Anal. Chem. 2007, 79, 6215-6221 Sample-First Preparation: A Method for Surface-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry Analysis of Cyclic Oligosaccharides Hsin-Pin Wu,† Chih-Lin Su,† Hui-Chiu Chang,‡, § and Wei-Lung Tseng*,†,§ Department of Chemistry, National Sun Yat-sen University, Kaohsiung, Taiwan, Graduate Institute of Medicine, College of Medicine, Kaohsiung Medical University, Kaohsiung, Taiwan, and National Sun Yat-sen UniversitysKaohsiung Medical University Joint Research Center, Kaohsiung, Taiwan A new sample preparation method for the analysis of cyclic oligosaccharides in surface-assisted laser desorption/ ionization mass spectrometry (SALDI-MS) is presented. We call this new technique “sample first method”, in which a sample is deposited first and then bare gold nanoparticles (AuNPs), which serve as the SALDI matrixes, are added to the top of the sample layer. The use of the sample first method offers significant advantages for improving shot-to-shot reproducibility, enhancing the ionization efficiency of the analyte, and reducing sample preparation time as compared to the dried-droplet method, wherein samples and bare AuNPs are mixed and dried together. The relative standard deviation (RSD) values of the signal intensity as calculated from 65 sample spots was 25% when the sample first methods were applied to the analysis of β-cyclodextrin. The results were more homogeneous as compared to the outcome using drieddroplet preparation of AuNPs (RSD ) 66%) and 2,5dihydroxybenzoic acid (RSD ) 209%). We also found out that the optimal concentration of AuNP for ionization efficiency is 7.4 nM (4.52 × 1012 particles/mL) while the lowest detectable concentration of cyclic oligosaccharides through this approach is 0.25 µM. Except for the cyclic oligosaccharide, the proposed method was also applied to the analyses of other biological samples, including neutral carbohydrate and steroid, aminothiols, and peptides as well as proteins. An effective matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) analysis is strongly dependent on the sample preparation.1-6 It is therefore not surprising that a wide variety of methods and protocols for sample preparation have been developed, such as the dried-droplet,1 fast evaporation,2 vacuum * To whom correspondence should be addressed. E-mail: tsengwl@ mail.nsysu.edu.tw. Fax: 011-886-7-3684046. † National Sun Yat-sen University. ‡ Kaohsiung Medical University. § National Sun Yat-sen UniversitysKaohsiung Medical University Joint Research Center. (1) Karas, M.; Hillenkamp, F. Anal. Chem. 1988, 60, 2299-2301. (2) Vorm, O.; Roepstorff, P.; Mann, M. Anal. Chem. 1994, 66, 3281-3287. (3) Papac, D. I.; Wong, A.; Jones, A. J. S. Anal. Chem. 1996, 68, 3215-3223. 10.1021/ac070847e CCC: $37.00 Published on Web 07/14/2007 © 2007 American Chemical Society drying,3 sandwich,4 two-layer,5 and even a three-layer method.6 So far, one of the easiest and most frequently used methods for detection of peptides and proteins by MALDI-MS is the drieddroplet method. In this method, the solid matrix, such as 2,5dihydroxybenzoic acid (DHB) and sinapinic acid, is mixed with a sample at an optimal ratio and then dried on the MALDI target plate. Unfortunately, the inhomogeneous cocrystallization of the analytes with the matrix requires sweet-spot searching and results in poor shot-to-shot and sample-to-sample reproducibility.7 This is the main factor that hampers the application of MALDI-MS for quantitative measurements. Other factors, such as solvent composition, pH, and the presence of matrix additives can alter the crystallization behavior and thus improve the quality of the preparations.8,9 The method applied for sample preparation still observes the sweet-spot effect. To overcome this problem, liquid matrixes such as