wertheim, Germany), the pumping rate of which was optimized for maximum ion detection while minimizing pressure in the first differentially pumped chamber. The mass spectrometer voltage settings were optimized for maximum transmission of low-mass (30−300 Da) ions (Table S1, Supporting Information). For accurate mass measurement purposes, the mass spectrometer was first calibrated using a 10 μM solution of PEG 400 and PEG 600 (50:50) in MeOH. A continuous supply of this calibration solution was delivered through a 10 μm i.d. silica capillary placed in front of the ion source at a flow rate of 3 μL min−1 using a KD Scientific syringe pump (New Hope, PA). The measured resolving power (fwhm) of the Q-TOF mass spectrometer was 9500 for m/z 152.0706 (acetaminophen) and 13 500 at m/z 430.9138 (PEG 600). Following calibration, a mass accuracy of 1−5 ppm was measured for multiple runs of the [M + H]+ acetaminophen ion. To extend the calibration into the 30−100 Da range, the instrument was recalibrated with a mixture of 0.1% trimethylamine (TMA) with 10 μM PEG 400 and PEG 600 dissolved in 50:50 MeOH/water. Though mass accuracies of 1−5 ppm were attainable for masses greater than 150 Da, mass measurement errors of up to 10 mDa were still observed for m/z values below this range. Bruker Daltonics DataAnalysis version 4.0 software package was used for processing all acquired data. In an alternate setup used during this study, the DART-100 ion source described above was coupled in-line to another orthogonal acceleration TOF mass spectrometer (JEOL AccuTOF, Tokyo, Japan). Ion source parameters were kept identical to those used in the DART-micrOTOF-Q I setup. The DART nozzle was placed 10 mm away from and concentric with the inlet of the AccuTOF mass spectrometer. In this setup an interface between the DART ion source and the mass spectrometer to prevent gas overload on the mass spectrometer vacuum system was not required. The AccuTOF

mass spectrometer provided a mass resolution range of ∼1700−4100 in the mass range of 18.02−121.10 Da. ETV Unit. The schematic of the DART-ETV-MS setup is presented in Figure 1A and a photograph in Figure 1B. The ETV unit was constructed using two glass tubes and a nichrome ribbon powered by a programmable dc power supply (G W Instec, PSM-3004). The ribbon was threaded through two 3mm-wide, 1-cm-long slot cuts on the inner tube placed 1.5 cm past the inlet. The ribbon was held securely by compressing it between the inner tube and two halves of an outer glass tube. These halves were held together using a flexible metal clamp. Inside the inner tube the ribbon was slightly curved, and an indent (1 mm diameter) was made on its surface for depositing a measured liquid sample drop. The ribbon was positioned in the upper half of the inner tube in such a way that the edge of the ribbon faced the front (inlet) side of the ETV unit. Holes (1 mm diameter) were made on the outer and inner glass tubes for sample introduction and were aligned with the ribbon indent. The close-up image of the ETV unit in Figure 1C shows the placement of the nichrome ribbon (3 mm wide, 50 mm long) within the ETV unit that consists of an inner glass tube (68 mm long, 10 mm o.d., 0.7 mm i.d.), an outer glass tube (20 mm long, 12 mm o.d., 10 mm i.d.), a clamp, a sample pit, and a sample injection port. The outlet end of the ETV unit was fitted over the 3-cm-long ceramic inlet tube of the VAPUR interface. During operation, the DART plasma plume enveloped the ribbon while simultaneously dragging the thermally desorbed analyte and reached the GIST interface, which directed ions into the mass spectrometer capillary inlet for chemical analysis. In this configuration, obstruction of plasma flow by the sample holder was minimized, thereby increasing ion transmission and reducing plasma-metal interaction, which can quench metastable ions.28 Enclosure of the vaporizer ribbon ensured minimum analyte losses to the atmosphere, confining the 9900

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reactant ion-analyte cloud to the internal volume of the ETV unit. This configuration is preferred over the typical transmission mode DART configuration, in which the plasma passes through a mesh on which sample is deposited.29,30 As less turbulence is generated, losses to the atmosphere are minimized and a smaller metal surface is exposed to the plasma before it interacts with the sample, thereby reducing neutralization of ions and metastable intermediates.28

