Mass Spectrometry-Based Proteomics (Methods in Molecular Biology, 2718) 1071634569, 9781071634561

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Methods in Molecular Biology 2718

Kris Gevaert Editor

Mass SpectrometryBased Proteomics

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Mass Spectrometry-Based Proteomics Edited by

Kris Gevaert VIB-UGent Center for Medical Biotechnology, Ghent, Belgium

Editor Kris Gevaert VIB-UGent Center for Medical Biotechnology Ghent, Belgium

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3456-1 ISBN 978-1-0716-3457-8 (eBook) https://doi.org/10.1007/978-1-0716-3457-8 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Selected state-of-the-art protocols for contemporary mass spectrometry-based proteomics are described in this book and can be used by both novices in the field of proteomics as well as specialists. One protocol describes how the overall proteome coverage is increased by using different proteases. Those interested in the cellular surfaceome will find here a protocol in which an alternative method for surface biotinylation is described. Proteins are organized in larger complexes, which can be studied by different means, including proximity-induced biotinylation, abduction of complexes in viral-like particles, and thermal proteome profiling, protocols for which are provided in this book. Protein functions and localization are very often determined by modifications, and the reader will find updated protocols for identifying protein N-terminal acetylation, protein processing by proteases, protein N-glycosylation, and protein phosphorylation in this book. Over recent years, researchers are increasingly focusing on clinical proteomics, and we have provided protocols for automated preparation of clinical samples, the analysis of formalin-fixed paraffin-embedded samples, and protocols for the isolation of extracellular vesicles and for the monitoring of selected protein modifications in clinical samples. Finally, we touch upon structural proteomics by providing a state-of-the-art protocol for hydrogen/deuterium exchange mass spectrometry. Kris Gevaert

Ghent, Belgium

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 Increasing the Overall Proteome Coverage by Combining Protein Digestion by Tryp-N and Trypsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jade I. Hawksworth, Lode Denolf, Evy Timmerman, and Kris Gevaert 2 Cell Surface Biotinylation Using Furan Cross-Linking Chemistry . . . . . . . . . . . . . Esperanza Ferna´ndez, Laia Miret-Casals, Annemieke Madder, and Kris Gevaert 3 Studying Cellular Dynamics Using Proximity-Dependent Biotinylation: Somatic Cell Reprogramming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reuben Samson, Francesco Zangari, and Anne-Claude Gingras 4 Virotrap: Trapping Protein Complexes in Virus-Like Particles . . . . . . . . . . . . . . . . George D. Moschonas, Margaux De Meyer, Delphine De Sutter, Evy Timmerman, Petra Van Damme, and Sven Eyckerman 5 Thermal Proteome Profiling for Drug Target Identification and Probing of Protein States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patricia Sauer and Marcus Bantscheff 6 Improved Coverage of the N-Terminome by Combining ChaFRADIC with Alternative Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xuehui Jiang, Ying Lao, Victor Spicer, and Rene´ P. Zahedi 7 Sensitive and High-Throughput Exploration of Protein N-Termini by TMT-TAILS N-Terminomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Konstantinos Kalogeropoulos, Louise Bundgaard, and Ulrich auf dem Keller 8 The Global Acetylation Profiling Pipeline for Quick Assessment of Protein N-Acetyltransferase Specificity In Cellulo . . . . . . . . . . . . . . . . . . . . . . . . . Thierry Meinnel, Jean-Baptiste Boyer, and Carmela Giglione 9 Systems-Wide Site-Specific Analysis of Glycoproteins . . . . . . . . . . . . . . . . . . . . . . . . Kathirvel Alagesan and Emmanuelle Charpentier 10 Targeted Profiling of Protein Phosphorylation in Plants . . . . . . . . . . . . . . . . . . . . . Xiangyu Xu, Kris Gevaert, Ive De Smet, and Lam Dai Vu 11 Automated Sample Preparation for Mass Spectrometry-Based Clinical Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ ller, Mauro A. Cremonini, Georg Kliewer, and Jeroen Krijgsveld Torsten Mu 12 Proteomics-Based Analysis and Diagnosis of Formalin-Fixed Paraffin-Embedded Amyloidosis Samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Delphi Van Haver, Ame´lie Dendooven, and Francis Impens

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Contents

A Robust and Clinically Applicable Sample Preparation Protocol for Urinary Extracellular Vesicle Isolation Suitable for Mass Spectrometry-Based Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Leyla A. Erozenci, Irene V. Bijnsdorp, Sander R. Piersma, and Connie R. Jimenez Density-Based Fractionation of Cell-Conditioned Medium to Prepare Proteomics Grade Extracellular Vesicles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Quentin Roux, Sarah Deville, and An Hendrix Mass Spectrometry-Based Analysis of Histone Posttranslational Modifications from Laser Microdissected Samples . . . . . . . . . . . . . . . . . . . . . . . . . . Roberta Noberini and Tiziana Bonaldi Phosphoproteomics After Guanidinium Thiocyanate Extraction of Tissue Biopsies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Frank Rolfs, Richard R. de Goeij-de Haas, Jaco C. Knol, Sander R. Piersma, and Connie R. Jimenez Probing Antibody Structures by Hydrogen/Deuterium Exchange Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zuzana Kalaninova´, Luka´ˇs Fojtı´k, Josef Chmelı´k, Petr Nova´k, Michael Volny´, and Petr Man

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors KATHIRVEL ALAGESAN • Max Planck Unit for the Science of Pathogens, Berlin, Germany ULRICH AUF DEM KELLER • Department of Biotechnology and Biomedicine, Technical University of Denmark, Lyngby, Denmark MARCUS BANTSCHEFF • Cellzome GmbH, GlaxoSmithKline (GSK), Heidelberg, Germany IRENE V. BIJNSDORP • Department of Urology, Amsterdam UMC, Location VUMC, Amsterdam, The Netherlands TIZIANA BONALDI • Department of Experimental Oncology, IEO, European Institute of Oncology IRCCS, Milan, Italy; Department of Oncology and Haematology-Oncology, University of Milano, Milan, Italy JEAN-BAPTISTE BOYER • Universite´ Paris Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), Gif-sur-Yvette, France LOUISE BUNDGAARD • Department of Biotechnology and Biomedicine, Technical University of Denmark, Lyngby, Denmark EMMANUELLE CHARPENTIER • Max Planck Unit for the Science of Pathogens, Berlin, Germany; Institute for Biology, Humboldt University, Berlin, Germany JOSEF CHMELI´K • Institute of Microbiology of the Czech Academy of Sciences, Prague, Czech Republic MAURO A. CREMONINI • Agilent Technologies Italy, Milan, Italy RICHARD R. DE GOEIJ-DE HAAS • Department Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUmc, Amsterdam, The Netherlands MARGAUX DE MEYER • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium; iRIP Unit, Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium IVE DE SMET • Ghent University, Department of Plant Biotechnology and Bioinformatics, Ghent, Belgium; VIB Center for Plant Systems Biology, Ghent, Belgium DELPHINE DE SUTTER • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium AME´LIE DENDOOVEN • Department of Pathology, Ghent University Hospital, Ghent, Belgium; Laboratory for Experimental Medicine and Pediatrics, Antwerp University, Edegem, Belgium LODE DENOLF • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium SARAH DEVILLE • Laboratory of Experimental Cancer Research, Department of Human Structure and Repair, Ghent University, Ghent, Belgium; Cancer Research Institute Ghent, Ghent, Belgium LEYLA A. EROZENCI • Department of Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUMC, Amsterdam, The Netherlands; Department of Urology, Amsterdam UMC, Location VUMC, Amsterdam, The Netherlands SVEN EYCKERMAN • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium

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ESPERANZA FERNA´NDEZ • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium LUKA´Sˇ FOJTI´K • BioCeV -Institute of Microbiology of the Czech Academy of Sciences, Vestec, Czech Republic; Department of Biochemistry, Faculty of Science, Charles University, Prague, Czech Republic KRIS GEVAERT • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium CARMELA GIGLIONE • Universite´ Paris Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), Gif-sur-Yvette, France ANNE-CLAUDE GINGRAS • Lunenfeld-Tanenbaum Research Institute, Sinai Health, Toronto, ON, Canada; Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada JADE I. HAWKSWORTH • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium AN HENDRIX • Laboratory of Experimental Cancer Research, Department of Human Structure and Repair, Ghent University, Ghent, Belgium; Cancer Research Institute Ghent, Ghent, Belgium FRANCIS IMPENS • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium; VIB Proteomics Core, Ghent, Belgium XUEHUI JIANG • Manitoba Centre for Proteomics and Systems Biology, Winnipeg, MB, Canada CONNIE R. JIMENEZ • Department of Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUMC, Amsterdam, The Netherlands ZUZANA KALANINOVA´ • BioCeV -Institute of Microbiology of the Czech Academy of Sciences, Vestec, Czech Republic; Department of Biochemistry, Faculty of Science, Charles University, Prague, Czech Republic KONSTANTINOS KALOGEROPOULOS • Department of Biotechnology and Biomedicine, Technical University of Denmark, Lyngby, Denmark GEORG KLIEWER • German Cancer Research Center (DKFZ), Heidelberg, Germany; Heidelberg University, Medical Faculty, Heidelberg, Germany JACO C. KNOL • Department Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUmc, Amsterdam, The Netherlands JEROEN KRIJGSVELD • German Cancer Research Center (DKFZ), Heidelberg, Germany; Heidelberg University, Medical Faculty, Heidelberg, Germany YING LAO • Manitoba Centre for Proteomics and Systems Biology, Winnipeg, MB, Canada ANNEMIEKE MADDER • Organic and Biomimetic Chemistry Research Group, Department of Organic and Macromolecular Chemistry, Ghent University, Ghent, Belgium PETR MAN • BioCeV -Institute of Microbiology of the Czech Academy of Sciences, Vestec, Czech Republic THIERRY MEINNEL • Universite´ Paris Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), Gif-sur-Yvette, France LAIA MIRET-CASALS • Organic and Biomimetic Chemistry Research Group, Department of Organic and Macromolecular Chemistry, Ghent University, Ghent, Belgium GEORGE D. MOSCHONAS • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium TORSTEN MU¨LLER • German Cancer Research Center (DKFZ), Heidelberg, Germany; Heidelberg University, Medical Faculty, Heidelberg, Germany

Contributors

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ROBERTA NOBERINI • Department of Experimental Oncology, IEO, European Institute of Oncology IRCCS, Milan, Italy PETR NOVA´K • BioCeV -Institute of Microbiology of the Czech Academy of Sciences, Vestec, Czech Republic SANDER R. PIERSMA • Department of Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUMC, Amsterdam, The Netherlands FRANK ROLFS • Department Medical Oncology, OncoProteomics Laboratory, Cancer Center Amsterdam, Amsterdam UMC, Location VUmc, Amsterdam, The Netherlands QUENTIN ROUX • Laboratory of Experimental Cancer Research, Department of Human Structure and Repair, Ghent University, Ghent, Belgium; Cancer Research Institute Ghent, Ghent, Belgium REUBEN SAMSON • Lunenfeld-Tanenbaum Research Institute, Sinai Health, Toronto, ON, Canada; Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada PATRICIA SAUER • Cellzome GmbH, GlaxoSmithKline (GSK), Heidelberg, Germany VICTOR SPICER • Manitoba Centre for Proteomics and Systems Biology, Winnipeg, MB, Canada EVY TIMMERMAN • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium; VIB Proteomics Core, Ghent, Belgium PETRA VAN DAMME • Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium; iRIP Unit, Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium DELPHI VAN HAVER • VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium; VIB Proteomics Core, Ghent, Belgium MICHAEL VOLNY´ • BioCeV -Institute of Microbiology of the Czech Academy of Sciences, Vestec, Czech Republic LAM DAI VU • Ghent University, Department of Plant Biotechnology and Bioinformatics, Ghent, Belgium; VIB Center for Plant Systems Biology, Ghent, Belgium; VIB-UGent Center for Medical Biotechnology, Ghent, Belgium; Department of Biomolecular Medicine, Ghent University, Ghent, Belgium XIANGYU XU • Ghent University, Department of Plant Biotechnology and Bioinformatics, Ghent, Belgium; VIB Center for Plant Systems Biology, Ghent, Belgium RENE´ P. ZAHEDI • Manitoba Centre for Proteomics and Systems Biology, Winnipeg, MB, Canada; Department of Internal Medicine, University of Manitoba, Winnipeg, MB, Canada; Department of Biochemistry and Medical Genetics, University of Manitoba, Winnipeg, MB, Canada; CancerCare Manitoba Research Institute, Winnipeg, MB, Canada FRANCESCO ZANGARI • Lunenfeld-Tanenbaum Research Institute, Sinai Health, Toronto, ON, Canada; Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada

Chapter 1 Increasing the Overall Proteome Coverage by Combining Protein Digestion by Tryp-N and Trypsin Jade I. Hawksworth, Lode Denolf, Evy Timmerman, and Kris Gevaert Abstract Mass spectrometry-based proteomics combining more than one protease in parallel facilitates the identification of more peptides and proteins than when a single protease is used. Trypsin cleaves proteins C-terminally to arginine and lysine, while its mirroring protease Tryp-N cleaves N-terminally to the same amino acids. Here, we combine trypsin and Tryp-N with the commercially available S-Trap columns, which purify protein samples and catalyze digestion. Comparison of trypsin or Tryp-N coupled with S-Trap columns demonstrates plasma and cell lysate proteins unique to one protease. We thus suggest the use of both proteases in a complementary manner to obtain deeper proteome coverage. Key words Proteases, Trypsin, Tryp-N, S-Trap, Mass spectrometry

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Introduction Bottom-up proteomics consists of proteome digestion using a (specific) protease, followed by liquid chromatography–tandem mass spectrometry (LC-MS/MS) analysis of the resulting peptide mixture, and identification of proteins through reconstitution of peptide sequences using bioinformatics [1]. By far, the most commonly used protease in bottom-up proteomics is trypsin, which cleaves C-terminally to arginine and lysine (Fig. 1). However, there is evidence that the use of multiple proteases with different specificities facilitates the identification of more peptides and more proteins, thus leading to a more complete proteome coverage [2]. The recently described protease Tryp-N, which cleaves N-terminally and thus not C-terminally to arginine and lysine (Fig. 1), was reported to be complimentary to trypsin [3, 4]. Comparison of digestions by trypsin or Tryp-N determined that more proteins could be identified using a combination of both proteases than with trypsin alone [4]. Tryp-N could particularly be useful for the study of N-terminally acetylated peptides, as non-N-terminal

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Cleavage sites for trypsin and Tryp-N. This exemplar protein Costars Family Protein ABRACL was identified in MCF7 cell lysate. Potential trypsin cleavage sites are denoted by a blue arrow, while potential Tryp-N cleavage sites are denoted with an orange arrow. In this exemplar protein, the Tryp-N N-terminal peptide (orange) was found, but the tryptic N-terminal peptide was not. R = arginine, K = lysine

peptides generated upon Tryp-N digestion are more basic due to the presence of both a primary alpha-amino group and a N-terminal basic amino acid, which are thereby efficiently separated from N-terminally acetylated peptides via a simple strong cation exchange (SCX) step at acidic pH [5]. N-terminal protein modifications such as N-terminal acetylation are of high biological relevance as these direct protein location, activity, and degradation. Not surprisingly, methods for N-terminal peptide enrichment have been published [6], but further protocols could still contribute to the field. Tryp-N has also been validated for use on the Alzheimer’s disease-related protein tau, and over 80% of peptides found were unique to either trypsin or Tryp-N [3]. Here, Tryp-N was combined with the S-Trap protocol [7, 8], a commercially available column that can be used on many sample types. While Tryp-N-digested peptides tend to contain more missed cleavage sites than tryptic peptides [3], the S-Trap facilitates more effective protease activity and fewer missed cleavages than digestion in solution [7], thus making it ideal to be used in conjunction with Tryp-N. In brief, in this method proteins are reduced and alkylated and then captured on the S-Trap column (Fig. 2). Using a simple on-column protocol, proteins are washed, digested, peptides eluted, and dried prior to analysis by LC-MS/MS. The S-Trap washing steps remove the need for further sample purification, and the S-Trap protocol is both quick and simple. MS analysis of Tryp-N-digested peptides identifies about 70% as many proteins as trypsin when digestion takes place in solution [4]; however, we found a similar number of proteins using trypsin and Tryp-N when digestion took place in conjunction with the S-Trap (Fig. 3). Both proteases identified unique proteins and peptides, while N-terminally acetylated peptides were only found with Tryp-N, supporting a role for Tryp-N to be used in parallel to trypsin.

Deeper Proteome Coverage by Combining Tryp-N and Trypsin

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Fig. 2 Experimental design. After reduction and alkylation in lysis buffer, full length proteins and proteases are captured in the pores within the S-Trap. This affinity is lost after digestion with trypsin or Tryp-N and the resulting peptides are eluted off the column in three stages

Fig. 3 Tryp-N and trypsin digestion of an MCF-7 cell lysate and human plasma identify shared and unique proteins and peptides. For a comparison of the proteins identified with trypsin (blue) or Tryp-N (yellow), proteins which were identified in at least two of three technical replicates are visualized in a Venn diagram. On a peptide level, both total peptides and unique N-terminally acetylated (Nt-ace) peptides with a start position of 1 or 2 were counted and visualized

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Materials All organic solvents and reagents are liquid chromatography grade and analytical grade, respectively. All water must be ultrapure, with a maximum resistance of 18.2 mΩ/cm at 25 °C. Wherever percentages are stated for buffers, the remainder of the buffer should be

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made up with water. Where solutions of two densities are combined, the buffer should be made up to its final stated volume in a measuring cylinder or volumetric flask. 2.1 Sample Preparation

1. 100 mM sodium hydroxide (NaOH): mix 3.99 mg NaOH powder with 1 mL water in a plastic container (see Note 1). 2. Reduction buffer: 120 mM tris(2-carboxyethyl)phosphine (TCEP). Mix 34.4 mg of TCEP powder into 1 mL of water. Use 100 mM NaOH and a pH meter to adjust the pH of the solution from 2.5 to pH 8.0–8.5 (see Note 2). Aliquot remaining TCEP out into small volumes and store at -20 °C. 3. Alkylation buffer: 625 mM iodoacetamide (IAA). Mix 57.8 mg IAA into 0.5 mL of water (see Note 3).

