Open Port Sampling Interface System for

Article Cite This: Anal. Chem. XXXX, XXX, XXX-XXX

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Combined Falling Drop/Open Port Sampling Interface System for Automated Flow Injection Mass Spectrometry Gary J. Van Berkel,*,† Vilmos Kertesz,*,† Matt Orcutt,‡ Adam Bentley,§ Jim Glick,§ and Jimmy Flarakos§ †

Mass Spectrometry and Laser Spectroscopy Group, Chemical Sciences Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee 37831, United States ‡ Resolution Laboratories, New Haven, Indiana 46745, United States § Novartis Institute for Biomedical Sciences Drug Metabolism & Pharmacokinetics, East Hanover, New Jersey 07936, United States S Supporting Information *

ABSTRACT: The aim of this work was to demonstrate and to evaluate the analytical performance of a combined falling drop/open port sampling interface (OPSI) system as a simple noncontact, no-carryover, automated system for flow injection analysis with mass spectrometry. The falling sample drops were introduced into the OPSI using a widely available autosampler platform utilizing low cost disposable pipet tips and conventional disposable microtiter well plates. The volume of the drops that fell onto the OPSI was in the 7−15 μL range with an injected sample volume of several hundred nanoliters. Sample drop height, positioning of the internal capillary on the sampling end of the probe, and carrier solvent flow rate were optimized for maximum signal. Sample throughput, signal reproducibility, matrix effects, and quantitative analysis capability of the system were established using the drug molecule propranolol and its isotope labeled internal standard in water, unprocessed river water and two commercially available buffer matrices. A sample-to-sample throughput of ∼45 s with a ∼4.5 s base-to-base flow injection peak profile was obtained in these experiments. In addition, quantitation with minimally processed rat plasma samples was demonstrated with three different statin drugs (atorvastatin, rosuvastatin, and fluvastatin). Direct characterization capability of unprocessed samples was demonstrated by the analysis of neat vegetable oils. Employing the autosampler system for spatially resolved liquid extraction surface sampling exemplified by the analysis of propranolol and its hydroxypropranolol glucuronide phase II metabolites from a rat thin tissue section was also illustrated.

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liquid vortex capture (LVC) probe with an OPSI-like design was placed below the sample to collect the particulates6−8 or whole sample microdissections8,9 formed from laser ablation. The soluble species were extracted from the captured material into the flowing solvent during transport to the ESI or APCI source and subsequent mass spectral analysis. Most notably, an in-house developed system utilizing a commercial laser microdissection (LMD) system for laser ablation was demonstrated for sub-micrometer resolution MS-based imaging of a blended polymer thin film7 and spatially resolved (20−40 μm) absolute quantitation of a targeted pharmaceutical.9 More recently, we have demonstrated the use of the OPSI in the noncontact capture and analysis of nanoliter volume droplets of analyte solution dispensed from specialized microtiter plate wells.10 Analysis of individual nanoliter volume droplets is a means to limit sample consumption, transfer all the sample material into an ionization source more efficiently, and increase sample analysis throughput. The device used in that work was the commercially available Immediate Drop on Demand Technology (I-DOT).11−13 The I-DOT provides fully automated noncontact dispensing of liquid samples from the individual wells of a specifically designed 96-well microtiter

e recently introduced and demonstrated an open port sampling interface (OPSI) as a simple, versatile, and self-cleaning system to rapidly introduce multiple types of samples into a solvent flow stream for subsequent ionization and analysis by mass spectrometry (MS).1,2 This sampling and rapid flow injection analysis (FIA) system3,4 makes use of a vertically aligned, continuous-flow, coaxial-tube sampling probe. Simply touching a sample to the liquid at the top of the OPSI introduces the sample into a flowing solvent stream that transports the material directly to the commercial ionization source of the mass spectrometer (e.g., atmospheric pressure chemical ionization (APCI) or electrospray ionization (ESI)). From there the resultant ions are drawn into the AP sampling inlet of the mass spectrometer and mass analyzed. With these “contact sampling” approaches, the OPSI has been shown to provide the means to manually introduce and quickly analyze unprocessed solid or liquid samples like plastics, ballpoint and felt tip ink pens, skin, and vegetable oils.1,2 In addition, Liu and Pawliszyn demonstrated the use of the OPSI (open port probe (OPP) in their terminology) as a robust interface to couple biocompatible solid phase microextraction (Bio-SPME) fibers with MS.5 What might be considered automated and “non-contact” sampling can also be performed with this basic OPSI probe design. One such use has been the capture and mass spectral analysis of laser ablated samples.6−9 In these experiments a © XXXX American Chemical Society

Received: September 22, 2017 Accepted: October 27, 2017

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DOI: 10.1021/acs.analchem.7b03899 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry

Figure 1. Schematic illustration of falling drop/OPSI-MS system showing (a) the pipet tip mandrel connecting to the 100 μL autosampler syringe with a Teflon tubing; (b) mandrel picking up a disposable pipet tip by pushing the mandrel into a pipet tip; (c) lowering the pipet tip into a selected sample well of a well plate and aspirating a given volume (∼10−20 μL) of sample solution into the pipet tip by drawing air into the syringe; (d) the pipet tip is positioned directly above the vertically aligned OPSI dispensing a single droplet of solution into the OPSI by pushing air into the pipet tip; and (e) the sample traveling through the inner capillary of the probe to the ESI or APCI source, where the sample material is ionized, and then mass analyzed (FIA peak with illustrative mass spectrum in the inset).