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RESULTS AND DISCUSSION DART-ETV-MS Operation. For typical DART-ETV-MS operation, a small sample volume (0.5−2 μL) is pipetted directly onto the nichrome ribbon of the electro-thermal vaporizer. The nichrome ribbon in the ETV unit is connected to a programmable power supply that outputs controlled current steps (1−5 s duration, in a user-selectable 0−15 V/7 A or 0−30 V/4 A output range). Through the current step sequence, the water solvent is evaporated, and target solutes are concomitantly (or sequentially) volatilized and exposed to the plasma ionizing stream, generating a transient ionic signal with amplitude proportional to the deposited analyte amount. Aided by the DART gas flow, the fluid dynamic focusing effect of the ETV unit enclosure and the combined suction of the GIST interface and mass spectrometer vacuum system, ions are transmitted into the spectrometer for detection and identification. Helium is usually chosen as the DART gas for its higher energy, longer lifetime metastables and for its better heat transfer to high molecular weight analytes.31 However, because of the low molecular weights (30−94 Da) of the test analytes chosen for this study and the gas availability constraints on board the ISS, N2 was used as the DART gas. The temperature of the DART gas was kept ambient to reduce gas-aided vaporization of sample during the deposition step. Sample was pipet-deposited on the sample ribbon pit through the sampling port using a disposable pipet tip, the end of which was modified by addition of a 1-in.-long 22-gauge stainless steel needle. This modification was required to avoid introduction of plastic contaminants onto the vaporizer. This was particularly important during repeat operation, when sampling port edges of the ETV unit might not have completely cooled between sample depositions. Typical DART-ETV-MS spectra of background and acetaminophen ions, in both positive and negative iondetection modes, are depicted in Figure 1D (1−4). A blank spectrum of background species was acquired by depositing 2 μL of Nanopure water on the sample pit, and acetaminophen spectra were acquired by depositing 2 μL of 10 μM acetaminophen solution in water (3 ng, 20 pmol). For acetaminophen measurements, the ETV power supply was programmed to ramp the current output from 2 to 7 A in three steps: (1) desolvation (1 s, 2 A current), (2) analyte vaporization (2 s, 5 A), and (3) ribbon decontamination to prevent carryover (2 s, 7 A). These stages provided optimum sensitivity for this test analyte. Temperature and duration of the heating steps could be set differently for applications involving other analytes or more complex sample matrixes. DART-ETV-MS Water Analysis. The applicability of DART-ETV-MS for analysis of low-molecular-weight organics in water was investigated using standard solutions of the target analytes. Optimum current required for efficient analyte vaporization was determined by monitoring the effect of this variable on extracted analyte ion chronograms. Figure 2A shows the temperature measured with a thermocouple at positions 1

Figure 2. Panel A shows the temperature (°C) measured at position 1 (nichrome ribbon just outside the ETV glass enclosure, ▲) and position 2 (near the sample pit indented on the nichrome ribbon,  ●), as a function of the ETV current applied. The other panels (B1, B2, B3, and B4) show extracted ion chronograms for the acetaldehyde response ion at m/z 115.0839 ± 0.009 at second stage (vaporization) currents of 7, 5, 3, and 1 A, respectively.

and 2 on the nichrome ribbon (shown in Figure 1C) at currents of 1, 3, 5, and 7 A. Figure 2B shows the extracted ion chronogram (EIC) at m/z 115.0839 ± 0.009 corresponding to the [M++ 3H2O + NH3] ion of acetaldehyde (m/z = 115.0754). The EIC was acquired for four successive manual injections of 180 nmol of acetaldehyde when a desolvation step (1 s, 2 A current), an analyte vaporization step (2 s, variable current), and a ribbon decontamination step (2 s, 7 A) were applied. Maximum temperature attained at the sample pit with a 7 A current was 85 °C even though the temperature of the ribbon just outside the glass body of the ETV was 250 °C. This shows that the ETV glass enclosure and flowing DART gas 9901

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Figure 3. (A) Background-subtracted DART-ETV mass spectrum of methyl ethyl ketone (MEK, 110 mM solution in water). (B) Background mass spectrum. (C) EICs of prominent MEK response ions at m/z 57.0334 ± 0.015 (C1), 73.0648 ± 0.006 (C2), 90.0913 ± 0.006 (C3), 145.1223 ± 0.005 (C4), and 127.1117 ± 0.005 (C5). The following are tentative identities for these response ions: C1, [M − CH4 + H]+; C2, [M + H3O]+; C3, [M + NH4]+; C4, [2M + H]+; C5, [2M − H2O + H]+. M = C4H8O (exact mass = 72.0575 Da).

