Liquid Phase Collection Surface Sampling

Feb 17, 2011 - The spatial sampling resolution afforded by the laser ablation, as well as the ability to use sample processing methods like HPLC betwe...
1 downloads 11 Views 2MB Size
Download PDF
LETTER pubs.acs.org/ac

Combining Laser Ablation/Liquid Phase Collection Surface Sampling and High-Performance Liquid Chromatography-Electrospray Ionization-Mass Spectrometry Olga S. Ovchinnikova,†,‡ Vilmos Kertesz,† and Gary J. Van Berkel*,†,‡ †

Organic and Biological Mass Spectrometry Group, Chemical Sciences Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee 37831-6131, United States ‡ Department of Physics and Astronomy, University of Tennessee, Knoxville, Tennessee 37996-1200, United States

bS Supporting Information ABSTRACT: This letter describes the coupling of ambient pressure transmission geometry laser ablation with a liquid phase sample collection method for surface sampling and ionization with subsequent mass spectral analysis. A commercially available autosampler was adapted to produce a liquid droplet at the end of the syringe injection needle while in close proximity to the surface to collect the sample plume produced by laser ablation. The sample collection was followed by either flow injection or a high-performance liquid chromatography (HPLC) separation of the extracted components and detection with electrospray ionization mass spectrometry (ESI-MS). To illustrate the analytical utility of this coupling, thin films of a commercial ink sample containing rhodamine 6G and of mixed isobaric rhodamine B and 6G dyes on glass microscope slides were analyzed. The flow injection and HPLC-ESI-MS analysis revealed successful laser ablation, capture, and with HPLC, the separation of the two compounds. The ablated circular area was about 70 μm in diameter for these experiments. The spatial sampling resolution afforded by the laser ablation, as well as the ability to use sample processing methods like HPLC between the sample collection and ionization steps, makes this combined surface sampling/ionization technique a highly versatile analytical tool.

The coupling of atmospheric pressure (AP) surface sampling and ionization with mass spectrometry (MS) dates back to at least the mid-1970s, but there has been a resurgence in interest, research and use of such techniques since the early 2000s.1-9 Numerous applications have been demonstrated or explored including, among others, the direct identification and sometimes quantification of small molecules, like pharmaceuticals, metabolites, and lipids, and larger molecules like peptides and proteins, from various analytically relevant surfaces including planar separation media,10 biological tissue,11 and surfaces of forensic interest.12 Among these many AP surface/sampling ionization approaches, our group has focused in large part on the use of direct liquid extraction methods that employ variations of what has been termed a liquid microjunction surface sampling probe (LMJ-SSP).1 These probes reconstitute or extract analytes from surfaces into a fluidic junction between the sampling end of the probe and the surface. These probes can be operated either in a continuous sampling mode13 or in a droplet dispensing/retrieving mode.14-16 A unique advantage of the LMJ-SSP approach to surface sampling is the ability, with some of the implementations, to process the materials extracted into solution postsampling. Relative to a direct mass spectrometric analysis, this provides the capacity to eliminate matrix effects or add an additional, r 2011 American Chemical Society

orthogonal dimension of detection selectivity or specificity. The importance of this capability was recently demonstrated by combining the use of the droplet sampling mode with a subsequent HPLC separation and targeted detection of a drug and its isomeric glucuronide metabolites from thin tissue sections of drug dosed mice using an autosampler.17 These isomers were not distinguished from a similar sample using simple MS or MS/MS detection methods.15 One limit of the LMJ-SSP approach is the lack of ability to robustly sample from some wettable and absorbent surfaces.1,18 With such surfaces, the extraction solvent can be drawn out from the probe into the surface and trapped, thus the sampled material cannot be efficiently aspirated back into the probe and analyzed. Another limit of the LMJ-SSP approach is the achievable spatial resolution of the technique, which is restricted by the size of the probe and the fluidic junction to the surface. Typically, reported spatial resolution has been about 400-650 μm for spot samples and lane scans,13,19 with one recent report of a nanoelectrospray version of such a probe achieving a single spot sample approximately 100 μm in diameter.20 One might envision constructing Received: January 7, 2011 Accepted: February 10, 2011 Published: February 17, 2011 1874