glycerol10 and ionic liquid11 have been developed and successfully applied in the MALDI-MS analysis of highly sulfated oligosaccharides, peptides, proteins, and poly(ethylene glycol).12-14 Although the use of liquid matrixes exhibits higher signal reproducibility, it is still limited to low mass resolution and poor ionization efficiency. Another possibility for improving the spot homogeneity of analyte/matrix preparation is the use of nanomaterials such as (4) (a) Whiteaker, J. R.; Fenselau, C. C.; Fetterolf, D.; Steele, D.; Wilson, D. Anal. Chem. 2004, 76, 2836-2841. (b) Canelle, L.; Pionneau, C.; Marie, A.; Bousquet, J.; Bigeard, J.; Lutomski, D.; Kadri, T.; Caron, M.; JoubertCaron, R. Rapid Commun. Mass Spectrom. 2004, 18, 2785-2794. (5) (a) Zheng, J.; Li, N.; Ridyard, M.; Dai, H.; Robbins, S. M.; Li, L. J. Proteome Res. 2005, 4, 1709-1716. (b) Dai, Y.; Whittal, R. M.; Li, L. Anal. Chem. 1999, 71, 1087-1091. (6) Keller, B. O.; Li, L. J. Am. Soc. Mass Spectrom. 2006, 17, 780-785. (7) Tholey, A.; Heinzle, E. Anal. Bioanal. Chem. 2006, 386, 24-37. (8) Cohen, S. L.; Chait, B. T. Anal. Chem. 1996, 68, 31-37. (9) (a) Distler, A. M.; Allison, J. Anal. Chem. 2001, 73, 5000-5003. (b) Kim, Y.; Hurst, G. B.; Doktycz, M. J.; Buchanan, M. V. Anal. Chem. 2001, 73, 2617-2624. (10) (a) Overberg, A.; Karas, M.; Bahr, U.; Kaufmann, R.; Hillenkamp, F. Rapid Commun. Mass Spectrom. 1990, 4, 293-296. (b) Strupat, K.; Karas, M.; Hillenkamp, F.; Eckerson, C.; Lottspeich, F. Anal. Chem. 1994, 66, 464470. (11) Armstrong, D. W.; He, L.; Liu, Y.-S. Anal. Chem. 1999, 71, 3873-3876. (12) Laremore, T. N.; Murugesan, S.; Park, T.-J.; Avci, F. Y.; Zagorevski, D. V.; Linhardt, R. J. Anal. Chem. 2006, 78, 1774-1779. (13) Li, Y. L.; Gross, M. L. J. Am. Soc. Mass Spectrom. 2004, 15, 1833-1837. (14) Mank, M.; Stahl, B.; Boehm, G. Anal. Chem. 2004, 76, 2938-2950. Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 6215 gold nanoparticles (AuNPs),15 silicon nanoparticles,16 and carbon nanotubes.17 As compared with the conventional organic matrix, the use of nanomaterials for the analysis of small molecules can reduce matrix ion interference. This approach, which is known as the surface-assisted laser desorption/ionization (SALDI)-MS, offers the advantages of simple sample preparation, high surface areas, flexibility in the sample deposition conditions. and independence from irradiation wavelength.18 Furthermore, the high molar matrix-to-analyte ratio (∼107-109 analytes/particle) is obtained using nanomaterials as the SALDI matrixes and thus contributing to high efficient ionization. Small organic compounds, peptides, and even proteins have all been detected with remarkable sensitivities using nanomaterials.15-19 For example, the applicability of Nile red-adsorbed AuNPs as the SALDI matrixes was successfully demonstrated in the analysis of aminothiols.20 Without enrichment, the limit of detection of glutathione obtained by SALDI-MS is down to 1.0 µM. However, the amount of analytes adsorbed on the particle surface is sensitive to the adsorption time. We found that detection sensitivity dramatically decreases with the decrease in contact time between analytes and particles. To achieve a faster analysis, a new sample preparation method should be developed for SALDI-MS. In this work, we report the application of a sample first preparation method for the analysis of cyclic oligosaccharides when using bare AuNPs as the assisted matrix. In the sample first method, the analyte is deposited first and then followed by the bare AuNPs. The cyclic oligosaccharides, including R-, β-, and γ-cycodextrin (CD), are difficult to ionize by MALDI-MS, but they could be cationized very efficiently using bare AuNPs as matrixes in combination with a sample first preparation method. As compared with the