target analytes in the absence of MS/MS experiments. In addition, summation of peak areas of known and reproducible response peaks was found to increase the sensitivity of detection for analytes (not shown). However, care was taken to confirm the presence of all response peaks chosen for summation at all concentrations in the calibration range. For example, multiple propylene glycol response ions were detected at concentrations above 1 μM; however, at concentrations below 1 μM, only ions corresponding to [M + NH3 + H] + at m/z 94.0909 and [3(M − H2O) + H] + at m/z 175.1332 were detected. Representative background-subtracted mass spectra of the remainder of the test analytes are shown in Figures S1 and S2 (Supporting Information) in positive and negative ion detection modes, respectively. With the exception of DMSD, which was detected only in negative ion mode, all tested analytes were successfully detected in both modes. Signal-tonoise ratios were at least 2× times better for positive ions. Table 1 lists tentative identifications of prominent response ion peaks for all target analytes. Extracted ion chronograms of response peaks at the m/z values listed in this table were used to confirm the identity and positive detection of the analytes for repeat sample injections. Figure S3 (Supporting Information) illustrates the DART-ETV negative ion mass spectrum acquired for (A) 100 ppm DMSD solution (1.08 mM) in water and (B) background/reactant ions. Panels C1−5 in Figure S3 in the Supporting Information show EICs of prominent DMSD response ions detected for 198 ng of DMSD deposited on the ETV sample pit. The DMSD response ion corresponding to the negatively charged species at m/z 91.0371 generated through deprotonation of the monomer and a dehydrated dimer ion at m/z 165.0498 were detected along with three other peaks tentatively identified as being cyclic DMSD products at m/z 185.9545, 188.9761, and 188.9761. Similar to the multiple response peaks observed for DMSD in negative mode, the other selected analytes also produced

acted as a significant heat sink, and if higher temperatures are required for efficient vaporization, the design of the ETV will require further modifications. These modifications could include decreasing the resistivity of the ribbon and reducing the direct contact of the ribbon with the ETV glass tube. However, the temperature attained at the pit was sufficient for rapid vaporization of these test analytes without carryover because of their low mass and high volatility. As depicted in Figure 2B, the EICs showed instantaneous vaporization of the analyte as sharp peaks at vaporization currents of 5−7 A, whereas peak shapes at 1 and 3 A showed delayed, stepwise, or insufficient vaporization, leading to jagged peaks and carryover. Unless indicated otherwise, all further measurements were made at an ETV vaporization current of 6 A. Figure 3 illustrates a background ion mass spectrum (A), a background-subtracted mass spectrum (B), and EICs (C1−5) of identified response ions for MEK. EICs of prominent MEK response ions at m/z 57.0201, 73.0552, 90.0848, 145.1215, and 127.1105 are shown. The detected response ions corresponded to ions tentatively identified as [M − CH4 + H]+, [M + H2O + H]+, [M + NH3 + H]+, [2M + H]+, and [2M − H2O + H]+ with exact m/z values of 57.0334, 73.0647, 90.0913, 145.1223, and 127.1117 Da. With successively increasing concentrations of MEK in water from 0.011 to 1.1 M, percent standard deviations in peak areas ranged from 4 to 12 (n = 6) with an average deviation of 8%. Multiple response ions for the analyte were detected, constituting mostly water and ammonia adducts, clusters, and protonated fragments. This was expected, as analyte ionization by proton transfer with reactant hydronium ions is the prevailing ionization pathway in the DART ionization process.19,21 Cation adducts (e.g., Na+, K+) usually observed in solution-based ionization techniques such as electrospray ionization and desorption electrospray ionization are never observed in DART.32 Though the presence of multiple response ions can complicate a mass spectrum, multiple ions are very useful in confirming the identity of 9902

