Article pubs.acs.org/ac
Polystyrene Spheres-Assisted Matrix-Assisted Laser Desorption Ionization Mass Spectrometry for Quantitative Analysis of Plasma Lysophosphatidylcholines Yanbo Wei,†,‡,⊥ Shumu Li,∥,⊥ Jingxia Wang,∥ Chunying Shu,∥ Jian’an Liu,‡ Shaoxiang Xiong,‡ Jianwen Song,§ Junjie Zhang,§ and Zhenwen Zhao*,†,‡ †
Key Laboratory of Analytical Chemistry for Living Biosystems, ‡Beijing Mass Spectrum Center, and ∥Key Laboratory of Molecular Nanostructure and Nanotechnology, Institute of Chemistry Chinese Academy of Science, Beijing 100190, China § Key Laboratory of Cell Proliferation and Regulation Biology, Ministry of Education, Institute of Cell Biology, College of Life Sciences, Beijing Normal University, Beijing 100875, China ABSTRACT: The quantitative analysis by matrix-assisted laser desorption ionization mass spectrometry (MALDI MS) is a challenge due to the poor reproducibility originating from the heterogeneity of the matrix−analyte crystals. Polystyrene (PS) colloidal spheres have superior monodispersed property and can self-assemble to form photonic crystals. With the assistance of PS spheres, a uniform matrix−analyte cocrystal was constructed for the quantitative analysis of plasma lysophosphatidylcholines (LPCs). The matrix and the solvent in MALDI MS analysis were optimized, and the reproducibility and the accuracy were investigated in detail. This is the first report about the very simple method of PS spheres-assisted MALDI MS for quantitative analysis, where it is believed that this approach will greatly expand the applications of MALDI MS.
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reduced pressure, was proposed.6,7 In addition, recent advances in nanotechnology and nanoscience made nanoparticles (NPs, most commonly Au, Ag, SiO2, TiO2, and Fe3O4) can be used instead of organic matrixes to prepare the analytical samples for LDI-MS. One of the advantages by using NPs as a matrix is to readily obtain a homogeneous crystallization for quantification of LDI MS.8−10 In this paper, a new strategy was presented to overcome the heterogeneity of the matrix−analyte crystals. Polystyrene (PS) colloidal spheres, characterized by their large specific surface area, agglutination, and uniform spherical morphology, have shown considerable potential application in optical devices, catalysis, and being used as a template in recent years.11−15 More importantly, PS colloidal spheres have a superior monodispersed property and can self-assemble to form photonic crystals, and according to particular demand, some other substances such as organic dye or polymer monomer could be mixed into the colloidal spheres.16,17 It inspired a new thought: when mixing matrix and analytes with PS colloidal spheres, along with PS colloidal spheres self-assembling, PS spheres may act as a crystal nucleus, which could combine with matrix-analytes, and finally result in a homogeneous crystal like photonic crystal. On the basis of this hypothesis, in this study, we examined the morphology of the crystal and the feasibility
atrix-assisted laser desorption ionization mass spectrometry (MALDI MS) has been applied to analyses of complex biological systems because of its benefits, such as simple operation and sample preparation, high sensitivity, highthroughput analyses, and reliable results. However, MALDI MS analysis has the disadvantages of rather poor reproducibility, mainly originating from heterogeneity of the matrix−analyte crystals, which leads to MALDI MS being heavily criticized for its quantitative analysis. Numerous efforts have been made to improve the performance of quantitative analysis by MALDI MS. Stable-isotope labeling of molecules as an internal standard is an attractive technique that enables the quantitative analysis of specific molecules in a complicated system.1,2 In addition, binary matrixes, such as the combination of 2,5-dihydroxybenzoic acid/N,N-dimethylaniline, were reported for the quantitative analysis of oligosaccharides.3,4 The observed highly reproducible MALDI spectra were explained as a result of a uniform crystal layer.5 In many cases, the best MALDI performance is usually