dried-droplet and matrix first methods, a sample first method not only provides high sensitivity for the measurement of neutral carbohydrate but also improves the spot homogeneity. This practical method was further validated by the analyses of biological samples, including neutral carbohydrate, neutral steroid, aminothiols, and peptides. EXPERIMENTAL SECTION Chemicals. R-CD, β-CD, γ-CD, ribose, glucose, raffinose, maltose, testosterone, progesterone, corticosterone, cysteine, homocysteine, glutathione, angiotensin I, substance P, and insulin were all obtained from Sigma (St. Louis, MO). Citric acid, hydrogen tetrachloroaurate(III) dehydrate, sodium borohydride, and DHB were purchased from Aldrich (Milwaukee, WI). Ethanol and ammonium hydroxide were obtained form Acros (Geel, Belgium). When 150 mM citric acid was used to prepare citrate buffer, NH4OH (25-30%) was used to adjust the pH to 4.0. Milli-Q ultrapure water was used in all of experiments. Apparatus. A double-beam UV-visible spectrophotometer (Cintra 10e, GBC Scientific Equipment Pty Ltd., Dandenong, (15) McLean, J. A.; Stumpo, K. A.; Russell, D. H. J. Am. Chem. Soc. 2005, 127, 5304-5305. (16) (a) Wen, X.; Dagan, S.; Wysocki, V. H. Anal. Chem. 2007, 79, 434-444. (b) Finkel, N. H.; Prevo, B. G.; Velev, O. D.; He, L. Anal. Chem. 2005, 77, 1088-1095. (17) (a) Xu, S.; Li, Y.; Zou, H.; Qiu, J.; Guo, Z.; Guo, B. Anal. Chem. 2003, 75, 6191-6195. (b) Ren, S.-F.; Zhang, L.; Cheng, Z.-H.; Guo, Y.-L. J. Am. Soc. Mass Spectrom. 2005, 16, 333-339. (18) Sunner, J.; Dratz, E.; Chen, Y.-C. Anal. Chem. 1995, 67, 4335-4342. (19) Chen, C.-T.; Chen, Y.-C. Anal. Chem. 2005, 77, 5912-5919. (20) Huang, Y.-F.; Chang, H.-T. Anal. Chem. 2006, 78, 1485-1493. 6216 Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 Victoria, Australia) was used to measure the absorbance of the AuNPs. The homemade dark-field scattering system is made of an Olympus IX70 inverted microscope (Tokyo, Japan), a DP70 digital camera (Olympus, Tokyo, Japan), and high-numericalaperture dark-field condenser (NA ) 1.2-1.4; U-DCW, Olympus).21 White light from a 100-W halogen lamp and a focusing lens within a condenser are angled with respect to the objective (40×; NAs ) 0.55) so that the illuminating light does not directly enter the objective; thus, this arrangement results in a low background. The scattering images for a AuNP-deposited glass slide were observed using dark-field scattering microscopy. We note that glass slide was used to replace a standard MALDI plate because the inverted microscope required a transparent substrate. The ImageJ program (http://rsb.info.nih.gov/ij/) was used to analyze the images. MS experiments were performed in the positive-ion mode on a reflectron-type time-of-flight (TOF) mass spectrometer (Autoflex, Bruker) equipped with a 3-m flight tube. Desorption/ionization was obtained by using a 337-nm-diameter nitrogen laser with a 3-ns pulse width. The available accelerating voltages existed in the range from +20 to -20 kV. To obtain good resolution and signal-to-noise (S/N) ratios, the laser power was adjusted to slightly above the threshold and each mass spectrum was generated by averaging 500 laser pulses. Synthesis of AuNPs with NaBH4. The bare (uncapped) AuNPs were prepared by the chemical reduction of metal salt precursor (hydrogen tetrachloroaurate) in a liquid phase.22 Typically, a 200-µL aliquot of 0.05 M aqueous HAuCl4 solution was added to 50 mL of deionized water. Subsequently, 600 µL of freshly prepared 0.05 M NaBH4 solution was added to the system, whereby the solution gradually changed color to ruby red. We note that these bare AuNPs are stable for a couple weeks because small ions such as sodium, potassium, and chloride are absorbed on the surface of bare AuNPs. The average size distribution of the AuNPs is 