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multiple response ions in positive mode. Protonated ions [M + H]+, their water and/or ammonia clusters, and protonated fragment ions ([M − nNL + H]+, where n = 1, 2, 3...., and NL = H2O, CH4, CH3CH2, CH2, etc.) were detected for most analytes. Interestingly, acetaldehyde and formaldehyde produced multiple species corresponding to molecular ions (M•+) clustered with or without water and/or ammonia. Ethylene glycol produced a molecular ion dimer of the 2M•+ type in addition to protonated fragments and other adduct ions. Generation of molecular ions (M•+), similar to that observed in electron impact ionization (internal energy 40−80 eV),33 has been previously reported in DART ionization.32 The process of M•+ ion formation in DART, though not fully elucidated, is primarily attributed to (1) Penning ionization, which results from the interaction of high-energy excited-state metastables with analyte neutrals, and (2) charge-transfer ionization between analyte molecules and O2•+ reactant ions, when the analyte’s ionization potential is lower than that of O2 and the proton affinity is lower than that of water. When helium is used as the DART working gas, the high-energy (19.8 eV) He metastables that are generated possess enough internal energy to ionize common atmospheric gases and analyte neutrals (with ionization energy less than 19.8 eV) through the Penning ionization mechanism to form molecular ions. Formation of oxygen or water molecular ions through Penning ionization cannot be anticipated, however, when nitrogen is used as the DART working gas. This is because the nitrogen metastables (internal energy of up to 11.6 eV)33,34 have lower energy than the ionization energy of water (12.62 eV) and oxygen (12.07 eV).35 With ionization energies of 10.22 eV, 10.88 eV, and 10.16 eV for AcA, FM, and EG, respectively,35 production of molecular ions of these analytes is possible through interaction between nitrogen metastables and analyte neutrals whereas protonated ions would be generated through charge-transfer reactions. As both protonated and molecular ions were detected simultaneously for FM and EG, it can be inferred that both ion formation mechanisms occurred concurrently. Though detection of molecular ions has been correlated with high grid voltages of the DART ion source and with excess oxygen in the ionization region,32 molecular ions of these analytes were observed even at low grid voltages and high water concentration in the ionization region. Another interesting observation was that m/z values of some FM response ions (Table 1) suggested the detection of both the aldehyde (CH2O) and its reduction product (CH4O). As the most common FM production process involves catalytic oxidation of MeOH to FM, the presence of MeOH with FM can be attributed to processing impurity. Despite the detection of MeOH species as reproducible response ions for FM, response ions for MeOH as an analyte were not reproducibly observed. This suggests that the FM response peaks at 51.0440 and 65.0597 Da (the MeOH species) were fragments of adduct ions with masses of 81.0546 and 95.0652 Da. When MeOH, EtOH, ACTN, and IPA (introduced as neat vapor between the ion source and mass spectrometer inlet or as liquid deposited on the ETV) were analyzed by DART-MS, either response ions were not detected in every repeat analysis and/or the intensities were inconsistent between injections. Low-intensity protonated EtOH and MeOH signals with signalto-noise ratios in the 2−6 range were observed but not in every run. In the case of ACTN and IPA, although response ions were detected in each sample run, their intensities fluctuated 2to 12-fold between injections. Although detection of MeOH

methanol (MeOH) CH4O 32.0262 ethanol (EtOH) C2H6O 46.0419 59.0364; 59.0491 [M − H2O + H]+ 77.0514; 77.0597 [M + H]+ 94.0809; 94.063 [M + NH3 + H]+ 117.0894; 117.0910 [2(M − H2O) + H]+ 175.1332; 175.1328 [3(M − H2O) + H]+ propylene glycol (PG, positive mode) C3H8O2 76.0518

73.0364; 73.0468 [M − H2O + H]+ 91.0504; 91.0573 [M + H]+ 167.0550; 167.0918 [2M − CH2 + H]+

63.0325; 63.0440 [M + H]+ 80.0631; 80.0706 [M + NH3 + H]+ 107.0683; 107.0702 [2M − H2O + H]+ 121.0859; 121.0859 [M + acetone + H]+ 124.0729; 124.0730 [2M]•+

91.0371; 91.0208 [M − H]− 165.0498; 165.0405 [2M − H2O − H] −, (C4H13O3Si2)− 185.9545; 185.9282 [H2O8Si2]− 188.9761; 188.9517 [H5O8Si2]− 203.9633; 203.9388 [H2O + H2O8Si2]− 62.0486; 62.0362 [M+ + H2O] 115.0753; 115.0839 [M+ + 3H2O + NH3] 132.1026; 132.1104 [M+ + 3H2O + 2NH3] 150.1142; 150.1210 [M+ + 4H2O + 2NH3] 194.1417; 194.1472 [2M+ + 4H2O + 2NH3] [M + H]+ not consistently detected on micrOTOF-Q1. Easily detected on JEOL AccuTOF