dx.doi.org/10.1021/ac200051y | Anal. Chem. 2011, 83, 1874–1878

Analytical Chemistry and using smaller probes than employed at present, but such implementations might be expected to limit the robustness of operation because of simple issues like precise probe positioning relative to the surface and probe plugging. Laser desorption (LD) or ablation (LA) based approaches provide a proven means to sample from a surface at AP with a spatial resolution substantially better than 100 μm and to create in the gas phase intact molecular species for mass analysis.1,6 Theoretically, the laser spot size is limited by the diffraction limit (D) of the light (D ∼ λ/2),21 or when using near field phenomena, to even smaller dimensions.22,23 To date, laserbased AP surface sampling and ionization approaches have reported spatial sampling resolutions from about 50 to 70 μm24 down to as small as a few micrometers.25-27 Like most of the other AP surface sampling/ionization approaches, however, these laser based methods do not provide the opportunity to process for analytical benefit the sampled material prior to ionization and mass analysis. We report here on an AP surface sampling/ionization combination that provides both the higher spatial resolution surface sampling of LA and the postsampling processing capabilities of liquid extraction-based sampling probe methods. This combination also circumvents the need for direct contact between the surface and the liquid extraction solvent, which is expected to Scheme 1. Structure and Mass-to-Charge Ratio for Rhodamine B (1) and Rhodamine 6G (2)

LETTER

eliminate problems in the analysis of unmodified absorbent surfaces that have previously limited liquid extraction-based sampling probe methods. Specifically, a commercially available autosampler was adapted to produce a liquid droplet at the end of the syringe injection needle while in close proximity to the surface to collect the sample plume produced by a vertically aligned transmission geometry LA system. The sample collection was followed by either flow injection or an HPLC separation of the extracted components. In this case, electrospray ionization (ESI) was used. However, other liquid introduction ionization sources like atmospheric pressure chemical ionization (APCI) or atmospheric pressure photoionization (APPI) could be used for analysis of more nonpolar species or potentially inductively coupled plasma (ICP) for elemental analysis of the captured materials.1 Analyses of thin films of a commercial ink sample containing rhodamine 6G (compound 2 in Scheme 1) and of mixed isobaric rhodamine B (compound 1 in Scheme 1) and rhodamine 6G dyes on glass microscope slides (all observed at m/z 443 in ESI-MS) were used to illustrate the analytical utility of this coupling. Details regarding the chemicals and experimental methods used in this work can be found in the Supporting Information. As shown in Figure 1, the vertically aligned transmission geometry laser ablation arrangement used in these experiments allowed for direct on-axis alignment of the laser beam with the also vertically aligned autosampler syringe needle for capture of the ablated material into the suspended liquid droplet. With this geometry, a 0.1 μL volume liquid droplet could be reproducibly produced at the end of the syringe needle and positioned to be about 500 μm above, but never touch, the surface. Furthermore, this transmission geometry provided a platform for the future from which to achieve very high spatial resolution laser ablation (e.g., 1-2 μm) similar, for example, to that possible with many existing laser capture microdissection devices.28-31 However, if neither the highest spatial resolution possible nor postprocessing of the sample are prime objectives, this approach can be used with a reflection geometry laser ablation system or with continuous flow versions of the LMJ-SSP (see the Supporting Information). The ability to capture some or all of the laser ablated sample/ plume into a suspended liquid droplet was first tested in a flow injection mode using a thin film of red Sharpie ink applied to a

Figure 1. (a) Schematic illustration of the experimental setup. (b) Zoomed in region around the sample and the collection syringe needle showing laser ablation of the sample and its consequent collection in the liquid phase. 1875

dx.doi.org/10.1021/ac200051y |Anal. Chem. 2011, 83, 1874–1878

Analytical Chemistry

Figure 2. (a) Total ion current (TIC) chronogram from replicate sampling (S1 and S2), double blank (DB1 and DB2), and blank (B1 and B2) experiments (see text for details). The inset shows the approximately 70 μm-diameter ablated area after the 10 Hz laser was turned on for 15 s with an output energy of 5 μJ/pulse during the sampling process (S1 and S2). (b) Extracted ion chronogram generated using the ion intensity for m/z 443 from the same experiment. The full scan mass spectra shown in panels c, d, and e were obtained by averaging over the time points in the TIC corresponding to the peaks labeled S1, DB1, and B1, respectively. The relative intensity in panels d and e is normalized to the intensity of m/z 443 in panel c. The sampling solution was 80/20/0.1 (v/v/v) ACN/water/FA. The sample was injected into a stream of the same solvent and sprayed at a flow rate of 50 μL/min.

