Easy, Robust, and Repeatable Online Acid Cleavage of Proteins in Mobile Phase for Fast Quantitative LC-MS Bottom-Up Protein Analysis─Application for Ricin Detection
Jazyk angličtina Země Spojené státy americké Médium print-electronic
Typ dokumentu časopisecké články, práce podpořená grantem
PubMed
37565982
PubMed Central
PMC10448442
DOI
10.1021/acs.analchem.3c01772
Knihovny.cz E-zdroje
- MeSH
- chromatografie kapalinová metody MeSH
- peptidy MeSH
- proteiny analýza MeSH
- proteomika metody MeSH
- ricin * MeSH
- tandemová hmotnostní spektrometrie metody MeSH
- Publikační typ
- časopisecké články MeSH
- práce podpořená grantem MeSH
- Názvy látek
- peptidy MeSH
- proteiny MeSH
- ricin * MeSH
Sample preparation involving the cleavage of proteins into peptides is the first critical step for successful bottom-up proteomics and protein analyses. Time- and labor-intensiveness are among the bottlenecks of the commonly used methods for protein sample preparation. Here, we report a fast online method for postinjection acid cleavage of proteins directly in the mobile phase typically used for LC-MS analyses in proteomics. The chemical cleavage is achieved in 0.1% formic acid within 35 s in a capillary heated to 195 °C installed upstream of the analytical column, enabling the generated peptides to be separated. The peptides generated by the optimized method covered the entire sequence except for one amino acid of trastuzumab used for the method development. The qualitative results are extraordinarily stable, even over a long period of time. Moreover, the method is also suitable for accurate and repeatable quantification. The procedure requires only one manual step, significantly decreasing sample transfer losses. To demonstrate its practical utility, we tested the method for the fast detection of ricin. Ricin can be unambiguously identified from an injection of 10 ng, and the results can be obtained within 7-8 min after receiving a suspicious sample. Because no sophisticated accessories and no additional reagents are needed, the method can be seamlessly transferred to any laboratory for high-throughput proteomic workflows.
Zobrazit více v PubMed
Vandermarliere E.; Mueller M.; Martens L. Getting intimate with trypsin, the leading protease in proteomics. Mass Spectrom. Rev. 2013, 32, 453–465. 10.1002/mas.21376. PubMed DOI
Capelo J. L.; Carreira R.; Diniz M.; Fernandes L.; Galesio M.; Lodeiro C.; Santos H. M.; Vale G. Overview on modern approaches to speed up protein identification workflows relying on enzymatic cleavage and mass spectrometry-based techniques. Anal. Chim. Acta 2009, 650, 151–159. 10.1016/j.aca.2009.07.034. PubMed DOI
Switzar L.; Giera M.; Niessen W. M. Protein digestion: an overview of the available techniques and recent developments. J. Proteome Res. 2013, 12, 1067–1077. 10.1021/pr301201x. PubMed DOI
Duong V.-A.; Lee H. Bottom-Up Proteomics: Advancements in Sample Preparation. Int. J. Mol. Sci. 2023, 24, 5350.10.3390/ijms24065350. PubMed DOI PMC
Peterson D. S.; Rohr T.; Svec F.; Frechet J. M. High-throughput peptide mass mapping using a microdevice containing trypsin immobilized on a porous polymer monolith coupled to MALDI TOF and ESI TOF mass spectrometers. J. Proteome Res. 2002, 1, 563–568. 10.1021/pr0255452. PubMed DOI
Nicoli R.; Gaud N.; Stella C.; Rudaz S.; Veuthey J. L. Trypsin immobilization on three monolithic disks for on-line protein digestion. J. Pharm. Biomed. Anal. 2008, 48, 398–407. 10.1016/j.jpba.2007.12.022. PubMed DOI