95.0121; 95.0161 [M + H]+ 112.0425; 112.0406 [M + NH3 + H]+ 61.0250; 61.0284 [M − CH2CH2 + H]+ 89.0541; 89.0597 [M + H]+ 106.0839; 106.0862 [M + NH3 + H]+

isopropanol (IPA, positive mode) C3H8O 60.0569 methyl sulfone (DMSO2, positive mode) C2H6O2S 94.0089 dimethylsilanediol (DMSD, negative mode) C2H8O2Si 92.0294 acetaldehyde (AcA, positive mode) C2H4O 44.0262 59.0353; 59.0491 [M + H]+ 76.0664; 76.0756 [M + NH3 + H]+

61.0523; 61.0647 [M + H]+ 121.1219; 121.1223 [2M + H]+ 258.08 unidentified 285.08 unidentified

methylethylketone (MEK, positive mode) C4H8O 72.0569 M = CH2O, M′ = CH4O 51.0289; 51.0440 [M′ + H2O + H]+ 65.0487; 65.0597 [2M′ + H]+ 81.0477; 81.0546 [M′ + M + H2O + H]+ 95.0652; 95.0702 [2M′ + M + H]+ 107.0684; 107.0577 [3M+ + NH3] 137.0811; 137.0682 [4M+ + NH3]

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formaldehyde (FM, positive mode) CH2O 30.0100 (positive) acetone (ACTN, positive mode) C3H6O 58.0419 ethyl acetate (EtOAc, positive mode) C4H8O2 88.0518 trimethylsilanol (TMS, positive mode) C3H10OSi 90.0494 ethylene glycol (EG, positive mode) C2H6O2 62.0362

analyte elemental formula exact mass analyte response ions measured m/z; exact m/z analyte elemental formula exact mass

Table 1. Response Ions Observed for Target Analytes

57.0201; 57.0334 [M − CH4 + H]+ 73.0586; 73.0647 [M + H]+ 90.0858; 90.0913 [M + NH3 + H]+ 127.1120; 127.1117 [2M − H2O + H]+ 145.1239; 145.1223 [2M + H]+

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once increasing TOC is detected in the WPA product water; the DART-EVT-MS instrument could identify the compound responsible for that increase. Most of the required detection limits for the target compounds are in the vicinity of 1 mg L−1. As it is possible that more than 1 compound could be responsible for an observed TOC increase, it is desirable to be able to monitor the compounds at concentrations below the equivalent to the TOCA method detection limit. To that end, the desired detection limit for each compound was set at roughly half that equivalent concentration or 500 μg L−1. This is approximately equivalent to an analyte concentration of 5−20 mM (5−15 μmol) depending on the analyte. The upper limits for detection were set to cover the allowable concentrations for compounds documented in the Spacecraft Water Exposure Guidelines (SWEGs).38−41 The following are the 10-day SWEGs for selected analytes: FM, 20 mg L−1; ACTN, 3500 mg L−1; EG, 140 mg L−1; PG, 8 g L−1; and MEK, 54 mg L−1. Analysis of analyte solutions with varying analyte concentrations was conducted to study their correlation with response ion signal intensities. Figure S5 (Supporting Information) illustrates the variation in peak areas of select mass spectral analyte response ions as a function of analyte concentration. Limits of detection (LoD) for each analyte were calculated as the signal equivalent to 3 times the background noise for the most prominent analyte response peak (Table S2, Supporting Information). Depending on the analyte identity, the LoD ranged between 2 and 2000 pmol. It was noticed that the background noise levels varied with each response ion and each analyte and ranged between ∼20 ion counts (DMSD, m/z 91.0371) and 1200 ion counts (acetaldehyde, m/z 115.0753). The presence of trace amounts of analytes of interest in the ambient laboratory atmosphere was suspected as the cause for higher background noise levels for certain analytes. Representative calibration curves for MEK (0.011−1.1 M), EG (0.0017−0.35 M), and PG (0.0001−0.01 M) are presented in Figure S5 (Supporting Information). These curves were plotted using peak areas of response ions at m/z values of 127.1120 ± 0.005 (MEK), 107.0683 ± 0.005 and 121.0859 ± 0.005 (EG), and 175.1332 ± 0.005 (PG). Detection limits for ACTN, EtOH, MeOH, and IPA could not be estimated as the response ion peaks were not detected consistently in all sample runs. Response ion signal intensities for the rest of the test analytes increased with increasing analyte concentration in water from sub-mM up to molar concentrations. In some cases the signal intensities almost reached the levels equivalent to the neat analytes without noticeable saturation. Volatility of the analytes, combined with the open-air sampling nature and subsequent dilution by DART gas, could be the reason why saturation was not attained for any analyte at the examined concentrations. The results in Table S2 in the Supporting Information indicate that this method can be used to detect picograms of analytes deposited on the ETV sample pit; this amount is well below the target detection limit of 5−15 μmol for analytes of interest. Moreover, as the ETV used in the study was limited by the maximum current output of 7 A by the programmable power supply unit and by the size of the sampling pit that could be indented on the nichrome ribbon, the DART-ETV-MS has the potential to be modified to attain even lower detection limits by depositing larger sample volumes that can be concentrated by in situ evaporation.42 One can speculate that a low-gas flow miniature plasma ionization source43 might provide lower detection limits than