achieved only at certain locations (sweet spots) of the matrix−analyte crystals. A uniform matrix−analyte crystal layer minimizes the need to search for sweet spots, and more importantly, it avoids the variability of signal intensity across different locations on the target surface due to the heterogeneous crystals and greatly improves spot-tospot reproducibility, which provides a basis for quantitative analysis by MALDI MS. To obtain a uniform crystallization for quantification of MALDI MS, a faster crystallization method, that is, the matrix is prepared in a volatile solvent and/or the MALDI probe is dried at an elevated temperature and/or © 2013 American Chemical Society
Received: February 11, 2013 Accepted: April 10, 2013 Published: April 10, 2013 4729
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acceleration voltage, 19 kV; reflectron voltage, 21 kV; delayed extraction voltage, 18.56 kV; mass range, m/z 400−1000. Data acquisition has been optimized using a random-walk routine under fuzzy logic control so that 600 laser shots are acquired per sample. Data processing was performed using the software “Flex Analysis” version 3.0 (Bruker Daltonics). Mass calibrations were performed externally using CHCA and a peptide mixture (angiotensin II, substance P, bombesin, and ACTH clip 1-17) (Sigma, St. Louis, MO) as mass standards. A mass spectrometer (Applied Biosystems Sciex 3200 Qtrap, Ontario, Canada) was used for electrospray ionization (ESI) MS analysis.19 Preparation of Polystyrene (PS) Spheres. Styrene (A.R. grade) was purchased from Tianjin Institute of Fine Chemicals, China. Before using, the styrene was washed with NaOH (0.1 M) and then water and repeated four times. Dodecyl sodium sulfate (SDS) and K2S2O7 (C.P. grade) were bought from Sinopharm Chemical Reagent Co., China. Polystyrene (PS) spheres with diameters of 150, 240, and 370 nm were prepared via emulsion polymerization.21,22 In a typical synthesis, styrene monomer (10 mL) and ultrapure water (80 mL) were mixed under magnetic stirring, and then SDS (60, 30, and 10 mg for preparation of 150, 240, and 370 nm PS spheres, respectively) was added to form an emulsion. The mixture was kept at 70 °C for 20 min while bubbling with N2 to remove O2 in the system. Then the polymerization was started when a K2S2O7 solution (0.15 g of K2S2O7 dissolved in 10 mL of water) was added dropwise into the mixture with a constant pressure dropping funnel in 10 min. The reaction was sustained for 24 h with the temperature kept at 70 °C. When the reaction was over, the latex was harvested and could be used without further purification. The final concentration of the polystyrene spheres was 11.1% (wt %). The size and morphology of the PS latex spheres were characterized by scanning electron microscopy (SEM JEOL 6701). Quantitiative Analysis of LPCs. Lysophosphatidylcholines (LPCs) are important intermediates as signaling molecules. Plasma LPCs levels were decreased in colorectal cancer (CRC) patients and have the potential as biomarkers for CRC.18,19 Developing a method to accurately quantify the LPC levels in plasma is important and significant. In this experiment, the photonic crystals were first generated by directly depositing 1 μL of polystyrene colloidal spheres (0.22% in water) on the MALDI MS probe. A TiO2 crystal layer was obtained by directly depositing 1 μL of TiO2 solution (0.22% in water) on the probe. The matrix solutions were prepared at a concentration of 50 mg/mL in CH3CN. A volume of 10 μL of matrix, 5 μL of plasma lipid extracts, 5 μL of CH3CN, and 20 μL of water were mixed, and 0.5 μL of the mixture was deposited on the photonic crystals formed by PS spheres or the TiO2 crystal layer for MALDI MS analysis. Standard curves were established for quantitative analyses of all lipids. In brief, different amounts of lipids standards (2.5−50 μmol/L) were mixed with internal standard (12:0 LPC, 12.5 μmol/L) and then mixed with the matrix solution. A volume of 0.5 μL of the mixture was analyzed by MALDI MS. The peak intensity ratios (a lipid/internal standard) versus the molar ratios (a lipid/internal standard) were plotted and fitted to a linear regression. For commercially unavailable lipids, the slopes of the closest structures were used. For example, we used the slope of 18:1 LPC for 18:2 LPC. Statistical Analysis. Correlation analyses were performed using Pearson’s correlation coefficient to assess the associations