2.2

Digestion Buffers

1. Trypsin digestion buffer: 0.25 μg/μL trypsin. Resuspend 20 μg trypsin gently on ice in 80 μL of 25 mM TEAB (see Note 4). 2. RapiGest solution: Dissolve 1 mg vial of RapiGest in 100 μL MQ water (see Note 5). 3. Tryp-N digestion buffer: 0.25 μg/μL Tryp-N, 0.1% RapiGest, 1 mM CaCl2, and 50 μM ZnCl2. Add 40 μL of 0.5 μg/μL Tryp-N and 8 μL 1% RapiGest to 32 μL MQ water (see Note 6).

2.3

S-Trap Protocol

1. Lysis buffer: 10% sodium dodecyl sulfate (SDS) and 100 mM triethylammonium bicarbonate (TEAB). Dissolve 10 μg SDS in 10 μL of 1 M TEAB and 80 μL of water. 2. Binding/wash buffer: 100 mM TEAB in 90% methanol. To make 8 mL of binding/wash buffer add 0.8 mL of 1 M TEAB into 7.2 mL of methanol (see Note 7). 3. Elution buffer 1: 50 mM TEAB. Combine 15 μL of 1 M TEAB with 285 μL of water. 4. Elution buffer 2: 0.2% formic acid. Mix 6 μL formic acid into 297 μL water. 5. Elution buffer 3: 50% acetonitrile (ACN). Mix ACN and water in equal volumes (see Note 8).

2.4 Mass Spectrometry

1. LC-MS/MS solvent A: 0.1% formic acid (FA) (see Note 9).

2.5

1. Consumables: S-Trap Micro (ProtiFi).

Equipment

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Methods

3.1 Sample Preparation and Loading

1. Calculate the volume of a liquid sample needed for 100 μg protein. Combine with 11.5 μL of lysis buffer. Make the sample up to 23 μL with MQ water. Alternatively, dissolve a cell pellet/ solid sample in 11.5 μL of lysis buffer and 11.5 μL of MQ water (see Note 10). 2. Add 1 μL of 120 mM TCEP. Incubate at 55 °C for 15 min. Cool sample down quickly on ice (see Note 11). The sample must return to room temperature (RT) before continuing with the protocol. 3. Add 1 μL of 625 mM IAA and incubate for 15 min at RT in the dark (see Note 3). 4. Add 2.5 μL of phosphoric acid to the sample and vortex to acidify the sample for a final pH of below 1 (see Note 12). 5. Put each S-Trap into a 2 mL Eppendorf tube for waste flowthrough. 6. Pipette 165 μL of binding/wash buffer onto the S-Trap column, pipetting to the top of the narrow stem (see Note 13). 7. Trap proteins in column by centrifugation at 4000 g for 30 s and recentrifuge again if not all sample enters the S-Trap. 8. Pipette 150 μL binding/wash buffer onto the S-Trap column, and centrifuge at 4000 g for 30 s. Repeat twice more, rotating 180° after each centrifugation step (see Note 14). 9. Centrifuge the S-Trap column at 4000 g for 1 min, which should remove all binding/wash buffer (see Note 15). 10. Move the S-Trap column to a clean 2 mL sample tube. 11. Add 20 μL of digestion buffer to each sample, containing 5 μg protease (see Note 16). 12. Place the cap on the S-Trap without fully tightening the screw top to limit evaporative buffer loss while preventing formation of a vacuum within the S-Trap (see Note 17). 13. Incubate overnight at 37 °C, without agitation (see Note 18).

3.2

Peptide Elution

1. Add 40 μL of elution buffer 1 to the top of the stem of the S-Trap. 2. Centrifuge at 4000 g for 1 min and collect eluate. 3. Add 40 μL of elution buffer 2 to the top of the stem of the S-Trap. 4. Centrifuge at 4000 g for 1 min and collect eluate. 5. Add 40 μL of elution buffer 3 to the top of the stem of the S-Trap.

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6. Centrifuge at 4000 g for 1 min and collect eluate. 7. Pool eluate from all three steps, dry down under vacuum, and resuspend in suitable MS solvent. 3.3 Mass Spectrometry

1. Determine the peptide concentration (see Note 19). 2. Load an appropriate amount of the peptide mixture onto the LC column. 3. For LC-MS/MS analysis, we coupled an Ultimate 3000 RSLC nanoLC (Thermo Fisher Scientific) in-line to an Orbitrap Elite (Thermo Fisher Scientific) mass spectrometer, equipped with a pneu-Nimbus dual ion source. However, many other MS systems optimized for proteomic analysis would be suitable.

3.4

Data Analysis

1. Analyze raw files (LC-MS/MS output files) using a suitable software. We use MaxQuant (Max Planck Institute of Biochemistry) and MASCOT (Matrix Science), with settings that are described below, but other software for MS data analysis can be used based on one’s data analysis requirements. 2. In MaxQuant, run two analyses, searching separately for trypsin and Tryp-N peptides (see Note 20). Missed cleavages should be set to 2, and label free quantification (LFQ) on. Set the number of processors to (ideally) the number of samples to allow simultaneous analysis of all samples. Select modifications, and set an appropriate database for peptide and protein identification (see Note 21). 3. MaxQuant output files are filtered in Perseus. After loading LFQ data into the main column, use the function “Filter rows based on categorical column” to remove contaminants and reverse sequences and proteins which were identified by site only. Transform data by log2 and complete statistical analysis depending on data characteristics (see Note 22). Both cell lysate and plasma demonstrate a pool of proteins shared between the proteases and proteins unique to each protease (Fig. 3). 4. N-terminally acetylated peptides can be identified from raw files using MASCOT. For trypsin, the protease is set to “Semi-Tryp,” so only one trypsin digestion site is needed per peptide, and set the parameters for the Tryp-N search similarly. Set missed cleavages to 2. Choose appropriate peptide modifications, MS search settings, and human database for peptide and protein identification (see Note 23). 5. MASCOT output data can be analyzed using any software with data processing and statistical capacity (see Note 24). Here, MASCOT analysis of the raw MS output data demonstrates 31 N-terminal acetylated peptides from the cell lysate proteome found only after Tryp-N digestion, and 3 N-terminal acetylated peptides found in plasma only with Tryp-N (Fig. 3).

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Notes 1. NaOH must be stored in plastic, at room temperature (RT) and in the dark. 2. A pH of 8.0–8.5 is required for alkylation, and even a small volume of unadjusted TCEP can affect sample pH. 3. IAA is sensitive to light so make up this solution fresh for each experiment, exposing the IAA to as little light as possible. To maintain IAA activity, we transport the fresh solution in an Eppendorf tube wrapped in tin foil. Pipette quickly, and place an inverted ice box/polystyrene box over the whole sample holder as soon as possible. 4. To ensure trypsin is fully suspended, pipette the digestion buffer slowly in and out of the trypsin vial. Freeze any leftover trypsin in aliquots at -20 °C for future use. Avoid freeze-thaw cycles wherever possible as protease activity decreases with each cycle. 5. RapiGest should be stored at 4 °C and used within a week. We have used RapiGest in accordance with the ProtiFi Tryp-N datasheet and as was demonstrated previously [4]. However, in another study, sodium deoxycholate reduced missed cleavages from 14% to less than 6% [5]. A third paper lists no detergent in the Tryp-N digestion buffer [3]. Optimize this step according to the generated data. 6. Tryp-N is a metalloprotease, and metal salts are thus required in the digestion buffer. If Tryp-N is sourced from ProtiFi, the only current commercial supplier, the metal salts CaCl2, and ZnCl2 are already present within the protease buffer at concentrations of 2 mM and 100 μM, respectively, and thus do not need to be added to the digestion buffer. 7. The densities of TEAB and methanol are different. Add roughly half of the TEAB, then the methanol, and make the final volume up to 8 mL with TEAB. 8. Put 100 μL ACN into a measuring cylinder or volumetric flask. Add 150 μL water and then make volume up to 300 μL with ACN. 9. Ultrafiltered water is recommended for MS solvents and samples which will be injected directly into the MS. Any impurities and volatile solvents from previous steps are removed by the S-Trap and drying down, respectively, but any contaminants at this stage can enter the MS instrument. 10. We worked with 100 μg of protein from both plasma and cell lysate. This is the maximum amount of protein recommended for the S-Trap Micro. Other sizes can be used when the mass of

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the sample falls outside of 1–100 μg. If a different S-Trap column is used, it is important to appropriately modify the volumes of each buffer when moving to an S-Trap column with a different capacity. When using the S-Trap Micro, it is important that all samples added to the column have a volume of 23 μL. Samples with a larger volume can be loaded in multiple centrifugation steps. 11. If SDS precipitates when the sample is cooled, warm to RT before continuing. 12. The pH must be below 1 for effective sample capture. Add more acid if the pH is above 1. We recommend optimizing this step with your sample type prior to starting an experiment. 13. Add all sample to the S-Trap column, including any insoluble sample. 14. Make a mark on the outside of the lip of the top of the S-Trap. Use this mark to rotate S-Traps by 180° between each centrifugation step. Wash should be optimized for different sample types, e.g., if sample is more viscous, further washing may be needed. The supernatant will be collected in the Eppendorf tube during each wash step, and this liquid needs to be discarded before it touches the bottom of the S-Trap. If the supernatant reaches the bottom of the S-Trap, this will prevent the sample from moving down the column. 15. Centrifuge the S-Trap column again if any liquid is visible above the filter. 16. Proteases and digestion buffers can be optimized, and ProtiFi has tested commonly used proteases for compatibility with the S-Trap. 17. Creating a vacuum prevents buffers from moving through S-Trap. The S-Trap Micro has no pressure vents, so it is important not to tightly lid the columns. Also, the S-Trap filter must not dry out. To prevent this, incubate the samples either in a water bath or in a stationary heat block with several unlidded Eppendorf tubes of water. 18. While the product sheet and one paper recommend shorter, higher temperature digestions for Tryp-N [4], we found 37 °C overnight to be optimal, providing the highest number of peptide and protein identifications. Digestion conditions of 37 °C overnight were also used in references [3, 5]. 19. Peptide concentrations can be measured with the Lunatic UV-Vis spectrophotometer (Unchained Labs) [9], our instrument of choice for determining peptide and protein concentration, however the Lunatic spectrophotometer was found be unreliable when preparing samples with the S-Trap protocol. The reason for this remains unknown, but we postulate it may

Deeper Proteome Coverage by Combining Tryp-N and Trypsin

9

be due to incompatibility of the S-Trap buffer with the Lunatic chip or instrumentation. Utilizing the LC-MS/MS and columns used within our lab, 1.5 μg peptide provides an optimal TIC of 1.0E08 to 9.9E09. 20. Do not set the number of processors to more than the number of cores available on the computer. When configuring protease searches, choose semi-trypsin or semi-Tryp-N as this allows to identify peptides with only one tryptic/Tryp-N cleavage site. In fact, some N- and C-terminal peptides would only have one tryptic/Tryp-N cleavage site, and not setting this setting to “semi” would remove these from the dataset. 21. In this case, the following protein modifications were set: oxidation (M), acetyl (protein N-term) and carbamidomethyl (C). Set the maximum number of modifications per peptide to 5. We used the Swiss-Prot human database 2022, UPhuman9609.fasta [10]. 22. After MaxQuant and Perseus, we compared the number and identity of proteins identified using each protease. 23. Modifications in MASCOT can be set as fixed or variable. In this analysis, carbamidomethylation (C) is a fixed modification, while N-terminal acetylation and oxidation (M) are variable modifications. Set appropriate MS settings according to the mass spectrometer used. In this case, this includes a maximum of two missed cleavages, a MS/MS tolerance of 20 mmu and a peptide tolerance of 10 ppm. An appropriate human database should be chosen for peptide and protein identification; here, this is the same Swiss-Prot human 2022 database as indicated above [10]. 24. We use RStudio to count modifications and visualize data from MASCOT. Any program with data visualization tools would be suitable.

Acknowledgments This work was supported by the Stichting Alzheimer Onderzoek (SAO-FRA, Belgium), awarded to Kris Gevaert. References 1. Aebersold R, Mann M (2016) Massspectrometric exploration of proteome structure and function. Nature 537(7620): 3 4 7 – 3 5 5 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature19949 2. Swaney DL, Wenger CD, Coon JJ (2010) Value of using multiple proteases for large-

scale mass spectrometry-based proteomics. J Proteome Res 9(3):1323–1329. https://doi. org/10.1021/pr900863u 3. Zhou M, Duong DM, Johnson ECB, Dai J, Lah JJ, Levey AI, Seyfried NT (2019) Mass spectrometry-based quantification of tau in human cerebrospinal fluid using a

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complementary tryptic peptide standard. J Proteome Res 18(6):2422–2432. https://doi. org/10.1021/acs.jproteome.8b00920 4. Wilson JP, Ipsaro JJ, Del Giudice SN, Turna NS, Gauss CM, Dusenbury KH, Marquart K, Rivera KD, Pappin DJ (2020) Tryp-N: a thermostable protease for the production of N-terminal argininyl and lysinyl peptides. J Proteome Res 19(4):1459–1469. https://doi. org/10.1021/acs.jproteome.9b00713 5. Chang CH, Chang HY, Rappsilber J, Ishihama Y (2020) Isolation of acetylated and unmodified protein N-terminal peptides by strong cation exchange chromatographic separation of TrypN-digested peptides. Mol Cell Proteomics 20:100003. https://doi.org/10.1074/mcp. TIR120.002148 6. Bogaert A, Gevaert K (2020) Protein aminotermini and how to identify them. Expert Rev Proteomics 17(7–8):581–594. https://doi. org/10.1080/14789450.2020.1821657 7. Ludwig KR, Schroll MM, Hummon AB (2018) Comparison of in-solution, FASP, and

S-Trap based digestion methods for bottom-up proteomic studies. J Proteome Res 17(7): 2480–2490. https://doi.org/10.1021/acs. jproteome.8b00235 8. HaileMariam M, Eguez RV, Singh H, Bekele S, Ameni G, Pieper R, Yu Y (2018) S-Trap, an ultrafast sample-preparation approach for shotgun proteomics. J Proteome Res 17(9): 2917–2924. https://doi.org/10.1021/acs. jproteome.8b00505 9. Maia TM, Staes A, Plasman K, Pauwels J, Boucher K, Argentini A, Martens L, Montoye T, Gevaert K, Impens F (2020) Simple peptide quantification approach for MS-based proteomics quality control. ACS Omega 5(12):6754–6762. https://doi.org/ 10.1021/acsomega.0c00080 10. Uniprot Consortium (2021) UniProt: the universal protein knowledgebase in 2021. Nucleic Acids Res 49(D1):D480–D489. https://doi. org/10.1093/nar/gkaa1100

Chapter 2 Cell Surface Biotinylation Using Furan Cross-Linking Chemistry Esperanza Ferna´ndez, Laia Miret-Casals, Annemieke Madder, and Kris Gevaert Abstract A detailed study of the cellular surfaceome poses major challenges for mass spectrometry analysis. Surface proteins are low abundant compared to intracellular proteins, and their inefficient extraction in aqueous medium leads to their aggregation and precipitation. To tackle such problems, surface biotinylation is frequently used to tag surface proteins with biotin, allowing for their enrichment, leading to a more sensitive mapping of surface proteomes. We here detail a new surface biotinylation protocol based on furan-biotin affinity purification to enrich plasma membrane proteins for proteomics. This protocol involves biotinylation of cell surface membrane proteins on viable cells, followed by affinity enrichment using streptavidin beads, trypsin digestion, peptide cleanup, and LC-MS/MS analysis. Key words Furan-biotin, Affinity purification, Plasma membrane proteins, LC-MS/MS, Proteomics, Surfaceome

1

Introduction Plasma membrane proteins that are exposed to the extracellular space, collectively called the surfaceome, are responsible for cell homeostasis and the communication of a cell with its environment. The surfaceome is thereby involved in a vast variety of dynamic cellular processes such as ion and molecular transport, signal reception, and transduction, giving cells their shape and intercellular communication. Glycosylation, a modification occurring on over 90% of the surface proteins, when aberrant, is associated with Alzheimer’s disease [1] and cancer [2, 3]. The extracellular matrix, which is in close contact with the surfaceome, is known to be associated with neurodegeneration, cancer metastasis, and other diseases [4–6], and the surfaceome also reflects intracellular changes consequent of disease or infection that may affect protein

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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abundance, expose alternative proteoforms, and present antigens [7–9]. Surface proteins are a major target for biomedical research due to their utility as cellular markers of pathophysiological changes and their accessibility for pharmaceutical delivery. Membrane proteins comprise about 30% of the mammalian proteome and account for more than 60% of drug targets [10]. However, the identification of surface proteins via mass spectrometry (MS) poses several challenges. Surface proteins have structural domains that span membranes and contain a considerable number of hydrophobic amino acids [11]. Such hydrophobic stretches may promote protein aggregation and precipitation, leading to incomplete protein extraction and underrepresentation of hydrophobic membrane proteins in total cellular lysates. Chaotropes and detergents are used to counter these challenges but are often noncompatible with downstream mass-spectrometric (MS) analysis. Intrinsic properties of membrane proteins may also hinder their identification by MS as long hydrophobic stretches that lack polar residues important for proper ionization, and globular parts and extracellular loops that are heavily modified (e.g., by glycosylations) sterically hinder the proteases’ access, yielding too few and/or too long peptides for identification upon LC-MS/MS analysis [12]. Due to their lower abundance compared with cytosolic proteins, enrichment of surface proteins either exploiting their physicochemical properties, applying affinity-based methods, or chemically labeling the extracellular regions is highly recommended to increase the overall sensitivity of detection. Biotin is one of the most widely used molecules for protein labeling prior to their downstream analysis [13]. It has strong affinity for avidin and streptavidin with a dissociation constant of about 10-15 M, and this interaction is highly resistant to chaotropic agents and high concentrations of detergents and salts [14]. Biotin can be coupled to several chemical moieties that target specific chemical groups, such as free amines of lysine residues and N-termini [15, 16], carboxyl groups of aspartate and glutamate residues and C-termini [17], sulfhydryl group of cysteine residues [18], and glycans [19, 20], all present in surface-exposed regions of membrane proteins. A broadly applied method to label the surfaceome is the use of an N-hydroxysuccinimide ester of biotin (NHS-biotin) that targets the accessible primary amines exposed on surface proteins [15]. Following cell lysis, biotinylated proteins are enriched by pull-down with streptavidin or captured by an anti-biotin antibody and digested into peptides for LC-MS/MS analysis [15, 21]. We here describe the development of a new labeling method for surface-exposed residues (Fig. 1). We explored the possibility of using D-biotin coupled to a furan moiety (furan-biotin) to target surface-exposed lysine, cysteine, and tyrosine residues exposed on membrane proteins. The furan moiety can act as chemical warhead

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13

Fig. 1 Scheme of the method to enrich surface proteins following their linkage to the furan-biotin compound. (1) Furan-biotin is cross-linked to surface proteins; (2) cells are lysed and proteins solubilized following standard procedures; (3) surface proteins, cross-linked to furan-biotin are captured by avidin beads; (4) captured proteins are digested on beads; (5) peptides are cleaned up from undigested proteins and digestion reagents; and (6) eluted peptides are analyzed by LC-MS/MS. NBS: N-bromosuccinimide (red dot, biotin; green dot, avidin; gray dot, bead)

that can be triggered into a keto-enol in a selective and spatiotemporally controlled way upon the production of singlet oxygen [22, 23] or by adding N-bromosuccinimide [24, 25]. Furanoxidation cross-linking has been extensively applied for several intermolecular cross-link applications in vitro: DNA-, RNA-, or PNA-interstrand cross-linking [22, 26], cross-linking DNA-peptide complexes [27], cross-linking of protein partners [28], as well as in vivo on living cells: ligand-cell surface receptor cross-linking [29]. More recently, we used the well-known heterodimeric E3/K3 coiled coil system to identify new nucleophilic partners to react with a furan moiety and form a covalent bond. We demonstrated that lysine, cysteine, and tyrosine are suitable nucleophiles for the keto-enol moiety generated upon furan oxidation [30]. We here show how furan-biotin can be used as a novel chemical label for affinity-based enrichment of plasma membrane proteins (Fig. 1).