transparent glass microscope slide. This ink contained compound 2, which is observed at m/z 443. Parts a and b of Figure 2 show the total ion current and the m/z 443 extracted ion current profile, respectively, of replicate experiments for flow injection of a 0.1 μL droplet (80/20/0.1 (v/v/v) acetonitrile (ACN)/water/ formic acid (FA)) that had been suspended (0.5 mm) above the ablation region and the 10 Hz laser fired for 15 s (∼150 laser shots, 5 μJ/shot) to ablate the dye from the surface (sample, S1 and S2), for a droplet simply suspended 31.5 mm over the dye surface, but with the laser turned off for the 15 s sampling time (double blank, DB1 and DB2), and for a droplet simply suspended 0.5 mm over the dye surface, but with the laser turned

LETTER

off for the 15 s sampling time (blank, B1 and B2). For each experiment, a different area of the thin film was positioned in the ablation region. Parts c, d, and e of Figure 2 show the corresponding averaged mass spectra for the flow injection peaks observed from the first sample (S1), double blank (DB1), and blank (B1) replicates, respectively. The delay between experiments was set at 2 min to provide sufficient time for complete washout of the injected rhodamine from the system given the solvent and flow rates used. Carryover between experiments was eliminated using the wash cycle of the autosampler to clean the sample collection/injection syringe. The data in Figure 2 clearly demonstrate that laser ablated rhodamine was successfully collected by the hanging liquid droplet and that no direct liquid extraction sampling took place. The mass spectrum obtained from sampling experiment S1 showed m/z 443 as the base peak with a minor abundance peak at m/z 415, the known major product ion from the dissociation of compound 2 (Figure 2c). This minor abundance ion may have been formed by fragmentation in the interface region or by the laser ablation process. Also, the replicate data in Figure 2 show that the sampling and analysis process was reproducible. The spatial resolution of the sampling was determined by the size of the focused laser beam spot on the surface. In this case, as the optical image insert in Figure 2a shows, the diameter of the ablation area was about 70 μm. This ablated area was somewhat smaller than the best spot sampling size reported for direct liquid extraction surface sampling (∼100 μm),20 but significantly larger than the 15 μm spot size that has been demonstrated with transmission geometry matrix assisted laser desorption ionization (MALDI).25 A smaller diameter fiber optic cable and improved laser focusing optics incorporated into the present apparatus would reduce the size of the ablation spot achieved. In addition to the simple flow injection type analysis just discussed, this laser ablation/liquid phase collection combination allowed for further processing of the collected sample before mass spectral analysis. To demonstrate this processing capability, we used a thin film of a mixture of the isobaric compounds 1 and 2 prepared on a glass slide. The sampling procedure was identical to that described above (except for the solvent droplet composition, 50/50/0.1 (v/v/v) ACN/water/FA) and the ablated area was also about 70 μm in diameter. Rather than the performance of a simple flow injection, an Hydro-RP 80A HPLC column (50 mm  2 mm, 4 μm particle size; Phenomenex, Torrance, CA) was positioned between the injector and ESI source and an isocratic separation (50/50/0.1 (v/v/v) ACN/water/FA) of the captured ablated material was carried out at a flow rate of 50 μL/ min. Figure 3a shows the extracted ion chromatogram recorded for m/z 443 from the 8 min HPLC-MS run. Two distinct peaks were observed in the chromatogram at 5.3 and 6.3 min corresponding to compounds 1 and 2, respectively. Using the surface concentration of the dyes (about 6.4 nmol/mm2 for each individual dye) and size of the ablated area (3.85  10-3 mm2), one calculates that about 25 pmol (11 ng) of each dye was ablated from the surface. The amount of that material collected into the drop has not been determined. However, visual observation of the plume of laser ablated material during the experiment made it apparent that not all of the material was captured. As a control experiment, Figure 3b shows the extracted ion chromatogram of m/z 443 under the same conditions except that the laser was not fired when the droplet was in the sampling position. No signal corresponding to the respective dyes was observed in that case. Note that other ions were observed eluting 1876

dx.doi.org/10.1021/ac200051y |Anal. Chem. 2011, 83, 1874–1878

Analytical Chemistry

Figure 3. Extracted ion chromatograms for m/z 443 recorded during a 8 min HPLC-MS run with the laser turned (a) ON and (b) OFF during the sampling process. Measured signal intensities in both figures were normalized to the highest signal in part a. Output fluence of the 10 Hz laser was 5 μJ/pulse ablating an approximately 70 μm-diameter area that corresponded to about 25 pmol (about 11 ng) of both rhodamines B (compound 1) and 6G (compound 2) being removed from the glass slide surface. Sampling and isocratic HPLC eluent solutions were both 50/50/0.1 (v/v/v) ACN/water/FA. Flow rate of the HPLC separation was 50 μL/min.

