Increasing Surface Capacity for On-Probe Affinity Capture MALDI-MS

Mar 9, 2011 - Department of Chemistry and Biochemistry, University of Texas at Arlington, 502 Yates Street, Arlington, Texas 76019-0065, United States...
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Increasing Surface Capacity for On-Probe Affinity Capture MALDI-MS via Gold Particle Attachment to Allyl Amine Plasma Polymers Zaneer M. Segu,†,§ Richard B. Timmons,‡ and Gary R. Kinsel*,† †

Department of Chemistry and Biochemistry, Southern Illinois University Carbondale, 1245 Lincoln Drive, Carbondale, Illinois 62901-4409, United States ‡ Department of Chemistry and Biochemistry, University of Texas at Arlington, 502 Yates Street, Arlington, Texas 76019-0065, United States ABSTRACT: In this study, we demonstrate that the protein binding capacity of a surface modified matrix-assisted laser desorption/ionization (MALDI) target can be increased significantly by architecturing the surface of the MALDI probe using gold microparticles. In the present approach, a MALDI target, initially modified via pulsed rf plasma deposition of an allyl amine polymer thin film, is subsequently architectured via reaction with 2-iminothiolane and surface attachment of gold microparticles. The modified probe is then exposed to thiolated biotin to introduce an avidin binding element on the surface of the gold beads. The protein binding capacity of this architectured target is compared with a similarly plasma polymer modified MALDI target that is directly biotinylated. Application of various surface concentrations of avidin to the two probes and MALDI-MS analysis of avidin contained in the solution removed from the probe reveals that saturation of the gold-particle architectured target occurs at a factor of 15-30 higher applied surface concentration, as compared with the unarchitectured target. Furthermore, MALDI-MS analysis of the avidin retained on the two probes reveals that the limit of detection is lowered by a factor of 15-20 on the gold-particle architectured target as compared with the unarchitectured target.

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atrix-assisted laser desorption/ionization (MALDI) is now broadly recognized as one of the mainstream tools for the mass spectrometric characterization of peptides and proteins.1 In its most common incarnation, MALDI analysis involves the deposition of the peptide/protein slated for analysis, together with the MALDI matrix, onto the surface of a stainless steel target. Since the original description of MALDI, however, there has been considerable interest in using other materials for the target construction.2 Originally these efforts were directed at improving the performance of the MALDI analysis (e.g., by controlling the crystallization process)3-6 or at addressing the translational needs of other bioanalytical methods (e. g., direct MALDI analysis of peptides and proteins on polymer membranes).7 More recently, surface chemically modified MALDI targets have been used to either selectively concentrate targeted biomolecules or to fractionate complex peptide/protein mixtures on the basis of broad chemical characteristics.8-15 This latter application is exciting in that it has shown promise as an approach to the identification of biomarkers for various human diseases.16 One of the significant limitations of any on-MALDI-probe based approach to either the selective enrichment of targeted biomolecules or the fractionation of complex protein mixtures, however, certainly lies in the limited capacity of the probe surface for protein binding. Indeed, a number of surface enhanced laser desorption/ionization (SELDI17) studies involving the detection of proteins in biological media have identified this as a major problem.18,19 Strictly speaking, the capacity of a modified region of a MALDI probe may be defined as the total number of targeted analyte molecules that can be explicitly r 2011 American Chemical Society

bound to the probe surface. For a protein such as lysozyme (∼14 400 Da, 7.9  10-12 mm2/molecule cross-sectional area20) deposited on a microscopically smooth probe surface, a maximum surface concentration of about 3 ng/mm2 can be obtained for a close-packed array of molecules. For a typical 1 mm diameter active area on a chemically modified MALDI probe, this translates into a maximum of ∼200 fmol of protein bound to the probe surface, a quantity that is within the limits of detection of most modern MALDI mass spectrometers. However, the problem of limited probe capacity will quickly emerge if many peptides/proteins are competing for the surface binding sites, as might be encountered in the analysis of a complex protein mixture, and/or the density of surface binding sites is substantially less than that needed for close-packed binding of protein. Either of these circumstances may be encountered in a typical application. In general, approaches to increasing the capacity of a surface chemically modified MALDI probe will involve increasing the morphological surface area of the probe and/or increasing the density of the binding element. In this investigation, an approach which effectively incorporates both modes of capacity enhancement has been employed wherein gold microparticles are covalently attached to a MALDI target coated with a thin polymer film of rf plasma polymerized allyl amine via reaction Received: October 17, 2010 Accepted: February 8, 2011 Published: March 09, 2011 2500