2

Materials

2.1 Furan-Biotin Synthesis

1. Furfurylamine, 99%. 2. D-biotin. 3. 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate. 4. N,N-Diisopropylethylamine. 5. CH2Cl2 99.9%, extra dry.

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6. N2 gas (inert atmosphere for the reaction). 7. Ethyl acetate. 8. Saturated aqueous NaHCO3. 9. Saturated aqueous NH4Cl. 10. Distilled H2O. 11. Brine. 12. Magnesium sulfate. 13. Acetonitrile (ACN). 14. Trifluoroacetic acid (TFA). 15. Agilent 218 SEMI-PREP system with a UV-VIS dual wavelength detector. 16. SEMI-PREP column: Prepak cartridge (Delta-pak C18 100A). 17. SEMI-PREP solvent system A (H2O + 0.1% TFA). 18. SEMI-PREP solvent system B (ACN + 0.1% TFA). 2.2 Surfaceome Labeling

1. Furan-biotin stock: 50 mM furan-biotin in dimethyl sulfoxide (see Note 1). 2. N-Bromosuccinimide (NBS). 3. Phosphate-buffered saline.

2.3

Biotin Pull-Down

1. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid. 2. Tri-ethyl ammonium bicarbonate (TEAB). 3. Sodium deoxycholate. 4. IGEPAL® CA-630. 5. Sodium dodecyl sulfate. 6. Sodium chloride. 7. Ethylenediaminetetraacetic acid (EDTA). 8. Iodoacetamide. 9. Tris(2-carboxyethyl)phosphine hydrochloride (TCEP-HCl). 10. cOmplete™ protease inhibitor cocktail, tablets. 11. Bio-Rad DC Protein Assay Kit. 12. Pierce™ Avidin agarose beads. 13. Cell lysis buffer: 50 mM HEPES pH 8.0, 150 mM NaCl, 0.5% sodium deoxycholate (w/v), 1% IGEPAL® CA-630 (v/v), 0.1% sodium dodecyl sulfate (w/v), 1 mM EDTA in water, cOmplete™ protease cocktail tablet. 14. Pull-down washing buffer: 50 mM HEPES pH 8.0, 150 mM NaCl, 0.5% sodium deoxycholate (w/v), 1% IGEPAL® CA-630 (v/v), 0.1% sodium dodecyl sulfate (w/v), 1 mM EDTA in water.

Furan-Biotin-Based Affinity Purification of the Surfaceome

15

15. Urea washing buffer: 50 mM HEPES pH 8.0, 150 mM NaCl, 2 M urea in water. 16. Reduction and alkylation buffer: 50 mM HEPES pH 8.0, 150 mM NaCl, 2 M urea, 20 mM iodoacetamide, 10 mM TCEP.HCl pH 8.0 in water (see Note 2). 17. TEAB washing buffer: 50 mM TEAB in water. 18. Sequencing-grade modified trypsin. 19. Sonicator. 2.4 Cleanup of Purified Peptides

1. Bond-Elut OMIX C18 tips (Agilent). 2. Pre-equilibration solvent: 80% acetonitrile, HPLC grade, 0.1% trifluoroacetic acid. 3. Solvent A: 0.1% trifluoroacetic acid. 4. Solvent B: 60% trifluoroacetic acid.

3

acetonitrile,

HPLC

grade,

0.1%

Methods

3.1 Furan-Biotin Synthesis

The scheme of the synthesis of the furan-biotin compound is shown in Fig. 2. 1. Weigh 879 mg of D-biotin (3.60 mmol, 1.4 equivalents), and transfer it to a three-neck round-bottom flask (capacity 250 mL). 2. Add a magnet to the three-neck round-bottom flask containing the D-biotin. 3. Add 40 mL of dry CH2Cl2 to D-biotin under inert atmosphere (N2 gas) in the three-neck round-bottom flask, and stir it on a magnetic stir plate. 4. Dissolve 227.5 μL of furfurylamine (250 mg, 2.57 mmol, 1 equivalent) in 40 mL of dry CH2Cl2 in a three-neck roundbottom flask (capacity 100 mL) under inert atmosphere (N2 gas).

Fig. 2 Reaction conditions between D-biotin and furfurylamine. A solution of furfurylamine in dry CH2Cl2 was added over a solution of D-biotin in dry CH2Cl2 under an inert atmosphere (N2 gas). Then, HBTU was added and slow addition of DIPEA followed

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5. Add the solution of furfurylamine in dry CH2Cl2 via the cannula over the solution of D-biotin in dry CH2Cl2 under inert atmosphere (N2 gas) while stirring. 6. Weigh 1.95 g of HBTU (5.14 mmol, 2 equivalents). 7. Add HBTU in one portion to the furfurylamine and D-biotin mixture in dry CH2Cl2 while stirring under inert atmosphere (N2 gas). 8. Add 1.76 mL of DIPEA (10.28 mmol, 4 equivalents) to the mixture slowly while stirring under inert atmosphere (N2 gas). 9. Stir the reaction mixture at room temperature for 3 h under inert atmosphere (N2 gas) and a white precipitate will appear (urea). 10. Remove the urea from the reaction using the Buchner filter funnel, and collect the filtered liquid in the filter flask. 11. Transfer the filtered liquid in a round-bottom flask, and evaporate CH2Cl2 under reduced pressure using the rotary evaporator. 12. Dilute the crude reaction mixture with ethyl acetate (200 mL), and wash subsequently with saturated aqueous NaHCO3 (150 mL), H2O (150 mL), saturated aqueous NH4Cl (twice 150 mL), H2O (150 mL), and finally brine (150 mL) using the separating funnel. 13. Dry the organic layer over MgSO4, filter, and concentrate under reduced pressure using the rotary evaporator. 14. Purify furan-biotin on an Agilent 218 SEMI-PREP system with a UV-VIS dual wavelength detector using a Prepak cartridge (Delta-pak C18 100A) and a two solvent system, solvents A and B, at a flow rate of 65 mL/min. Elute the column with a linear gradient from 100% solvent A to 100% solvent B over 100 min. Collect and lyophilize the fractions containing furan-biotin (which elute at 32% solvent B, 32 min, see Note 3). 3.2 Surfaceome Labeling

The labeling of the surfaceome can be performed on a wide range of mammalian cells (see Note 4). Here, we provide a method to label MCF7 human breast cancer cells (#HTB-22™, ATCC). 1. Grow mammalian cells in 100 mm Petri dishes until they reach a confluency of about 80%. 2. Remove the growth media, and wash the cells with 10 mL of PBS. Repeat this step twice. 3. Dilute 50 μL of the furan-biotin stock solution in 5 mL of ice-cold PBS, and add it to the cells (see Notes 5 and 6). 4. Immediately add 50 μL of 50 mM NBS in PBS and mix gently.

Furan-Biotin-Based Affinity Purification of the Surfaceome

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5. Incubate the plates for 15 min on ice with gentle agitation. 6. Remove the labeling solution and wash three times with 10 mL of ice-cold PBS (see Note 7). 3.3

Biotin Pull-Down

1. Add 1 mL of cell lysis buffer to the plates. 2. Harvest the cells with cell scrapers and collect them in a 1 mL Eppendorf tube. 3. Sonicate at 30 kHz for 2 s and repeat nine times (see Note 8). 4. Put the tubes in an end-over-end rotator for 30 min at 4 °C. 5. Centrifuge the cell lysates for 10 min at 14,000 × g at 4 °C. 6. Collect the supernatant and determine the protein concentration using the Bio-Rad DC Protein Assay Kit. 7. Take a volume of supernatant corresponding to 2 mg of protein material and adjust the volume to 2 mL with lysis buffer. 8. Add the lysate to 50 μL of avidin resin that was washed three times with ice-cold cell lysis buffer (see Note 9). 9. Incubate the lysate with the beads in an end-to-end rotator for 2.5 h at 4 °C. 10. Pellet the beads for 3 min at 800 × g at 4 °C. 11. Discard the supernatant (unbound material) and wash the beads with 1 mL of pull-down washing buffer. Incubate the washing buffer with the beads in an end-over-end rotator for 15 min at 4 °C. 12. Repeat step 11 twice. 13. Wash the beads with 1 mL of urea washing buffer for 15 min at room temperature. 14. Pellet the beads for 3 min at 800 × g at 4 °C, and discard the supernatant. 15. Add 1 mL of reduction and alkylation buffer and incubate the beads in an end-over-end rotator for 20 min at room temperature in the dark (see Notes 10 and 11). 16. Pellet the beads for 3 min at 800 × g at 4 °C and discard the supernatant. 17. Add 1 mL of pull-down washing buffer and mix the beads by inversion. 18. Pellet the beads for 3 min at 800 × g at 4 °C and discard the supernatant. 19. Repeat step 17 with 1 mL of TEAB washing buffer (see Note 12). 20. Pellet the beads for 3 min at 800 × g at 4 °C and discard the supernatant.

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21. Resuspend the beads in 200 μL of TEAB buffer and add 1.5 μg of trypsin. 22. Incubate the mixture overnight at 37 °C in a thermo shaker with constant agitation. 23. Pellet the beads and collect the supernatant (containing the peptides) into a LoBind 1.5 mL Eppendorf tube and dry them completely under vacuum. 3.4 Cleanup of Peptides

1. Redissolve the peptides in 120 μL of solvent A. 2. Pre-equilibrate the Bond-Elut OMIX C18 tips with 150 μL of the pre-equilibration solvent by pipetting fresh solvent for five times. 3. Remove the pre-equilibration solvent by pipetting 150 μL of solvent A for 5 times (see Note 13). 4. Pipet the peptide solution for 20 times to let the peptides bind to the C18 beads. 5. Wash the peptides by pipetting 150 μL of solvent A for three times. 6. Elute the peptides by pipetting 70 μL of solvent B for ten times. 7. Repeat step 6, and combine both eluates. 8. Transfer the peptides to an HPLC vial and dry completely under vacuum (see Note 14).

3.5

Data Analysis

3.5.1 Search Settings for Peptides and Proteins

Data analysis includes the creation of peak list files from the recorded mass spectra and their search against a protein sequence database. In our lab, we typically use the MaxQuant search engine with the MaxLFQ algorithm, a generic method for label-free protein quantification [31]. Any other search engine and data analysis package able to perform these tasks can also be used. 1. Set fixed modifications to S-carbamidomethylation of cysteine residues. 2. Set variable modifications to methionine oxidation (sulfoxide) and N-terminal protein acetylation. 3. Set enzyme setting to Tryp/P to identify trypsin cleavages at Arg or Lys that is followed by Pro residue. 4. Set missed cleavages to two (see Notes 15 and 16). 5. Label-free quantification of proteins was set to a minimum of two peptides (razor + unique).

Furan-Biotin-Based Affinity Purification of the Surfaceome

4

19

Notes 1. Furan-biotin can be dissolved in dimethyl sulfoxide and kept at -20 ° C until further use. 2. TCEP and iodoacetamide can be diluted from a stock solution. Always prepare a fresh iodoacetamide stock solution. The TCEP stock solution has to be adjusted to pH 8 with 5 M NaOH by adding about 60 μL of 5 M NaOH to 200 μL of 750 mM TCEP-HCl (check the pH with indicator strips), further lowering the TCEP stock concentration to 570 mM. Adjusting this pH, and if necessary also the pH of the reduction and alkylation buffer, is critical to ensure efficient and complete alkylation of cysteine residues. 3. Analyze the final product by reverse phase high-performance liquid chromatography-mass spectrometry (RP-HPLC-MS) and 1H and 13C nuclear magnetic resonance spectroscopy (NMR). 4. The protocol has been set up to label mammalian cells. 5. Prepare the amount of labeling solution according to the number of replicates and use immediately. 6. Add equivalent amounts of DMSO to vehicle samples that can be used as control samples to evaluate the efficiency of biotinbased pull-down of plasma membrane proteins. 7. Remove PBS from the last wash as much as possible. Leftovers of PBS will dilute the cell lysis buffer. 8. Keep the lysate on ice during the sonication steps. 9. The volume of resin points to the bed volume and not the slurry volume. 10. The reaction should proceed in the dark as iodoacetamide is a light-sensitive reagent. 11. Longer incubation times should be avoided as these will promote alkylation at other groups, for instance, of the epsilonamines of lysine side chains [32]. 12. The TEAB washing buffer step aims to remove chemicals from previous steps. 13. All solvents are discarded once pipetted except for the sample that will be continuously pipetted for 20 times. 14. Samples are analyzed by LC-MS/MS. Any LC-MS/MS method and mass spectrometer can be applied. 15. Other settings such as the mass tolerance of the precursor and fragment ions are specific for the mass spectrometer used. 16. For peptide identifications, we suggest only peptides scoring above the threshold of a 99% confidence level are retained.

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References 1. Akasaka-Manya K, Manya H (2020) The role of APP O-glycosylation in Alzheimer’s disease. Biomolecules 10(11):1569. https://doi.org/ 10.3390/biom10111569 2. Fang J, Sheng X, Bao H, Zhang Y, Lu H (2021) Comparative analysis of intact glycopeptides from mannose receptor among different breast cancer subtypes using mass spectrometry. Talanta 223(Pt 1):121676. https://doi.org/10.1016/j.talanta.2020. 121676 3. Gupta R, Leon F, Rauth S, Batra SK, Ponnusamy MP (2020) A systematic review on the implications of O-linked glycan branching and truncating enzymes on cancer progression and metastasis. Cell 9(2):446. https://doi.org/10. 3390/cells9020446 4. Pinter P, Alpar A (2022) The role of extracellular matrix in human neurodegenerative diseases. Int J Mol Sci 23(19):11085. https:// doi.org/10.3390/ijms231911085 5. Sonbol HS (2018) Extracellular matrix remodeling in human disease. J Microsc Ultrastruct 6(3):123–128. https://doi.org/10.4103/ JMAU.JMAU_4_18 6. Bonnans C, Chou J, Werb Z (2014) Remodelling the extracellular matrix in development and disease. Nat Rev Mol Cell Biol 15(12): 7 8 6 – 8 0 1 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nrm3904 7. Chen R, Fulton KM, Twine SM, Li J (2021) Identification of MHC peptides using mass spectrometry for neoantigen discovery and cancer vaccine development. Mass Spectrom Rev 40(2):110–125. https://doi.org/10. 1002/mas.21616 8. Sendoel A, Dunn JG, Rodriguez EH, Naik S, Gomez NC, Hurwitz B, Levorse J, Dill BD, Schramek D, Molina H, Weissman JS, Fuchs E (2017) Translation from unconventional 5′ start sites drives tumour initiation. Nature 541(7638):494–499. https://doi.org/10. 1038/nature21036 9. Vialaret J, Schmit PO, Lehmann S, Gabelle A, Wood J, Bern M, Paape R, Suckau D, Kruppa G, Hirtz C (2018) Identification of multiple proteoforms biomarkers on clinical samples by routine Top-Down approaches. Data Brief 18:1013–1021. https://doi.org/ 10.1016/j.dib.2018.03.114 10. Overington JP, Al-Lazikani B, Hopkins AL (2006) How many drug targets are there? Nat Rev Drug Discov 5(12):993–996. https://doi. org/10.1038/nrd2199

11. Ulmschneider MB, Sansom MS, Di Nola A (2005) Properties of integral membrane protein structures: derivation of an implicit membrane potential. Proteins 59(2):252–265. https://doi.org/10.1002/prot.20334 12. Savas JN, Stein BD, Wu CC, Yates JR 3rd (2011) Mass spectrometry accelerates membrane protein analysis. Trends Biochem Sci 36(7):388–396. https://doi.org/10.1016/j. tibs.2011.04.005 13. Busch G, Hoder D, Reutter W, Tauber R (1989) Selective isolation of individual cell surface proteins from tissue culture cells by a cleavable biotin label. Eur J Cell Biol 50(2): 257–262 14. Bayer EA, Wilchek M (1980) The use of the avidin-biotin complex as a tool in molecular biology. Methods Biochem Anal 26:1–45. h t t p s : // d o i . o r g / 1 0 . 1 0 0 2 / 9780470110461.ch1 15. Zhao Y, Zhang W, Kho Y, Zhao Y (2004) Proteomic analysis of integral plasma membrane proteins. Anal Chem 76(7):1817–1823. https://doi.org/10.1021/ac0354037 16. Roesli C, Borgia B, Schliemann C, Gunthert M, Wunderli-Allenspach H, Giavazzi R, Neri D (2009) Comparative analysis of the membrane proteome of closely related metastatic and nonmetastatic tumor cells. Cancer Res 69(13):5406–5414. https:// doi.org/10.1158/0008-5472.CAN-08-0999 17. Ozkan Kucuk NE, Sanal E, Tan E, Mitchison T, Ozlu N (2018) Labeling carboxyl groups of surface-exposed proteins provides an orthogonal approach for cell surface isolation. J Proteome Res 17(5):1784–1793. https://doi. org/10.1021/acs.jproteome.7b00825 18. Hill BG, Reily C, Oh JY, Johnson MS, Landar A (2009) Methods for the determination and quantification of the reactive thiol proteome. Free Radic Biol Med 47(6):675–683. https:// doi.org/10.1016/j.freeradbiomed.2009. 06.012 19. Wollscheid B, Bausch-Fluck D, Henderson C, O’Brien R, Bibel M, Schiess R, Aebersold R, Watts JD (2009) Mass-spectrometric identification and relative quantification of N-linked cell surface glycoproteins. Nat Biotechnol 27(4):378–386. https://doi.org/10.1038/ nbt.1532 20. Matta C, Boocock DJ, Fellows CR, Miosge N, Dixon JE, Liddell S, Smith J, Mobasheri A (2019) Molecular phenotyping of the surfaceome of migratory chondroprogenitors and mesenchymal stem cells using biotinylation,