from the column as peaks in addition to the expected m/z 443. However, those ions observed were the same as those observed in the chromatogram when direct liquid microjunction surface sampling17 was used to sample the same surface. Thus, no significant sample decomposition or alteration due the laser ablation process appeared to have taken place. The data presented here prove the ability to sample analyte from a surface using laser ablation and to capture that ablated material into a suspended liquid droplet which can then be directly analyzed, or further processed, in this case by HPLC, prior to mass spectral analysis. Relative to direct liquid extraction based surface sampling approaches, the present combination offers superior sampling resolution and also avoids issues that can arise when attempting to analyze wettable surfaces. Relative to direct AP laser-based surface sampling methods, this combination provides the ability to process the sample collected prior to mass spectral analysis for enhanced analytical benefit (e.g., separation of isobaric components in a mixture). Work is underway to quantify and improve various analytical figures of merit for this new methodology including capture efficiency of the laser ablated material into the droplet, minimum sample spot size that provides detectable signal, and detection levels for various compound classes. As additional data in the Supporting Information demonstrates, this sampling approach can be used, if desired, with a reflection geometry laser ablation or with a continuous flow LMJSSP. The ability to achieve the best possible spatial sampling resolution may be more difficult to obtain in reflection geometry versus the transmission geometry. Also, the flow probe does not allow as simple a coupling for sample processing as the syringe droplet capture and injection procedure described above. However, because it is continuously sampling and ionizing collected material, it might be used as a means to increase sampling throughput or even be used to provide this technique the ability

LETTER

for chemical imaging. Nonetheless, the fundamentals of the ablation and sample capture process are the same with either laser ablation geometry or sampling probe type. Therefore, analyses that are successful in one configuration should be achievable in another. Thus, in addition to the rhodamine dye data focused on here, we can also report that the proteins cytochrome c and bovine insulin have been sampled and multiply charged, and small fragile molecules like propranolol glucuronide successfully sampled and detected (see the Supporting Information). In general, the laser ablation and liquid phase collection approach to surface sampling can be applied to materials that can be ablated and captured and, for the most part, dissolved in the capture solvent, then effectively ionized with the ionization source in use. Different laser wavelengths, power, pulse widths, and repetition rates and the use of surface chemistries or chemical matrices as in MALDI might be among the techniques used to improve the effectiveness of ablation for different analytes from diverse sample matrices. More nonpolar solvents might be used to capture less polar analytes in conjunction with ionization using APCI or APPI. In cases where elemental analysis is desired, one can imagine the use of acidic aqueous solutions to capture ablated materials, including particulates, which then can be delivered to an ICP for atomization and ionization prior to mass analysis.

’ ASSOCIATED CONTENT

bS

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

’ AUTHOR INFORMATION Corresponding Author

*Phone: 865-574-1922. Fax: 865-576-8559. E-mail: vanberkelgj@ ornl.gov.

’ ACKNOWLEDGMENT Dr. Mariam ElNaggar (ORNL) is thanked for creating the schematics of the experimental setups. Mr. Joe Stankovich (ORNL) is thanked for assistance in acquiring data presented in the Supporting Information. This work was supported by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences, United States Department of Energy. ORNL is managed by UT-Battelle, LLC for the U.S. Department of Energy under Contract DE-AC05-00OR22725. ’ REFERENCES (1) Van Berkel, G. J.; Pasilis, S. P.; Ovchinnikova, O. J. Mass Spectrom. 2008, 43, 1161–1180. (2) Harris, G. A.; Nyadong, L; Fernandez, F. Analyst 2008, 133, 1297–1301. (3) Venter, A.; Nefliu, M.; Cooks, R. G. Trends Anal. Chem. 2008, 27, 284–290. (4) Weston, D. J. Analyst 2010, 135, 661–668. (5) Chen, H.; Gamez, G.; Zenobi, R. J. Am. Soc. Mass Spectrom. 2009, 20, 1947–1963. (6) Huang, M. Z.; Yuan, C. H.; Cheng, S. C.; Cho, Y. T.; Shiea, J. Ann. Rev. Anal Chem. 2010, 3, 43–65. (7) Ifa, D. R.; Wu, C.; Ouyang, Z.; Cooks, R. G. Analyst 2010, 135, 669–681. 1877