and EtOH by MS is common, the inability to reproducibly detect these analytes by DART-ETV-MS could be attributed to ion-molecule reactions occurring within the long transfer capillary used in the mass spectrometer interface, suggesting that quantification might still be possible with MS systems having shorter transfer capillaries or pinhole inlets. To verify this hypothesis, the DART-100 ion source was coupled to the orthogonal TOF mass spectrometer (JEOL AccuTOF, Tokyo, Japan) with a pinhole inlet (400 μm i.d.). Figure S4 (Supporting Information) shows the mass spectra of MeOH, EtOH, ACTN, and IPA acquired using the DART−AccuTOF platform. The analyte was introduced in the ionization region as a vapor. Depending on the voltages applied on the ring lens, orifice lens 1, and orifice lens 2, ion populations of different identities were instantly and consistently detected. As shown in Figure S4a in the Supporting Information, molecular ions of type [M + H2O]•+ were detected for all four target ions at lower lens voltages (ring lens at 5 V, orifice 1 at 1 V, orifice 2 at 1 V, ion guide bias at 29 V, pusher bias voltage at −28 V). The possibility that the ions detected were the ammonium adducts was ruled out because ammonium reactant ions were absent from the background mass spectrum. This hypothesis was also supported by the increase in the intensities of the peaks at 18.02 and 36.03 Da with the introduction of water into the ionization region. At higher lens voltages (ring lens at 8 V, orifice 1 at 25 V, orifice 2 at 2 V, ion guide bias at 29 V, pusher bias voltage at −28 V) [2M + H]+, [M + H2O + H]+, and/or [M + H]+ type ions were detected, Figure S4b (Supporting Information). Under low ring lens voltage settings, the reactant ion population was dominated by high-intensity signals at m/z 18.02 and 36.03 and relatively less intense signals at 19.02, 37.03, and 55.04, corresponding to water [M]+, [2M]+, [M + H]+, [2M + H]+, and [3M + H]+ ions, respectively. Interestingly, increasing the ring lens voltage resulted in an increase in protonated water and water clusters in the reactant ion population and the subsequent increase in protonated analyte species. Given the high reactivity of molecular cations,36,37 it can be speculated that, even though molecular cations of analytes are generated by the DART ion source, only long-lived species would be detected by the mass spectrometer provided they are efficiently transferred to the detector. One other factor that might also be contributing to specific ion losses in the micrOTOF-Q I instrument is the relatively high potential used on the reverse geometry of the capillary inlet biasing, which is required to transport ions from the ionization region into the first funnel region. Despite these findings for these specific analytes, our investigations did support the suitability of plasma ion sources for the detection of lowmolecular-weight organic volatiles when a suitable atmospheric pressure interface is implemented. These tests were conducted as preliminary experiments; with more in-depth studies are now underway. Analytical Figures of Merit. Currently, the only instrument available for monitoring organic contaminants in water samples on the ISS is the U.S. TOCA, which measures the total amount of carbon present in a sample; it does not have the capability to monitor specific organic compounds. The method detection limit for the TOCA is 475 μg L−1 of carbon. The required detection limit for each of the target analytes by the DART-ETV-MS method was calculated as the concentration of that compound that is equivalent to a total organic carbon (TOC) concentration of 475 μg L−1. The DART-ETV-MS system would thus be complementary to the TOCA in that 9904