plate vertically down toward specified locations on a target plate. Short gas (nitrogen) pulses are applied to the fluid containing well releasing a defined volume of liquid in the nanoliter range through a small diameter orifice in the bottom of the well. The I-DOT system had been used for several noncontact droplet dispensing applications as reported in the literature,11−13 but had never before been coupled directly with a mass spectrometer. The I-DOT system was shown using moderate molecular weight pharmaceutical compounds to be a means to provide individual analyte containing drops for single transient analyses or a train of drops providing a steady state continuous infusion like signal for extended time sample analysis. Maximum sample analysis throughput was determined to be as fast as 5 s per sample. Low nanoliter volumes of nanomolar analyte solutions provided adequate signal levels for either qualitative analysis (e.g., analyte identification) or quantitation using isotope labeled internal standards. The ability to analyze macromolecules like the protein cytochrome c was also demonstrated.10 A very simple, reproducible, noncontact drop based sample injection method, namely, the “falling drop interface”, was introduced 20 years ago by Liu and Dasgupta for capillary zone electrophoresis (CZE).14 In this approach, a sample drop falls from a capillary attached to a conventional syringe directly to a CZE capillary inlet resulting in repeatable low nanoliter volume injection into that CZE capillary. In the present report, we build on this basic falling drop interface concept combining it with the OPSI to provide a simple noncontact, no-carryover, automated system for FIA with MS. A widely available autosampler platform using low cost disposable pipet tips and conventional disposable microtiter well plates was employed to deliver samples drops to the OPSI. The volume of the drops that fell onto the OPSI was in the 7−15 μL range resulting in injected sample volumes of several hundred nanoliters. Sample throughput, signal reproducibility, matrix effects and quantitative analysis capability of the system are established using small molecule drugs and their isotope labeled internal standards in

various commercially available solvent matrices. Quantitation with minimally prepared samples was demonstrated with statins in rat plasma and the direct characterization capability of unprocessed samples was demonstrated by the analysis of neat vegetable oils. We also show that the autosampler system can be used for spatially resolved liquid extraction surface sampling that is exemplified by the analysis of a drug and its metabolite from a rat thin tissue section.

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EXPERIMENTAL SECTION Chemicals, Materials, and Sample Preparation. LC-MS Chromasolv grade water, acetonitrile and acetonitrile with 0.1% formic acid by volume, methanol and methanol with 0.1% formic acid by volume, chloroform (Chromasolv Plus for HPLC), and ACS grade dimethyl sulfoxide (DMSO) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Vegetable oils were purchased locally. Rat plasma, K2EDTA was purchased from BioreclamationIVT (Hicksville, NY, USA). Water from the Clinch River (Oak Ridge, TN, USA) was sampled locally just below the water surface into a polyethylene bottle and used without further preparation. Hanks’ balanced salt solution (HBSS)15 1×, with calcium and magnesium, without phenol red (0.14 g/L CaCl2 (anhydrous (anh.)), 0.4 g/ L KCl, 0.06 g/L KH2PO4, 0.0977 g/L MgSO4 (anh.), 8.00 g/L NaCl, 0.0477 g/L Na2HPO4 (anh.), 0.35 g/L NaHCO3 and 1.00 g/L D-glucose in water) was obtained from Mediatech, Inc. (Herndon, VA, USA). Phosphate buffered saline (PBS) tablets were obtained from Fisher Scientific (Fair Lawn, NJ, USA) and PBS buffer was prepared per instructions resulting in a pH = 7.4, 0.01 M aqueous phosphate buffer containing 0.0027 M KCl and 0.137 M NaCl. Propranolol and propranolol-d7 were obtained from Sigma-Aldrich. Atorvastatin, rosuvastatin, and fluvastatin, and deuterated stable isotope label (SIL) internal standards atorvastatin-d5, rosuvastatin-d6, and fluvastatin-d7 were obtained from Toronto Research Chemicals (Toronto, ON, Canada). Scheme S1 shows structures and mass-to-charge ratios of these compounds. Details of the preparation of stock, B


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parts with larger internal diameters (530 and 150 μm, respectively). The ion source nebulizing gas (nitrogen) was set to its maximum level (i.e., GS1 = 90 in the Analyst software) creating a venturi pump/effect that aspirated solvent from the sampling end of the OPSI through the inner tube into the ionization source. The total internal volume of the PEEK transfer capillary plus the emitter capillaries was ∼8.6 and 6.9 μL for the ESI and APCI setups, respectively. Ion source settings and detection conditions for specific analytes are given in the main text.