12 ( 1.9 nm, which is confirmed by TEM measurements (data not shown).23 To further determine the concentration of the nanoparticles, we assumed that the reduction from gold(III) to gold atoms was 100% complete.24 The concentration of the original bare AuNPs is ∼3.7 nM (2.26 × 1012 particles/ mL), which we denote in this study as 1×. Preparation of Samples. Three sample preparation methods for analysis of cyclic oligosaccharides were tested before SALDIMS measurements. Although the drying speed can be increased by rising the temperature, fast evaporation of sample solution will result in an increase in aggregation degree of AuNPs.25 Thus, three sample preparation methods were all performed at room temperature. (a) Dried-Droplet cmethod. The cyclic oligosaccharides (500 µL) were added separately to 1× AuNP solutions (500 µL). Immediately, the resulting mixtures (1 µL) were pipetted into a stainless steel 384-well target (Bruker Daltonics) and dried in air. We note that the drying time for the resulting mixtures is ∼30 min. (21) Tseng, W.-L.; Lee, K.-H.; Chang, H.-T. Langmuir 2005, 21, 10676-10683. (22) Gole, A.; Murphy, C. J. Chem. Mater. 2004, 16, 3633-3640. (23) Su, C.-L.; Tseng, W.-L. Anal. Chem. 2007, 79, 1626-1633. (24) Neiman, B.; Grushka, E.; Lev, O. Anal. Chem. 2001, 73, 5220-5227. (25) (a) Sen, D.; Spalla, O.; Tache´, O.; Haltebourg, P.; Thill, A. Langmuir 2007, 23, 4296-4302. (b) Wang, W.-N.; Lenggoro, I. W.; Okuyama, K. J. Colloid Interface Sci. 2005, 288, 423-431. Figure 1. Mass spectra of β-CD obtained by using three sample preparation methods: (A) dried-droplet, (B) sample first, and (C) matrix first method. The full scan mass spectrum was acquired from m/z 200 to 2000 (inset). The 1× bare AuNPs were used as the matrixes while the sample concentration was 10 µM. The peaks at m/z 1157.95 and 1173.95 are assigned to the [β-CD + Na]+ and [β-CD + K]+, respectively. A total of 500 pulsed laser shots were applied under a laser fluence of 60 µJ. (b) Matrix First Method. The 1× AuNP solutions (1 µL) were deposited first onto the sample plate and let dry. Then, l µL of the cyclic oligosaccharides was deposited onto the first layer and allowed to dry in air. (c) Sample First Method. The cyclic oligosaccharides (1 µL) were deposited first onto the sample plate and let dry in air. To search for an optimal sample first method for SALDI-MS measurements, different concentrations (0.5-3.0×; 1 µL each) of AuNPs solutions were deposited onto the first layer and allowed to dry in air. We note that the analysis of carbohydrates, steroids, aminothiols, peptide, and proteins was conducted by use of the sample first method. In the cases where DHB was used as a comatrix for MALDIMS analysis, DHB (1.0 µL) was mixed with the cyclic oligosaccharides (1.0 µL). Saturated matrix solution was prepared by dissolving DHB in 50% ethanol. The resulting mixtures were then pipetted into the wells of the steel plate and dried in air at room temperature prior to MALDI-MS measurements. RESULTS AND DISCUSSION Comparison of Sample Preparation Methods for SALDIMS Analysis. Although a variety of sample preparation methods have been proposed for MALDI-MS analysis,1-6 their impact on SALDI-MS has not been carefully studied. Our initial investigations using the dried-droplet methods revealed some inherent problems.21 In particular, the ionization efficiency is determined by the amounts of analyte adsorbed on a particle. Thus, high ionization efficiency was obtained when the contact time between the analyte of interest and particles reached the adsorption equilibrium time. For example, the adsorption equilibrium time of neutral carbohydrate on the bare AuNP (12 ( 1.9 nm) was ∼2 h. On the other hand, we showed in an earlier publication21 that the use of AuNPs as the SALDI matrixes