Cingöz A.; Hugon-Chapuis F.; Pichon V. Evaluation of various immobilized enzymatic microreactors coupled on-line with liquid chromatography and mass spectrometry detection for quantitative analysis of cytochrome c. J. Chromatogr. A 2008, 1209, 95–103. 10.1016/j.chroma.2008.08.120. PubMed DOI
Zhong X.; Chen H.; Zare R. N. Ultrafast enzymatic digestion of proteins by microdroplet mass spectrometry. Nat. Commun. 2020, 11, 104910.1038/s41467-020-14877-x. PubMed DOI PMC
Rivera-Albarran M. E.; Ray S. J.; Novel A. Combined Microstrip Resonator/Nanospray Ionization Source for Microwave-Assisted Trypsin Digestion of Proteins. J. Am. Soc. Mass Spectrom. 2020, 31, 1684–1696. 10.1021/jasms.0c00115. PubMed DOI
Bekker-Jensen D. B.; Martinez-Val A.; Steigerwald S.; Ruther P.; Fort K. L.; Arrey T. N.; Harder A.; Makarov A.; Olsen J. V. A Compact Quadrupole-Orbitrap Mass Spectrometer with FAIMS Interface Improves Proteome Coverage in Short LC Gradients. Mol. Cell. Proteomics 2020, 19, 716–729. 10.1074/mcp.TIR119.001906. PubMed DOI PMC
Messner C. B.; Demichev V.; Bloomfield N.; Yu J. S. L.; White M.; Kreidl M.; Egger A. S.; Freiwald A.; Ivosev G.; Wasim F.; Zelezniak A.; Jurgens L.; Suttorp N.; Sander L. E.; Kurth F.; Lilley K. S.; Mulleder M.; Tate S.; Ralser M. Ultra-fast proteomics with Scanning SWATH. Nat. Biotechnol. 2021, 39, 846–854. 10.1038/s41587-021-00860-4. PubMed DOI PMC
Demichev V.; Szyrwiel L.; Yu F.; Teo G. C.; Rosenberger G.; Niewienda A.; Ludwig D.; Decker J.; Kaspar-Schoenefeld S.; Lilley K. S.; Mulleder M.; Nesvizhskii A. I.; Ralser M. dia-PASEF data analysis using FragPipe and DIA-NN for deep proteomics of low sample amounts. Nat. Commun. 2022, 13, 394410.1038/s41467-022-31492-0. PubMed DOI PMC
Wang Z.; Mulleder M.; Batruch I.; Chelur A.; Textoris-Taube K.; Schwecke T.; Hartl J.; Causon J.; Castro-Perez J.; Demichev V.; Tate S.; Ralser M. High-throughput proteomics of nanogram-scale samples with Zeno SWATH MS. eLife 2022, 11, e8394710.7554/eLife.83947. PubMed DOI PMC
Ishikawa M.; Konno R.; Nakajima D.; Gotoh M.; Fukasawa K.; Sato H.; Nakamura R.; Ohara O.; Kawashima Y. Optimization of Ultrafast Proteomics Using an LC-Quadrupole-Orbitrap Mass Spectrometer with Data-Independent Acquisition. J. Proteome Res. 2022, 21, 2085–2093. 10.1021/acs.jproteome.2c00121. PubMed DOI PMC
Lenčo J.; Semlej T.; Khalikova M. A.; Fabrik I.; Svec F. Sense and Nonsense of Elevated Column Temperature in Proteomic Bottom-up LC-MS Analyses. J. Proteome Res. 2021, 20, 420–432. 10.1021/acs.jproteome.0c00479. PubMed DOI
Lenčo J.; Vajrychova M.; Pimkova K.; Proksova M.; Benkova M.; Klimentova J.; Tambor V.; Soukup O. Conventional-Flow Liquid Chromatography-Mass Spectrometry for Exploratory Bottom-Up Proteomic Analyses. Anal. Chem. 2018, 90, 5381–5389. 10.1021/acs.analchem.8b00525. PubMed DOI
Partridge S. M.; Davis H. F. Preferential release of aspartic acid during the hydrolysis of proteins. Nature 1950, 165, 62.10.1038/165062a0. PubMed DOI
Schultz J.; Allison H.; Grice M. Specificity of the cleavage of proteins by dilute acid. I. Release of aspartic acid from insulin, ribonuclease, and glucagon. Biochemistry 1962, 1, 694–698. 10.1021/bi00910a024. PubMed DOI
Tsung C. M.; Fraenkel-Conrat H. Preferential Release of Aspartic Acid by Dilute Acid Treatment of Tryptic Peptides. Biochemistry 1965, 4, 793–801. 10.1021/bi00881a001. PubMed DOI
Schultz J.[28] Cleavage at aspartic acid. In Methods Enzymology; Academic Press, 1967; Vol. 11, pp 255–263.