standard, and quality control samples of propranolol and the statins in different matrices are detailed in the Supporting Information. Thin tissue sections for liquid extraction surface sampling were prepared from a male rat (HanWistar, 150−225 g) dosed orally at a level of 40 mg/kg with propranolol. Details of the dosing and tissue preparation can be found in the Supporting Information. Experimental Setup. Figure 1 is an illustration of the overall autosampler/OPSI system and its operation in liquid sampling mode. A main component of the setup was an HTS PAL autosampler (CTC Analytics AG, Zwingen, Switzerland) equipped with an air displacement disposable pipet tip mandrel, a sample plate holder, and a pipet tip rack (see Figure S1 for a photograph of the system). The pipet tip mandrel was connected to the 100 μL autosampler syringe with a 13.5 cm long, 1/16” o.d., 254 μm i.d. Teflon tubing that penetrated ∼1 cm into a 7.7 cm long, 1/16” diameter hole drilled through the mandrel (Figure 1a). An automated analysis with the system started with the pickup of a disposable pipet tip on the mandrel by pushing the mandrel into a racked pipet tip (Figure 1b). This was followed by lowering the pipet tip into a selected sample well of a well plate and aspirating a volume of sample solution (∼10−20 μL) into the pipet tip by drawing air into the syringe (by moving the syringe plunger up, not shown) (Figure 1c). The sample containing pipet tip was then positioned at a given distance (∼2 mm in general) directly above the vertically aligned OPSI. The syringe plunger pushed air into the pipet tip to dispense a single droplet of solution that fell onto the OPSI (Figure 1d). A small fraction of the total sample droplet volume was injected in the flow stream of the OPSI, and traveled through the inner capillary of the probe to the ESI or APCI source, where the sample material was ionized, and then mass analyzed (Figure 1e). The used tip was ejected into a waste tip bin (not shown) and the process repeated with the same or a new sample. The autosampler was controlled via the regular CTC PAL autosampler software package (FlexSampler by Resolution Laboratories) and custom methods were created using the regular CTC autosampler commands (atoms). The current OPSI probe coaxial tube sampling end design was similar to one version we have described previously.1,2 The outer tube (304 stainless steel, 0.84 mm i.d. × 1.28 mm o.d. × ∼9 cm long, Grainger, Lake Forest, IL, USA) was tapered to the inner diameter and connected to the mass spectrometer electrical ground. The inner tube was a PEEK capillary (0.18 mm i.d. × 0.794 mm o.d. × ∼20 cm long, IDEX Health & Science, Lake Forest, IL, USA). Solvent was delivered by an HPLC pump (model 1100, Agilent Technologies, Santa Clara, CA, USA) into the annulus region of the two tubes where it flowed to the top of the vertically mounted probe. When the solvent reached the top of the tubes it was aspirated down the inner capillary, which exited the manifold through the second horizontal port, into the Turbo V ion source of a TripleTOF 5600+ (Sciex, Concord, Ontario, Canada). Solvent flow in excess of that which could be aspirated poured out over the top washing the outside of the outer capillary on the way down into a waste basin. The OPSI and wash basin were mounted to the arm that normally held the PAL system wash ports. The autosampler system was positioned on top of the mass spectrometer so the OPSI was in near vertical alignment with the ESI/APCI probe on the mass spectrometer. To increase the maximum self-aspiration flow rate of the system, the nebulizer and emitter capillaries (375 and 100 μm i.d., respectively) of the standard ESI and APCI probes were replaced by equivalent

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RESULTS AND DISCUSION Optimization of Operational Parameters. Figure S1 shows a photograph of the instrumental components of this liquid sample drop and analysis system. Figure 1a−e illustrates the sequential steps in the sampling and analysis process. Video S1 shows an actual sampling and analysis process using a neat vegetable oil as the sample of interest. Note that although open wells are shown in the photo and video, standard foil cover well plates were used without issue. For the present application, the OPSI was operated in the solvent spillover mode distinguished by a convex liquid meniscus, or solvent dome, at the sampling end of the probe.1 This mode was achieved by a solvent flow rate into the probe set higher than the aspiration rate of liquid from the probe to the ionization source. The liquid in excess of the aspiration rate spilled over the top of the probe and down the sides of the outer capillary. This mode provided a stable flow rate to the ionization source and a continuous washing of the top surfaces of the probe resulting in a reproducible sample injection and rapid elution peak with minimal tailing. Compared to our other OPSI report,2 the OPSI coaxial capillary configuration used here had a significantly smaller sized annulus (see Figure S2). This was achieved using an outer capillary with both a significantly smaller outer and inner diameter compared to our previous systems. Exploratory studies with the original OPSI capillary configuration showed that the large annular area and resulting volume at the top of the OPSI was a source for peak tailing when a sample drop was dropped onto the probe. This tailing could be overcome by increasing the solvent flow rate into OPSI well beyond the typical solvent flow rates (e.g., ν = 1 mL/min versus a typical ν = 400 μL/min). As solvent flow rate was increased the linear velocity up and out of the annulus also increased, flushing this region and the solvent dome at the top of the OPSI more quickly. However, to minimize solvent consumption, we chose a geometry that physically reduced the annulus size to mitigate this issue at our typical solvent flow rate. To further optimize the operation of the system, the effect of the height from which the droplet was dropped onto the OPSI (hdropping, distance between bottom of the droplet and top of the OPSI solvent dome), the flow rate of solvent (ν) into the OPSI, and the inner capillary positioning within the OPSI (dcapillary) on sample wash through times and peak heights/areas were investigated. For these experiments, ∼15 μL droplets of 50 nM aqueous propranolol solution were dropped into the OPSI using 100/0.1 (v/v) methanol/formic acid as the OPSI solvent. Figure 2a shows a SRM ion current chronogram of propranolol recorded at with hdropping = 1 mm, ν = 400 μL/ min, and dcapillary= −500 μm (capillary recessed by 500 μm inside the outer capillary of the OPSI). The peak profile observed from the wash through of the injected sample exhibited the expected asymmetric flow injection profile with an C