Furan-Biotin-Based Affinity Purification of the Surfaceome glycocapture and quantitative LC-MS/MS proteomic analysis. Sci Rep 9(1):9018. https://doi.org/10.1038/s41598-01944957-y 21. Udeshi ND, Pedram K, Svinkina T, Fereshetian S, Myers SA, Aygun O, Krug K, Clauser K, Ryan D, Ast T, Mootha VK, Ting AY, Carr SA (2017) Antibodies to biotin enable large-scale detection of biotinylation sites on proteins. Nat Methods 14(12): 1167–1170. https://doi.org/10.1038/ nmeth.4465 22. Op de Beeck M, Madder A (2012) Sequence specific DNA cross-linking triggered by visible light. J Am Chem Soc 134(26):10737–10740. https://doi.org/10.1021/ja301901p 23. Antonatou E, Hoogewijs K, Kalaitzakis D, Baudot A, Vassilikogiannakis G, Madder A (2016) Singlet oxygen-induced furan oxidation for site-specific and chemoselective peptide ligation. Chemistry 22(25):8457–8461. https://doi.org/10.1002/chem.201601113 24. Kobayashi Y, Nakano M, Kumar GB, Kishihara K (1998) Efficient conditions for conversion of 2-substituted furans into 4-oxygenated 2-enoic acids and its application to synthesis of (+)aspicilin, (+)-patulolide A, and (-)-pyrenophorin. J Org Chem 63(21):7505–7515. https://doi.org/10.1021/jo980942a 25. Halila S, Velasco T, Clercq PD, Madder A (2005) Fine-tuning furan toxicity: fast and quantitative DNA interchain cross-link formation upon selective oxidation of a furan containing oligonucleotide. Chem Commun (Camb) 7:936–938. https://doi.org/10. 1039/b415092a 26. Manicardi A, Cadoni E, Madder A (2020) Visible-light triggered templated ligation on surface using furan-modified PNAs. Chem Sci

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Chapter 3 Studying Cellular Dynamics Using Proximity-Dependent Biotinylation: Somatic Cell Reprogramming Reuben Samson, Francesco Zangari, and Anne-Claude Gingras Abstract Assessing the reorganization of proteins and organelles following the induction of reprogramming and differentiation programs is crucial to understand the mechanistic underpinning of morphological and fate changes associated with these processes. The advent of proximity-dependent biotinylation (PDB) methods has overcome some of the limitations of biochemical purification methods, enabling proteomic characterization of most subcellular compartments. The first-generation PDB enzyme, the biotin ligase BirA* used in BioID, has now been used in multiple studies determining the cellular context in which proteins reside, typically under standard growth conditions and using long labeling (usually 8–24 h) times. Capitalizing on the generation of more active PDB enzymes such as miniTurbo that can generate strong biotinylation signals in minutes rather than hours, as well as the development of an inducible lentiviral toolkit for BioID, we define here protocols for time-resolved PDB in primary cells. Here, we report the optimization and application of lentivirally delivered miniTurbo constructs to a mouse fibroblast model of somatic cell reprogramming, allowing the study of this dynamic process. This detailed protocol also provides a baseline reference for researchers who wish to adapt these techniques to other dynamic cellular processes. Key words Proximity-dependent biotinylation, BioID, Proximity labeling, Proteomics, Protein-protein interaction, Lentivirus, Reprogramming, Fibroblast, Mass spectrometry, Biotin, Streptavidin

1

Introduction Associations between proteins (and association of proteins with other macromolecules, including nucleic acids and lipids) are essential for proper cellular function. Traditionally, protein-protein interactions have been identified using a variety of methods [1], with cellular fractionation and affinity purification (AP) assays coupled with mass spectrometry (MS) among the most common [2]. These approaches rely on maintaining native protein interactions, complexes, organelles, or other structures to determine their composition and have been used to successfully define a knowledge base of protein-protein interactions [3–6] and the organellar organization of cells [7]. However, the gentle lysis conditions required

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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to maintain typical interactions make less soluble cellular compartments, including the cytoskeleton and plasma membrane, refractory to these strategies [2]. Another critical drawback of the mild lysis used in AP-MS and other biochemical fractionation experiments is the loss of spatial information associated with the process. The mixing of cell compartments after lysis can also promote artificial protein contacts, resulting in false-positive interactions [3, 8]. The emergence of proximity-dependent biotinylation (PDB) methods using peroxidases (APEX) or biotin ligases (BioID and related methods) has overcome some of these limitations [9, 10]. Briefly, in PDB, an enzyme that promotes the covalent attachment of biotin to proteins is fused to a protein of interest (referred to as a “bait”) and expressed within a relevant cellular context. Upon biotin (or biotin-phenol plus peroxide for peroxidase enzymes) supplementation, proximal proteins (referred to as “preys”) are modified with biotin moieties. As this linkage is covalent, there is no need to maintain native protein interactions, complexes, or organelles, and harsher lysis conditions (typically including ionic detergents) can be used, solubilizing previously refractory cellular compartments. Preys are purified using streptavidin (or other biotin-affinity) beads and are then either eluted from the beads or subjected to trypsin digestion to elute peptides that are subsequently identified by MS. Quantitative or semiquantitative assessment (e.g., using spectral counting, label-free intensity measurements in data dependent of independent acquisition modes, or isotopic/isobaric quantification) is usually performed to reveal the relative abundances of enriched biotinylated proteins. Performing experiments in parallel with proper negative controls enables the confident identification of proteins specifically detected in the bait’s proximity. These proteins include direct protein-protein interactors, indirect interactors bridged by other proteins or macromolecules, or proteins that simply reside within the neighborhood of the bait. PDB, and in particular BioID, has become an important approach for resolving the protein interactions that maintain cellular substructures. However, pioneering BioID studies used a single point mutant of the Escherichia coli BirA enzyme (R118G, known as BirA*) that typically required ≥6 h to generate sufficient signals for MS analysis. These long labeling times meant the acquired signal often encompassed the totality of a bait’s life cycle, limiting the information gleaned regarding dynamic processes, including cell signaling and rapid changes in protein associations or localization (e.g., due to morphological changes). To address this, many improved enzymes have been developed, including BioID2 [11], BASU [12], and miniTurbo and TurboID (which are 6- and 23-fold more active than BirA*, respectively [13]). In the case of miniTurbo and TurboID, this has enabled robust biotinylation

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within minutes, improving sensitivity and permitting studying dynamic processes [14]. However, it is necessary to optimize the labeling time for various enzymes and biological questions and to include proper controls [8, 15, 16]. While PDB approaches have now been used in hundreds of studies to identify proximity partners for baits of interest and reveal the composition of many membrane-bound and membraneless organelles [15–17], most have been performed in model cell lines (primarily HEK293 and HeLa cells) that do not represent all biological contexts. Recently, BioID was adapted for more generic delivery methods, including through lentiviral delivery, allowing the approach to be applied to more technically challenging cell types, including primary cells [18–24]. This has allowed for profiling of proteins in various cellular contexts. Somatic cell reprogramming, or the acquisition of pluripotency, is a highly complex stepwise resetting of the cell toward a naive state. Somatic cell reprogramming has been achieved in laboratory settings using various techniques, from somatic cell nuclear transfer (SCNT) and cellular fusion to the ectopic expression of a set of defined transcription factors (OCT4, KLF4, SOX2, and MYC; known collectively as OKSM, or the Yamanaka factors) [25, 26] to generate induced pluripotent stem cells (iPSCs), which could have broad applications in the fields of regenerative medicine and personalized therapeutics. Moreover, characterizing the molecular events that occur during this dedifferentiation process will expand our understanding of cell fate decisions, cellular identity, and cellular plasticity. However, the process remains poorly understood. Over the past two decades, many researchers have focused on characterizing the complex changes that occur during somatic cell reprogramming. Transcriptomic analysis of secondary (derived from a primary line) reprogramming mouse embryonic fibroblasts (MEFs) identified temporal switches in gene expression that led to the classification of three phases: initiation, maturation, and stabilization [27]. Quantitative and temporal proteomic analysis has also been performed on this secondary MEF reprogramming system, highlighting stage-specific proteome changes [28]. Notably, these studies reported profound morphological changes with hallmarks of a mesenchymal-to-epithelial transition (MET) that transformed the fibroblasts into tightly packed colonies during the initiation phase [27, 29]. This MET-like process has been identified as the rate-limiting step in the early acquisition of pluripotency, and manipulating it can enhance both reprogramming speed and efficiency [27, 29, 30]. Though many studies have aimed to understand the molecular mechanisms involved in cellular reprogramming, key technological limitations have hindered their scopes of inquiry. Performing proximity-dependent biotinylation analysis (BioID) in the context of early cellular reprogramming has

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the potential to capture the dynamics of cellular reorganization following the induction of cell fate transitions. In this chapter, we discuss the application of our lentivirusmediated BioID system to the study of early somatic cell reprogramming (see Fig. 1 for overview). We chose to use the secondary somatic reprogramming MEF system first described in [31] based on its relatively high cellular reprogramming efficiency, with ~50% of cells displaying rapid activation of early reprogramming markers. While we describe the system in some detail here, we note that other somatic cell reprogramming (or cellular differentiation) systems can also be used, as long as the following conditions are achieved: (1) the cells must be compatible with lentivirus-mediated transgene expression and (2) either changes in cellular state must occur in a high fraction (typically >50%) of the population or a selection system should be used to enrich for the desired population. We also outline the generation, validation, and use of our lentiviral BioID toolkit [24], focusing specifically on the use of the miniTurbo enzyme for temporal labeling. With a few modifications, this strategy is applicable to other dynamic cellular processes. Finally, we outline an updated protocol for proximity-dependent biotinylation followed by mass spectrometry (PDB-MS).

2

Materials

2.1 Generating Lentiviral Expression Constructs

1. Gateway entry vector (e.g., pDONR223) for the protein(s) of interest with either “open” (no stop codon) or “closed” (with stop codon) open reading frames (ORFs, depending on whether C- or N-terminal fusions are desired, respectively). 2. Gateway-compatible lentivirus PDB destination vector (described in [24]). BirA*, BioID2, TurboID, and miniTurbo versions can be requested from the Gingras lab. 3. Gateway LR Clonase II enzyme mix (Invitrogen). 4. TE buffer. 5. Competent bacteria (e.g., DH5α, NEB10B). 6. Lysogeny broth (LB). 7. LB-ampicillin (100 μg/mL) agar plates. 8. Ampicillin stock (100 mg/mL). 9. DNA miniprep kit. 10. BsrGI restriction enzyme. 11. Standard DNA agarose gel system (used for clone validation).

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Fig. 1 Lentivirus-mediated BioID in secondary reprogramming MEFs. (a) Generation of secondary reprogramming MEFs expressing a BioID-tagged bait. Secondary MEFs are isolated from E14.5 PB-TET chimeric embryos and simultaneously expanded and verified for chimeric contributions by fluorescent activated cell sorting (FACS) analysis. Specific pooled populations are then frozen at early passages. To generate a lentivirus that can mediate BioID-tagged bait expression in MEFs, HEK293TN cells are co-transfected with a secondgeneration packaging vector (psPAX2), coat protein vector (VSV.G), and the open reading frame transfer vector (ORF-BioID). Purified lentivirus is then used to transduce secondary MEFs prior to simultaneous reprogramming and bait expression. Upon biotin supplementation, proximal proteins (preys) are labeled (see (b)). Cells are then processed for streptavidin capture (see (c)), Western or streptavidin-HRP blot analysis, and/or fluorescence microscopy. (b) Proximal labeling during biotin supplementation. The enzyme tag on the bait (miniTurbo here) activates free biotin, covalently labeling neighboring proteins within a short radius (~10 nm). (c) General BioID workflow. After harvesting, cells are lysed, clarified, and subjected to streptavidinagarose bead capture. Biotinylated proteins are captured, stringently washed, and trypsin-digested on the beads. Samples (alongside controls) are subjected to LC-MS/MS followed by data analysis and visualization (Image created with BioRender.com)

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2.2 Generating Secondary Reprogramming MEFs

1. MEF culture medium: 500 mL Dulbecco’s modified Eagle’s medium (DMEM; high glucose with L-glutamine and sodium pyruvate), 50 mL fetal bovine serum (FBS; heat-inactivated at 56 °C for 30 min), 2.5 mL Penicillin/Streptomycin (100× solution), 5 mL MEM Non-Essential Amino Acids Solution (100× solution). 2. Phosphate-buffered saline without calcium and magnesium (PBS). 3. Embryo washing solution: 500 mL Hanks Balanced Salt Solution, 5 mL Penicillin/Streptomycin (100×). 4. 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA). 5. Gelatin coating solution: 1 g gelatin from porcine skin in 1 L ddH2O, autoclaved (see Note 1). 6. Tissue culture-grade water (autoclaved ddH2O). 7. 2 mL cryo-tubes. 8. MEF cell freezing medium: 25 mL MEF culture medium, 5 mL sterile dimethyl sulfoxide (DMSO), 20 mL FBS (heatinactivated at 56 °C for 30 min). 9. Sterilized and autoclaved surgical instruments: tweezers, forceps, scissors, scalpel, and scalpel blades. 10. Dissecting microscope. 11. 75% ethanol. 12. Flow cytometry solution: 2% FBS in PBS. 13. Flow cytometry tubes with cell strainer caps (Corning).

2.3 Generating Lentivirus for Bait Expression

1. HEK293TN cells (System BioSciences; HEK293T cells can also be used (ATCC)).

alternatively,

2. Standard cell culture medium: 500 mL DMEM, 50 mL FBS, 5 mL Penicillin/Streptomycin (100×). 3. Virus production medium: 500 mL DMEM, 25 mL FBS (heatinactivated at 56 °C for 30 min), 5 mL Penicillin/Streptomycin (100×). 4. psPAX2 (second-generation lentivirus packaging vector; Addgene). 5. VSV.G (envelope protein vector; Addgene). 6. Lentiviral BioID Subheading 3.1).

expression

vector

(generated

7. JetPRIME transfection kit (Polyplus-transfection). 8. Syringes (10 mL). 9. 0.45 μm filters. 10. 2 mL cryo-tubes.

in

Proximity-Dependent Biotinylation to Study Cell Reprogramming

2.4 Generating BaitExpressing Reprogramming MEFs

1. Secondary reprogramming Subheading 3.2).

MEFs

2. Frozen and titered Subheading 3.3).

lentivirus

BioID

(generated (generated

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in in

3. MEF culture medium. 4. Protamine sulfate stock solution (for 10 mL of 1 mg/mL (a 1000× stock)): dissolve 10 mg of protamine sulfate (Sigma-Aldrich) in 10 mL ddH2O, filter through a 0.45 μm filter, and store in 1 mL aliquots at 4 °C. 5. Reprogramming cell culture medium: 500 mL DMEM (high glucose with L-glutamine and sodium pyruvate), 93 mL FBS, 3 mL Penicillin/Streptomycin (100×), 6 mL MEM Non-Essential Amino Acid Solution (100×), 3 mL GlutaMAX (Thermo Fisher Scientific), 4 μL β-mercaptoethanol, and leukemia inhibitory factor (final concentration 1000 U/mL, Sigma-Aldrich). 6. Gelatin coating solution (see Subheading 2.2). 7. Doxycycline stock solution (for 10 mL of 1 mg/mL (a 1000× stock)): dissolve 10 mg of doxycycline in 10 mL ddH2O, filter through a 0.45 μm filter, and store in 1 mL aliquots at -20 °C, protected from light (see Note 2). 8. Biotin stock solution (for 20 mL of 20 mM (a 400× stock)): add 2 mL of 30% NH4OH to 100 mg biotin in a 50 mL centrifuge tube, and place on ice. Slowly add 18 mL of 1 N HCl, with the final 5 mL added dropwise with agitation to prevent biotin from precipitating. Store at 4 °C, protected from light (see Note 3). 9. 0.25% trypsin-EDTA. 10. Glass coverslips (12 mm). 11. PBS with calcium and magnesium (hereafter: PBS++). 12. PBS without calcium and magnesium (PBS). 13. Fixing solution: 4% paraformaldehyde (PFA) in PBS++. 14. Permeabilization solution: 0.25% Triton X-100 in PBS. 15. Fluorescence microscopy bovine serum albumin (BSA) blocking solution: 2.5% BSA in PBS. 16. Fluorescence microscopy milk blocking solution: 2.5% skim milk powder in PBS. 17. Fluorescence microscopy wash buffer: 0.05% Triton X-100 in PBS. 18. 6× Laemmli sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE) sample buffer (for 10 mL): add 1.2 g SDS, 6 mg bromophenol blue, 4.7 mL glycerol, 1.2 mL 0.5 M Tris–HCl pH 6.8, and 2.1 mL ddH2O to a 15 mL centrifuge

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tube and invert until dissolved, warming throughout in a 42 °C water bath to help with dissolving. Add 0.93 g of dithiothreitol (DTT), dissolve, aliquot, and store at -20 °C. 19. Ponceau S stain (for 500 mL): Combine 25 mL glacial acetic acid, 475 mL ddH2O, and 0.5 g Ponceau S dye. 20. Protein blot wash buffer (TBS-T): 0.1% Tween-20 in Trisbuffered saline (TBS). 21. Protein blot BSA blocking solution: 4% BSA in TBS-T. 22. Protein blot milk blocking solution: 4% skim milk powder in TBS-T. 23. Primary antibodies: monoclonal anti-FLAG M2 mouse antibody (1:2000, Sigma-Aldrich, for Western blotting and immunofluorescence), anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH; G-9) antibody (1:1000, Santa Cruz Biotechnology, for Western blotting). 24. Secondary antibodies: sheep anti-mouse IgG-horseradish peroxidase (HRP, 1:10,000, GE Healthcare Life Science), HRP-conjugated streptavidin (1:3000, GE Healthcare Life Science), goat anti-mouse Alexa Fluor 488 (1:1000, Thermo Fisher Scientific), Alexa Fluor 594 streptavidin conjugate (1: 2500, Thermo Fisher Scientific). 25. 4′,6-diamidino-2-phenylindole (DAPI) nuclear counterstain (1:20,000 of 20 mg/mL in PBS, Sigma-Aldrich). 26. ProLong Gold Antifade Mountant (ThermoFisher Scientific). 27. 10% SDS-PAGE gels and standard electrophoresis and membrane transfer systems. 28. Standard Western blot detection system (e.g., electrochemiluminescent reagent, autoradiography film, exposure cassette, and film developer). 29. Confocal microscope system capable of at least 400× total magnification and image processing software (e.g., Volocity or ImageJ). 2.5

