Organosilica Microspheres

Enrichment and Detection of Peptides from Biological Systems Using Designed Periodic Mesoporous Organosilica Microspheres

Kun Qian, Wenyi Gu, Pei Yuan, Fang Liu, Yunhua Wang, Michael Monteiro, and Chengzhong Yu*

Nanoporous materials with uniform and adjustable pore sizes (1–100 nm) have important applications in modern biotechnology.[1] Many biomolecules, such as peptides and proteins, have crucial functions in biological systems and exist in nanoscale sizes (1–100 nm) at extremely low con-centrations. In order to separate or identify biomolecules in biological systems, nanoporous materials with carefully designed pore structures and pore sizes are ideal hosts. By further manipulating the composition, surface chemistry, and morphology of nanoporous materials, their applications in diverse biological areas are promising but as yet require fur-ther exploration.[2,3]

Enrichment of peptidome fractions is a key to identify which peptides control molecular and cellular pathways in life science.[4,5] Mass spectrometry (MS)-based peptide map-ping is one of the fundamental tools for current peptidome research.[6,7] Detection of functional peptides in biosys-tems (e.g., in serum and on the cell surface) is currently an important but extremely challenging topic. Serum consists of a complex biological fluid, in which the target peptide may exist in a very low abundance and is mixed with a large number of other peptides and proteins, making detection in such a biosystem difficult. For one example, the enrichment and detection of the cytotoxic lymphocyte (CTL) epitope of E7 (a functional peptide residue) in biosystems is of signifi-cant importance in the immunotherapy of human papilloma-virus (HPV)-related diseases, and has consequently attracted intensive attention in pathological and clinical studies.[8–10] Very recently, the E7 peptide has been shown effective as a preventive therapeutic vaccine to generate immune responses and eliminate tumour growth in clinical-phase trials.[9] For

© 2012 Wiley-VCH Verlag Gmsmall 2012, 8, No. 2, 231–236

DOI: 10.1002/smll.201101770

K. Qian, W. Gu, P. Yuan, Prof. M. Monteiro, Prof. C. YuAustralian Institute for Bioengineering and Nanotechnology The University of Queensland Brisbane, QLD 4072, Australia Fax: (+)61-7-33463973 E-mail: c.yu@uq.edu.au

F. Liu, Prof. Y. WangDepartment of Chemistry Fudan University Shanghai 200433, China

future clinical applications, the detection of E7 epitope in serum is important as the serum level of E7 peptides is highly associated with the resultant immune response, however there are no reports in the direct detection of E7 epitope in serum. For another example, in order to detect cell surface peptides, large amounts of cells (≈108–109) were necessary and expen-sive equipments were used in order to separate and elute the peptides before analysis.[6,11]

Numerous materials have been developed to isolate pep-tides modified by specific chemical groups, such as the phos-phor group[12] and the glycol group.[13] Nevertheless, many important peptides like the E7 epitope do not have such modifications naturally and thus become difficult to enrich through chemo-affinities. In this regard, materials with a high general peptide enrichment capacity will be good can-didates.[14,15] In order to detect functional peptides from bio-logical systems with high sensitivity, it is important to achieve high efficient peptide enrichment while excluding proteins. For example, the poly(methyl methacrylate) (PMMA)-type material is reported to enrich both proteins and peptides,[16,17] but the proteins can strongly interfere with the peptide signal and thus affect the analysis of peptides.[18,19]

Herein, we report the enrichment and detection of the E7 peptide from biological systems by using a carefully designed periodic mesoporous organosilica microsphere (PMOM) material. The PMOM is designed in our study on the basis of the following hypothesis: 1) Compared to silica materials generally used in the enrichment of peptides,[18] PMO with homogenously dispersed organic groups in its framework has been demonstrated to show good peptide enrichment efficiency;[20,21] 2) PMOMs have a small pore size of 3.1 nm and a cubic Pm-3n mesostructure,[22] and both parameters were previously shown to be advantageous for the size-exclusive enrichment of peptides;[18,19,23] and 3) PMOMs have a spherical morphology with micrometer diameters, and thus undesired nonspecific adsorption towards proteins can be minimized due to a reduced outer surface area. The combination of the above advantages of PMOMs has led to its excellent size-exclusive peptide enrichment performance. By using PMOMs, E7 can be successfully detected in serum and protein mixtures; moreover, this approach is also suc-cessfully employed in the analysis of protein digests and a cell-washing solution to identify peptides from the surface of TC1 cells.[8]

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Figure 1. A) XRD pattern, B) TEM image, C) SEM image, and D) pore-size distribution curve (inset is the nitrogen sorption isotherm) of PMOM.