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∼4.5 s base-to-base wash through time. By varying hdropping, we found that the closer the drop to the probe when released from the pipet tip, without actually touching the solvent dome on the OPSI, the larger were both the peak area and peak height recorded (Figure 2b). To ensure that the drop did not touch the OPSI solvent dome before being released from the pipet the minimum hdropping was set at 1 mm. Touching the drop to the OPSI before release results in a completely variable volume injection and is to be avoided. Larger distances resulted in lesser peak heights and areas, while above hdropping = 11 mm there was no change. Larger distances increased the possibility that something might alter the path of the droplet into the probe (e.g., air flows), thus in this regard the closer spacing for sample dropping was considered more robust. With the OPSI configuration and aspiration setting of the ionization source, a solvent flow rate of at least ∼250 μL/min was required to maintain a consistent solvent spillover condition. Solvent flows less than this value might allow the system to periodically slip into a one of the vortex modes1 of operation. No matter the solvent flow rate into the OPSI above this value, the aspiration rate remained relatively constant at an estimated flow of 240 μL/min. At ν = 400 μL/min and above the signals were relatively stable as solvent flow rate increased further (Figure 2c). Thus, for all other work ν = 400 μL/min was chosen. The absolute sample volume injected into the solvent flow stream by dropping a large drop onto the OPSI was estimated for different positioning of the internal capillary on the sampling end of the probe. This estimation was done by dropping a drop of aqueous 50 nM propranolol/50 nM propranolol-d7 solution onto the OPSI from a height above probe solvent dome of about 1 mm (hdropping = 1 mm) at a solvent flow rate into the probe of 400 μL/min (ν = 400 μL/ min) (see Supporting Information for calculation details). The absolute volume injected was estimated to be about (340 ± 19) nL (5.6% relative standard deviation (RSD)) with the coaxial tubes positioned flush (dcapillary = 0 μm) at the sampling end. The injection volume increased as the inner capillary was recessed from dcapillary = −200 μm ((400 ± 24) nL, 6.0% RSD) to dcapillary = −800 μm ((500 ± 55) nL, 11.0% RSD) into the outer capillary. Our typical assembly of the OPSI positioned the inner capillary recessed by ∼500 μm resulting in an injection volume of about 450 nL. That volume was ∼3% of the total ∼15 μL volume dropped onto the OPSI. Most importantly, for any one capillary positioning, the absolute volume injected was found to be reproducible within ∼6−11% RSD as illustrated by the variance of the calculated volumes shown above. Matrix Effects on Analyte Signal Levels and Quantitation. The quantitative analysis capability of this sampling and analysis approach was evaluated using the analyte propranolol and its stable isotope labeled internal standard propranolol-d7. Use of an isotope labeled internal standard addressed potential detrimental effects on quantitation from sample matrices (e.g., ionization suppression) and possible variations in the sample volumes injected into the OPSI. Matrix effects were examined using this analyte/standard pair prepared in HPLC grade water, untreated river water, HBSS and PBS). The selected reaction monitoring (SRM) ion current chronograms for propranolol obtained by dropping a drop of 100 nM propranolol /250 nM propranolol-d7 prepared in each of the four different solvent systems are shown in Figure S3. Analyte peak widths observed were largely unaffected by the nature of the solution matrix, but peak heights were varied.

Figure 2. (a) Extracted ion current chronogram obtained from sampling a drop (∼15 μL) of a 50 nM solution of propranolol in water into the OPSI. Drop was dropped from 1 mm and flow rate was 400 μL/min. Bar graphs showing the (black filled) mass spectral peak area and (gray filled) peak height recorded after dropping a ∼15 μL of a 50 nM solution of propranolol in water into the OPSI (b) from different heights above the OPSI (flow rate was 400 μL/min, inner capillary position ∼−500 μm (recessed); (c) using different solvent flow rates (drop height was 1 mm, inner capillary position ∼−500 μm); and (d) using different inner capillary positions (negative value means recessed inside the outer capillary, flow rate was 400 μL/min, drop height was 1 mm). The OPSI solvent was 100/0.1 (v/v) methanol/formic acid. The m/z 260.1 → 116.1 signal was collected in positive-ion mode ESI with an emitter voltage of +5.5 kV, a collision energy of 25 eV, a declustering potential of 60 V, an accumulation time of 100 ms with a Turbo V heater temperature of 300 °C. D


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Table 1. Nominal (cpropranolol) Concentrations, Precision (prec. %CV, n = 3), and Accuracy (acc., %bias) of Propranolol Quality Control (QC) Samples Prepared in the Different Matrices Based on a Linear Fit of Calibration Standards (STDs) water

river water

HBSS

PBS

cpropranolol (nM)

prec. (% CV)

acc. (% bias)

prec. (% CV)

acc. (% bias)

prec. (% CV)

acc. (% bias)

prec. (% CV)

acc. (% bias)

1.5 3.5 7.5 15

6.4 2.3 −4.5 −7.0

1.4 4.2 1.0 1.0

15.2 4.1 −3.9 −5.6

4.1 7.1 3.1 3.9

−9.7 7.0 3.9 −1.6

32.3 8.2 3.1 6.3

−4.8 −3.1 −2.7 −4.1

22.8 5.0 3.9 5.0

statin drugs in the 10−4000 ng/mL range. After spiking, 100 μL of rat plasma was combined with 400 μL of methanol containing 250 ng/mL internal standard of the corresponding drug, centrifuged, and 200 μL supernatant was collected and analyzed. Important to note that the samples sat in covered wells allowing the successful analysis of these samples with volatile solvent methanol where evaporation might pose an issue using other drop dispensing methods.10 For analysis 3 replicate ∼6.6 μL drops of the methanolic samples were dropped onto the OPSI at 8 standard (STD) (0, 10, 20, 50, 100, 250, 500, 1000, and 4000 ng/mL) and 4 quality control (QC) (30, 200, 750, and 3000 ng/mL) concentrations. The resulting individual calibration curves and corresponding equations are shown in the Figure S5. Table 2 lists the calculated accuracy and %CV values for the QC samples. The best performance was obtained for