of using PS spheres-assisted MALDI MS for quantitative analysis. To examine the feasibility of using this approach, we employed the lipid extracts from plasma as our model samples. Plasma lysophosphatidylcholines (LPCs) levels were decreased in colorectal cancer (CRC) patients and have been identified as the potential diagnostic biomarker for CRC by electrospray ionization (ESI) MS.18,19 In this study, we confirmed the result by MALDI MS for quantitative analysis of plasma LPCs. The accuracy of MALDI MS analysis of LPCs was investigated by comparing the data obtained by ESI MS. Because of the low molecular weight of LPCs (∼400−600 Da), to avoid the chemical noise in low-mass regions caused by excess matrixes in MALDI analysis, the matrix and the solvent were carefully investigated as well to ensure the good-quality and reliable MALDI MS measurements.
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EXPERIMENTAL SECTION Materials and Reagents. Standard lipids, including 1dodecanoyl-sn-glycero-3- phosphocholine PC (12:0 LPC), 1hexadecanoyl-sn-glycero-3-phosphocholine PC (16:0 LPC) and 1-(9Z-octadecenoyl)-sn-glycero-3-phosphocholine PC (18:1 LPC) were purchased from Avanti Polar Lipids (Birmingham, AL). The matrix α-cyano-4-hydroxycinnamic acid (CHCA) and 9-aminoacridine (9-AA) were purchased from Acros (New Jersey). The matrix 2,5-dihydroxybenzoic acid (DHB) was purchased from Lancaster (Morecambe, U.K.). HPLC-grade methanol (MeOH), acetonitrile (CH3CN), and chloroform (CHCl3) were purchased from Fluka Feinchemikalien GmbH (part of Sigma−Aldrich Chemie GmbH, Taufkirchen, Germany). Titanium dioxide (TiO2; CAS, 13463-67-7; SKU, 634662) was purchased from Sigma-Aldrich (St. Louis, MO). Ultrapure water from Milli-Q purification system (Millipore Corporation) was used. All of the above materials were used as received without further purification. Participants. Plasma samples were a gift from Dr. Junjie Zhang,19 including 10 patients with CRC, 10 patients with adenoma, and 10 unaffected controls enrolled between May 2007 and 2010 at the General Hospital of Second Artillery Force of Chinese PLA, Beijing Hospital, and People’s Hospital of Peking University. The project was approved by the Institutional Review Board, and written informed consent forms were signed by participants. Blood Processing and Lipid Extraction. Blood samples were collected in EDTA-containing tubes and centrifuged at 1 750g for 15 min at room temperature. Plasma samples were aliquoted into siliconized eppendorf tubes (SafeSeal microcentrifuge tubes; PGC Scientifics, Frederick, MD) and frozen at −80 °C until use. Lipids were extracted by the Blight and Dyer method with minor modifications.20 Briefly, 20 μL of each plasma sample were added into 80 μL of CHCl3/MeOH (2:1, v/v) with 2 nmol of 12:0 LPC as the internal standard, followed by vortexing for 30 s and incubating for 5 min at room temperature, vortexed again for 30 s, and then centrifuged at 10 000g for 5 min. The upper phase was discarded, and the 50 μL of the lower phase was collected and evaporated to dryness under a nitrogen stream. The lipid extract was resuspended with 25 μL of CH3CN for MALDI MS analysis. MS Instrument Parameters. MALDI-TOF mass spectra were acquired on a Bruker Autoflex mass spectrometer (Bruker Daltonics, Bremen, Germany) in positive ion mode. The device parameters were chosen as follows: laser wavelength, 337 nm; 4730
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between the data obtain by MALDI and ESI MS. The R value at the range from 0.5 to 1.0 means that the variables are highly positive related. If P < 0.05, it means the correlation is significant. The one-way ANOVA (nonparametric) statistical analysis was performed to compare the differences of total plasma LPCs in three groups. P < 0.05 and F > 10 were considered to be statistically significant.