BioID and MS

Non-autoclaved pipette tips and tubes are used throughout the procedure (see Note 4). Reagents and buffers are made using high-performance liquid chromatography (HPLC)-grade water (this is most important from the ammonium bicarbonate (ABC) washes and elution steps onward). 1. Complete modRIPA lysis buffer (for 50 mL; make fresh before use (see Note 5)): 50 mM Tris–HCl pH 7.5 (2.5 mL of 1 M stock), 150 mM NaCl (1.5 mL of 5 M stock), 0.1% SDS (500 μL of 10% stock), 1% Nonidet P-40 (5 mL of 10% stock), 0.4% sodium deoxycholate (4 mL of 5% stock), 1 mM MgCl2 (50 μL of 1 M stock), 1 mM triethylene glycol diamine

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tetraacetic acid (EGTA; 200 μL of 0.25 M stock), 0.5 mM EDTA (50 μL of 0.5 M stock), and HPLC-grade water up to 50 mL. Add 1:500 protease inhibitors and 1:100 phenylmethylsulfonyl fluoride (PMSF; 0.1 M in isopropanol) immediately before lysis. 2. Benzonase nuclease (250 U/μL, EMD-Millipore; alternatively, TurboNuclease (BioVision) may be used). 3. RNase A (10 mg/mL). 4. RIPA wash buffer (for 20 mL): 50 mM Tris–HCl pH 7.5 (1 mL of 1 M stock), 150 mM NaCl (600 μL of 5 M stock), 0.1% SDS (200 μL of 10% stock), 1% Nonidet P-40 (2 mL of 10% stock), 0.4% sodium deoxycholate (4 mL of 5% stock), 1 mM EDTA (40 μL of 0.5 M stock), and HPLC-grade water up to 50 mL. 5. SDS wash buffer (for 10 mL): 2% SDS (2 mL of 10% stock), 25 mM Tris–HCl pH 7.5 (250 μL of 1 M stock), and HPLCgrade water up to 10 mL. 6. TENN wash buffer (for 20 mL): 50 mM Tris–HCl pH 7.5 (1 mL of 1 M stock), 150 mM NaCl (600 μL of 5 M stock), 1 mM EDTA (40 μL of 0.5 M stock), 0.1% Nonidet P-40 (200 μL of 10% stock), HPLC-grade water up to 20 mL. 7. Streptavidin Sepharose High Performance beads (GE Healthcare). Different lots should be tested for quality control as in reference [32]. 8. ABC wash buffer (for 50 mL of 50 mM, pH 8.0): 200 mg of ABC in 50 mL HPLC-grade water (see Note 6). 9. Trypsin from porcine pancreas (proteomics grade; see Note 7): resuspend in ABC wash buffer to a final concentration of 100 ng/μL. 10. Formic acid (MS grade): 5% and 50% stocks freshly prepared in HPLC-grade water and stored in clean glass vials. 11. Autosampler vials and caps.

3

Methods

3.1 Generating Lentiviral BioID Expression Constructs

The lentiviral vectors engineered with PDB/BioID enzymes (BirA*, BioID2, miniTurbo, and TurboID) are designed for Gateway Cloning and can be used directly in LR reactions with entry clones (sequence-verified Gateway donor vectors containing the Open Reading Frame, ORF, of a protein of interest). Certain versions are also available in a multiple cloning site design for standard restriction enzyme/ligation cloning. The selection of the terminus that should be tagged for each bait protein should take known protein structures and interactions, including membrane topology when relevant (noting that N-terminal fusions should

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contain a stop codon while C-terminal fusions should not [33]); alternatively, the enzyme may be attached to each terminus in parallel experiments. Lentiviral backbone vector selection should also consider the ORF size (because of lentivirus packaging limits) and reflect the needs of the experiment, leveraging their unique features (e.g., selectable markers, an integrated reverse tetracyclinecontrolled transactivator (rtTA), or an integrated fluorescent tag) [24]. The choice of BioID enzyme should also reflect the biological question and consider their advantages and limitations [15, 16]. Here, we used the pSTV2-miniTurbo lentiviral destination vector to maximize ORF size and increase the temporal specificity of labeling (using 1 h of biotinylation). Fusion protein doxycycline-inducible expression will be driven by the rtTA transgene in the secondary reprogramming MEFs, negating the need to supply the rtTA on the same or a different lentiviral vector. 1. For the Gateway LR reaction, combine 0.5 μL entry vector (150 ng/μL), 0.5 μL destination vector (150 ng/μL), and 3.5 μL TE buffer in a microcentrifuge tube. 2. Thaw the LR Clonase on ice, briefly vortex (twice for 2 s each), add 0.5 μL to the reaction tube, and vortex briefly to mix well. Immediately return the LR Clonase to the freezer. 3. Incubate the reaction mixture at room temperature for 1 h to overnight (see Note 8). 4. Add 1 μL of Proteinase K to the reaction, mix briefly, and incubate at 37 °C for 10 min to inactivate the LR Clonase. Add 5 μL TE buffer to dilute the reaction mixture prior to transformation. 5. Briefly thaw 30 μL competent bacteria on ice, add 1 μL of the LR reaction, and incubate on ice for 10 min. 6. Heat-shock the bacteria at 42 °C for 45 s, and return to ice for 1 min. 7. Add 500 μL LB to bacteria, and recover for 1 h at 37 °C with shaking. Meanwhile, pre-warm an LB-ampicillin agar plate to 37 °C (all lentiviral vectors contain the ampicillin resistance cassette). 8. Plate 50–75 μL transformed bacteria solution, and incubate at 37 °C overnight. 9. Pick single colonies for standard mini-prep, midi-prep, or maxi-prep protocols as needed (see Note 9). 10. Verify purified plasmid DNA by complete sequencing and restriction enzyme digest (use BsrGI for Gateway plasmids, as the flanking recombination sites each contain a BsrGI cut site) followed by DNA gel electrophoresis to confirm the presence of the ORF (the insert should migrate to the appropriate band size).

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3.2 Generating a Chimeric Secondary Reprogramming MEF Line

It is important to generate a large homogeneous batch of early passage secondary MEFs with high reprogramming efficiency, because (1) since these are primary cultures, the cells will begin to senesce within five to six passages, (2) reprogramming efficiency diminishes with each passage, and (3) to perform most PDB-MSbased experiments, a significant quantity of starting material is required (typically 100 mg). The following protocols describe the large-scale generation of highly efficient, early passage secondary reprogramming MEFs.

3.2.1 Chimeric Embryo Generation

Embryo aggregation and implantation procedures should be performed in local mouse facilities with ethical approval from an institutional Animal Care Committee. For this work, embryo aggregation and implantation were performed in-house through the Model Production Core at the Centre for Phenogenomics (Sinai Health). The procedure is not described here (but has been extensively documented in [34]), as each mouse facility has specific requirements and may use different protocols for chimera generation. For the purposes of replicating this specific chimeric generation, we did the following (adapted from [31]). 1. Chimeric embryos were produced through the aggregation of PB-TET iPSC clumps (5–7 cells, derived from the Gt(ROSA) 26Sortm1.1(rtTA, EGFP)Nagy strain) with 2.5 days postcoitum (dpc) diploid Hsd:ICR (CD-1) embryos. 2. Embryos were transferred into pseudopregnant recipient females the next day. 3. At 14.5 dpc, females were sacrificed by cervical dislocation, and embryos were harvested for MEF isolation.

3.2.2

MEF Isolation

Not all embryos will lead to sufficient chimeric contribution; therefore, to attain a large population of secondary MEFs that reprogram with high efficiency (~50%), embryos are harvested and treated separately until characterized by flow cytometry for percentage of GFP positivity. Once chimeric contributions are confirmed (see Subheading 3.2.4), MEFs can be pooled and frozen in batches with known reprogramming efficiencies. The following protocol isolates MEFs from each embryo separately until chimeric contribution is confirmed. Use newly sterilized forceps and scissors to dissect the female and process the embryos. Alternatively, ethanol and flame sterilize each instrument before each step. 1. Lay a female mouse on its back in a clean laminar flow hood. Spray the abdomen with 75% EtOH and let dry for 1 min. 2. Using clean forceps, lift only the fur at the base of the abdomen (near the midline), and make a small snip. Pull and separate the fur vertically to expose the abdominal membrane and cavity.

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3. In the same position, lift only the abdominal membrane, and make a small snip (be careful not to puncture the intestine). Continue cutting the membrane toward the arms and legs, exposing the organs. 4. Carefully push the intestines aside to locate the cervical attachment point midway between the base of the abdomen and the uterine horns and snip to detach while slowly pulling the uterine horns away from the body. Continue excising the uterine horns by snipping away attachment points and fat pads until they are released at the ovarian ends. 5. Immediately place the uterine horns into a 100 mm dish filled with enough embryo washing solution to cover (see Note 10). Continue this process until all females are dissected. 6. Dispose of the carcasses and thoroughly sterilize the laminar flow hood and dissecting microscope. 7. Place uterine horns into a new 100 mm dish with embryo washing solution, and carefully dissect out each embryo from the horn wall and amniotic sac by making small incisions into each layer until the embryo is released. Cut the umbilical cord as close to the embryo as possible and let the blood drain. Continue this process with all harvested embryos. 8. In a new 100 mm dish with embryo washing solution, decapitate the embryo following the jaw line, snip open the chest cavity, and scrape to remove all internal organs (see Note 11). Wash the carcass once in embryo washing solution, and transfer to a numbered 60 mm dish with just enough embryo washing solution to keep it wet (~ 2 mL). Repeat for all embryos. 9. Process each embryo separately and mince 20–25 times using a scalpel blade. Add 2 mL of pre-warmed 0.25% trypsin-EDTA, spread tissue pieces evenly, and incubate in a humidified 37 °C incubator at 5% CO2 for 5 min (see Note 12). 10. Add an additional 2 mL of trypsin-EDTA solution, mix by trituration, and incubate for an additional 5 min. 11. Using a 5 mL pipette, mix the contents thoroughly (if cells do not resuspend easily, incubate them for an additional 2 min), and place them in a 15 mL conical tube with an equal volume of MEF culture medium. Allow the mixture to stand for 1 min, so large bone fragments can settle at the bottom of the tube. 12. Carefully transfer the contents (avoiding the larger tissue pieces) into a numbered and gelatinized 150 mm dish, and add pre-warmed MEF medium to 20 mL. Incubate overnight in a humidified 37 °C incubator at 5% CO2. Cells should adhere within the first 24 h, at which point they are at passage 0. 13. Monitor the cells daily until 85% confluent (typically 1–2 days).

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MEF Expansion

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Prior to freezing MEFs, they can be expanded to simultaneously assess the chimeric contributions of the iPSCs in each embryo (see Subheading 3.2.4) and to generate sufficient material for proteomic experiments. When cells are ~85% confluent, expand them into a corresponding set of numbered (to keep track of each embryo’s chimerism until flow cytometry) 150 mm dishes (now at passage 1) for further freezing and a 60 mm dish for characterization by flow cytometry, using pre-warmed solutions and the following passaging protocol. 1. Aspirate the medium and wash the cells with 10 mL PBS. Gently tap and swirl the plate to remove cellular debris and non-adherent cells. 2. Carefully aspirate large fragments (i.e., bone, cartilage) that may have carried over from the initial plating and all remaining PBS. 3. Add 2 mL of 0.25% trypsin-EDTA to the plate, and incubate for 1–2 min until the cells begin to lift off (see Note 13). This can be monitored using an inverted microscope. 4. Resuspend the cells in 8.5 mL of MEF medium (total volume: 10.5 mL), and pipette up and down to create a single-cell suspension. 5. In a gelatinized 60 mm dish containing 3 mL MEF medium, dispense 150–200 μL of the single cell resuspension, and incubate overnight in a humidified 37 °C incubator at 5% CO2 (to analyze chimeric contributions the next day, see Subheading 3.2.4 and Note 14). 6. In four gelatinized 150 mm dishes containing 18 mL MEF medium each, dispense 2.5 mL of the single cell resuspension, and incubate overnight in a humidified 37 °C incubator at 5% CO2 (for pooling and freezing, see Subheading 3.2.5). Monitor the cells daily until 85% confluent (1–3 days), replacing the medium as necessary.

3.2.4 Analysis of Chimeric Contributions

We have found that a chimeric contribution of 40–60% is sufficient to produce adequate material and reprogramming potential for PDB-MS experiments (as described in Subheading 3.4 onward). To determine the chimeric contributions of the PB-TET iPSCs in each embryo, the constitutive low expression of GFP (from the ROSA.26 locus of the contributing cells) can be analyzed by flow cytometry. Embryo-derived MEFs can then be pooled to generate batches with specific reprogramming potentials. A negative control set of CD-1 MEFs is required to determine appropriate flow cytometry parameters and gating. 1. For each 60 mm dish, aspirate the medium and wash cells with 2 mL PBS.

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2. Aspirate the PBS, add 0.25 mL of 0.25% trypsin-EDTA, and incubate for 1–2 min. 3. Resuspend to single cells using an additional 0.8 mL of MEF medium, and transfer to a 1.5 mL Eppendorf tube. 4. Gently spin down the cells at 400 × g for 4 min and aspirate the supernatant. 5. Gently resuspend the cells in 200 μL of flow cytometry solution, and pass the mixture through a cell strainer cap attached to a round-bottom polystyrene tube suitable for flow cytometry at 400 × g for 30 s. Briefly vortex to resuspend cells. 6. Perform flow cytometric analysis as per standard protocols to determine the chimeric contributions of each embryo (% chimerism = (GFP+ cells/total cells) × 100). Staining for cellular viability is recommended. 3.2.5 Large-Scale Batch Generation

Once the chimeric contributions of each embryo have been determined, embryo-derived MEFs can be pooled to freeze a large, highly efficient reprogramming MEF set at passage 2. It is also useful to freeze a negative (non-reprogramming) set and a low/medium reprogramming set (for follow-up experiments). From each confluent 150 mm plate, five cryovials can be frozen down. The following freezing protocol assumes that 20 confluent plates are mixed and frozen down in one batch: 1. Working in sets of 4–5 confluent 150 mm dishes (to avoid drying out the cells), aspirate the medium, wash the cells with 8 mL PBS, and aspirate the PBS. 2. Add 2 mL of 0.25% trypsin-EDTA to the plate, and incubate for 1–2 min. 3. Resuspend the cells in 8 mL of MEF medium until a single cell suspension is created, and transfer into a 50 mL centrifuge tube. To save medium, the resuspension medium from one plate can be reused to resuspend the cells on an additional plate. Continue until all plates in a desired set are harvested. 4. Gently spin down the cells at 400 × g for 5 min and aspirate the supernatant. 5. Gently resuspend the cells in 10 mL MEF freezing medium (pooling the contents of each 50 mL tube if more than one was used), and split them evenly into two 50 mL centrifuge tubes (each containing 5 mL resuspended cells). Add 45 mL MEF freezing medium to each tube and gently resuspend. 6. Transfer 1 mL aliquots of resuspended cells into pre-labeled cryo-tubes, resuspending frequently to ensure proper distribution, and freeze slowly in an insulated cryo-container at -80 °C. Store at -80 °C for up to 1–2 months, or for longterm storage, store in liquid nitrogen (see Note 15).

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3.3 Generating Lentiviral Particles to Transduce BioID Baits

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Lentiviral particles should be generated and used in an approved biological safety cabinet/facility following local safety protocols. The protocol below generates ~3 mL of lentivirus per BioID construct, which is typically enough for titer testing and the infection of two 10 cm plates of cells. The virus should be titered prior to use in large-scale BioID experiments. Titer can be estimated by transducing a known number of seeded cells with a diluting range of each virus, then performing fluorescence analysis (see Subheading 3.4.2). Use a titer that transduces 75–80% of cells. 1. Count and seed HEK293TN (or HEK293T) cells in 2 mL of standard cell culture medium per well in a 6-well plate so they are 80–85% confluent at the time of transfection (see Note 16), and incubate overnight in a humidified 37 °C incubator at 5% CO2. 2. The following day, set up the lentivirus transfection mixture: add 200 μL of JetPrime buffer, 1.3 μg of psPAX2 lentiviral packaging vector, 1.3 μg of lentiviral BioID expression vector, and 0.8 μg of VSV.G envelope vector to a 1.5 mL microfuge tube. Briefly vortex and spin down. 3. Add 8 μL of JetPrime, vortex for 1 s, and briefly spin down. Incubate the transfection mixture for 10 min. 4. Add the transfection mixture dropwise to a single well. Repeat as needed for all lentiviral BioID constructs, and return the cells to the incubator. 5. 4 h post-transfection, carefully replace the medium in each well with 3 mL of pre-warmed virus production medium, ensuring cells do not detach (see Note 17). 6. 48 h post-transfection, carefully collect the viral supernatant into 15 mL centrifuge tubes, spin down at 500 × g for 5 min, and pass the supernatant through a syringe attached to a 0.45 μm filter into a new 15 mL centrifuge tube (see Note 18). 7. Use virus immediately or freeze into one-time use (0.5–1 mL) aliquots at -80 °C (see Note 19).

3.4 Generating and Validating BaitExpressing Secondary Reprogramming MEFs

3.4.1 Generating BioID Bait-Expressing MEFs

The following protocols describe the generation and reprogramming of bait-expressing secondary MEFs, bait expression validation, and labeling of a single bait across four timepoints in early reprogramming (days 1, 2, 4, and 6). A single vial of highly efficient secondary reprogramming MEFs at passage 2 is sufficient to generate enough material for a single-replicate analysis of these four timepoints by MS. 1. Thaw one vial of secondary reprogramming MEFs rapidly in 37 °C water for 2–3 min until almost completely thawed (see Note 20).