The PMOM material is prepared by using cetyltrimethy-lammonium bromide (CTAB) surfactant as a template and 1,2-bis(trimethoxysilyl)ethane (BTME) as a silica source under basic conditions (see the Supporting Information (SI) for details). The X-ray diffraction (XRD) pattern of PMOM is shown in Figure 1A. Three well-resolved peaks are observed in the range of 1–2° (2θ), which can be assigned to the (200), (210), and (211) reflections for a cubic Pm-3n structure.[22,24] Another five diffraction peaks in the range of 2–4° are indexed as (222), (321), (400), (420), and (431) reflec-tions. The unit cell, a, is calculated to be 10.18 nm. Transmis-sion electron microscopy (TEM) images show the typical ordered mesopore array recorded along the [100] direction (Figure 1B) and [111] direction (inset of Figure 1B), similar to previous literature reports.[22,24] The scanning electron micro-scopy (SEM) images in Figure 1C and the inset display that PMOM has a spherical morphology with particle sizes of 2–5 μm. The nitrogen adsorption analysis depicts a Type IV isotherm, as shown in the inset of Figure 1D. PMOM exhibits a uniform pore size distribution centred at 3.1 nm (Figure 1D). The sur-face area and pore volume of PMOM are calculated to be 612 m2 g−1 and 0.56 cm3 g−1, respectively.

The Fourier-transform infrared (FTIR) spectrum of PMOM is shown in Figure 2A, which displays typical bands at ≈2900, 1450, and 1680 cm−1 (marked with o), corresponding to the aliphatic C–H stretching, bending vibrations, and carbonyl stretching vibration, respectively.[20,25] There are also two other bands present at 1260 and 700 cm−1 (marked with #), which are typically attributed to the Si–C bond.[26] The presence

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of the C–H and Si–C peaks provides evidence of ethane attached to Si. The 29Si magic-angle spinning nuclear magnetic resonance (MAS-NMR) spectrum features two signals at –57 and –64 ppm, which are indexed as the T2 [CSi(OSi)2OH] and T3 [CSi(OSi)3] silicon resonances, respectively (Figure 2B). In addition, a relatively weak signal at –47 ppm can also be observed and assigned to the T1 [C–Si(OH)2(OSi)] species. No Q1–Q4 peaks (Si–(OSi)x(OH)4-x, x = 1–4) at lower ppm are observed, indicating that the Si–C bond cleavage does not occur in the preparation process of PMOM.[21,24,25]

Previously, we prepared periodic mesoporous organo-silica nanoparticles (PMONs) with the same composition and structure as PMOMs but smaller in particle size (see SI, Table S1).[23] We further compared the protein immobi-lization performance of two materials using bovine serum albumin (BSA) as an example. The BSA immobilization capacity is 21.7 mg g−1 for PMOMs, less than one third of the adsorbed BSA amount in PMONs (69.6 mg g−1). BSA has a large molecular weight (66 kDa) and large size (≈10 nm) in buffer solution,[27] thus cannot enter the mes-opores of PMONs or PMOMs with pore sizes smaller than 3.2 nm. The adsorption of BSA should preferentially occur at the external surface of PMO materials. PMOMs with larger particle size has smaller external surface area compared to PMONs, leading to reduced adsorption amount of BSA. It is suggested that size-exclusive adsorbents with a larger par-ticle size (i.e., PMOMs) will reduce surface adsorption, espe-cially of large proteins, and advantageously aid in isolating peptides from protein/peptide mixtures.

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Detection of Peptides Using Organosilica Microspheres

Figure 2. A) FTIR and B) 29Si NMR spectra of PMOM.