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RESULTS AND DISCUSSION Effect of Matrix. The analyses of organic synthetic compounds, peptides, and proteins using MALDI MS have
Figure 2. Morphologies of DHB (12.5 mg/mL) crystal (a1), DHB− sample cocrystal (a2), TiO2 crystal (b1), TiO2−DHB−sample cocrystal (b2), polystyrene spheres (size, 240 nm) crystal (c1), and polystyrene spheres (size, 240 nm)−DHB−sample cocrystal (c2) on the MALDI probe. Pictures were obtained by the camera equipped in the MS instrument. Pictures of a3, b3, c3, d, e, and f were highresolution images of DHB−sample cocrystal, TiO2−DHB−sample cocrystal, polystyrene spheres (size, 240 nm)−DHB−sample cocrystal, polystyrene spheres (size, 150 nm)−DHB−sample cocrystal, polystyrene spheres (size, 370 nm)−DHB−sample cocrystal, and mixed polystyrene spheres−DHB−sample, respectively, which were obtained by scanning electron microscopy (SEM JEOL 6701). The mixing ratio of PS spheres with different sizes is 1:1:1 in mixed polystyrene spheres in part f. The insert in part c1 was the high-resolution image of the polystyrene spheres (size, 240 nm) crystal. The solvent was CH3CN/ H2O (5:5, v/v).
Figure 1. Mass spectra of plasma LPCs from a CRC patient using DHB (a), CHCA (b), and 9-AA (c) as a matrix, respectively. The insert in part a was an enlarged mass spectrum with the m/z range from 517 to 527. a.i., absolute intensity.
Table 1. m/z Values in Figure 1 with Corresponding QuasiMolecular Ions of LPCsa m/z 440.2 462.2 478.2 496.3 518.3 534.3 520.3 542.3 558.3 522.3 544.3 560.3 524.3 546.3 562.3 544.3 582.3
quasi-molecular ions [12:0 [12:0 [12:0 [16:0 [16:0 [16:0 [18:2 [18:2 [18:2 [18:1 [18:1 [18:1 [18:0 [18:0 [18:0 [20:4 [20:4
LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC LPC
+ + + + + + + + + + + + + + + + +
H]+ Na]+ K]+ H]+ Na]+ K]+ H]+ Na]+ K]+ H]+ Na]+ K]+ H]+ Na]+ K]+ H]+ K]+
Figure 3. Mass spectra of plasma LPCs from a healthy control using DHB (a), polystyrene spheres (size, 150 nm) /DHB (b), and polystyrene spheres (size, 240 nm)/DHB (c) as the matrix, respectively.
a
The numbers, like 12:0, 16:0, and 18:2, stand for the numbers of carbon atom and double bond in the fatty acyl chain. For example, 18:2 LPC means that there are 18 carbon atoms and 2 double bonds in the fatty acyl chain of LPC structure.
matrix. Although the molecular weights of the different matrixes are in the range of about 150−200 g/mol, photoreactions such as trimerizations23 occurring upon laser irradiation as well as
been widely developed. However, the lipid identification using MALDI MS has been limited due to less existence of a proper 4731
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Figure 7. Correlation of total plasma LPC between the results obtained by MALDI MS and ESI MS (P < 0.0001, R2 = 0.8189).