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2. Gently resuspend the cells, and add to a 15 mL centrifuge tube; add warm MEF medium to a total volume of 4 mL. Gently pellet cells at 400 × g for 5 min (see Note 21). 3. Aspirate the supernatant, and gently resuspend the cells in 5 mL MEF medium, and add to a gelatin-coated 100 mm tissue culture plate containing 5 mL MEF medium. Swirl to evenly distribute cells, and incubate overnight in a humidified 37 °C incubator at 5% CO2. 4. When cells are 60–70% confluent, replace the medium with 10 mL MEF medium containing 1 μg/mL protamine sulfate to facilitate viral transduction. 5. Add the appropriate amount of freshly thawed pre-titered virus (from Subheading 3.3) dropwise to transduce the cells, and incubate overnight in a humidified 37 °C incubator at 5% CO2. 6. To prepare for reprogramming and subsequent harvest, expand the cells when they are 85–90% confluent. Trypsinize and resuspend (as in Subheading 3.2.3) in 11 mL total volume MEF medium, and replate into ten gelatinized 100 mm plates containing 10 mL MEF medium total using the following amounts: four plates at 1.2 mL each (for day 1), three plates at 1 mL each (for day 2), two plates at 0.9 mL each (for day 4), and one plate at 0.8 mL (for day 6; see Note 22). Optionally, single drops of resuspended cells (in 1 mL of MEF medium) can be plated onto gelatin-coated coverslips in a 24-well plate to examine bait expression, localization, and proximity labeling (by streptavidin-fluorescence) on each reprogramming day. These can be reprogrammed similarly to the 100 mm dishes (using scaled-down amounts) before fixation for fluorescence analysis (see Subheading 3.4.3). 7. Once the cells have reached 80% confluency, initiate reprogramming by replacing the medium with reprogramming medium containing 1 μg/mL doxycycline (this will also begin the expression of the bait-enzyme fusion). Continue reprogramming cells in their initial plate, refeeding them with fresh medium daily until harvest (see Note 23). 8. To provide sufficient signal for biotinylation capture, replace the medium one final time with medium containing a final concentration of 50 μM biotin for the intended labeling period (see Note 23). For miniTurbo labeling of proximal proteins in this experimental setup, 1 h of labeling is sufficient prior to harvest (see Note 24); however, the labeling time should be optimized depending on the biological question and application of the lentiviral system to other cellular contexts.

Proximity-Dependent Biotinylation to Study Cell Reprogramming 3.4.2 Preparing Cell Pellets for BioID

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Cell pellets can be prepared for BioID via (1) cold inactivation and cell scraping, (2) on-plate cell lysis, or (3) cold trypsin harvest. This protocol describes a gentle cold trypsin harvest as an ideal method to tackle both the adherent colonies formed during early reprogramming (which are difficult to collect by cell scraping) and that enables a simple pellet weight method for lysis buffer normalization. Because miniTurbo is a more kinetically active enzyme than BirA*, ensure harvesting is performed quickly to maintain consistent biotinylation across samples. 1. Shortly before harvest, pre-weigh a 1.5 mL centrifuge tube for each sample. This will be subtracted from the postharvest weight to attain the mass of the cell pellet. 2. Working quickly, remove the plates from the incubator, aspirate the medium, and add 8 mL cold PBS to wash the cells and remove debris and excess biotin (see Note 25). 3. Aspirate the PBS and add 1 mL cold 0.25% trypsin-EDTA for 1 min. 4. Resuspend cells in 8 mL cold MEF medium (to inactivate the trypsin), and transfer them to a 15 mL centrifuge tube on ice. The resuspension mixture can be reused to pool and resuspend all plates for a particular sample. 5. In a cooled centrifuge, gently spin down cells at 400 × g for 5 min. 6. Carefully aspirate all media, leaving the cell pellet undisturbed. Resuspend the cells in 1 mL cold PBS (to wash away excess media) and transfer to a pre-weighed 1.5 mL tube. 7. Spin down cells at 400 × g for 5 min. Carefully aspirate all PBS, leaving the cell pellet undisturbed. Weigh the tube again and determine the weight of the cell pellet. 8. Snap freeze pellets on dry ice and store at -80 °C for further processing.

3.4.3 Parallel Quality Control of Bait Expression, Labeling, and Localization

To quickly assess bait expression, labeling, and proper subcellular localization before investing efforts toward BioID sample preparation, we recommend performing protein blots (blotting with Streptavidin-HRP or FLAG antibodies) and fluorescence (streptavidin-fluor or bait immunofluorescence) analyses. A small subset of cells can be seeded, transduced, induced, and labeled within smaller multi-well plates for both assays prior to large-scale BioID. For protein blotting, on-plate cell lysis can be used to allow rapid collection of cellular material (see Note 26). Reagents for on-plate lysis should be chilled, and lysis should occur on ice. All other steps can be performed at room temperature. During largescale BioID, an aliquot of clarified lysate can also be reserved for protein blotting analysis (see Subheading 3.5.1). The staining

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protocols below should be used with your established standard SDS-PAGE, membrane transfer, and fluorescence microscopy methods. For streptavidin-HRP and Western blot analysis, we recommend verifying even loading and clear protein transfer to the membrane (i.e., no bubbles) before blotting using a Ponceau S solution. This can be removed with three washes with TBS-T. 1. Blotting for biotin: block membrane in 4% BSA in TBS-T for 1 h with shaking (see Note 27). Remove BSA blocking solution, and incubate membrane with HRP-conjugated streptavidin (1:2500) in BSA blocking solution. This can be done at room temperature for 1 h or overnight at 4 °C with constant agitation. Wash the membrane three times with TBS-T for 5 min with shaking to remove excess streptavidin. 2. Blotting for FLAG-tagged bait proteins: block membrane in 4% milk in TBS-T for 1 h with shaking. Remove milk blocking solution, and incubate the membrane with mouse anti-FLAG M2 antibody (1:2000) in milk blocking solution for 4 h at room temperature or overnight at 4 °C with constant agitation. Optionally, membranes can be simultaneously blotted for a loading control (e.g., rabbit anti-GAPDH antibody, 1:1000). Wash membrane three times with TBS-T for 5 min with shaking to remove excess antibody. 3. Incubate membrane with HRP-conjugated anti-mouse secondary antibody (1:5000) in milk blocking solution for 1 h at room temperature with constant agitation (see Note 28). If blotting for a loading control, simultaneously incubate the membrane with the appropriate HRP-conjugated secondary antibody. 4. Wash three times with TBS-T for 5 min with shaking to remove excess antibody. 5. Visualize the proteins using your detection system of choice. For fluorescence microscopy staining, the following protocol is performed at room temperature and can be paused and kept at 4 °C at any of the PBS wash steps after fixation. Storage for longer than 1 week (especially after cells are stained with fluorophoreconjugated antibodies) is not recommended. The following procedure follows the induction and labeling steps of transduced cells on multi-well plates containing gelatin-coated glass coverslips. All wash steps and incubations can be performed in 1 mL and 500 μL volumes, respectively. 1. After 1 h of labeling, aspirate the medium, and wash the cells once in PBS++. 2. Aspirate the PBS++ from cells, and fix them in 4% paraformaldehyde in PBS++ for 10 min. Collect and dispose of paraformaldehyde appropriately, and wash cells twice with PBS.

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3. Aspirate the PBS from the cells, and permeabilize them using 0.25% Triton X-100 (or NP-40) in PBS for 10 min (see Note 29). Aspirate and wash the cells twice with PBS. 4. For streptavidin staining, block the cells in 2.5% BSA in PBS for 1 h (see Notes 27 and 30). Aspirate the BSA blocking solution, and add a streptavidin-conjugated fluorophore (e.g., Alexa Fluor 594 (1:3000) to visualize in a red channel) in BSA blocking solution for from 1 h at room temperature up to overnight at 4 °C. From this point forward, keep the cells away from light to preserve fluorophore activity (see Note 31). Aspirate the streptavidin staining solution, and wash cells twice with wash buffer (0.05% Triton X-100 in PBS). 5. For FLAG-tagged bait protein staining, block the cells in 2.5% milk in PBS for 1 h. Aspirate the milk blocking solution, and add monoclonal anti-FLAG M2 antibody (1:2000) in milk blocking buffer for from 1 h at room temperature up to overnight at 4 °C. Aspirate the anti-FLAG staining solution, and wash cells twice with wash buffer. 6. Aspirate the wash buffer, and stain the cells with a fluorophoreconjugated anti-mouse secondary antibody (e.g., Alexa Fluor 488 (1:1000) to visualize in a green channel) in milk blocking buffer for 1 h at room temperature up to overnight at 4 °C (see Note 32). 7. Aspirate the secondary antibody solution, and wash cells twice with wash buffer. 8. To counterstain the nucleus, aspirate the wash buffer and stain cells with DAPI (1:20,000 of a 20 mg/mL stock) in PBS for 15 min. 9. Aspirate the DAPI stain and wash the cells twice with wash buffer. 10. Mount the coverslips onto glass slides with a mounting reagent (e.g., ProLong Gold AntiFade Mountant) and store horizontally in a dry dark place overnight (see Note 33). 11. Image and process slides using a standard fluorescent microscope and imaging software. 3.5

BioID

We have adapted our BioID protocol to use a modified RIPA lysis buffer with higher detergent concentrations to improve the capture of proximal proteins from both soluble and insoluble cellular compartments. This protocol requires 75–150 mg of cellular material, which typically corresponds to ~40 million cells or, for example, four confluent 100 mm plates of MEFs per replicate on the first day of reprogramming (see Subheading 3.4.1 for plating distribution). This input typically allows at least four injections into an AB Sciex TripleTOF 6600 mass spectrometer for data-dependent

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acquisition. For practical purposes, the protocol can be split into 2 or 3 days at the streptavidin bead capture and overnight trypsin digestion steps. 3.5.1 Lysis and Streptavidin Bead Capture

Steps 1–9 should be performed at 4 °C and/or on ice using prechilled solutions. All remaining steps can be performed at room temperature. This protocol assumes a 100 mg cell pellet and 1 mL of lysis buffer. Volumes need to be adjusted accordingly. 1. Using the frozen cell pellets derived in Subheading 3.4.2, determine the masses of the cell pellets (tube with pellet weight subtracted by empty tube weight). 2. Thaw the pellets on ice and add modRIPA lysis buffer (with freshly added protease inhibitors) at a ratio of 10:1 v/w (e.g., a 100 mg pellet receives 1 mL of lysis buffer). Resuspend the cell pellet completely by pipetting up and down. 3. Sonicate the lysate at 30% amplitude for 15 s (5 s “on,” 3 s “off”) using a probe sonicator (see Note 34). 4. Add 1 μL of RNase A and 1 μL of Benzonase (or Turbonuclease) and rotate end-over-end at 4 °C for 15 min to degrade nucleic acids. 5. Bring the total SDS concentration up from 0.1% to 0.5%, and mix end-over-end for 5 min to further solubilize proteins (see Note 35). 6. Clarify lysates at 15,000 × g for 15 min in a precooled (4 °C) centrifuge. 7. Meanwhile, prepare the streptavidin beads. Streptavidin Sepharose High Performance beads are supplied as a 60% slurry in 20% ethanol solution, and 15 μL of beads (~25 μL slurry volume) is used per sample. Transfer the desired amount of beads (accounting for pipetting loss) into a 1.5 mL centrifuge tube, and wash with 1 mL modRIPA three times (all bead centrifugation steps are performed at 400 × g for 30 s). Resuspend washed beads at a 1:3 ratio of beads to modRIPA (e.g., 15 μL beads in 45 μL modRIPA) to facilitate reproducible pipetting to new 1.5 mL centrifuge tubes. 8. Transfer each clarified lysate to a new 1.5 mL tube (see Note 36). Small aliquots (60 μL) can be taken here to assess the total lysates by Western or streptavidin-HRP blots (see Subheading 3.4.3). 9. Resuspend and distribute 45 μL of the bead mixture to another new set of 1.5 mL tubes (see Note 37). Add 1 mL (or the lowest volume among the samples) of each lysate to the beads and incubate for a minimum of 3 h to overnight at 4 °C, mixing end-over-end (see Note 38).

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1. After incubation, spin down the beads, and discard the supernatants (see Note 39), leaving ~20 μL above the beads to prevent loss. Optionally, take an aliquot of each supernatant to assess the unbound fractions by Western and streptavidinHRP blots. 2. Add 250 μL of RIPA wash buffer and transfer the beads to a new 1.5 mL tube. Wash the inside of the old tube with 500 μL of RIPA wash buffer to collect the remaining beads and add them to the new tube (see Note 40). 3. Centrifuge the beads at 400 × g for 30 s, and aspirate the supernatant. 4. Perform seven wash steps in this order (see Notes 41 and 42): once with SDS wash buffer, twice with RIPA wash buffer, once with TENN wash buffer, and three times with ABC wash buffer. If beads are still in suspension after centrifugation (particularly for the final ABC wash), increase the centrifugation time to 1–2 min before removing the supernatant. 5. Once all washes are complete, using a manual pipette, remove all residual ABC solution without disturbing the beads. 6. Add 60 μL ABC containing 1 μg of proteomics-grade trypsin to the beads, and incubate overnight (at least 12 h) at 37 °C with constant shaking/rotation (see Note 43). 7. Add 5 μL ABC containing 0.5 μg of proteomics-grade trypsin, and incubate for an additional 3 h. 8. Centrifuge beads and carefully transfer 50 μL of the tryptic peptides to a new 1.5 mL tube (see Note 44). 9. Add 80 μL of HPLC-grade H2O to the beads, and gently vortex to resuspend. Centrifuge and add 75 μL to the previously transferred peptides. Repeat for a total of two transfers (the pooled peptides should now be at 200 μL). 10. Spin the pooled peptides at 10,000 × g for 5 min to pellet any potential beads and/or debris that may have been transferred in the previous steps. 11. Carefully transfer 180 μL of peptides, leaving the bottom 20 μL undisturbed, into a new 1.5 mL tube. 12. Add 20 μL 50% formic acid (to a final concentration of 5%) to acidify the sample and inhibit trypsin activity and vortex. 13. Dry peptides in a SpeedVac concentrator or similar apparatus and store peptides at -40 °C until ready for MS analysis.

3.5.3 MS and Data Analysis

The volume in which to resuspend dried peptides will depend on many factors, not limited to but including the overall peptide abundance post-digestion (considering the starting material, bait expression level, and labeling time) and the LC-MS/MS setup

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(including the acquisition method). Typically, this method enables at least four 5 μL injections into an AB Sciex TripleTOF 6600 mass spectrometer for data-dependent acquisition. 1. Resuspend peptides in 10 μL 5% formic acid, and vortex the tube rigorously for 15–20 s. 2. Centrifuge sample at 12,000 × g for 5 min. 3. Add 3.5 μL 5% formic acid to an autosampler tube, then add 2.5 μL of resuspended sample, and mix by pipetting up and down. This gives a total volume of 6 μL, with 5 μL taken for injection. 4. Store unused samples at -40 °C (do not redry). If a sample needs to be rerun, thaw it at room temperature, vortex, and begin this sample injection procedure again at step 2. LC-MS/MS setups vary, and as such, the machine used, the acquisition method, and the analysis pipeline must be determined by the user. We have described standard LC-MS/MS setups previously (see examples in [35–37]) and only provide a brief outline here. Samples are placed into an autosampler and loaded onto fused silica columns (0.75 μm) packed with 10 cm C18 (3.5 μm) reversed phase material. Peptides are ionized using a nanoelectrospray ion source in line with a nano-HPLC system. A minimum of two biological replicates of each sample are analyzed using datadependent acquisition, alongside negative controls (to assist in identifying true proximal interactors and common contaminants). The sample acquisition order is randomized within batches (to minimize abundant peptides contaminating the same subsequent samples), and the samples are separated by long wash cycles. Data are stored, tracked, annotated, and processed using ProHits [38], an open-source software developed and maintained in-house. High-confidence proximity interactors can be identified by comparison with negative controls using the integrated statistical tool SAINT [8] or through the Contaminant Repository for Affinity Purification (CRAPome) website [39], which contains a wide set of negative controls for BioID experiments and also enables the use of user controls. After analysis, results can be visualized using ProHits-viz [40], a suite of web tools designed to support visualizing quantitative differences between baits and generate publication-ready figures.

4

Notes 1. To coat tissue culture plates, add gelatin coating solution (~5 mL/100 mm dish), swirl to cover the surface, and allow to coat for 1 h at room temperature in a biological safety

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cabinet. Aspirate the coating solution, and wash the plate once with an equal volume of tissue culture-grade water. Allow to dry for 1 h prior to use. Plates can be stored covered for up to 1 month. 2. Once a doxycycline aliquot is thawed, use within 1 month and then discard. If a precipitate forms or the solution darkens, discard. 3. Biotin can easily precipitate during its preparation if the pH decreases too rapidly (note that this is reversible to some extent with adding a few drops of ammonium hydroxide solution). Slow and careful addition of 1 N HCl while the mixture is cold is important. Biotin is stable at 4 °C for 6–12 months if protected from light. 4. As sterility is not essential for these protocols (other than during cell culture), we recommend using tips and tubes directly as received from the manufacturer to prevent potential contamination during autoclaving. It is also important to use gloves for all steps and protect all plasticware and reagents from dust and other environmental contaminants. 5. A stock of modRIPA lysis buffer can be kept at 4 °C, with protease inhibitors and nucleases added fresh to the working volume required for lysis. 6. Freshly made ammonium bicarbonate solution (ABC) has a pH of ~7.8 in our hands, but this may vary depending on the manufacturer. The pH increases over time (to pH ~8.5 in a few weeks). Mixing freshly prepared ABC with an older stock will readjust the pH to near 8.0 for optimal trypsin digestion. 7. We have not observed a difference in the digestion capabilities of trypsin supplied in 20 μg (Cat #T6567, Sigma-Aldrich) and 1 μg (Cat #T7575, Sigma-Aldrich) aliquots. The main differences between these formats are the cost per sample and the ease of use. 8. LR reactions can be done in as little as 10 min, though longer ORFs often require careful molarity calculations and longer incubation times. 9. Typically, a single colony grown overnight in a shaking culture of 2–4 mL LB containing ampicillin (100 μg/mL) is sufficient for a standard mini-prep. High-quality DNA is critical for downstream high-titer virus production. We recommend ethanol-precipitating eluates from commercial midi/maxiprep production to desalt and concentrate the DNA prior to transfection. 10. During embryo harvest, continue to transfer the uterine horns or embryos to new 100 mm dishes with embryo washing solution as needed, to remove excess debris and blood.