dx.doi.org/10.1021/ac200051y |Anal. Chem. 2011, 83, 1874–1878

Analytical Chemistry

LETTER

(8) Alberici, R. M; Simas, R. C.; Sanvido, G. B.; Rom~ao, W.; Lalli, P. M.; Benassi, M.; Cunha, I. B. S.; Eberlin, M. N. Anal. Bioanal. Chem. 2010, 398, 265–294. (9) Huan-Wen, C.; Bin, H.; Xie, Z. Chin. J. Anal. Chem. 2010, 38, 1069–1088. (10) Pasilis, S. P.; Van Berkel, G. J. J. Chromatogr., A 2010, 1217, 3955–3965. (11) Wiseman, J. M.; Ifa, D. R.; Zhu, Y.; Kissinger, C. B.; Manicke, N. E.; Kissinger, P. T.; Cooks, R. G. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 18120–18125. (12) Ifa, D. R.; Jackson, A. U.; Paglia, G.; Cooks, R. G. Anal. Biol. Chem. 2009, 394, 1995–2008. (13) Van Berkel, G. J.; Sanchez, A. D.; Quirke, J. M. E. Anal. Chem. 2002, 74, 6216–6223. (14) Van Berkel, G. J.; Kertesz, V.; King, R. C. Anal. Chem. 2009, 81, 7096–7101. (15) Kertesz, V.; Van Berkel, G. J. J. Mass Spectrom. 2010, 45, 252–260. (16) Marshall, P.; Toteu-Djomte, V.; Bareille, P.; Perry, H.; Brown, G.; Baumert, M.; Biggadike, K. Anal. Chem. 2010, 82, 7787–7794. (17) Kertesz, V.; Van Berkel, G. J. Anal. Chem. 2010, 82, 5917–5921. (18) Walworth, M. J.; Stankovich, J. J.; Van Berkel, G. J.; Schulz, M.; Minarik, S.; Nichols, J.; Reich, E. Anal. Chem. 2011, 83, 591–597. (19) Van Berkel, G. J.; Kertesz, V.; Koeplinger, K. A.; Vavrek, M.; Kong, A.-N. T. J. Mass Spectrom. 2008, 43, 500–508. (20) Roach, P. J.; Laskin, J.; Laskin, A. Analyst 2010, 135, 2233–2236. (21) Hecht, E. Optics, 4th ed.; Addison Wesley: San Fransico, CA, 2002; p 595. (22) Novotny, L.; Stranick, S. J. Annu. Rev. Phys. Chem. 2006, 57, 303–331. (23) Zeisel, D.; Dutoit, B.; Deckert, V.; Roth, T.; Zenobi, R. Anal. Chem. 1997, 69, 749–754. (24) Shrestha, B.; Vertes, A. Anal. Chem. 2009, 81, 8265–8271. (25) Inutan, E. D.; Richards, A. L.; Wager-Miller, J.; Mackie, K.; McEwen, C. N.; Trimpin, S. Mol. Cell. Proteomics 2011, 10, M110.000760. (26) Koestler, M.; Kirsch, D.; Hester, A.; Leisner, A.; Guenther, S.; Spengler, B. Rapid Commun. Mass Spectrom. 2008, 22, 3275–3285. (27) Bradshaw, J. A.; Ovchinnikova, O. S.; Meyer, K. A.; Goeringer, D. E. Rapid Commun. Mass Spectrom. 2009, 23, 3781–3786. (28) Emmert-Buck, M. R.; Bonner, R. F.; Smith, P. D.; Chuaqui, R. F.; Zhuang, Z.; Goldstein, S. R.; Weiss, R. A.; Liotta, L. A. Science 1996, 274, 998–1001. (29) Bonner, R. F.; Emmert-Buck, M. R.; Cole, K.; Pohida, T.; Chuaqui, R. F.; Goldstein, S. R.; Liotta, L. A. Science 1997, 278, 1481–1483. (30) http://www.zeiss.de/microdissection/, accessed January 4, 2011. (31) http://www.leica-microsystems.com/products/light-microscopes/life-science-research/laser-microdissection/, accessed January 4, 2011.

1878

dx.doi.org/10.1021/ac200051y |Anal. Chem. 2011, 83, 1874–1878