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Figure 1. Schematic diagram illustrating the fabrication of the biotinylated flat MALDI probe and the biotinylated gold microparticle architectured MALDI probe.

with 2-iminothiolane. Gold particles have been widely used in architecturing of surfaces because of their small size, inertness, visible colors, and ease of subsequent chemical modification through gold-thiol chemistry.21,22 In this study, we utilize the gold microparticle modified MALDI target surface as a scaffold for subsequent attachment of biotin. We show, through comparisons with flat biotinylated MALDI target surfaces, that the goldmicroparticle architecturing of a MALDI probe leads to significant increases in the binding capacity of the probe and concurrent reductions in the limit of detection for the protein avidin.

’ EXPERIMENTAL SECTION Chemicals and Materials. Polyethylene terephthalate (PET, 0.125 mm thickness) was obtained from Goodfellow Corp. (Berwyn, PA). Gold particles (1.5-3.0 μm diameter), allyl amine, 2-iminothiolane (2IT), triethylamine (TEA), trifluoroethanol (TFE), trifluoroacetic acid (TFA), phosphate buffered saline (PBS), biocytin, biotin 3-sulfo-N-hydroxysuccinimide ester sodium salt, avidin, and sinapinic acid (SA) were purchased from Sigma (St. Louis, MO). Absolute ethyl alcohol (200 proof) was purchased from AAPER Alcohol and Chemical Co (Shelbyville, KY). PBS was prepared as directed to achieve a concentration of 10 mM phosphate and a solution pH of 7.4. TEA and TFA were diluted with Milli-Q water to make 0.02 M and 10% (v/v) solution concentrations, respectively. Modification of the MALDI Probe. In general, 22 mm diameter aluminum disks were used as the substrates in the MALDI studies described here. These disks were placed inside a stainless steel mask which revealed an array of twelve 3 mm diameter spots. For the production of the gold-microparticle modified MALDI probe, the initial modification of the aluminum substrates involved the deposition of a thin film of pulsed radio frequency plasma polymerized allyl amine (ppAA).23,24 Specifically, the masked aluminum disk was placed in the plasma reactor, and allyl amine (flow rate 2.3 cm3/min,

standard temperature and pressure (STP)) was plasma polymerized/deposited at a constant power input of 200 W and a pulsing duty cycle of 3 ms on/45 ms off for a total of 12 min. The ppAA modified aluminum disk was then removed from the mask and exposed to 0.01 M 2IT in Milli-Q water in the presence of 0.02 M TEA for 1 h to incorporate thiol groups on the ppAA film coated surface.25 Following washing of the surface with Milli-Q water, the 2IT modified films were exposed to gold particles (d = 1.5-3.0 μm, c = 2 mg Au/10 mL of ethanol) and shaken for 1 h. The modified aluminum disks sections were subsequently washed thoroughly with ethanol to remove uncoupled gold particles. Finally, the gold particle coated aluminum disks were immersed in a 0.01 M solution of thiolated biotin (produced by reacting biocytin with 2IT26) in TFE for 3 h at room temperature. The resulting chemically modified MALDI probe was washed sequentially with ethanol and water and dried in air. The production of the flat biotinylated control MALDI probe proceeded via a similar set of procedures. Specifically, the ppAA modified aluminum disk was directly subjected to a 0.01 M solution of biotin 3-sulfo-N-hydroxysuccinimide ester sodium salt for 2 h at room temperature to incorporate biotin directly on the plasma polymer surface. The resultant probe was then washed sequentially with water and dried in air. A schematic diagram of the two methods of probe preparation is shown in Figure 1. Mass Spectral Analysis. Avidin was used to investigate the capacity of both the flat and gold-microparticle modified MALDI probes. Specifically, a 1 mg/mL stock solution of avidin in 0.01 M PBS solution was subjected to serial dilution to yield a number of solutions having a range of avidin concentrations from 0.005 to 1.0 mg/mL. In a typical experiment, a 2 μL aliquot of a given avidin solution was applied to the MALDI probe surface and allowed to incubate for 45 min. Subsequently, the solution was withdrawn from the modified MALDI probe surface and spotted on a conventional unmodified stainless steel MALDI target. Finally, a 0.5 μL aliquot of the 10% TFA solution and a 1 μL aliquot of SA (20 mg/mL in 5:5:1 Milli-Q water-acetonitrile2501

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Figure 2. Optical microscope images of gold microparticle retention on (A) a 2IT modified ppAA surface and (B) a ppAA surface, following washing of the surfaces with ethanol.