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11. It is crucial to remove all organs, especially the kidneys, which can be tucked away in the base of the intestinal cavity and difficult to see. 12. Separating the embryo sections in trypsin allows for proper dissociation and better recovery of MEFs over time. 13. Do not over-trypsinize the cells. As soon as they round up and lift off the plate, begin resuspending them. If insufficient detachment is detected, the plate can be placed at 37 °C for 1–2 min. 14. Analyzing chimeric contributions is best performed 1 day after plating the cells and not directly after embryo dissociation. This decreases the identification of non-adherent cells in the total population, enables the removal of dead cells and large debris, and ensures that the cells analyzed more closely resemble the frozen stock at the next passage. 15. It is important to thaw and test one vial of frozen cells to verify their viability and ensure that their reprogramming potential is maintained. 16. We find that seeding 650,000–700,000 HEK293TN cells in the wells of a 6-well plate the evening before transfection provides ~85% confluency the following morning. 17. Cells begin to round up during the process of viral production and will detach easily from the plate (reducing overall virus production). Ensure that the medium is added slowly and against the side of the well to prevent detachment. 18. Viral supernatants should be filtered slowly to avoid shearing the particles. When working with large volumes, the filter may become clogged. Do not force the supernatant through; rather, change the filter. 19. Filtered virus can be kept at 4 °C for up to 2 weeks with minimal loss to infection potential or at -80 °C for up to 6 months. If the virus is to be frozen, determine its titer using a frozen aliquot. 20. Thaw cells quickly and work with only four to five vials at a time so the cells are not exposed to high DMSO levels for too long. It is important to avoid overheating the cells during thawing. 21. Although cells can be added directly to a tissue culture plate containing fresh medium (with a medium change the next day), we recommend removing the DMSO (by gently pelleting the cells and resuspending in fresh medium) used during freezing as this can enhance cellular recovery and viability. 22. The described plating setup has been optimized for seeding density and accounts for cell growth over the required days.

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The number of plates specified will provide similar amounts of starting materials (i.e., cell pellet weights) for BioID-MS. 23. As cells proliferate and proceed through reprogramming, more total medium may be required to support them. Adding an extra 1–2 mL each day is recommended. Many cells may proceed to apoptosis and removing these is important. When refeeding the cells, gently tap the plate before removing the medium to dislodge and aspirate dead/dying cells. It is especially important that all debris and dead cells are removed during the final medium change prior to labeling. 24. Depending on the PDB enzyme used, the bait, the experimental design, and biological question being asked, a labeling test should be done to verify a sufficient signal compared to a control. For example, miniTurbo can provide a sufficient biotinylation signal for MS in as little as 5 min; however, for this experimental design, 1 h of labeling provided a robust signalto-noise ratio and high-quality protein identifications by MS. If working with enzymes with strong affinities to biotin (e.g., BioID2 and TurboID), the use of biotin-depleted serum is strongly recommended to minimize spurious biotinylation, as free biotin is commonly found in FBS. Biotin can be depleted by incubating 50 μL of streptavidin Sepharose beads (washed twice with sterile water) with 50 mL FBS and rotating it endover-end for 3 h at 4 °C. Pellet the beads by centrifugation at 1000 × g for 5 min, filter the supernatant (0.45 μm), and add it to the medium. 25. It is important to remove any dead cells and cellular debris before harvest. Gently tap the plate prior to removing the PBS to dislodge, and aspirate dead/dying cells. 26. If the cells are similarly confluent at harvest and lysis is performed correctly, the samples should be similar in overall protein concentration and can be directly processed for Western and streptavidin-HRP blot analysis. Otherwise, we recommend quantifying total protein concentration using a standard kit (e.g., DC Protein Assay kit, Cat: #5000122, Bio-Rad) and normalizing across the samples. 27. Use BSA when incubating with streptavidin-HRP conjugate to assess biotinylation, as milk will interfere with the biotinstreptavidin interaction. 28. Limit the incubation time with the HRP-conjugated antimouse secondary antibody to 1–2 h to prevent background binding. 29. Due to the tightly formed colonies that appear during reprogramming, more time for permeabilization may be needed (1–2 h), along with gentle rocking.

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30. Clarify BSA and milk blocking solutions at 500 × g for 1 min prior to use for fluorescence microscopy, as small particulates may be visible during imaging. 31. For all incubations with antibodies, the further along in reprogramming the cells are, the more time will be needed for the incubation step to ensure the antibody can penetrate the center of the colony. The addition of 0.05% Triton X-100 and gentle rocking overnight at 4 °C will enhance immunostaining. 32. Another subcellular marker can be added to visualize compartments or proteins within the cells in another channel (e.g., phalloidin in the 647 nm far-red channel to visualize the actin cytoskeleton), depending on the ability of the fluorescence microscope. 33. Slides can be kept for up to 1 week with minimal signal loss and stored at 4 °C for up to a few weeks away from light. If stored at 4 °C, remove at least 1 h before imaging to allow the slides to return to room temperature and completely redry. 34. Ensure the sonication probe is held ¾ down into the sample to maximize shearing. 35. The addition of SDS will be proportional to the initial lysis volume (e.g., to 1 mL lysis volume, add 40 μL of 10% SDS to get 0.5% SDS). 36. When transferring to a new tube, mix the lysate properly, in case any gradients form during centrifugation. 37. It is important to keep the beads well suspended through the aliquoting process, so they are evenly distributed across the samples. Using precut tips (cut with a clean razor blade) can help when aliquoting, along with frequently mixing the bead suspension. Distributing the bead mixture rapidly into new tubes instead of adding them directly to the lysate ensures accurate and reproducible bead volumes. This is preferred as beads can stick to the pipette tip (both inside and outside) during aliquoting, leading to variations between samples. 38. We find that 3 h is sufficient to capture the majority of the biotinylation signals in most samples using the suggested beadlysate ratio. Regardless of the incubation period chosen, it is important to use a consistent binding time when comparing different samples. 39. Following a quick centrifugation, rapidly invert the tube once to detach any beads still stuck to the sides/lid of the tube and centrifuge again. Allow beads to settle for 1 min prior to aspiration (a small portion may stick to the side of the tube if using a fixed rotor centrifuge). We suggest to leave ~20 μL volume above the beads during aspiration to avoid bead loss.

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40. Careful attention needs to be paid for this step to minimize bead loss. To remove residual contaminants that may adhere to the tube, the beads are placed into a new tube. Beads may adhere to the transfer pipette tip and the tube; therefore, do not discard the transfer tip. Use a separate pipette to add the RIPA wash buffer to the original tube. Then, using the initial transfer pipette, wash the tube and tip by pipetting up and down to transfer beads for minimal loss. 41. To minimize bead loss, avoid pipetting the beads up and down during further wash steps. Instead, add the buffer and invert until beads are resuspended (see Note 39). 42. A stepwise increase (50 μL) in wash volume can be performed for each successive wash (e.g., 800 μL for the SDS wash, 850 μL for the RIPA wash, etc.) to ensure that the detergent is completely removed prior to proteolysis. 43. For on-bead trypsin digestion, we use a “rotating drum” style rotator (a Cel-Gro Tissue Culture Rotator from Thermo Fisher Scientific) placed inside a 37 °C incubator or a thermomixer set to ~1100 rpm and 37 °C. Ensure beads are resuspended well in the trypsin mixture to maximize digestion. 44. Changing the tube at this step helps eliminate residues that may have adhered to the tube walls. A further tube transfer will prevent any remaining beads from contaminating the sample.

Acknowledgments The authors wish to acknowledge the Model Production Core at the Centre for Phenogenomics, Sinai Health, for generating iPSC embryo chimeras and Jeffrey L. Wrana for the PB-TET iPSC cells used for chimera production. We thank Payman SamavarchiTehrani for assistance in optimizing lentivirus generation, BioID, MEF generation, and reprogramming methodologies as well as for providing the lentiviral miniTurbo vectors; William Rod Hardy, Queenie Hu, and Dan Trcka for assistance in generating MEFs; Ugo Dionne for helpful comments; all members of the Gingras lab for helpful discussions; and High-Fidelity Science Communications for manuscript editing. This work was supported by a Canadian Institutes of Health Research Foundation Grant to A.-C.G. (FDN143301). Proteomics work was performed at the Network Biology Collaborative Centre at the Lunenfeld-Tanenbaum Research Institute, a facility supported by Canada Foundation for Innovation funding, by the Government of Ontario and by Genome Canada and Ontario Genomics (OGI-139). Anne-Claude Gingras holds the Canada Research Chair in Functional Proteomics.

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40. Knight JDR, Choi H, Gupta GD, Pelletier L, Raught B, Nesvizhskii AI, Gingras A-C (2017) ProHits-viz: a suite of web tools for visualizing interaction proteomics data. Nat Methods 14: 645–646

Chapter 4 Virotrap: Trapping Protein Complexes in Virus-Like Particles George D. Moschonas, Margaux De Meyer, Delphine De Sutter, Evy Timmerman, Petra Van Damme, and Sven Eyckerman Abstract The discovery of protein-protein interactions can provide crucial information on protein function by linking proteins into known pathways or complexes within the cell. Mass spectrometry (MS)-based methods, such as affinity purification (AP)-MS and proximity-dependent biotin identification (BioID), allowed for a vast increase in the number of reported protein complexes. As a more recent addition to the arsenal of MS-based methods, Virotrap represents a unique technology that benefits from the specific properties of the human immunodeficiency virus-1 (HIV-1) Gag polyprotein. More specifically, Virotrap captures protein complexes in virus-like particles budded from human embryonic kidney (HEK293T) cells, bypassing the need for cell lysis and thus supporting identification of their content using MS. Being intrinsically different to its two main predecessors, affinity purification MS (AP-MS) and biotin-dependent identification (BioID), Virotrap was shown to complement data obtained with the existing MS-based toolkit. The proven complementarity of these MS-based strategies underlines the importance of using different techniques to enable comprehensive mapping of protein-protein interactions (PPIs). In this chapter, we provide a detailed overview of the Virotrap protocol to screen for PPIs using a bait protein of interest. Key words Virotrap, Protein-protein interaction, Interaction network, Mass spectrometry, Protein complex

1

Introduction Mass spectrometry (MS)-based strategies to analyze protein complexes [1], such as affinity purification (AP)-MS [2], BioID [3], and Virotrap [4], enable interactome screening of bait proteins against complete cellular proteomes. MS-based methods have greatly contributed to our current understanding of intracellular protein complexes [5, 6] and have proven to complement each other vastly [7– 9]. In the context of binary protein interaction assays [1], complementarity among different assays—in part due to intrinsic method

George D. Moschonas and Margaux De Meyer contributed equally. Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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biases—is generally known and has therefore been supported by a standardized interaction confidence-scoring system [10]. The Virotrap technology is based on the viral particle-forming properties of the human immunodeficiency virus-1 (HIV-1) Gag polyprotein [4]. Expression of an N-terminal fusion of Gag to a bait protein of interest gives rise to budding of virus-like particles (VLPs) from HEK293T cells with the bait protein lining the inside of the particles (Fig. 1). As a result, proteins interacting with the C-terminal portion of the fusion, i.e., the bait, are abducted in the cytosol and subsequently trapped inside the VLP lumen. For downstream particle purification, FLAG-tagged vesicular stomatitis virus glycoprotein (VSV-G) is expressed alongside the Gag-bait fusion to mark the VLP surface with an affinity tag. We noticed that sole expression of FLAG-VSV-G expression did not result in robust presentation of the tagged marker. Co-expression of untagged VSV-G protein was required to circumvent this issue, likely explained by facilitating complexation of (a) tagged version(s) in the trimeric VSV-G complex. Upon budding of the VLPs along with their protein content in the cell culture medium, a straightforward anti-FLAG purification of the VLPs in collected, centrifuged, and filtered supernatant allows for isolation of the particles. Further sample processing involves specific elution from the purification matrix and a proteomic workflow consisting of particle lysis, detergent removal, and trypsin treatment for bottom-up shotgun proteomics. Virotrap was successfully applied on Harvey Rat Sarcoma viral oncogene homolog (HRAS) revealing Leucine-zipper-like transcriptional regulator 1 (LZTR1) [11] and TNF receptor-associated factor 7 (TRAF7) [12] as novel interaction partners, highlighting the complementary nature of Virotrap with existing technologies. In Masschaele et al., the method was compared side by side with AP-MS and BioID for ring finger protein 41 (RNF41) revealing overlapping and complementary data [7]. Due to Gag oligomerization, Virotrap captures interactions of clustered bait molecules, thereby increasing the avidity and allowing binding of low-affinity preys. The latter is clearly illustrated in the study of Thery and colleagues where Virotrap identified ring finger protein 213 (RNF213) as a novel ISG15 interacting protein. This interaction was only confirmed using GST pull-down where a similar avidity effect occurs while classic co-immunoprecipitation (co-IP) with antibodies failed [13]. In addition to its proven success in disclosing novel human PPIs, Virotrap has also been used in the context of host-pathogen interactions to screen for human targets of the Salmonella effector protein SspH2 [9]. The original paper reporting on the Virotrap method relied on the use of a “black list” strategy for elimination of false-positive interaction partner candidates that were co-sorted in the particles. While this strategy proved a powerful but highly stringent approach

Quantitative Interactomics Using Virotrap

1

Transfection

GAG-POI(bait) or

GAG-eDHFR(control)

VSV-G-FLAG

VSV-G

55

GAG HIV

VS

VS

2

VLPs Harvest

supernatant

intracellular

3

(WB Pilot Study)*

Sample Preparation for MS pH: 3

pH: 3

scale

pH: 7-8

APols

Trypsin

5

LC- MS/MS

Data Analysis

-LogP

IIntencity t it ((counts) t )

4

m/z

Log2 Difference

Fig. 1 Overview of the Virotrap procedure for protein-protein interaction screening. In Virotrap, a Gag-tagged bait protein, i.e., the protein of interest, is overexpressed in HEK293T cells along with VSV-G-(FLAG). In parallel, Gag-eDHFR is overexpressed in HEK293T along with VSV-G-(FLAG) as a control setup. Gag multimerizes at the plasma membrane and evokes budding of virus-like particles (VLPs) that are internally lined with the bait protein. As a result, bait-interacting proteins are trapped inside the VLPs (1). VLPs can be isolated

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for identification of relevant protein partners [4], we developed the straightforward filtering index (SFINX; sfinx.ugent.be) based on peptide count probabilities between bait and control experiments being the result of random events following a binomial distribution [14, 15]. To further enhance sensitivity, we found that label-free quantification (LFQ) using a pairwise differential comparison between experimental condition (i.e., bait of interest) versus control bait provided good results when particle levels are similar. The Andromeda search engine combined with quantification implemented in the MaxQuant computational platform [16], followed by Perseus statistical analysis and visualization [17] are ideally suited for Virotrap data analysis. It is fundamental to note that the experimental design, i.e., the number of replicate samples of both control and bait setup, plays a crucial part in the outcome of the Virotrap experiment. In fact, we typically use four replicates, which allows for retention of more protein identifications after filtering for valid values (i.e., allowing one missing value per setup) over an experiment with only three replicates per setup. This implies that less false-negative interaction candidates will occur. In this chapter, we lay out a detailed description of the experimental procedure for conducting a Virotrap screen using a bait protein of interest (POI), encompassing HEK293T transfection, VLP harvest, MS sample preparation, LC-MS/MS, and data analysis (Fig. 1). Further, we present a precursory optimization experiment to fine-tune VLP levels for optimal downstream pairwise comparison between experimental and control setup (Fig. 1). Note that more advanced designs involving multiple baits or bait variants are possible, but these require alternative analysis pipelines (e.g., limma which we will not address in detail in the protocol below).

2

Materials All HEK293T cell culture manipulations should be performed under aseptic conditions using laminar flow cabinets and strict procedures. Master cultures are best maintained without antibiotics to ensure aseptic procedures. All solutions are prepared with ultrapure water (resistivity of 18.2 MΩ.cm at 25 °C) and analytical-grade or MS-grade reagents. Solutions are typically stored at 4 °C unless stated otherwise.

ä Fig. 1 (continued) from the cell culture medium using anti-FLAG-coated beads (2). To disrupt the VLPs and precipitate the proteins in the VLP samples, amphipols (APols) is added to the immune-purified fraction. Subsequent trypsinization and acidification allow for collection of the resulting peptides from the supernatant (3). Following liquid chromatography (LC)-MS/MS (4), analysis of the raw files enables identification of the VLP protein contents and detection of candidate preys upon pairwise analysis of bait versus control setup (5). *A pilot Western blot (WB) experiment is conducted for optimization of Gag-bait versus Gag-eDHFR levels

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Table 1 Overview of plasmids used in Virotrap Name

Application

Addgene no.

pMET7-GAG-EGFP

Transfection control

80605

pMET7-GAG-HRAS/GW

Cloning of bait construct: cut and paste/ 80604/ Gateway (GW) 194634

pMET7-GAG-eDHFR

Gag-tagged eDHFR (control setup)

194635

pMET7-GAG-GOI

Gag-tagged protein of interest (bait)

–

pMD2.G

VSV-G envelope-expressing plasmid

12259

pcDNA3-FLAG-VSV-G

FLAG-VSV-G

80606

pSV-SPORT or any empty vector backbone (mock) with a mammalian promoter other than CMV or SRalpha (e.g., EF1-alpha)

Mock plasmid (no expression protein)

15386014 or other

2.1 Transfection for VLP Production

1. Human embryonic kidney 293T (HEK293T) cells (see Note 1).

2.1.1

2. Dulbecco’s Modified Eagle Medium (high glucose, glutamine, and pyruvate; DMEM).

Reagents

3. Culture medium: DMEM, 10% Fetal Bovine Serum, 100 U/ mL Penicillin-Streptomycin. 4. Transfection medium: DMEM, 2% Fetal Bovine Serum. 5. Dulbecco’s phosphate-buffered saline, without calcium and magnesium. 6. Polyethylenimine (PEI) transfection reagent (1 mg/mL): Dissolve 10 mg of polyethylenimine (PEI) (linear, MW 25,000) in 9.5 mL of 5 mM HCl. Adjust the pH to 7 with diluted NaOH, and add 5 mM HCl to reach a PEI concentration of 1 mg/mL. Sterilize the PEI solution by passing it through a 0.22 μm filter. Store the solution at 4 °C for 1–2 months or at -20 °C for up to 1 year. 7. Plasmids required for VLP production are listed in Table 1. 2.1.2 Materials and Equipment

1. 75 cm2 cell culture flasks (T75) with filter cap. 2. 6-well cell culture plate, TC-treated surface. 3. 1.5 mL laboratory tubes. 4. Cell culture facility (Biosafety level 1). 5. CO2 incubator at 37 °C with a humidified atmosphere containing 5% CO2.