provided high detection sensitivity for small neutral carbohydrates but not oligosaccharides. The lower ionization efficiency for oligosaccharides can be attributed to larger steric hindrance, which led to less adsorption between analyte and nanoparticles. To speed up the analysis time and improve the detection sensitivity in SALDI-MS, we tested different sample preparation methods that include dried-droplet, sample first, and matrix first methods. Figure 1A displays the SALDI mass spectrum of β-CD obtained from the dried-droplet method whereby the sample and bare AuNP (1×) are mixed and immediately deposited on the sample target. As expected, the lowintensity ions at m/z 1157.95 were observed because only small amounts of β-CD were adsorbed on the surface of AuNPs within a short contact time. The peaks at m/z 1157.95 and 1173.95 correspond to [β-CD + Na]+ and [β-CD + K]+, respectively. No MH+ signal obtained by SALDI-MS was due to the low proton affinity of saccharides. By using the sample first method, a similar mass spectrum was found but with a high intensity as shown in Figure 1B. When the AuNPs are deposited on top of the sample to form the second layer, they would be distributed on the entire sample surface. On the basis of this phenomenon, the matrix-toanalyte ratio was relatively increased as compared to the drieddroplet method. More importantly, the resultant sample spots were smooth and caused a significant improvement in spot homogeneity. Interestingly, the ion intensity of β-CD obtained by the sample first method was still relatively high as compared to our previous results using dried-droplet methods wherein the analytes were equilibrated with bare AuNPs for 2 h. Inthe sample first method, the upper layer of the AuNPs was irradiated with the laser beam and resulted in an increase in the local temperature. The heat was then transferred to the bottom layer of the analyte molecules Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 6217 Figure 2. Normalized [β-CD + Na]+ ion intensities obtained from 65 different sample spots. The samples are prepared by the sample first (black squares) and dried-droplet (gray squares) methods, respectively. The RSD of the data series are presented as bar graphs; i.e., the black and gray bars indicate the RSD values obtained using the sample first and dried-droplet methods, respectively. A total of 50 pulsed laser shots were applied under a laser fluence of 60 µJ. The other conditions were the same as those in Figure 1. and became well desorbed/ionized. By contrast, the analytes in the dried-droplet methods adsorbed on the surface of the particles were irradiated with the laser beam before the AuNPs. Thus, the local temperature of the AuNPs was relatively lower and resulted in poor ionization efficiency for β-CD. To support our hypothesis, we first deposited the AuNPs on the sample stage, followed by the samples (Figure 1C). Apparently, this preparation method was not effective for ionization of β-CD. To investigate possible improvements in shot-to-shot reproducibility using the sample first method, we compared it with the dried-droplet method in SALDI-MS. Figure 2 shows that the use of the sample first method decreased the variability of the signal intensities of β-CD that were collected from 65 different sample spots. The relative standard deviation (RSD) values of the signal intensity were 25% for the sample first method and 66% for the dried-droplet method. By contrast, these variations were greater than 209% when using the dried-droplet preparation of DHB with β-CD (Supporting information, Figure S1). The homogeneity of the AuNP distribution was further investigated using the UVvis absorption spectrometer and dark-field microscopy when the dried-droplet and sample first methods were conducted in the