Peak heights for the samples prepared in HPLC grade water and river water were virtually identical. However, signal levels for the sample prepared in the high salt HBSS and PBS were suppressed by ∼75−80%. Signal suppression in such matrices was not unexpected. Importantly, quantitation could still be achieved albeit with poorer detection limits than for the samples without added matrices. The accuracy and precision of the quantitative analysis of propranolol in each of these solvent systems was determined. Data obtained from three dispenses at propranolol concentrations of 0, 1, 2, 5, 10, and 20 nM were selected to be part of a standard (STD) data set, while data obtained from an additional 3 dispenses of 1.5, 3.5, 7.5, and 15 nM propranolol solutions were part of the quality control (QC) sample set. The calibration curves constructed using the ratio of background corrected integrated SRM signal of propranolol and that of the internal standard propranolol-d7 (Apropranolol/Apropranolol‑d7) for the STD data points (n = 3, black filled circles) as a function of propranolol concentration (cpropranolol) are plotted for each solvent system in Figure S4 along with the corresponding equations. In all cases, the coefficient of determination, r2 > 0.992, was considered reasonable as propranolol was being detected with a more than adequate signal-to-noise ratio even at the lowest (1 nM) propranolol concentration (∼10 (buffers) to ∼100 (water and river water matrices)). The averaged data (magenta filled squares) and associated error bars obtained from the separate analysis of the QC samples (n = 3) are also plotted in Figure S4. As summarized in Table 1, the precision of measurements (i.e., %CV) was better than ∼6% for all samples made using HPLC grade water, and %CV decreased in the order of HBSS > PBS > river water matrix. The back calculated concentrations for the QC samples generally showed an accuracy within 10% of the true value (15.2% for the 1.5 nM river water sample). From the linear calibration curves, limit of quantitation (LOQ) was estimated (10 sx/y/slope, where sx/y, the standard error of the y value estimates, is assumed to approximate the standard deviation of the blank, sblank).16 These LOQ values were about 0.1 nM for both the water and river water samples, and increased to ∼0.3 nM and ∼0.7 nM using HBSS and PBS matrices, respectively. Quantitation of Statins from Rat Plasma. “Statins” are a class of drugs that lower the level of cholesterol in the blood.17 They reduce the production of cholesterol by the liver by blocking the hydroxy-methylglutaryl-coenzyme A reductase (HMG-CoA reductase) enzyme that is responsible for making cholesterol.18 Here we demonstrate the quantitative determination of three different statins in a rat plasma sample with minimal sample preparation prior to analysis. The three statins used here were atorvastatin, rosuvastatin and fluvastatin along with their deuterated internal standards atorvastatin-d5, rosuvastatin-d6, and fluvastatin-d7, respectively. Rat plasma samples were spiked individually with the different

Table 2. Nominal (cstatin) Concentrations, Precision (prec. % CV, n = 3), and Accuracy (acc., % bias) of Given Statin Quality Control (QC) Samples Prepared in the Different Matrices Based on a Linear Fit of Calibration Standards (STDs) atorvastatin

rosuvastatin

fluvastatin

cstatin (ng/mL)

prec. (% CV)

acc. (% bias)

prec. (% CV)

acc. (% bias)

prec. (% CV)

acc. (% bias)

30 200 750 3000

8.2 −3.3 −2.4 8.2

1.1 3.8 6.0 4.3

5.8 −1.4 −3.4 −1.1

26.5 1.1 1.8 4.1

26.8 −0.9 −3.3 6.1

16.9 11.4 6.4 8.6

atorvastatin, where the accuracy and %CV of the quantitation were better than 6% and 8.2%, respectively, down to the 30 ng/ mL level. For the other two statins, the accuracy and %CV values were also acceptable at or above 200 ng/mL. These performance metrics worsened for the 30 ng/mL samples of these two drugs with accuracy and %CV values being slightly over 25%. In addition, the overall statistics on rosuvastatin were better than that of fluvastatin. This trend could be understood by close inspection of the data showing signal-to-noise (S/N) ratios decreasing in the order of S/Natorvastatin > S/Nrosuvastatin > S/Nfluvastatin at the same concentration. In summary, even with the minimal sample preparation utilized here, the analysis method provided the required accuracy and precision within internationally recognized acceptance criteria for assay validations19 down to 30 ng/mL for atorvastatin and 200 ng/ mL for rosuvastatin and fluvastatin. Direct Characterization of Vegetable Oils. An increasing need to characterize and authenticate various vegetable oils has encouraged the pursuit of relatively rapid and simple analytical methods to do so.20 For determining vegetable oils triacylgyceride (TAG) distributions these analytical methods have included, among other approaches, the use of DARTE


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Analytical Chemistry MS,21 DESI-MS,22 and easy ambient sonic spray ionization (EASI)-MS.23 These approaches typically require either a sample dilution or placing a droplet of the oil on a surface to be positioned precisely in the ionization source region for analysis. In previous work with the OPSI, the ability to analyze and differentiate various vegetable oils was demonstrated.1,2 In that case, the sampling of the vegetable oils was carried out by first dipping the closed end of a melting point capillary in a respective oil. The oil was wiped from the end of the capillary with a tissue and then the end of the capillary touched to the liquid dome of OPSI for about 1 s. Though simple, fast, and effective, this OPSI method involved a manual contact sampling procedure that might not be considered optimal for work requiring a heavy sample load or work requiring multiple replicate analyses. In the present case, we illustrate the use of the drop capture method with the OPSI as an automated alternative means to analyze neat liquid vegetable oils. For illustration, Figure 3a shows the positive ion mode APCI extracted ion current of the most abundant peak m/z 599.5 (a diacylglyceride (DAG)) from sampling a drop of grape seed oil. The averaged, background subtracted full scan mass spectrum from this sampling peak is shown in Figure 3b. Because of the nonpolar