Figure 4. Representative mass spectra of plasma LPCs from an adenoma patient when a laser irradiated different spots using polystyrene spheres (size, 240 nm)/DHB as a matrix, which showed a good spot-to-spot reproducibility.
incomplete matrix cluster decomposition and adduct formation, may generate a multitude of matrix peaks at higher m/z values (100−500 Da), which suppressed or hid the lipid signals with a molecular weight lower than 500. In addition, the lipid extracts from a biological sample are usually a complex system, where the interferences and discriminations of different molecules make it more difficult to be analyzed. The choice of matrix, one of the important issues for MALDI MS, becomes complicated. In this study, three matrixes, DHB, CHCA, and 9-AA, were chosen to screen for the best matrix for LPCs analysis. The concentration of matrix is optimized, and the mass spectra were shown in Figure 1. The m/z values in the mass spectra with corresponding quasi-molecular ions were indicated in Table 1. There is a quaternary amine in the LPC structure, which leads to LPC being detected conveniently in positive ion mode. 9-AA is recently developed for negative charged lipids detection.24 In this study, it was found that 9-AA (3 mg/mL) could be used for positively charged LPCs detection and gave rise to hydrogenated quasi-molecular ions (Figure 1c). However, the signal intensity was weak. CHCA and DHB are classic matrixes, and both of them are widely used for
Figure 5. Standard curves for 16:0 LPC and 18:1 LPC. Cs, the concentration of standard (16:0 LPC and 18:1 LPC); CIs, the concentration of internal standard (12:0 LPC); Is, the MS signal intensity of standard; IIs, the MS signal intensity of internal standard.
Figure 6. Data points of plasma LPCs from a healthy control, adenoma, and CRC. 4732
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polystyrene spheres because of the phenyl group may have the capability of attracting DHB, which leads to DHB attaching to the polystyrene spheres and coself assembling and finally forming a uniform cocrystal. In contrast, the surface of TiO2 is hydrophilic, which may be the reason that TiO2 cannot combine DHB into a uniform crystal. Polystyrene (PS) colloidal spheres with the diameter at around 150 and 370 nm were further tested. On the surface of the photonic crystal formed by PS colloidal spheres with the diameter at around 150 nm, DHB and analyte distribute evenly, and the cocrystal stick together in muddy texture (Figure 2d). In contrast, on the surface of the photonic crystal formed by PS colloidal spheres with the diameter at around 370 nm, the DHB−analyte cocrystal did not distribute evenly (Figure 2e). Interestingly, on the surface formed by mixed polystyrene spheres with different sizes (the mixing ratio, 1:1:1, v/v/v), the cocrystal became homogeneous again (Figure 2f). These results suggested that the morphology of cocrystal is related with the size of polystyrene spheres, and it seemed that the smaller the spheres are the more homogeneous the cocrystal will be. The presence of small spheres in mixed polystyrene spheres explained the reason for the homogeneous cocrystal in Figure 2f. The results of MALDI MS analysis of plasma lipid extracts on these surfaces were shown in Figure 3. DHB alone and all the polystyrene spheres−DHB as matrix gave the LPCs signals. Compared with DHB alone as the matrix (Figure 3a), it is clear that the sodiated quasi-molecule ions of LPCs are decreased when PS spheres were used together with DHB as the matrix for MALDI MS analysis (shown in Figure 3b,c), which simplify the