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The efficiency of PMOMs in enriching the immunopep-tide E7 (sequence RAHYNIVTF, molecular weight of 1120.6 Da, isoelectric point of 8.75)[8,9,28] in various mixtures was detected via matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS). Figure 3a displays the MS spectrum obtained from an E7 peptide solu-tion (8.9 nm). At such a low concentration, no E7 signals can be detected. After enriched by PMOM, well-resolved peak with signal strength over 12 000 and S/N ratio over 2000 is observed (Figure 3b). The MS/MS spectrum confirms the sequence of RAHYNIVTF residues as shown in SI, Figure S1a. When the concentration of E7 is decreased to 0.89 nm (SI, Figure S1b), E7 is still detected after enrichment. E7 has an optimized linear conformation with a length of ≈3 nm and a diameter of 1.2 nm in water (data not shown), thus can enter PMOMs with a pore size of 3.1 nm. The total surface area is 612 m2 g−1 while the outer surface area is estimated to be ≈1 m2 g−1 for PMOMs (assuming a mean spherical particle size of 3 μm and a density of 2 g cm−3), thus it can be con-cluded that E7 peptides are predominately adsorbed inside the mesopores.

In order to study the enrichment and recovery efficiency of E7 peptide by PMOMs, further experiments were car-ried out at a high concentration of 8.9 μm. As shown in SI, Figure S1b, a strong E7 peptide peak is viewed in the bulk solution, however, no E7 peptide signal can be observed from the solution after enrichment by PMOMs (the inset of

© 2012 Wiley-VCH Verlag Gmbsmall 2012, 8, No. 2, 231–236

SI, Figure S1c), suggesting that the E7 peptide concentration has been deceased to <8.9 nm (see Figure 3a) and >99% E7 peptides have been absorbed into the PMOMs. The enriched E7 peptide was then eluted by aqueous acetonitrile solution (see Experimental Section)[18] in order to study the elution efficiency. It is noted that the concentration of E7 peptide cannot be directly determined from the peak intensity in MALDI-TOF spectra due to the unpredictable analyte-dependent ion-production behaviour. Therefore, a synthetic signature peptide was prepared as an internal standard (IS) to allow label-free quantification of the E7 peptide concen-tration by MALDI-TOF-MS.[29] The signature peptide has a sequence of RAHYNIATF (molecular weight 1092.2 Da), only one amino-acid residue difference compared to E7 (RAHYNIVTF); thus it should have a similar desorption/ion-ization performance with E7. SI, Figure S1d shows the mass spectrum from the mixture of E7 elution added with equal volume of 3 μg mL−1 of IS. By preparing a series of standard E7 peptides mixed with IS, the standard curve was obtained by measuring the signal ratio of E7/IS, which shows a linear relationship (R2, the regression coefficient, was 0.987) with the E7 concentration (SI, Figure S1e). From SI, Figure S1d,e, it can be calculated that ≈67% of enriched E7 peptides can be recovered by elution. Although the enrichment and elu-tion efficiency at a much lower concentrations of 0.89–8.9 nm (equivalent to 0.45–4.5 fmol of E7) in our experiments cannot be quantified by this method, it is concluded that PMOMs can enrich peptides and the adsorbed peptides can be effec-tively recovered by elution, thus the detection of limit can be enhanced.

To further demonstrate the advantage of PMOMs, size-selective extraction of E7 from E7/protein mixtures was performed. In a mixture of E7/BSA (1/10 000, w/w), no E7 peptide peak can be detected in the low MW range at 800–2000 Da (Figure 3c), while signals from BSA are observed in the high MW range (10–80 kDa, inset of Figure 3c). In parallel, after size-exclusive enrichment by PMOMs, the E7 peptide peak can be seen at 1120.60 Da with a high S/N ratio with no protein signals in the high MW range (Figure 3d). Even when the concentration of BSA is raised to 1 mg mL−1 (E7/BSA weight ratio of 1/100 000), PMOM can still extract E7 from the protein-overloaded mixture (SI, Figure S1f). Considering the amount of PMOMs (10 μg) used for enrichment in our approach and the measured adsorption ability of PMOMs (21.7 mg g−1, SI, Table S1), it is estimated that only ≈0.2 μg of BSA can be absorbed at the outer surface of PMOMs. By a further washing procedure after enrichment, it is suggested that the amount of proteins adsorbed at the outer surface of PMOMs can be significantly reduced, thus no signals of BSA are observed (Figure 3d).