Figure 4. Integrated avidin MALDI ion signal for avidin contained in the solutions withdrawn from the biotinylated gold microparticle architectured MALDI probe (9) and from the biotinylated flat MALDI probe (() plotted as a function of the initial applied avidin surface concentration. The figure inset is an expansion of the low applied surface concentration region.

Figure 3. MALDI mass spectrum of avidin retained on the biotinylated gold microparticle architectured MALDI probe after washing with 0.01 M PBS and after addition of a 10% TFA/SA MALDI matrix solution.

methanol) was added to the solution deposited on the conventional unmodified MALDI target and to the surface of the biotinmodified MALDI probe. After allowing the samples to air-dry, positive ion MALDI mass spectra were acquired from both samples using a Bruker AutoFLEX mass spectrometer operated in the linear mode.

’ RESULTS AND DISCUSSION Characterization of the MALDI Probe. Initially, several experiments were carried out to characterize the gold microparticle architectured MALDI probes. First, high-resolution X-ray photoelectron spectroscopy (XPS) analysis of the ppAA surface following exposure to 2IT confirmed the incorporation of sulfur, presumably as thiol groups, on the probe surface (0.8% atom content). Second, covalent attachment of the gold microparticles through the thiol group to the probe surface was confirmed by comparing the retention of gold microparticles on a 2IT modified ppAA surface with the retention of the gold microparticles on an unmodified ppAA surface. For these studies, PET was used as the substrate rather than the aluminum disks. Figure 2 shows the results of this comparison recorded using a light microscope. It is seen in part A of Figure 2 that a relatively high density of gold microparticles remain attached to the 2IT modified ppAA surface following washing in ethanol. This result

is in stark contrast with the number of gold microparticles retained by the unmodified ppAA surface shown in part B of Figure 2 where nearly all of the gold microparticles are removed by the ethanol wash. From Figure 2, it is apparent that 2IT modification of the ppAA surface to incorporate thiol groups is necessary for binding of the gold microparticles. Third, activation of the surface for avidin capture through attachment of biotin to the gold microparticles was confirmed by MALDI mass spectrometry. Figure 3 is a typical MALDI mass spectrum of the captured protein (after washing the surface with PBS solution) following exposure of the biotinylated gold microparticle architectured MALDI probe to an avidin containing solution. As noted in an earlier publication, the primary MALDI ion signal observed is the monomeric protein at 15 918 Da, rather than the tetrameric structure, and this result is likely due to the strong denaturing conditions used during MALDI matrix deposition.13,27 In contrast, it should be noted that no avidin MALDI ion signal is detected when the unbiotinylated gold microparticle architectured MALDI probe is exposed to an avidin containing solution (after washing the surface with PBS solution). Probe Capacity Studies. Figure 4 shows the integrated avidin MALDI ion signals obtained from the solutions withdrawn from the biotinylated flat and biotinylated gold microparticle architectured MALDI probes plotted versus the initial applied avidin surface concentration. (It should be noted that surface concentrations are calculated by dividing the moles of deposited avidin by the area over which the sample spreads. While this approach assumes that the area interacted with by the avidin is microscopically smooth, it should be recognized that errors in this assumption (e.g., the surface area is much larger due to morphologically roughness at the microscopic level) will be similar for both the biotinylated flat and biotinylated gold microparticle architectured MALDI probes. Thus, comparison of the relative MALDI ion signal trends remains valid even if the absolute avidin surface concentration is subject to error.) In general, in 2502

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Figure 5. Integrated avidin MALDI ion signal for avidin retained on the biotinylated gold microparticle architectured MALDI probe (9) and retained on the biotinylated flat MALDI probe (() plotted as a function of the initial applied avidin surface concentration. The figure inset is an expansion of the low applied surface concentration region.