MS spectra. As a matter of fact, the quasi-molecule ions of [18:1 LPC + Na]+ and [20:4 LPC + H]+ are all 544.3 (seen in Table 1), so that we cannot quantitatively analyze 20:4 LPC when using DHB alone as a matrix due to the interference of 18:1 LPC. However, the sodiated quasi-molecule ions of LPC almost disappear after using PS spheres, so the signal of 544.3 only comes from hydrogenated 20:4 LPC, which can be used for quantitative analysis of 20:4 LPC. The surface of PS spheres is negative charged,28,29 so positive Na+ could be captured by the PS spheres, which may inhibit the sodiated quasi-molecule ions formation. In addition, the excess K+ added (0.15 g of K 2 S 2 O 7 ) when synthesizing PS spheres increased the potassiated quasi-molecule ions of LPCs (Figure 3b,c, for example, [12:0 LPC + K]+ at m/z 478.2 and [16:0 LPC + K]+ at m/z 534.3). Furthermore, on the surface formed by polystyrene spheres with a size of 150 nm, 240 nm, and mixed, where the crystals were uniform; undoubtedly, a satisfied spot-to-spot reproducibility was obtained (shown in Figure 4) with the standard derivation (SD) lower than 4.1%. However, on the surface formed by polystyrene spheres with a size of 150 nm (Figure 3b) and mixed (similar with Figure 3b, data not shown), the MS signals were relatively low compared with those in Figure 3c. Their cocrystals are smaller (Figure 2d, f, compared with Figure 2c3) and could be embedded more deeply by PS spheres, which could decrease the MS signals. Therefore, PS spheres with a size at 240 nm were used for the following experiments to form the uniform cocrystal surface. The concentration of PS spheres (size, 240 nm) was optimized as well. Below the concentration of 0.22% (wt %), the cocrystal showed an uneven surface. Above the concentration of 0.22% (wt %), the MS spectra showed low S/N ratios. The most appropriate concentration is 0.22% (wt
identification of peptides or carbohydrates, etc. When DHB (12.5 mg/mL) is the matrix, besides hydrogenated quasimolecular ions, sodiated and potassiated quasi-molecule ions were also important ionized species for LPCs detection. For example, the hydrogenated (m/z at 440.2), sodiated (m/z at 462.2), and potassiated (m/z at 478.2) quasi-molecular ions of 12:0 LPC were all very strong in the mass spectrum (Figure 1a). However, when CHCA (5 mg/mL) is the matrix, the hydrogenated quasi-molecular ions were dominated (Figure 1b). From the results (Figure 1a,b), it seemed that DHB gave rise to a much stronger MS signal. Although the CHCA matrix produces relatively homogeneous crystals, a higher laser energy was necessary for desorption and ionization of LPCs in the MS ion source, which leads to a lower S/N ratio. Taken them together, DHB is the best matrix for analysis of plasma LPCs among these three matrixes, characterized by high sensitivity and high S/N ratio. PS Spheres-Assisted MALDI MS Analysis. One obvious drawback of DHB (12.5 mg/mL) as a MALDI matrix is its tendency to form large crystals (shown in Figures 2a1 and 3), where the DHB−analyte cocrystal is not evenly distributed, leading to poor spot-to-spot reproducibility. To change the DHB crystal morphology, increasing the ratio of organic solvent in the solution for dissolving DHB was suggested.6 In this study, different dual solvent systems, including CH3CN/ H2O, MeOH/H2O, and MeOH/CHCl3, their different ratios, and many additives, such as NH4OH, ammonium formate, and ammonium acetic were tested. However, all failed to obtain a uniform DHB−analyte cocrystal layer. CH3CN/H2O (5:5, v/v) gave rise to a strong MS signal and low interference signal; therefore, in