nature of these oils, the solvent system used was 50/50 (v/v) methanol/chloroform to improve dissolution and the time to wash the sample through the system. Washout was completed within about 5 s. Successive sampling of the same oil showed very consistent signal levels and mass spectra (Figure S6), which matched well with APCI-MS spectra of grape seed oil that appear in the literature.24 The main TAGs in grape seed oil correspond to a series of C55 species the most abundant being the trilinoleic acid triglyceride (LLL) observed as the protonated molecule at m/z 879.7 ((C57H98O6 + H)+, m/z (calcd) = 879.7, see LLL structure in Scheme S1) in the zoomed in region of m/z 878− 888 of the full mass spectra in Figure 3c.24,25 Peaks consistent with the protonated molecules of a more minor C55 series of TAGs were also observed in Figure 3b including the linoleic− palmitic−linoleic acid triglyceride (LPL) observed at m/z 855.7 ((C55H98O6 + H)+, m/z (calcd) = 855.7, see LPL structure in Scheme S1). Also observed, as typical for APCI-MS of these types of molecules,26 are diacylglyceride (DAG) (m/z 550− 650) and monoacylglyceride (MAG) and other fragments (m/z 250−350). The mass spectra of vegetable oils in APCI-MS differ in appearance because of the different TAGs present in the sample, but also because of the differences in fragmentation that occurs, which depend on the degree of unsaturation in the TAG fatty acid chains. To further illustrate this difference, the mass spectra of all nine different vegetable oils examined here and their description can be found in Figure S7 and Table S1, respectively. Liquid Extraction Surface Sampling. Direct liquid extraction based surface sampling probes27,28 are used to reconstitute or extract an analyte from a defined spatial location on a surface by contacting that surface with a confined liquid stream or a wall-less liquid microjunction. The extract is then transferred to an ionization source with subsequent mass analysis.29−32 In the one commercial implementation of these type of devices, termed Liquid Extraction Surface Analysis (LESA), a robotic pipettor is employed to both form and withdraw a liquid microjunction for sampling from a surface.29 The special conductive pipet tip is then engaged with a chipbased nanoESI emitter for a continuous infusion nanoESI mass spectrometric analysis of the extract. A one use pipet tip and one use nanoESI emitter for each sample eliminates sample to sample carryover, but results in a relatively high consumable cost for each measurement. This same basic surface sampling process was repeated here with the autosampler, falling drop/ OPSI system using conventional disposable pipet tips. In this case the sample appears as a rapid flow injection peak, rather than an extended continuous infusion. As such the present methodology is more suited for a targeted analysis rather than discovery, but the same advantages of eliminating sample to sample carryover exist without the high sample to sample consumable cost. The steps of this surface sampling and analysis methodology are shown schematically in Figure S8 and an actual analysis is shown in Video S2. Briefly, after securing a pipet tip on the mandrel (as shown in Figure 1b above), 20 μL of extraction solvent was aspirated into the tip. The pipet tip was then positioned ∼200 μm above the surface location to be sampled and 2 μL of extraction solvent was dispensed onto the sample, while maintaining a solvent connection between the surface and the pipet tip. After a 1 s extraction period, the dissolved sample was aspirated back into the tip. Two dispense/aspirate cycles were executed. After that, the sample in the pipet tip was

Figure 3. (a) Extracted ion current chronogram of m/z 599.5 (a diacylglyceride (DAG), most abundant peak) from the sampling of grape seed oil with the OPSI. (b) Averaged, background subtracted full scan mass spectrum from the sampling in panel a. (c) Zoomed in region (m/z 878−888) of mass spectrum in panel b. The oil sample was dropped from 1 mm, flow rate was 400 μL/min with an inner capillary position of ∼−500 μm (recessed). The OPSI solvent was 1/1 (v/v) methanol/chloroform. Positive ion mode APCI, corona discharge current = 3 μA, declustering potential = 100 V, accumulation time = 250 ms, Turbo V heater temperature = 400 °C. F


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Figure 4. (a) Photograph of a part of the 3” × 4” propranolol dosed rat (40 mg/kg, oral gavage, sacrificed 2 h after dose) whole-body thin tissue section showing six sampled propranolol spiked blood spots (0, 1, 5, 10, 50, and 100 ng/mL, respectively) and four discrete points on the tissue section (liver, lung, kidney and testis) analyzed using the autosampler/OPSI system. (b) Ratio of background corrected integrated SRM signal of propranolol and that of the internal standard propranolol-d7 (Apropranolol/Apropranolol‑d7) as a function of spiked propranolol concentration (cpropranolol). (c) Integrated peak area recorded for (filled bar) propranolol and for (empty bar) hydroxypropranolol glucuronide for the various organs. Extraction solvent was 50/50/0.1 (v/v/v) ACN/water/formic acid with 250 nM propranolol-d7. Extraction solvent volume was 2 μL using two extraction cycles. The extracted sample was dropped from 1 mm, flow rate was 400 μL/min with an inner capillary position of ∼−500 μm (recessed). The OPSI solvent was 100/0.1 (v/v) methanol/formic acid. The m/z 260.1 → 116.1 (propranolol), m/z 267.1 → 116.1 (propranolol-d7) and m/z 452.1 → 276.1 (hydroxypropranolol glucuronides) signals were collected in positive-ion mode ESI with an emitter voltage of +5.5 kV, a declustering potential of 60 V, an accumulation time of 100 ms with a Turbo V heater temperature of 300 °C using collision energies of 25, 25, and 35 eV, respectively.