these experiments it is expected that at applied surface concentrations below the biotinylated surface-avidin binding capacity, no avidin MALDI ion signals will be obtained from the solutions withdrawn from the biotinylated probe (i.e., avidin will remain bound to the probe surface). In contrast, at applied surface concentrations above the probe capacity, avidin MALDI ion signals will be obtained from the solutions withdrawn from the biotinylated probe. The impact of the increased surface capacity afforded by the gold microparticle architecturing of the MALDI probe is immediately apparent in the comparison shown in Figure 4. In the expansion of the low surface concentration region of the plot, it is seen that the avidin MALDI ion signal begins to appear in the solution withdrawn from the biotinylated flat MALDI probe at applied surface concentrations as low as ∼100 fmol/mm2. This experimentally observed response is consistent with expectations based on monolayer coverage of a flat surface by the avidin molecules (i.e., if a crosssectional area for avidin of ∼4.2  10-11 mm2 is assumed, based upon a linear extrapolation of molecular volume with increase in molecular mass from that of lysozyme, monolayer coverage of a flat surface would occur at a surface concentration of ∼40 fmol/mm2). For the gold microparticle architectured MALDI probe, the avidin MALDI ion signal does not appear in the solution withdrawn from the probe until an initial applied surface concentration of ∼1500 fmol/mm2 is reached. The avidin MALDI ion signal then increases gradually until an initial surface concentration greater than ∼3300 fmol/mm2 is applied, at which point the avidin concentration in the withdrawn solution increases dramatically. This observed behavior is symptomatic of a Langmuir type adsorption isotherm and is consistent with the expected strong equilibrium binding interaction of the avidin with the biotinylated gold microparticle architectured MALDI probe. Specifically, the region of the plot between ∼1500 and 3300 fmol/mm2, where the avidin concentration in the withdrawn solution increases gradually, is governed by the equilibrium between the surface bound and solution phase avidin in the initially

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deposited solution. Above 3300 fmol/mm2, the avidin binding sites become saturated and the avidin concentration in the withdrawn solution increases linearly with additional applied avidin. This behavior is consistent with previous MALDI-based investigations of protein binding to various polymer substrates performed in our laboratory.28,29 Regardless of the details dictating the avidin concentration in the withdrawn solution, it is apparent from Figure 4 that the capacity of the gold microparticle architectured MALDI probe is at least a factor of 15-30 greater than that of the flat MALDI probe. Figure 5 shows the integrated avidin MALDI ion signals for the protein retained by the biotinylated flat MALDI probe and by the biotinylated gold microparticle architectured MALDI probe plotted versus the initial applied avidin surface concentration (i.e., the MALDI mass spectra are taken directly from the probe surfaces). The influence of the increased surface capacity of the gold microparticle architecturing of the MALDI probe is again apparent in this comparison. Avidin MALDI ion signals first appear on the flat MALDI probe at applied surface concentrations above ∼350 fmol/ mm2 (see the inset of Figure 5). This lower limit of detection can be contrasted with the gold microparticle architectured MALDI probe where MALDI ion signals for avidin can be detected at applied surface concentrations as low as 20 fmol/mm2. Thus, similar to the probe capacity increase indicated by the data from the withdrawn solutions shown in Figure 4, the gold microparticle architecturing of the MALDI probe surface lowers the limit of detection for avidin by a factor of 15-20. It can also be seen in Figure 5 that the avidin MALDI ion signal obtained from the biotinylated flat MALDI probe never rises to the same intensity as the avidin MALDI ion signal obtained from the biotinylated gold microparticle architectured MALDI probe. This behavior is consistent with a reduced total avidin binding capacity of the flat probe as compared with the gold microparticle architectured probe. However, it is also apparent that the avidin MALDI ion signal on the gold microparticle architectured probe is not a similar factor of 15-20 greater than that obtained from the flat probe, as might be expected from the other results obtained in these studies. This apparent discrepancy may be more a reflection of the limitations of ion formation in MALDI than any indication of the capacity of the MALDI probe itself, i.e., the dynamic range for protein MALDI ion signals is known to be extremely narrow and it is possible that the maximum avidin MALDI ion signal obtained from the gold microparticle architectured MALDI probe is limited by saturation of the MALDI ionization process rather than the total amount of avidin bound to the probe surface.

’ CONCLUSIONS The approach to increasing the protein binding capacity of surface modified MALDI targets described here is relatively simple and appears to offer a straightforward approach to increasing the capacity of the binding element by at least an order of magnitude. Indeed, through the use of smaller gold particles and surface modification methods designed to increase the binding element scaffold density, it seems likely that the capacity of surface modified MALDI targets could be increased by 2 orders of magnitude or more. While the specific approach described here focuses on biotin-avidin binding interactions, the general utility of gold-thiol chemistry offers opportunities for increasing the binding element density for a wide variety of analyte targets. Thus, for emerging applications wherein capture of targeted species directly on the surface of a MALDI target is desired, the approach described here offers a viable approach to achieving a significant increase in the 2503

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’ AUTHOR INFORMATION Corresponding Author

*Phone: 618-453-6471. Fax: 618-453-6408. E-mail: gkinsel@ chem.siu.edu.