In addition to the high-molecular-weight BSA, the influ-ence of proteins with smaller molecular weights was further tested. It is shown that E7 can be extracted and detected from two standard protein/E7 mixtures including ovalbumin (MW 43 kDa, SI, Figure S2a,b) and beta-casein (MW 26 kDa, SI, Figure S2c,d). We also prepared a complex system containing the above three proteins mixed together with E7 (E7/protein ratio of 1/10 000 for each protein, w/w). No peptide can be detected and only strong signals from proteins can be seen in

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Figure 3. The mass spectra obtained from a) 8.9 nm E7 peptide solution, b) 8.9 nm E7 after enrichment by PMOMs; c) a mixture of 8.9 nm E7 in 0.1 mg mL−1 BSA, d) the same mixture as in (c) after enrichment by PMOMs; e) mice serum after injection with E7, and; f) the same serum as in (e) after enrichment by PMOMs. The insets of (c–f) show the high molecular-weight range (10–80 kDa) in the MS. The * represent for E7 peptide signal. K1, K2, and K3 are the single, double, and triple-charged protein peaks.

the bulk solution (SI, Figure S2e), whereas only E7 peptide signal is observed after PMOMs enrichment (SI, Figure S2f).

In the next step, this approach was employed to examine a biological serum sample from the E7 injected mice that

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mimics E7 peptide immunisation in mice (See SI, Experi-mental Section). As displayed in Figure 3e, the mice serum sample contains quantities of proteins, which suppress the E7 signal. On the other hand, by using PMOMs, the E7 peptide

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Detection of Peptides Using Organosilica Microspheres

Figure 4. Comparison of mass spectra achieved from a) digests of casein protein (5 nm) after enrichment by the PMOMs and before enrichment (the inset) and b) the TC1 cell surface washing buffer directly (the inset) and after enrichment by the PMOMs. The marked stars stand for the typical identified peptides (SI, Table S4).

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is detected with high resolution and the protein signals are greatly reduced. The results indicate that these nanoporous microspheres can be used effectively to analyze peptides from complicated biological systems such as serum, which otherwise cannot be directly detected.

Salts usually exist in biological samples and disturb the cocrystallization of analyte and matrix molecules, which is undesirable in the MALDI process and tends to decrease the sensitivity. It is noted that PMOMs can be effectively used to isolate and detect the E7 peptides (8.9 nm) in a salt-containing solution in SI, Figure S3a (66 mm NaCl and 100 nm urea), suggesting that PMOMs with hydrophobic walls possess a high affinity for peptides even in salty solutions, advantageous for the enrichment application of PMOMs in biological systems. The influence of surfactant on the enrich-ment process is also examined using a commercial surfactant (Brij 78). After enrichment by PMOM (8.9 nm of E7 peptide in 8.7 μm of Brij78), the E7 peptide can be detected together with the signals from Brij 78 (SI, Figure S3b), indicating that Brij 78 can also be absorbed by PMOMs and the surfactants used in the protein extraction may have negative influence towards the peptide enrichment effect.

To demonstrate another application potential of PMOM, E7 peptides were detected in the widely used cell-culture media. As displayed in SI, Figure S4ab, E7 peptides have been isolated by PMOMs for detection at a low concentration of 8.9 nm in Dulbecco’s Modified Eagles Medium (DMEM), which is used for mammalian cell culture. On the contrary, no peptides were observed without enrichment by PMOMs. Similar results are observed for E7 detection in another cell culture medium (Clicks’ medium for suspension culture, SI, Figure S4cd). The above experiments show that our strategy can be applied in both molecular and cellular peptidome research in cell experiments.[8,9,28]

Besides the standard peptide, PMOMs can enrich peptide mixtures as well. In the case of beta-casein protein digests at 5 nm (Figure 4a), the protein is identified with a sequence cov-erage of only 8% before enrichment, but the sequence cov-erage increases sharply to 40% after treating with PMOMs. In details, only 2 weak peptides signals at m/z 830 (AVPYPQR) and 1384 (LLYQEPVLGPVR) are identified in the spectrum before enrichment with the S/N ratio of 55 and 194 (316 and 1156 for intensity, SI, Table S2, and inset of Figure 4a), respec-tively. For comparison, 11 peptides are identified (SI, Table S3) and 9 fine-resolved peptides are observed (Figure 4a), where the MS signals at m/z 830, 1384 are highly enhanced with the S/N ratio of 194 and 2913 (2382 and 24799 for intensity, Figure 4a), respectively, after enrichment by PMOMs.