Inglis A. S.Cleavage at aspartic acid. In Methods Enzymology; Hirs C.H.W.; Timashef S. N., Eds.; Academic Press, 1983; Vol. 91, pp 324–332. PubMed
Li A.; Sowder R. C.; Henderson L. E.; Moore S. P.; Garfinkel D. J.; Fisher R. J. Chemical cleavage at aspartyl residues for protein identification. Anal. Chem. 2001, 73, 5395–5402. 10.1021/ac010619z. PubMed DOI
Zhong H.; Marcus S. L.; Li L. Microwave-assisted acid hydrolysis of proteins combined with liquid chromatography MALDI MS/MS for protein identification. J. Am. Soc. Mass Spectrom. 2005, 16, 471–481. 10.1016/j.jasms.2004.12.017. PubMed DOI
Hua L.; Low T. Y.; Sze S. K. Microwave-assisted specific chemical digestion for rapid protein identification. Proteomics 2006, 6, 586–591. 10.1002/pmic.200500304. PubMed DOI
Swatkoski S.; Russell S. C.; Edwards N.; Fenselau C. Rapid chemical digestion of small acid-soluble spore proteins for analysis of Bacillus spores. Anal. Chem. 2006, 78, 181–188. 10.1021/ac051521d. PubMed DOI
Swatkoski S.; Gutierrez P.; Ginter J.; Petrov A.; Dinman J. D.; Edwards N.; Fenselau C. Integration of residue-specific acid cleavage into proteomic workflows. J. Proteome Res. 2007, 6, 4525–4527. 10.1021/pr0704682. PubMed DOI
Swatkoski S.; Gutierrez P.; Wynne C.; Petrov A.; Dinman J. D.; Edwards N.; Fenselau C. Evaluation of microwave-accelerated residue-specific acid cleavage for proteomic applications. J. Proteome Res. 2008, 7, 579–586. 10.1021/pr070502c. PubMed DOI
Hauser N. J.; Basile F. Online Microwave D-Cleavage LC-ESI-MS/MS of Intact Proteins: Site-Specific Cleavages at Aspartic Acid Residues and Disulfide Bonds. J. Proteome Res. 2008, 7, 1012–1026. 10.1021/pr700596e. PubMed DOI
Chen L. C.; Rahman M. M.; Hiraoka K. High pressure super-heated electrospray ionization mass spectrometry for sub-critical aqueous solution. J. Am. Soc. Mass Spectrom. 2014, 25, 1862–1869. 10.1007/s13361-014-0974-0. PubMed DOI
Chen L. C.; Kinoshita M.; Noda M.; Ninomiya S.; Hiraoka K. Rapid Online Non-Enzymatic Protein Digestion Analysis with High Pressure Superheated ESI-MS. J. Am. Soc. Mass Spectrom. 2015, 26, 1085–1091. 10.1007/s13361-015-1111-4. PubMed DOI
Wang Y.; Zhang W.; Ouyang Z. Fast protein analysis enabled by high-temperature hydrolysis. Chem. Sci. 2020, 11, 10506–10516. 10.1039/D0SC03237A. PubMed DOI PMC
Powell T.; Bowra S.; Cooper H. J. Subcritical Water Processing of Proteins: An Alternative to Enzymatic Digestion?. Anal. Chem. 2016, 88, 6425–6432. 10.1021/acs.analchem.6b01013. PubMed DOI
Lord J. M.; Roberts L. M.; Robertus J. D. Ricin: structure, mode of action, and some current applications. FASEB J. 1994, 8, 201–208. 10.1096/fasebj.8.2.8119491. PubMed DOI
Audi J.; Belson M.; Patel M.; Schier J.; Osterloh J. Ricin poisoning: a comprehensive review. JAMA 2005, 294, 2342–2351. 10.1001/jama.294.18.2342. PubMed DOI
Worbs S.; Kohler K.; Pauly D.; Avondet M. A.; Schaer M.; Dorner M. B.; Dorner B. G. Ricinus communis intoxications in human and veterinary medicine-a summary of real cases. Toxins 2011, 3, 1332–1372. 10.3390/toxins3101332. PubMed DOI PMC
Janik E.; Ceremuga M.; Saluk-Bijak J.; Bijak M. Biological Toxins as the Potential Tools for Bioterrorism. Int. J. Mol. Sci. 2019, 20, 118110.3390/ijms20051181. PubMed DOI PMC
Ler S. G.; Lee F. K.; Gopalakrishnakone P. Trends in detection of warfare agents. Detection methods for ricin, staphylococcal enterotoxin B and T-2 toxin. J. Chromatogr. A 2006, 1133, 1–12. 10.1016/j.chroma.2006.08.078. PubMed DOI
Griffiths G. D. Understanding ricin from a defensive viewpoint. Toxins 2011, 3, 1373–1392. 10.3390/toxins3111373. PubMed DOI PMC