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(24) Savage, C. R.; Timmons, R. B.; Lin, J. W. Adv. Chem. Ser. 1993, 236, 745–768. (25) Singh, R.; Kats, L.; Bl€attler, W. A.; Lambert, J. M. Anal. Biochem. 1996, 236, 114–125. (26) Pradier, C. M.; Salmain, M.; Lie, Z.; Methivier, C. Surf. Inteface Anal. 2002, 34, 67–71. (27) Zehl, M.; Allmaier, G. Anal. Chem. 2005, 77, 103–110. (28) Zhang, J.; Kinsel, G R. Langmuir 2002, 18, 4444–4448. (29) Zhang, J.; Kinsel, G. R. Langmuir 2003, 19, 3531–3534.

Present Addresses §

Cook Pharmica, Bloomington, IN.

’ ACKNOWLEDGMENT The support of NSF Grant CHE0719426 is gratefully acknowledged. In addition, Dr. Mary Kinsel is acknowledged for fruitful discussions and Michael Coviello, UTA Materials Science and Engineering, is acknowledged for his assistance in obtaining the light microscope pictures of the gold microparticle modified surfaces. ’ REFERENCES (1) Karas, M.; Hillenkamp, F. Anal. Chem. 1988, 60, 2299–2301. (2) Vestling, M. M.; Fenselau, C. Mass Spectrom. Rev. 1995, 14, 169–178. (3) Landry, F.; Lombardo, C. R.; Smith, J. W. Anal. Biochem. 2000, 279, 1–8. (4) Luque-Garcia, J. L.; Zhou, G.; Sun, T.; Neubert, T. A. Anal. Chem. 2006, 78, 5102–5108. (5) Wu, Y.; Hsieh, C.; Tam, M. F. Rapid Commun. Mass Spectrom. 2006, 20, 309–312. (6) Albrethsen, J. Clin. Chem. 2007, 53, 852–858. (7) Dufresne-Martin, G.; Lemay, J.; Lavigne, P.; Klarskov, K. Proteomics 2005, 5, 55–66. (8) Hutchens, T. W.; Yip, T. T. Rapid Commun. Mass Spectrom. 1993, 7, 576–580. (9) Lubman, D. M.; Liang, X. Anal. Chem. 1998, 70, 498–503. (10) Bundy, J. L.; Fenselau, C. Anal. Chem. 2001, 73, 751–757. (11) Hobbs, S. K.; Shi, G.; Bednarski, M. D. Bioconjugate Chem. 2003, 14, 526–531. (12) Li, M.; Timmons, R. D.; Kinsel, G. R. Anal. Chem. 2005, 77, 350–353. (13) Eriksson, A.; Bergquist, J.; Edwards, K.; Hagfeldt, A.; Malmstroem, D.; Agmo Hernandez, V. Anal. Chem. 2010, 82, 4577–4583. (14) Najam-ul-Haq, M.; Rainer, M.; Schwarzenauer, T.; Huck, C. W.; Bonn, G. K. Anal. Chim. Acta 2006, 561, 32–39. (15) Hashir, M.; Stecher, G.; Mayr, S.; Bonn, G. K. Int. J. Mass Spectrom. 2009, 279, 15–24. (16) O’Gorman, D.; Howard, J. C.; Varallo, V. M.; Cadieux, P.; Bowley, E.; McLean, K.; Pak, B. J.; Gan, B. S. Clin. Invest. Med. 2006, 29, 136–145. (17) Tang, N.; Tornatore, P.; Weinberger, S. R Mass Spectrom. Rev. 2004, 23, 34–44. (18) Merchant, M.; Weinberger, S. R. Electrophoresis 2000, 21, 1164–1167. (19) Forde, C. E.; Gonzales, A. D.; Smessaert, J. M.; Murphy, G. A.; Shields, S. J.; Fitch, J. P.; McCutchen-Maloney, S. L. Biochem. Biophys. Res. Commun. 2002, 290, 1328–1335. (20) Gao, J.; Whitesides, G. M. Anal. Chem. 1997, 69, 575–580. (21) Teng, C.; Ho, K.; Ho, Y.; Lin, Y.; Chen, Y. Anal. Chem. 2004, 76, 4337–4342. (22) Mirkin, C. A. Inorg. Chem. 2000, 39, 2258–2272. (23) Panchalingam, V.; Chen, X.; Savage, C. R.; Timmons, R. B.; Eberhart, R. C. J. Appl. Polym. Sci.: Appl. Polym. Symp. 1994, 54, 123–141. 2504

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