Finally, PMOM was employed to examine the cell surface peptidome in the TC1 cell line.[8] In the mass spectrum of the cell washing solution, no peptides can be detected and only very weak signals with signal intensity <15 are observed (inset of Figure 4b). For comparison, when using PMOM for enrich-ment, a range of peptides are identified with signal intensity up to 2000, and 22 peptides have been successfully identified from the TC1 cell surface (see details in SI, Table S4). It is noted that compared with previous reports,[6,11] our approach uses less amount of cells (≈106), and PMOMs can be used to replace expensive equipment and avoid time-consuming

© 2012 Wiley-VCH Verlag Gmbsmall 2012, 8, No. 2, 231–236

procedures in the enrichment and detection of low-abun-dance peptides on cell surfaces.

In summary, we have designed a PMOM material with integrated structural features, including a cubic mesostruc-ture, hybrid organosilica wall composition, a uniform pore size of ≈3 nm, and a spherical morphology in micrometers, all advantageous for size-selective and highly efficient capture of peptides from protein/peptide mixtures. Consequently, the PMOM material provides a simple, cheap, and efficient method to capture peptides in a range of complex biological systems, including mice serum and cell-washing buffers for a peptidome of cell surfaces, which are important in many bio-logical applications.

Supporting Information

Supporting Information is available from the Wiley Online Library or from the author.

Acknowledgements

This work was supported by the Australia Research Council and 973 Program (2010CB226901) of China. We thank Wan JJ for providing the signature peptide.

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[1] J. Rabone, Y. F. Yue, S. Y. Chong, K. C. Stylianou, J. Bacsa,

D. Bradshaw, G. R. Darling, N. G. Berry, Y. Z. Khimyak, A. Y. Ganin, P. Wiper, J. B. Claridge, M. J. Rosseinsky, Science 2010, 329, 1053.

[2] C. Sanchez, P. Belleville, M. Popall, L. Nicole, Chem. Soc. Rev. 2011, 40, 696.

[3] N. Nassif, J. Livage, Chem. Soc. Rev. 2011, 40, 849. [4] K. R. Jordan, R. H. McMahan, C. B. Kemmler, J. W. Kappler,

J. E. Slansky, Proc. Natl. Acad. Sci. USA 2010, 107, 4652. [5] C. Y. Janda, J. Li, C. Oubridge, H. Hernandez, C. V. Robinson,

K. Nagai, Nature 2010, 465, 507. [6] M. H. Fortier, E. Caron, M. P. Hardy, G. Voisin, S. Lemieux,

C. Perreault, P. Thibault, J. Exp. Med. 2008, 205, 595. [7] C. C. Oliveira, P. A. van Veelen, B. Querido, A. de Ru, M. Sluijter,

S. Laban, S. H. van der Burg, R. Offringa, T. van Hall, J. Exp. Med. 2010, 207, 207.

[8] W. Y. Gu, M. Cochrane, G. R. Leggatt, E. Payne, A. Choyce, F. Zhou, R. Tindle, N. A. J. McMillan, Proc. Natl. Acad. Sci. USA 2009, 106, 8314.

[9] M. J. P. Welters, G. G. Kenter, P. J. D. van Steenwijk, M. J. G. Lowik, D. M. A. Berends-van der Meer, F. Essahsah, L. F. M. Stynenbosch, A. P. G. Vloon, T. H. Ramwadhdoebe, S. J. Piersma, J. M. van der Hulst, A. Valentijn, L. M. Fathers, J. W. Drijfhout, K. Franken, J. Oostendorp, G. J. Fleuren, C. J. M. Melief, S. H. van der Burg, Proc. Natl. Acad. Sci. USA 2010, 107, 11895.