Tevell Åberg A.; Bjornstad K.; Hedeland M. Mass spectrometric detection of protein-based toxins. Biosecur. Bioterror. 2013, 11, S215–26. 10.1089/bsp.2012.0072. PubMed DOI
Bozza W. P.; Tolleson W. H.; Rivera Rosado L. A.; Zhang B. Ricin detection: tracking active toxin. Biotechnol Adv 2015, 33, 117–123. 10.1016/j.biotechadv.2014.11.012. PubMed DOI
Duracova M.; Klimentova J.; Fucikova A.; Dresler J. Proteomic Methods of Detection and Quantification of Protein Toxins. Toxins 2018, 10, 9910.3390/toxins10030099. PubMed DOI PMC
Liu C. C.; Liang L. H.; Yang Y.; Yu H. L.; Yan L.; Li X. S.; Chen B.; Liu S. L.; Xi H. L. Direct Acetonitrile-Assisted Trypsin Digestion Method Combined with LC-MS/MS-Targeted Peptide Analysis for Unambiguous Identification of Intact Ricin. J. Proteome Res. 2021, 20, 369–380. 10.1021/acs.jproteome.0c00458. PubMed DOI
Oyler J. M.; Tran B. Q.; Kilgour D. P. A. Rapid Denaturing Organic Digestion Method for Targeted Protein Identification and Characterization. Anal. Chem. 2021, 93, 5046–5053. 10.1021/acs.analchem.0c04143. PubMed DOI
Soler-Rodriguez A. M.; Uhr J. W.; Richardson J.; Vitetta E. S. The toxicity of chemically deglycosylated ricin A-chain in mice. Int. J. Immunopharmacol. 1992, 14, 281–291. 10.1016/0192-0561(92)90041-I. PubMed DOI
Antoine M. C. Nouvelle relation entre les tensions et les temperatures. C. r. held Seanc. Acad. Sci. Paris 1888, 107, 681–684.
Onken U.; Rarey-Nies J.; Gmehling J. The Dortmund Data Bank: A computerized system for retrieval, correlation, and prediction of thermodynamic properties of mixtures. Int. J. Thermophys. 1989, 10, 739–747. 10.1007/BF00507993. DOI
Carman P. C. Fluid flow through a granular bed. Trans. Inst. Chem. Eng. London 1937, 15, 150–156.
Lenčo J.; Jadeja S.; Naplekov D. K.; Krokhin O. V.; Khalikova M. A.; Chocholous P.; Urban J.; Broeckhoven K.; Novakova L.; Svec F. Reversed-Phase Liquid Chromatography of Peptides for Bottom-Up Proteomics: A Tutorial. J. Proteome Res. 2022, 21, 2846–2892. 10.1021/acs.jproteome.2c00407. PubMed DOI
Bern M.; Kil Y. J.; Becker C., Byonic: advanced peptide and protein identification software. Curr. Protoc. Bioinf. 20124010.1002/0471250953.bi1320s40,Chapter 13, Unit13 20. PubMed DOI PMC
MacLean B.; Tomazela D. M.; Shulman N.; Chambers M.; Finney G. L.; Frewen B.; Kern R.; Tabb D. L.; Liebler D. C.; MacCoss M. J. Skyline: an open source document editor for creating and analyzing targeted proteomics experiments. Bioinformatics 2010, 26, 966–968. 10.1093/bioinformatics/btq054. PubMed DOI PMC
Dorfer V.; Pichler P.; Stranzl T.; Stadlmann J.; Taus T.; Winkler S.; Mechtler K. MS Amanda, a Universal Identification Algorithm Optimized for High Accuracy Tandem Mass Spectra. J. Proteome Res. 2014, 13, 3679–3684. 10.1021/pr500202e. PubMed DOI PMC
Hulsen T.; de Vlieg J.; Alkema W. BioVenn – a web application for the comparison and visualization of biological lists using area-proportional Venn diagrams. BMC Genom. 2008, 9, 488.10.1186/1471-2164-9-488. PubMed DOI PMC
Bobály B.; Lauber M.; Beck A.; Guillarme D.; Fekete S. Utility of a high coverage phenyl-bonding and wide-pore superficially porous particle for the analysis of monoclonal antibodies and related products. J. Chromatogr. A 2018, 1549, 63–76. 10.1016/j.chroma.2018.03.043. PubMed DOI
Khalikova M. A.; Skarbalius L.; Naplekov D. K.; Jadeja S.; Svec F.; Lenco J. Evaluation of strategies for overcoming trifluoroacetic acid ionization suppression resulted in single-column intact level, middle-up, and bottom-up reversed-phase LC-MS analyses of antibody biopharmaceuticals. Talanta 2021, 233, 12251210.1016/j.talanta.2021.122512. PubMed DOI