the following experiments, it was used as a solution to dissolve the matrix and analyte. To obtain a uniform crystal layer to improve spot-to-spot reproducibility for quantitative analysis by MALDI MS, a hypothesis, which is nanoparticles may act as a crystal nucleus to assist and accelerate DHB crystallization, to prevent DHB from forming a large crystal, and finally help DHB form a uniform crystal, was proposed. It was reported that TiO2 nanoparticles can be used as an inorganic matrix for LDI MS analysis,25,26 and when using nanoparticles as a matrix, it has an advantage to tend to form a homogeneous crystallization.27 TiO2 itself cannot be used alone for quantitative analysis of LPC in plasma due to the low sensitivity. Therefore, it was expected that TiO2 could assist DHB to form a uniform crystal and then perform quantitative analysis of LPCs. TiO2 with a diameter less than 100 nm were first tested, and the morphologies of crystal were shown in Figure 2b. TiO2 itself can form a uniform crystal (Figure 2b1); unfortunately, after dropping the mixed solution of DHB and analyte, the morphology of the cocrystal is very similar with that when DHB is alone as the matrix (Figure 2b2,a2). Polystyrene (PS) colloidal spheres with the diameter at around 240 nm were then tested. PS colloidal spheres have a superior monodispersed property and can self-assemble to form photonic crystals, a uniform crystal layer, shown in Figure 2c1. When we drop the mixed solution of DHB and analyte on the polystyrene spheres crystal layer and after drying, a relatively uniform crystal layer was obtained (shown in Figure 2c2). The higher resolution image (Figure 2c3) showed the details of the cocrystal: there was not a large DHB crystal, the cocrystal became much smaller, and the polystyrene spheres inserted into the cocrystal to separate them and finally form a regular surface. Considering the reason, the hydrophobic surface of 4733
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(5) David, J. H. Mass Spectrom. Rev. 2011, 31, 183−311. (6) Fujiwaki, T.; Tasaka, M.; Yamaguchi, S. J. Chromatogr., B 2008, 870, 170−176. (7) Nicola, A. J.; Gusev, A. I.; Proctor, A.; Jackson, E. K.; Hercules, D. M. Rapid Commun. Mass Spectrom. 1995, 9, 1164−1171. (8) Kailasa, S. K.; Wu, H. F. Analyst 2012, 137, 1629−1638. (9) Lin, P. C.; Tseng, M. C.; Su, A. K.; Chen, Y. J.; Lin, C. C. Anal. Chem. 2007, 79, 3401−3408. (10) Ho, Y. C.; Tseng, M. C.; Lu, Y. W.; Lin, C. C.; Chen, Y. J.; Fuh, M. R. Anal. Chim. Acta 2011, 697, 1−7. (11) Li, H.; Wang, J. X.; Lin, H.; Xu, L.; Xu, W.; Wang, R. M.; Song, Y. L.; Zhu, D. B. Adv. Mater. 2010, 22, 1237−1241. (12) Chai, G. S.; Shin, I. S.; Yu, J. S. Adv. Mater. 2004, 16, 2057− 2061. (13) Solodukhina, N. M.; Zlydneva, L. A.; Levshenko, E. N.; Myagkova, M. A.; Gritskova, I. A. Appl. Biochem. Micro. 2012, 48, 740−745. (14) Duan, G. T.; Cai, W. P.; Li, Y.; Li, Z. G.; Cao, B. Q.; Luo, Y. Y. J. Phys. Chem. B 2006, 110, 7184−7188. (15) Liu, G. Q.; Cai, W. P.; Kong, L. C.; Duan, G. T.; Li, Y.; Wang, J. J.; Zuo, G. M.; Cheng, Z. X. J. Mater. Chem. 2012, 22, 3177−3184. (16) Zhang, Z. B.; Shen, W. Z.; Ye, C. Q.; Luo, Y. M.; Li, S. H.; Li, M. Z.; Xu, C. H.; Song, Y. L. J. Mater. Chem. 2012, 22, 5300−5303. (17) Li, M. Z.; Xia, A. D.; Wang, J. X.; Song, Y. L.; Jiang, L. Chem. Phys. Lett. 2007, 444, 287−291. (18) Zhao, Z.; Xiao, Y.; Elson, P.; Tan, H.; Plummer, S. J.; Berk, M.; Aung, P. P.; Lavery, I. C.; Achkar, J. P.; Li, L.; Casey, G.; Xu, Y. J. Clin. Oncol. 