positioned above the OPSI and a drop of the extract dispensed onto the OPSI (as illustrated on Figure 1d). The surface sampled in this case was a portion of a wholebody thin tissue section of a rat that had been administered propranolol. To enable the quantitative analysis of this drug the extraction solvent (50/50/0.1 ACN/water/formic acid) was spiked with 250 nM propranolol-d7. In addition, the thin section contained blood thin sections spiked with 0−100 ng/ mL of the propranolol. SRM chronograms were recorded for propranolol (m/z 260.1 → 116.1), propranolol-d7 (m/z 267.1 → 116.1), and phase II metabolites aliphatic and aromatic hydroxypropranolol glucuronides (see structures in Scheme S1) (both m/z 452.1 → 276.1) from the sampling of the areas marked in the optical image of the tissue (Figure 4a). Figure 4b shows the ratio of background corrected integrated SRM signal of propranolol and that of the internal standard propranolol-d7 (Apropranolol/Apropranolol-d7) as a function of spiked propranolol concentration (cpropranolol) in the blood spots analyzed. A firstorder fit of the samples resulted in a R2 = 0.993 indicating a good linear correlation between concentration and obtained signal suggesting an (at least) semiquantitative performance of this sampling method. Filled and empty bars in Figure 4c show

integrated peak areas collected for dosed drug propranolol and phase II metabolites hydroxypropranolol glucuronides, respectively, for the four different organs (liver, lung, kidney and testis) and the blank blood spot (nonspiked, 0 ng/mL) sampled. The four organs were selected to be sampled due to known accumulation of the drug and/or metabolite in them.29,33 Propranolol was detected above background level in liver, lung and kidney, as expected. In addition, the metabolite was detected in all organs. Signal levels for hydroxypropranolol glucuronide were highest in the liver and kidney. Glucuronidation occurs mainly in the liver, making the drug more water-soluble to enhance subsequent elimination from the body by urination via the kidneys.34 Lower hydroxypropranolol glucuronide signal was observed in the lung and even lower in the testis. The blank blood spot showed no signal for either the parent drug or the metabolites. These general observations are in line with the previously reported liquid extraction based sampling29−31 and QWBA analyses33 of these tissues. G



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CONCLUSIONS

Article

ASSOCIATED CONTENT

S Supporting Information *

In this report, we have demonstrated the falling drop/OPSI combination as a simple and automated noncontact, nocarryover, flow injection mass spectrometry system for liquid analytes and analyte solutions. This data also highlights some of the benefits of this drop injection approach versus of previous IDOT/OPSI combination. (A comparison of the performance metrics of the falling drop/OPSI-MS and I-DOT/OPSI-MS approaches, and conventional HPLC/MS, can be found in Table S2.) The falling sample drops were provided via a widely available autosampler using inexpensive disposable pipet tips and microtiter well plates. While only analysis from a single 96well plate was demonstrated, the present autosampler system has the capability to handle multiple plates, different well plate configurations and foil sealed plates. Optimization of operational parameters for maximum signal and robust, reproducible operation resulted using a 1 mm spacing for sample dropping (bottom of drop to top of the OPSI) at an OPSI solvent flow rate of 400 μL/min with the internal capillary recessed by ∼500 μm. Under these conditions, the volume of the drops that fell onto the OPSI was in the 7 (methanolic solution) to 15 (aqueous solution) μL range with an injected sample volume of about 450 nL. The sample-to-sample throughput was ∼45 s with a ∼4.5 s base-to-base FIA profile. Throughput could be reduced (by 50% or more) by use of a commercial autosampler with, for example, multiple, individually addressable pipettors.35 Signal reproducibility, matrix effects and quantitative analysis capability of the system were established using the drug molecule propranolol and its isotope labeled internal standard in water, unprocessed river water, and HBSS and PBS buffer matrices. Peak heights for the samples prepared in HPLC grade water and river water were virtually identical. However, signal levels for the sample prepared in the high salt HBSS and PBS were suppressed by ∼75−80%. LOQ values of propranolol were ∼0.1 nM for both the water and river water samples, and increased to ∼0.3 nM and ∼0.7 nM using HBSS and PBS matrices, respectively. The precision of measurements was better than ∼6% for all samples made using HPLC grade water, and %CV decreased in the order of HBSS > PBS > River water matrix. The back calculated concentrations for the QC samples generally showed an accuracy within 10% of the true value (15.2% for the 1.5 nM river water sample). Quantitation with minimally processed rat plasma samples was demonstrated with three different statin drugs and the analysis method provided the required accuracy and precision within internationally recognized acceptance criteria for assay validations19 down to 30 ng/mL for atorvastatin and 200 ng/mL for rosuvastatin and fluvastatin. Despite these positive results further evaluation is needed in regard to the matrix tolerance of the falling drop/ OPSI-MS system. The direct characterization capability of unprocessed samples was demonstrated by the analysis of nine neat vegetable oils. A particular example included grape seed oil that showed the expected MAG, DAG, and TAG regions when using APCI-MS. Finally the system was shown to be capable of spatially resolved liquid extraction surface sampling. Both the dosed drug propranolol and its hydroxypropranolol glucuronide phase II metabolites were detected in the various organs of rat thin tissue section at levels in line with the previous liquid extraction based sampling reports for these tissues.

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.7b03899. Chemical structures, sample preparation details, injection volume calculations, table comparing falling drop/OPSIMS and related technologies, table showing vegetable oils used, photograph of the falling drop/OPSI-MS system, OPSI size details, data showing suppression effects and calibration curves, and vegetable oil mass spectra (PDF) Oil sampling video (AVI) Surface sampling video (AVI)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. Phone: 865-574-1922. *E-mail: [email protected]. Phone: 865-574-3469. ORCID

Gary J. Van Berkel: 0000-0001-5224-3969 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The work of G.J.V.B and V.K. on the development of the concept for noncontact liquid sample capture with the OPSI, the assembly of prototype interfaces for that purpose, and droplet capture optimization studies were supported by the United States Department of Energy, Office of Science, Basic Energy Sciences, Chemical Sciences, Geosciences and Biosciences Division. Support of G.J.V.B and V.K. for the particular applications presented, and the loan of the TripleTOF 5600+ mass spectrometer, was provided through a Cooperative Research and Development Agreement (CRADA NFE-1002966) with Sciex.