analysis of β-CD. After the AuNPs were mixed with the analyte in solution, the plasmon absorption of AuNPs exhibited a slight red shift from 526 to 529 nm (Supporting Information, Figure S2). This is because the resonance wavelength of the surface plasmon in metallic nanoparticles is highly dependent on the surrounding medium.26 However, when the mixture of the AuNPs and β-CD was dried on the target, the aggregation of AuNPs occurred and thus led to an increase in absorbance at longer wavelengths. Compared with the sample first method, the plasmon band for the dried-droplet method was considerably red-shifted and broadened (Supporting Information, Figure S2). On the other hand, dark-field microscopy is a useful imaging technique for distinguishing individual and aggregated nanoparticles.27 As soon as (26) McFarland, A. D.; Van Duyne, R. P. Nano Lett. 2003, 3, 1057-1062. 6218 Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 Figure 3. Dark-field images of AuNPs obtained from (A) dried-droplet and (B) sample first methods. The 1× bare AuNPs were used as matrixes, and the concentration of β-CD was 1 mM. Exposure times, 50 ms; scattering detection area using a 40× objective, 860 µm × 610 µm corresponding to 4080 (horizontal) × 3072 (vertical) pixels. the AuNPs were aggregated, the red shift of scattering would occur, and hence, the aggregation of the AuNP was easily Figure 4. Effect of AuNP concentration on the ionization efficiency of cyclic oligosaccharides: (A) 1×, (B) 1.5×, (C) 2×, (D) 2.5×, and (E) 3×. The samples were prepared by the sample first method. The sample concentration was 2 µM. Peak identities: m/z 995.55, [R-CD + Na]+; m/z 1157.95, [β-CD + Na]+; m/z 1320.45, [γ-CD + Na]+. The other conditions were the same as those in Figure 1. Figure 5. SALDI mass spectra of R- (0.25 µM), β- (0.25 µM), and γ-CD (0.5 µM) obtained using 2× AuNPs as the matrix. The samples were prepared by the sample first method. Peak identities: m/z 995.55, [R-CD + Na]+; m/z 1011.55, [R-CD + K]+; m/z 1157.95, [β-CD + Na]+; m/z 1173.95, [β-CD + K]+; m/z 1320.45, [γ-CD + Na]+. The other conditions were the same as those in Figure 1. recognized by conducting scattering measurements.28 Figure 3A reveals that the AuNPs significantly aggregated using the drieddroplet method. The orange and red spots corresponded to the scattering images of the AuNPs aggregates. It should be noted (27) Xu, X.-H. N.; Brownlow, W. J.; Kyriacou, S. V.; Wan, Q.; Viola, J. J. Biochemistry 2004, 43, 10400-10413. (28) So ¨nnichsen, C.; Reinhard, B. M.; Liphardt, J.; Alivisatos, A. P. Nat. Biotechnol. 2005, 23, 741-745. that the orange spot came from an aggregate of more than two particles. In comparison, Figure 3B shows that some spots displayed a green color, which corresponded to the scattering images of single AuNPs.29 It was suggested that an increase in homogeneity of the analyte could be achieved by using the sample first method (Figure 3B). These finding support the (29) Orendorff, C. J.; Sau, T. K.; Murphy, C. J. Small 2006, 2, 636-639. Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 6219 notion that the use of the sample first method offers a better reproducibility as a result of the improvement in the sample homogeneity. Effect of AuNP Concentration. When the AuNPs were used as the SALDI matrixes in the dried-droplet method, the ionization efficiency of the analyte was greatly affected by the particle concentration. Thus, the effect of the AuNP concentrations on the signal intensities of cyclic oligosaccharides was also studied in the sample first method. Figure 4 shows that the signal intensities of a mixture comprising equal concentrations (2 µM) of the R-, β-, and γ-CD were maximized while the AuNP concentration increased by up to 2×. The enhancement was likely due to the increase