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(10) Castro, V. A.; Thrasher, A. N.; Healy, M.; Ott, C. M.; Pierson, D. L. Microb. Ecol. 2004, 47, 119−126. (11) Straub, J. E., II; Plumlee, D. K.; Schultz, J. R. In 40th International Conference on Environmental Systems, Barcelona, Spain, 2010. (12) Straub, J. E., II; Plumlee, D. K.; Schultz, J. R.; McCoy, J. T. In 41st International Conference on Environmental Systems; American Institute of Aeronautics and Astronautics, 2011. (13) Straub, J. E., II; Rutz, J.; Kuo, C. M.; Cole, H.; Manuel, S.; Curtis, M.; Patrick, J.; Sparkman, O. D.; McCoy, J. T. In 41st International Conference on Environmental Systems; American Institute of Aeronautics and Astronautics, 2011. (14) Clements, A. L.; Stinson, R. G.; Van Wie, M.; Warren, E. SAE Int. J. Aerospace 2011, 4, 219−226. (15) Gazda, D. B.; Schultz, J. R.; Siperko, L. M.; Porter, M. D.; Lipert, R. J.; Carrizales-Flint, S. M.; McCoy, J. T. In 41st International Conference on Environmental Systems; American Institute of Aeronautics and Astronautics, 2011. (16) Macatangay, A. V.; Townsend, S. M.; Prokhorov, K. S. SAE Int. J. Aerospace 2009, 1, 420−428. (17) Granahan, J. E.; Thoresen, S. M. SAE Int. J. Aerospace 2009, 1, 25−32. (18) Guharay, S. K.; Dwivedi, P.; Hill, H. H. IEEE Trans. Plasma Sci. 2008, 36, 1458−1470. (19) Harris, G. A.; Nyadong, L.; Fernandez, F. M. Analyst 2008, 133, 1297−1301. (20) Monge, M. E.; Harris, G. A.; Dwivedi, P.; Fernandez, F. M. Chem. Rev. 2013, 113, 2269−2308. (21) Cody, R. B.; Laramee, J. A.; Durst, H. D. Anal. Chem. 2005, 77, 2297−302. (22) Horning, E. C.; Horning, M. G.; Carroll, D. I.; Dzidic, I.; Stillwel, R. N. Anal. Chem. 1973, 45, 936−943. (23) Chernetsova, E. S.; Bochkov, P. O.; Ovcharov, M. V.; Zhokhov, S. S.; Abramovich, R. A. Drug Testing Anal. 2010, 2, 292−294. (24) Kim, H. J.; Jee, E. H.; Ahn, K. S.; Choi, H. S.; Jang, Y. P. Arch. Pharmacal Res. 2010, 33, 1355−1359. (25) Zhou, M.; McDonald, J. F.; Fernandez, F. M. J. Am. Soc. Mass Spectrom. 2010, 21, 68−75. (26) Domin, M. A.; Steinberg, B. D.; Quimby, J. M.; Smith, N. J.; Greene, A. K.; Scott, L. T. Analyst 2010, 135, 700−704. (27) Cody, R. B.; Laramee, J. A. JEOL USA; 2005. (28) Cho, G.; Lim, H.; Kim, J. H.; Jin, D. J.; Kwon, G. C.; Choi, E. H.; Uhm, H. S. IEEE Trans. Plasma Sci. 2011, 39, 1234−1238. (29) Harris, G. A.; Kwasnik, M.; Fernandez, F. M. Anal. Chem. 2011, 83, 1908−1915. (30) Perez, J. J.; Harris, G. A.; Chipuk, J. E.; Brodbelt, J. S.; Green, M. D.; Hampton, C. Y.; Fernandez, F. M. Analyst 2010, 135, 712−719. (31) Harris, G. A.; Hostetler, D. M.; Hampton, C. Y.; Fernandez, F. M. J. Am. Soc. Mass Spectrom. 2010, 21, 855−863. (32) Cody, R. B. Anal. Chem. 2009, 81, 1101−1107. (33) Hunt, D. F.; Stafford, G. C.; Crow, F. W.; Russell, J. W. Anal. Chem. 1976, 48, 2098−2104. (34) Shi, H.; Cloutier, P.; Gamache, J.; Sanche, L. Surf. Sci. 1997, 380, 385−396. (35) National Institute of Standards and Technology. http:// webbook.nist.gov/chemistry/. (36) Jacox, M. E. Chem. Soc. Rev. 2002, 31, 108−115. (37) Snow, T. P.; Le Page, V.; Keheyan, Y.; Bierbaum, V. M. Nature 1998, 391, 259−260. (38) National Research Council. Spacecraft Maximum Allowable Concentrations for Selected Airborne Contaminants, Vol. 2; The National Academies Press: Washington, DC, 1996. (39) National Research Council. Spacecraft Maximum Allowable Concentrations for Selected Airborne Contaminants, Vol. 3; The National Academies Press: Washington, DC, 1996. (40) National Research Council. Spacecraft Water Exposure Guidelines for Selected Contaminants, Vol. 2; The National Academies Press: Washington, DC, 2007.