2007, 25, 2696−2701. (19) Li, S.; Guo, B.; Song, J.; Deng, X.; Cong, Y.; Li, P.; Zhao, K.; Liu, L.; Xiao, G.; Xu, F.; Ye, Y.; Zhao, Z.; Yu, M.; Xu, Y.; Sang, J.; Zhang, J. Metabolomics 2013, 9, 202−212. (20) Bligh, E. G.; Dyer, W. J. Can. J. Biochem. Physiol 1959, 37, 911− 917. (21) Du, X.; He, J. J. Appl. Polym. Sci. 2008, 108, 1755−1760. (22) An, G .; Gao, L.; Zhao, X. J. Funct. Mater. 2010, 41, 1571−1574. (23) Hoyer, T.; Tuszynski, W.; Lienau, C. Chem. Phys. Lett. 2007, 443, 107−112. (24) Fuchs, B.; Bischoff, A.; Suss, R.; Teuber, K.; Schuerenberg, M.; Suckau, D.; Schiller, J. Anal. Bioanal. Chem. 2009, 395, 2479−2487. (25) Shrivas, K.; Hayasaka, T.; Sugiura, Y.; Setou, M. Anal. Chem. 2011, 83, 7283−7289. (26) Torta, F.; Fusi, M.; Casari, C. S.; Bottani, C. E.; Bachi, A. J. Proteome Res. 2009, 8, 1932−1942. (27) Chiang, C. K.; Chen, W. T.; Chang, H. T. Chem. Soc. Rev. 2011, 40, 1269−1281. (28) Deng, T. S.; Marlow, F. Chem. Mater. 2012, 24, 536−542. (29) Sun, B.; Mutch, S. A.; Lorenz, R. M.; Chiu, D. T. Langmuir 2005, 21, 10763−10769.
%), where the crystal appearance is uniform and MS signal is strong enough (Figure 3c). Quantitative Analysis of LPC in Plasma. The standard curves for 16:0 LPC and 18:1 LPC were shown in Figure 5, plasma LPCs data were shown in Figure 6, and the correlation analysis of total LPCs between MALDI MS and ESI MS analysis were shown in Figure 7. The total plasma LPCs decreased in adenoma patients (171 ± 35) and decreased further in CRC patients (127 ± 34), compared with healthy controls (281 ± 82), consistent with the previous results.19 The one-way ANOVA (nonparametric) statistical analysis showed the P < 0.0001 with F = 18.62, meaning total LPCs in these three groups were significantly different. The data obtained by MALDI MS were significantly correlated with those obtained by ESI MS by Pearson’s Correlation Coefficient analysis (Figure 7, P < 0.0001, R2 = 0.8189), suggesting this approach of using polystyrene spheres-assisted MALDI MS for quantitative analysis of plasma LPCs was effective.
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CONCLUSIONS With the assistance of PS spheres (size, 240 nm), a uniform DHB−analyte crystal layer was realized, which can be used for the quantitative analysis of plasma LPCs by MALDI MS. The spot-to-spot reproducibility was satisfied with SD lower than 4.1%. The plasma LPCs data obtained by MALDI MS were significantly correlated with those obtained by ESI MS. Interestingly, the DHB−analyte crystals were related to the size of the PS spheres, and it seemed that the smaller the spheres were, the more homogeneous the cocrystals would be. However, the DHB−analyte crystals in a muddy texture when using a PS sphere with the size at 150 nm did not give rise to a strong MS signal. In our experiments, the PS spheres with a size at 240 nm were appropriate. With the assistance of PS spheres, the sodiated quasi-molecule ions of LPCs were clearly decreased, which simplified the MS spectra. The approach, photonic crystals formed by PS colloidal spheres-assisted MALDI MS, is first reported, which provides a simple way for quantitative analysis, and it was believed that this approach would greatly extend the uses of MALDI MS.
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AUTHOR INFORMATION
Corresponding Author
*Phone: +86-10-62561239. Fax: +86-10-62561285. E-mail:
[email protected]. Author Contributions ⊥
Y.W. and S.L. contributed equally to this work.
Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by NSF Grant 91127029/B040306, 973 Program (No. 2011CB302100) and MOST (Grant No. 2011IM030200).
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REFERENCES
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