in the number of the deposited AuNPs on top of the sample. Under this condition, we ensured that the entire sample surface was covered with AuNPs. However, a decrease in the signal was observed above 2× the level of concentration. This finding indicates that a thicker layer was formed when a higher concentration of AuNPs was deposited. This phenomenon would increase the isolated AuNPs and thereby result in the decrease of the local temperature. In comparison with the use of 1× AuNPs, the optimization of the concentration of AuNPs provided more than a 3-fold enhancement in the S/N ratio for cyclic oligosaccharides. Figure 5 shows the SALDI mass spectrum of a mixture containing R- (0.25 µM), β-(0.25 µM), and γ-CD (0.5 µM). However, no signals were identifiable when their concentrations were lowered to 0.1 µM, which indicated that the detection limits were somewhere between 100 and 250 fmol. By contrast, this method was 105 times more sensitive for the analysis of cyclic oligosaccharides than with the use of DHB as MALDI matrixes.30 Although the ionization efficiency of MALDI-MS can be further enhanced by reacting cyclic oligosaccharides with succinic anhydride, a long reaction time (24 h) is unduly required.30 Analysis of Different Biomolecules. The successful analysis of neutral, cyclic oligosaccharides, as described earlier, motivated us to apply the sample first method to the analysis of other biological compounds. The neutral carbohydrates and steroid, aminothiols, and peptides were tested by SALDI-MS in combination with the sample first method. Figure 6A displays the SALDI mass spectrum of the carbohydrate mixture (5 µM). The peaks that appear at m/z 172.86, 202.91, 364.74, and 526.76 corresponded to the sodiated molecular ions of ribose, glucose, maltose, and raffinose, respectively. In contrast to the SALDI-MS spectra obtained with the dried-droplet method, the use of the sample first method provided better sensitivity (except for glucose) and decreased the analysis time. To further explore the capability of the sample first method to analyze more hydrophobic compounds, a mixture of steroids (50 µM) was chosen in this study. It should be noted that steroids are ionized poorly by both MALDI and electrospray ionization MS.31 The successful ionization of neutral steroids is shown in Figure 6B. The peaks appearing at m/z 311.13, 337.14, and 369.15 were the sodiated molecular ions of testosterone, progesterone, and corticosterone, respectively. We were also interested in determining the thiol-containing amino acids, which were successfully ionized by using Nile red-absorbed (30) Ahn, Y. H.; Yoo, J. S.; Kim, S. H. Anal. Sci. 1999, 15, 53-56. (31) (a) Wang, Y.; Hornshaw, M.; Alvelius, G.; Bodin, K.; Liu, S.; Sjo¨vall, J.; Griffiths, W. J. Anal. Chem. 2006, 78, 164-173. (b) Griffiths, W. J.; Liu, S.; Alvelius, G.; Sjo ¨vall, J. Rapid Commun. Mass Spectrom. 2003, 17, 924935. 6220 Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 Figure 6. Sample first method for SALDI-MS analysis of biomolecules: (A) neutral carbohydrates (5 µM), (B) neutral steroids (50 µM), and (C) aminothiols (200 µM). Peak identities: (A) m/z 172.86, [ribose + Na]+; m/z 202.91, [glucose + Na]+; m/z 364.74, [maltose + Na]+; m/z 526.76, [raffinose + Na]+. (B) m/z 311.13, [testosterone + Na]+; m/z 337.14, [progesterone + Na]+; m/z 369.15, [corticosteron + Na]+. (C) m/z 144.00, [cysteine + Na]+; m/z 158.00, [homocysteine + Na]+; m/z 329.98, [glutathione + Na]+. The sample concentration was 10 µM while the other conditions were the same as those in Figure 5. AuNPs as the SALDI matrixes.20 Figure 6C displays the SALDI mass spectra of a mixture containing cysteine, homocysteine, and glutathione. It was suggested that the sample first method could be directly used for the detection of