[10] D. Cai, L. Ren, H. Z. Zhao, C. J. Xu, L. Zhang, Y. Yu, H. Z. Wang, Y. C. Lan, M. F. Roberts, J. H. Chuang, M. J. Naughton, Z. F. Ren, T. C. Chiles, Nat. Nanotechnol. 2010, 5, 597.

[11] S. Hofmann, M. Gluckmann, S. Kausche, A. Schmidt, C. Corvey, R. Lichtenfels, C. Huber, C. Albrecht, M. Karas, W. Herr, Mol. Cell. Proteomics 2005, 4, 1888.

[12] M. Mazanek, G. Mituloviae, F. Herzog, C. Stingl, J. R. A. Hutchins, J. M. Peters, K. Mechtler, Nat. Protoc. 2007, 2, 1059.

[13] J. Nilsson, U. Ruetschi, A. Halim, C. Hesse, E. Carlsohn, G. Brinkmalm, G. Larson, Nat. Methods 2009, 6, 809.

[14] W. T. Jia, X. H. Chen, H. J. Lu, P. Y. Yang, Angew. Chem. Int. Edit. 2006, 45, 3345.

6 www.small-journal.com © 2012 Wiley-VCH V

[15] Y. H. Zhang, X. Y. Wang, W. Shan, B. Y. Wu, H. Z. Fan, X. J. Yu, Y. Tang, P. Y. Yang, Angew. Chem. Int. Edit. 2005, 44, 615.

[16] H. M. Xiong, X. Y. Guan, L. H. Jin, W. W. Shen, H. J. Lu, Y. Y. Xia, Angew. Chem. Int. Edit. 2008, 47, 4204.

[17] H. M. Chen, C. H. Deng, X. M. Zhang, Angew. Chem. Int. Edit. 2010, 49, 607.

[18] R. J. Tian, H. Zhang, M. L. Ye, X. G. Jiang, L. H. Hu, X. Li, X. H. Bao, H. F. Zou, Angew. Chem. Int. Edit. 2007, 46, 962.

[19] S. S. Liu, H. M. Chen, X. H. Lu, C. H. Deng, X. M. Zhang, P. Y. Yang, Angew. Chem. Int. Edit. 2010, 49, 7557.

[20] J. J. Wan, K. Qian, J. Zhang, F. Liu, Y. H. Wang, P. Y. Yang, B. H. Liu, C. Z. Yu, Langmuir 2010, 26, 7444.

[21] S. Inagaki, S. Guan, T. Ohsuna, O. Terasaki, Nature 2002, 416, 304.

[22] Y. C. Liang, E. S. Erichsen, M. Hanzlik, R. Anwander, Chem. Mat. 2008, 20, 1451.

[23] F. Liu, P. Yua, J. J. Wan, K. Qian, G. F. Wei, J. Yang, B. H. Liu, Y. H. Wang, C. Z. Yu, J. Nanosci. Nanotechnol. 2011, 11, 5215.

[24] Y. C. Liang, M. Hanzlik, R. Anwander, J. Mater. Chem. 2005, 15, 3919.

[25] L. Zhang, J. Liu, J. Yang, Q. H. Yang, C. Li, Microporous Mesopo-rous Mat. 2008, 109, 172.

[26] H. Ohmi, Y. Hamaoka, D. Kamada, H. Kakiuchi, K. Yasutake, Thin Solid Films 2010, 519, 11.

[27] D. M. Togashi, A. G. Ryder, D. Mc Mahon, P. Dunne, J. McManus, in Diagnostic Optical Spectroscopy in Biomedicine IV, Vol. 6628 (Eds: D. Schweitzer, M. Fitzmaurice), SPIE Int. Soc. Optical Engi-neering, Bellingham, 2007, p. K6281.

[28] W. Y. Gu, L. N. Putral, A. Irving, N. A. J. McMillan, Curr. Opin. Mol. Ther. 2007, 9, 126.

[29] M. Cuollo, S. Caira, O. Fierro, G. Pinto, G. Picariello, F. Addeo, Rapid Commun. Mass Spectrom. 2010, 24, 1687.

Received: August 30, 2011 Revised: September 26, 2011 Published online: December 2, 2011

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