Kim M. H.; Kim C. S.; Lee H. W.; Kim K. Temperature dependence of dissociation constants for formic acid and 2,6-dinitrophenol in aqueous solutions up to 175 °C. J. Chem. Soc., Faraday Trans. 1996, 92, 4951–4956. 10.1039/FT9969204951. DOI
Capasso S.; Kirby A. J.; Salvadori S.; Sica F.; Zagari A. Kinetics and mechanism of the reversible isomerization of aspartic acid residues in tetrapeptides. J. Chem. Soc., Perkin Trans. 2 1995, 437–442. 10.1039/p29950000437. DOI
Hauser N. J.; Han H.; McLuckey S. A.; Basile F. Electron transfer dissociation of peptides generated by microwave D-cleavage digestion of proteins. J. Proteome Res. 2008, 7, 1867–1872. 10.1021/pr700671z. PubMed DOI PMC
Lenčo J.; Khalikova M. A.; Svec F. Dissolving Peptides in 0.1% Formic Acid Brings Risk of Artificial Formylation. J. Proteome Res. 2020, 19, 993–999. 10.1021/acs.jproteome.9b00823. PubMed DOI
Oses-Prieto J. A.; Zhang X.; Burlingame A. L. Formation of epsilon-formyllysine on silver-stained proteins: implications for assignment of isobaric dimethylation sites by tandem mass spectrometry. Mol. Cell Proteom. 2007, 6, 181–192. 10.1074/mcp.M600279-MCP200. PubMed DOI
Goodlett D. R.; Armstrong F. B.; Creech R. J.; van Breemen R. B. Formylated peptides from cyanogen bromide digests identified by fast atom bombardment mass spectrometry. Anal. Biochem. 1990, 186, 116–120. 10.1016/0003-2697(90)90583-U. PubMed DOI
Doucette A. A.; Vieira D. B.; Orton D. J.; Wall M. J. Resolubilization of precipitated intact membrane proteins with cold formic acid for analysis by mass spectrometry. J. Proteome Res. 2014, 13, 6001–6012. 10.1021/pr500864a. PubMed DOI
Hara T.; Huang Y.; Ito A.; Kawakami T.; Hojo H.; Murata M. Trifluoroethanol-containing RP-HPLC mobile phases for the separation of transmembrane peptides human glycophorin-A, integrin alpha-1, and p24: analysis and prevention of potential side reactions due to formic acid. J. Pept. Sci. 2015, 21, 61–70. 10.1002/psc.2717. PubMed DOI
Li J.; Shefcheck K.; Callahan J.; Fenselau C. Extension of microwave-accelerated residue-specific acid cleavage to proteins with carbohydrate side chains and disulfide linkages. Int. J. Mass Spectrom. 2008, 278, 109–113. 10.1016/j.ijms.2008.04.030. PubMed DOI PMC
Basile F.; Zhang S.; Kandar S. K.; Lu L. Mass Spectrometry Characterization of the Thermal Decomposition/Digestion (TDD) at Cysteine in Peptides and Proteins in the Condensed Phase. J. Am. Soc. Mass Spectrom. 2011, 22, 1926–1940. 10.1007/s13361-011-0222-9. PubMed DOI PMC
Han J. C.; Han G. Y. A procedure for quantitative determination of tris(2-carboxyethyl)phosphine, an odorless reducing agent more stable and effective than dithiothreitol. Anal. Biochem. 1994, 220, 5–10. 10.1006/abio.1994.1290. PubMed DOI
Suttapitugsakul S.; Xiao H.; Smeekens J.; Wu R. Evaluation and optimization of reduction and alkylation methods to maximize peptide identification with MS-based proteomics. Mol. BioSyst. 2017, 13, 2574–2582. 10.1039/C7MB00393E. PubMed DOI PMC
Müller T.; Winter D. Systematic Evaluation of Protein Reduction and Alkylation Reveals Massive Unspecific Side Effects by Iodine-containing Reagents. Mol. Cell. Proteomics 2017, 16, 1173–1187. 10.1074/mcp.M116.064048. PubMed DOI PMC
Wiśniewski J. R.; Zettl K.; Pilch M.; Rysiewicz B.; Sadok I. ’Shotgun’ proteomic analyses without alkylation of cysteine. Anal. Chim. Acta 2020, 1100, 131–137. 10.1016/j.aca.2019.12.007. PubMed DOI
Harris J. I.; Cole R. D.; Pon N. G. The kinetics of acid hydrolysis of dipeptides. Biochem. J. 1956, 62, 154–159. 10.1042/bj0620154. PubMed DOI PMC