aminothiols without the need for contact time between aminothiols and AuNPs. Since the successful implementation of SALDI-MS for the analysis of small biomolecules was well established, we also extend this method for peptides and proteins. Panels A and B in Figure 7 show that substance P and insulin were successfully ionized by using AuNPs as the SALDI matrixes. It should be noted that our proposed method cannot be applied to the analysis of angiotensin I, which is the standard peptide for MALDI-MS. It is believed that the ionization of angiotensin I can be achieved by searching the appropriate sample solution. The peaks that appeared at m/z 1370.64 and 5779.94 corresponded to [substance P + Na]+ and [insulin + 2Na]+, respectively. The peak shape and ionization efficiency of insulin were further improved when the sample was prepared in a citrate buffer at pH 4.0 (Supporting Information, Figure S3).32 However, difficulties were encountered in the (32) Chen, W.-Y.; Chen, Y.-C. Anal. Bioanal. Chem. 2006, 386, 699704. Figure 7. Sample first method for SALDI-MS analysis of (A) substance P (10 µM) and (B) insulin (100 µM). Peak identities: (A) m/z 1370.64, [substance P + Na]+; (B) m/z 5779.94, [insulin + 2Na]+ The other conditions were the same as those in Figure 5. analysis of cytochrome c (12 300 Da), which indicated that the upper detectable mass limit was ∼6000 Da. Moreover, the detection sensitivity decreased dramatically with increase in mass. The same phenomena mentioned above were also reported by Hillenkamp’s group.33 CONCLUSIONS The sample first method for SALDI-MS analysis of neutral and charged biomolecules using bare AuNPs provides a number of (33) Schurenberg, M.; Dreisewerd, K.; Hillenkamp, F. Anal. Chem. 1999, 71, 221-229. (34) Harvey, D. J. Mass Spectrom. Rev. 1999, 18, 349-351. (35) Stubiger, G.; Belgacem, O. Anal. Chem. 2007, 79, 3206-3213. (36) Yu, Y. Q.; Fournier, J.; Gilar, M.; Gebler, J. C. Anal. Chem. 2007, 79, 17311738. (37) Zhao, J.; Qiu, W.; Simeone, D. M.; Lubman, D. M. J. Proteome Res. 2007, 6, 1126-1138. distinctive advantages. First, it allows the sensitive detection of neutral oligosaccharides and steroid in their underivatized form wherein the detection limits for R-, β-, and γ-CD are in the highfemtomole levels. Second, this method is simple and time-efficient when nanomaterials are used as the SALDI matrixes. The analytes can be detected directly without any adsorption steps. Third, reproducibility of signal intensities is appreciably improved by the sample first method mainly because of the improvement in the spot homogeneity. Last, it can be applied in the analysis of peptides and proteins, whose the upper detectable mass limit was ∼ 6000 Da. On the basis of these advantages, we believe that the proposed methods can be applied not only to the analysis of many types of carbohydrate and carbohydrate-containing compounds34 but also in the detection of other neutral biomolecules, such as steroids31 and lipids.35 For example, we should be able to use the proposed method for identifying glycosylation sites of proteins after the appropriate enzyme digestion.36 Also, the method can be helpful in the comparative studies of the carbohydrate chains of glycoproteins produced by malignant cells and normal cells.37 ACKNOWLEDGMENT We thank the National Science Council (NSC 95-2113-M-110020-) of Taiwan and Aim for the Top University Plan, Ministry of Education, Taiwan for the financial support of this study. We thank Professor J. Shiea for allowing us to use the MALDI-TOF instrument. SUPPORTING INFORMATION AVAILABLE Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review April 26, 2007. Accepted June 13, 2007. AC070847E Analytical Chemistry, Vol. 79, No. 16, August 15, 2007 6221
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