those reached with the current DART-ETV-MS setup by minimizing sample dilution by the ion source working gas.

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CONCLUSIONS We here introduce a proof-of-principle electro-thermal vaporizer coupled to a plasma ionizer as a sampling and ionization unit for environmental monitoring applications to quantify nmol (or lower) amounts of low-molecular-weight organics in both positive and negative ion detection modes. With the current setup, protonated forms of most analytes were detected, indicating that the major process of ionization is protontransfer reactions. On the other hand, acetaldehyde and FM were mainly detected as molecular ions. These ions were most likely generated through the Penning ionization mechanism between N2 metastables and neutral analytes. It was evident that the mass spectrometer inlet design and parameter settings play a major role in determining the type of ionic species detected. Under identical DART conditions, molecular ions were primarily observed in a pinhole-inlet mass spectrometer under ion lens settings that resulted in minimum-energy collisions, whereas protonated ions were observed at higher lens potentials. The findings presented in this article clearly demonstrate the suitability of plasma ion sources for direct ionization and quantitation of low-molecular-weight organic volatiles in water samples. These experiments were conducted as part of a feasibility assessment, and additional studies are now underway. These studies will include efforts to miniaturize the plasma source and develop a prototype ionization system amenable for use in the spacecraft environment.

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: 404 385 4432. Fax: 404 385 3399. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by NASA Award Number NNX11AR50G to F.M.F.. REFERENCES

(1) Chary, N. S.; Fernandez-Alba, A. R. TrAC, Trends Anal. Chem. 2012, 32, 60−75. (2) Richardson, S. D. Anal. Chem. 2006, 78, 4021−4046. (3) Richardson, S. D. Anal. Chem. 2008, 80, 4373−4402. (4) Wang, Q.; O’Reilly, J.; Pawliszyn, J. J. Chromatogr., A 2005, 1071, 147−154. (5) Saison, D.; De Schutter, D. P.; Delvaux, F.; Delvaux, F. R. J. Chromatogr., A 2009, 1216, 5061−5068. (6) Kiba, N.; Sun, L.; Yokose, S.; Kazue, M. T.; Suzuki, T. T. Anal. Chim. Acta 1999, 378, 169−175. (7) Al Rayes, L.; Saliba, C. O.; Ghanem, A.; Randon, J. Food Addit. Contam., Part B 2012, 5, 221−227. (8) Nishikawa, H.; Sakai, T. J. Chromatogr., A 1995, 710, 159−165. (9) Cancho, B.; Ventura, F.; Galceran, M. T. J. Chromatogr., A 2002, 943, 1−13. 9905

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(41) National Research Council. Spacecraft Water Exposure Guidelines for Selected Contaminants, Vol. 3; The National Academies Press: Washington, DC, 2008. (42) Navare, A.; Nouzova, M.; Noriega, F. G.; Hernandez-Martinez, S.; Menzel, C.; Fernandez, F. M. Rapid Commun. Mass Spectrom. 2009, 23, 477−486. (43) Symonds, J. M.; Galhena, A. S.; Fernandez, F. M.; Orlando, T. M. Anal. Chem. 2010, 82, 621−627.

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