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REFERENCES

(1) Van Berkel, G. J.; Kertesz, V. Rapid Commun. Mass Spectrom. 2015, 29, 1749−1756. (2) Van Berkel, G. J.; Kertesz, V. Rapid Commun. Mass Spectrom. 2017, 31, 281−291. (3) Fan, H.; Schug, K. A. Spectroscopy 2012, 10, 26−33. (4) Ruzicka, J.; Hansen, E. H. Flow Injection Analysis, 2nd ed.; Wiley: New York, NY, 1988. (5) Gómez-Ríos, G. A.; Liu, C.; Tascon, M.; Reyes-Garces, N.; Arnold, D. W.; Covey, T. R.; Pawliszyn, J. Anal. Chem. 2017, 89, 3805−3809. (6) Ovchinnikova, O. S.; Bhandari, D.; Lorenz, M.; Van Berkel, G. J. Rapid Commun. 2014, 28, 1665−1673. (7) Cahill, J. F.; Kertesz, V.; Van Berkel, G. J. Anal. Chem. 2015, 87, 11113−11121. (8) Cahill, J. F.; Kertesz, V.; Van Berkel, G. J. Rapid Commun. Mass Spectrom. 2016, 30, 611−619. (9) Cahill, J. F.; Kertesz, V.; Weiskittel, T. M.; Vavrek, M.; Freddo, C.; Van Berkel, G. J. Anal. Chem. 2016, 88, 6026−6034. (10) Van Berkel, G. J.; Kertesz, V.; Boeltz, H. Bioanalysis 2017, DOI: 10.4155/bio-2017-0104. (11) Traube, A.; Brode, T. U.S. Patent 8,759,113 B2, June 24, 2014. (12) Arenas da Silva, L. F.; Schober, L.; Sloff, M.; Traube, A.; Hart, M. L.; Feitz, W. F. J.; Stenzl, A. Cytotherapy 2015, 17, 1655−1661. (13) Schober, L.; Büttner, E.; Laske, C.; Traube, A.; Brode, T.; Traube, A. F.; Bauernhansl, T. J. Lab Autom. 2015, 20, 154−163. (14) Liu, H.; Dasgupta, P. K. Anal. Chem. 1997, 69, 1211−1216. (15) Hanks, J. H.; Wallace, R. E. Exp. Biol. Med. 1949, 71, 196−200.

H


Article

Analytical Chemistry (16) Miller, J. C.; Miller, J. N. Statistics for Analytical Chemistry, 2nd ed.; Ellis Horwood, Ltd.: Chichester, 1988; pp 110−115. (17) Chou, R.; Dana, T.; Blazina, I.; Daeges, M.; Jeanne, T. L. J. Am. Med. Assoc. 2016, 316, 2008−2024. (18) Levine, T. B.; Levine, A. B.. Lipids, Atherogenic Dyslipidemia, and Therapy. In Metabolic Syndrome and Cardiovascular Disease, 2nd ed.; Blackwell Publishing, Ltd.: Oxford, UK, 2012; Chapter 13, pp 347−408. (19) Shah, V. P.; Midha, K. K.; Findlay, J. W. A.; Hill, H. M.; Hulse, J. D.; McGilveray, I. J.; McKay, G.; Miller, K. J.; Patnaik, R. N.; Powell, M. L.; Tonelli, A.; Viswanathan, C. T.; Yacobi, A. Pharm. Res. 2000, 17, 1551−1557. (20) Azadmard-Damirchi, S.; Torbati, M. J. Food Quality Hazards Control 2015, 2, 38−44. (21) Vaclavik, L.; Cajka, T.; Hrbek, V.; Hajslova, J. Anal. Chim. Acta 2009, 645, 56−63. (22) Gerbig, S.; Takats, Z. Rapid Commun. Mass Spectrom. 2010, 24, 2186−2192. (23) Simas, R. C.; Catharino, R. R.; Cunha, I. B. S.; Cabral, E. C.; Barrera-Arellano, D.; Eberlin, M. N.; Alberici, R. M. Analyst 2010, 135, 738−744. (24) De Marchi, F.; Seraglia, R.; Molin, L.; Traldi, P.; De Rosso, M.; Panighel, A.; Dalla Vedova, A.; Gardiman, M.; Giust, M.; Flamini, R. J. Mass Spectrom. 2012, 47, 1113−1119. (25) Jakab, A.; Nagy, K.; Héberger, K.; Vékey, K.; Forgács, E. Rapid Commun. Mass Spectrom. 2002, 16, 2291−2297. (26) Neff, W. E.; Byrdwell, W. C.; List, G. R. J. Liq. Chromatogr. Relat. Technol. 2001, 24, 837−854. (27) Laskin, J.; Lanekoff, I. Anal. Chem. 2016, 88, 52−73. (28) Venter, A. R.; Douglass, K. A.; Shelley, J. T.; Hasman, G., Jr.; Honarvar, E. Anal. Chem. 2014, 86, 233−249. (29) Kertesz, V.; Van Berkel, G. J. J. Mass Spectrom. 2010, 45, 252− 260. (30) Van Berkel, G. J.; Kertesz, V. Anal. Chem. 2009, 81, 9146−9152. (31) Kertesz, V.; Van Berkel, G. J. Bioanalysis 2013, 5, 819−826. (32) Van Berkel, G. J.; Kertesz, V. Rapid Commun. Mass Spectrom. 2013, 27, 1329−1334. (33) Kertesz, V.; Van Berkel, G. J.; Vavrek, M.; Koeplinger, K. A.; Schneider, B. B.; Covey, T. R. Anal. Chem. 2008, 80, 5168−5177. (34) Sten, T.; Qvisen, S.; Uutela, P.; Luukkanen, L.; Kostiainen, R.; Finel, M. Drug Metab. Dispos. 2006, 34, 1488−1494. (35) http://lifesciences.tecan.com/products/liquid_handling_and_ automation/freedom_evo_series (accessed August 30, 2017).

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