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VLP Harvest Reagents

1. Tris-buffered saline washing buffer (TBS): 20 mM Tris–HCl, pH 7.5, 150 mM sodium chloride in water. 2. Magnetic beads decorated for antibody coating (see Note 2). 3. Anti-FLAG antibody (see Note 2). 4. Anti-Gag antibody (see Note 3). 5. Protein loading dye.

2.2.2 Materials and Equipment

1. Sterile syringe filters with PVDF membrane, 0.45 μm, 33 mm. 2. 10 mL syringes compatible with filter. 3. 15 mL polypropylene lab tubes. 4. 14 mL round-bottom polypropylene tube with snap cap. 5. Protein LoBind tubes (1.5 mL). 6. Rotator with rotator holders for microtubes and 15 mL tubes. 7. Magnet holder for 1.5–2 mL tubes. 8. Magnet holder for 15 mL tubes. 9. Thermomixer for 1.5–2 mL tubes. 10. Vortex mixer. 11. Microcentrifuge. 12. Centrifuge for 15 mL tubes (at 400 × g). 13. 5 mg/mL DYKDDDDK FLAG peptide dissolved in TBS. Aliquots stored at -20 °C. Avoid repeated freeze-thaw cycles.

2.3 Sample Preparation for Mass Spectrometry 2.3.1

Reagents

1. 1 mg/mL amphipol A8–35 (APols) freshly prepared in water. 2. 50 mM MS-grade triethylammonium bicarbonate buffer (TEAB) freshly prepared from commercial stock solution in water. 3. Sequencing-grade modified trypsin: Dissolved sequencinggrade modified trypsin in resuspension buffer (see Note 4). 4. MS-grade formic acid (FA).

2.3.2 Materials and Equipment

1. pH indicator strips. 2. Centrifuge for 1.5–2 mL tubes (with minimal centrifugation speed of 12.000 × g). 3. Vials, crimp/snap top, polypropylene, certified for MS, 250 μL, 1000/pk. Vial size: 12 × 32 mm (11 mm cap). 4. 1 mm Combination Seal: PE Snap Ring Cap, transparent, center hole, soft version; silicone white/PTFE blue, 55° shore A, 1.0 mm, cross-slitted.

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1. Centrifuge for crimp/snap-top vials. 2. Trapping column (see Note 5). 3. Analytical column (see Note 6). 4. Acetonitrile. 5. Trifluoroacetic acid. 6. Fused silica emitter with pulled tip (see Note 7). 7. Mass spectrometer (see Note 8).

2.5

Data Analysis

1. Spreadsheet program (e.g., Microsoft Excel). 2. Peptide search engine (e.g., MASCOT, Andromeda). 3. Quantification software for MS data (e.g., MaxQuant, Proteome Discoverer). 4. Software for data handling (e.g., Perseus, https://maxquant. net/perseus/; R Studio, https://www.rstudio.com/).

3

Methods We here describe the detailed methodology for Virotrap, a state-ofthe-art MS-based approach that relies on trapping proteins inside bait-core VLPs. In a typical Virotrap design, these bait-enriched VLP proteomes are compared to proteins trapped inside control VLPs, i.e., particles formed by the Gag-tagged control protein Escherichia coli dihydrofolate reductase (eDHFR). To allow highly sensitive analysis, a similar amount of VLPs must be produced between replicates as well as the bait and the control conditions. In our experience, the number of produced VLPs can vary among different Gag-tagged bait proteins. Several reasons may cause this effect including a difference in stability between translational Gag-fusions, a difference in localization that affects or even prevents budding, and binding of the bait protein to (large) structures that may interfere with the budding process. To limit the number of MS runs required to align VLP production levels, we recommend performing a small-scale pilot study experiment for quality control first, i.e., a Western blot (WB) experiment of Virotrap samples, before performing the Virotrap experiment for MS. Virotrap for WB will allow an estimate of VLP production (usually between bait and control) and can aid in optimizing VLP production at the transfection step by choosing between two different (high vs. low) amounts of the Gag-bait plasmid to be transfected (Table 2 and Fig. 2). This will enable a better comparison between the experimental conditions and the control conditions in Virotrap for MS as transfection conditions can be adjusted

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Table 2 Constitution of low and high Virotrap transfection DNA mixtures Plasmid

Low mix (μg)

High mix (μg)

pSVsport (mock)

2.68 (0.35) *

0

pcDNA3-FLAG-VSV-G

0.36 (0.04) *

0.36 (0.04) *

pMD2.G-VSV-G

0.71 (0.09) *

0.71 (0.09) *

pMET7-GAG-bait

3.75 (0.48) *

6.43 (0.84) *

Cell lysate eDHFR 73 kD

bait 1

130 kD

Supernatant/VLPs bait 2

eDHFR 73 kD

90 kD

bait 1 130 kD

bait 2 90 kD

250 kD

150

100 75

50

37

25 20 15

Scaling eDHFR and Bait .1 eDHFR-High g Bait .1-High Scaling eDHFR and Bait .2 eDHFR-High Bait .2-Low

Fig. 2 Optimization of Gag-bait VLP levels for Virotrap analysis. To ensure similar VLP levels of Gag-bait versus Gag-eDHFR (control) for downstream pairwise analysis, a pilot experiment can be performed to adapt the amount of transfected DNA to low or high (see Table 2) accordingly. If the Western blot analysis proves lower levels of the Gag-bait VLPs compared to the Gag-eDHFR setup, the high DNA transfection mix is selected (bait 1). Analogously, if the Gag-bait VLP levels show to be higher than the Gag-eDHFR VLPs, the low DNA transfection mix is opted (bait 2). The figure represents the WB results for cell lysate and VLP enrichment on the same blot, and protein abundances can thus be compared relatively

accordingly, concomitantly maximizing the sensitivity of the labelfree quantification and statistical analysis of the MS study. Evaluation of VLP levels by WB analysis follows the same steps as the Virotrap protocol for MS but requires specific modifications which are indicated in bold type interface by “( ) *” in the text below.

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3.1 Transfection for VLP Production for MS (and WB Pilot Study) *

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1. Clone the bait sequence of interest into the Virotrap plasmid (see Note 9). 2. Seed 1 × 107(1.28 × 106) * low-passage HEK293T cells (below ten passages) in a 75 cm2 flask (6-well plate) * in 8–10 mL (2 mL) * of culture medium for each sample (see Note 10). Culture the cells for 24 h at 37 °C in a humidified atmosphere with 5% CO2. 3. On the day of transfection, check whether cells are at adequate density (~80% confluence) using a light microscope. 4. For transfection, prepare two tubes with 625 μL (80 μL) * of DMEM without additives for each sample. Add Virotrap DNA mixture including bait plasmid as indicated in Table 2 to the first tube. Add 37.5 μL (4.8 μL) * of PEI solution to the second tube. 5. Add the contents of the second tube with the PEI mixture to the first tube with the DNA mixture. Vortex the DNA-PEI mixture briefly, and incubate for 10 min at room temperature. 6. During the 10 min incubation, replace the medium of the HEK293T cells with 9.5 mL (1 mL) * of transfection medium (see Note 11). 7. After incubation, add the DNA-PEI mixture dropwise to the corresponding flask. Tilt the flask gently to distribute the DNA-PEI mixture in the medium. 8. Culture the cells with the DNA-PEI mixture for 6–8 h at 37 °C in a humidified incubator with 5% CO2. 9. Remove the DNA-PEI mixture from the cells by replacing the medium with 8 mL (2 mL) * of culture medium. 10. Culture the cells for 40–42 h at 37 °C in a humidified incubator with 5% CO2.

3.2 VLP Harvest for MS (and WB Pilot Study) *

1. Verify that the transfected cell monolayers are confluent using a light microscope. 2. Transfer 8 mL (2 mL) * of medium from the cell cultures to a 15 mL tube (2 mL tube) *, and centrifuge at 400 × g for 3 min at room temperature to remove cellular debris. 3. Filter the centrifuged supernatant through a 0.45 μm filter. Collect the filtered supernatant in a 14 mL round-bottom tube (see Note 12). (This step can be omitted for WB pilot study (see Note 13).) * 4. For each Virotrap sample, transfer 20 μL (10 μL) * of magnetic bead suspension to a 1.5 mL tube, and resuspend them in 300 μL of TBS. Mix the bead stock gently by pipetting/ tapping or by tilting and rotation until reaching a homogeneous suspension before use. Do not vortex the beads.

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5. Place the tube on a magnetic stand, wait for 1 min, and remove the TBS by pipetting. 6. Remove the tube form the magnetic stand, and resuspend the beads in 200 μL of TBS and 2 μL (1 μL) * of anti-FLAG antibody. 7. Incubate the mixture for 10 min with end-over-end rotation at room temperature. 8. Place the tube on a magnetic stand, wait for 1 min, and remove the TBS. 9. Remove the tube from the magnetic stand, and resuspend the beads in 250 μL of TBS. 10. Place the tube on a magnetic stand, wait for 1 min, and remove the TBS. 11. Resuspend the beads in 50 μL of TBS (see Note 14). 12. Add 50 μL of bead suspension to each 14 mL round-bottom tube (2 mL tube) * with filtered supernatant. 13. Incubate the tubes for 2 h with end-over-end rotation at room temperature. 14. Place the tubes on a 15 mL magnetic stand (1.5 mL magnetic stand) *, wait for 1 min, and remove the supernatant. 15. Resuspend the beads in 1 mL of TBS, and transfer the contents to a protein low protein-binding 1.5 mL tube. 16. Place the tubes in a 1.5 mL magnetic stand, wait for 1 min, and remove the supernatant. 17. Resuspend the beads in 200 μL of TBS. 18. Place the tubes on a magnetic stand, wait for 1 min, and remove the supernatant. Remove all residual TBS by careful pipetting. 19. Remove the tube from the magnetic stand, and elute particles by resuspending the beads in 20 μL of TBS and 0.8 μL (200 μg/mL) of FLAG peptide. While resuspending, avoid sample loss on the tube wall. (Elute particles using 40 μL 1× WB Loading buffer.) * 20. Incubate the tubes for 30 min at 37 °C in a thermomixer at 400 rpm. (Incubate for 5–10 min at 65 °C at 400 rpm in a thermomixer) * 21. Place the tubes on a magnetic stand and wait for 1 min. 22. Transfer 20 μL of TBS with eluted particles to a new protein low protein-binding 1.5 mL tube. The volume of TBS with eluted particles should be about 20 μL. If the volume is higher (e.g., residual TBS from Subheading 3.2, step 19), transfer only 20 μL to ensure a correct concentration of APols in

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Subheading 3.3, step 1. (Bind beads to magnet and transfer supernatant to new tube, heat samples for 5 min at 95 °C and put on ice or store at -20 °C.) * 23. *Run samples on Western blot and probe the Gag-bait and Gag-eDHFR using an anti-Gag antibody. In Fig. 2, an example of a representative WB is shown for optimization two different baits for transfection as described above. 3.3 Sample Preparation for Mass Spectrometry

1. Add 2 μL of the APols solution to the sample (final APols concentration; 1 μg/μL) (see Note 15). 2. Vortex and incubate samples for 10 min at room temperature. 3. Acidify the samples to pH 3 by adding 1.6 μL of 2.5% (v/v) FA in water. Check the pH of the sample by spotting a small volume on a pH indicator strip. Lowering the pH will result in the coprecipitation of APols and proteins. If the pH is still higher than 3, add small extra volumes of 2.5% (vol/vol) FA (0.5 μL) until pH 3. 4. Centrifuge the acidified samples for 10 min at full speed (>12,000 × g) at room temperature. 5. Remove the supernatant leaving the precipitate on the tube wall untouched (see Note 16). 6. Dissolve the precipitated APols-protein pellets in 20 μL of freshly prepared 50 mM TEAB buffer. Vortex the sample to ensure proper resuspension. The pH of the sample should be between 7 and 8. Check the pH by spotting a small volume on a pH indicator strip. 7. Heat the samples at 95 °C for 5 min, and place them on ice for 3 min. 8. Short-spin the samples to collect evaporated liquid from the lids and walls of the tubes. 9. Add 0.5 μg of the sequencing-grade modified trypsin solution to the samples. 10. Perform overnight trypsin digestion by sample incubation at 37 °C in an incubator (see Note 17). 11. Acidify the sample to pH 3 by adding 1.5 μL of 5% (v/v) FA in water. 12. Centrifuge the acidified sample for 10 min at full speed (>16,000 × g) at room temperature (see Note 18). 13. Transfer the supernatant to a crimp/snap-top vial for LC-MS/ MS analysis (see Note 19).

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3.4 LC-MS/MS Analysis

Peptides are analyzed using LC-MS/MS. 1. Inject 7.5 μL of sample (see Note 20) followed by trapping at 10 μL/min for 4 min in loading solvent (0.1% trifluoroacetic acid in water/acetonitrile (ACN) (98:2, v/v)) on a 20 mm trapping column. Flush the peptides from the trapping column, and separate on an analytical column kept at a constant temperature of 45 °C. Elute the peptides by a nonlinear gradient starting at 1% MS solvent B (0.1% FA in water/acetonitrile (2:8, v/v)) reaching 55% MS solvent B in 80 min, 97% MS solvent B in 90 min followed by a 5 min wash at 97% MS solvent B, and re-equilibration with MS solvent A (0.1% FA in water) at a flow rate of 300 nL/min (see Note 21). 2. Operate the mass spectrometer in data-dependent mode, automatically switching between MS and MS/MS acquisition for the 16 most abundant ion peaks per MS spectrum. Acquire full-scan MS spectra (375–1500 m/z) at a resolution of 60,000 in the Orbitrap analyzer after accumulation to a target value of 3,000,000. The 16 most intense ions above a threshold value of 15,000 are isolated with a width of 1.5 m/z for fragmentation at a normalized collision energy of 30% after filling the trap at a target value of 100,000 for maximum 80 ms. MS/MS spectra (200–2000 m/z) are acquired at a resolution of 15,000 in the Orbitrap analyzer. The polydimethyl cyclosiloxane background ion at 445.120025 Da is used for internal calibration (lock mass).

3.5

Data Analysis

1. Use MaxQuant for peptide and protein identification on the acquired raw files (see Note 22). Include the sequences of all Swiss-Prot human proteins, the eDHFR protein (NCBI Gene ID 944790), HIV-1 Gag protein, the VSV-G protein, and the FLAG-VSV-G fusion protein in the search database as FASTA files (see Note 23). Set methionine oxidation and N-terminal acetylation as variable modifications. Indicate trypsin/P (include R/P and K/P cleavages) as protease, and allow for one missed cleavage. Set the false discovery rate at 1% on the PSM, peptide, and protein level. Enable matching between runs using a match time window of 0.7 min and an alignment time window of 20 min (see Note 24). 2. Perform further data analysis using the proteinGroups.txt MaxQuant output file in the Perseus software [17] or R Studio. Remove reversed hits, potential contaminants, and proteins only identified by site by filtering the dataset (see Note 25). 3. Apply log2 transformation on all valid LFQ intensities for protein identifications in bait and control replicates. Filter protein groups for at least three valid values in at least one group (in case of four replicate samples). Impute missing values from the lower part of the normal distribution, representing the detection limit of the instrument (see Note 26).

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4. Different statistics can be applied to identify enriched proteins in the bait samples. In Perseus, perform a t-test for each candidate protein of the experimental setup in a pairwise comparison with the (eDHFR) control condition to reveal specific bait interaction partners (see Note 27). 5. Visualize the results of these t-tests in a volcano plot (see Fig. 3). For each protein, the log2 (bait/control) fold change value is indicated on the X-axis, while the statistical significance (-log p-value) is indicated on the Y-axis. Correction for multiple testing is implemented and based on the number of randomizations. Proteins outside the curved lines represent candidate proteins co-enriched with the bait protein at a defined FDR (see Note 28).

4

Notes 1. For our Virotrap screenings, we use HEK293T cells obtained from [18]. 2. It is imperative to evaluate the compatibility of the beads with downstream mass spectrometric analysis. We recommend using Dynabeads MyOne Streptavidin T1 (Thermo Fisher Scientific cat. no. 65601) as these have been routinely used in Virotrap combined with the BioM2 monoclonal anti-FLAG antibodies (Sigma Aldrich cat. no. F9291). In case other beads are used (e.g., pre-coated anti-FLAG beads), adaptations to the bead and/or antibody quantity, related to the binding capacity, and handling conditions (e.g., wash buffer) may be required for successful/quantitative VLP capturing. 3. We recommend detection of Gag-tagged proteins on Western blot using mouse anti-HIV1 p24 antibody from Abcam (1/5000 dilution; cat no. 9071). 4. We generally use sequencing-grade trypsin (modified trypsin of which lysine residues have been modified by reductive methylation yielding a stable protease resistant to proteolytic digestion) from Promega. Preferentially store dissolved trypsin at 20 °C in aliquots tailored to the size of future experiments. 5. We typically use trapping column with 100 μm internal diameter × 20 mm, packed in-house with Reprosil-Pur Basic-C18HD, 5 μm, Dr. Maisch. Alternative trapping columns with similar properties can be used. 6. We typically use a 250 mm Waters nanoEase M/Z HSS T3 Column, 100 Å, 1.8 μm, 75 μm inner diameter, Waters Corporation. Alternative analytical columns with similar properties can be used. 7. We typically use a fused silica emitter with pulled tip: 10 μm, length—12 cm (CoAnn Technologies, LLC).

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George D. Moschonas et al. Filter: i. Identified in Reverse ii. Identified by site iii. Remove contaminants

Log2 transformation LFQ intensity

Filter based on valid values: - Number in minimum 3 reps - In at least one group

POI(bait).3

POI(bait).2

POI(bait).1

eDHFR(control).3

eDHFR(control).2

eDHFR(control).1

5

Group replicates

4

4.5

GAG/Control/Bait enrichment between samples

Imputation replace from normal distribution

FRD:0.05 S0:1 Number of randomisation:1000

POI(bait)

Statistical Analysis t-test / pairwise comparison

-LogP

Bait vs eDHFR

eDHFR

Visualisation e.g Volcano Plot

Difference (eDHFR-POI)

Fig. 3 Workflow for Virotrap data analysis. Following LC-MS/MS analysis of the Virotrap samples, raw files are searched in MaxQuant, and the output tables are subsequently processed as presented in the figure. LFQ intensities are log2 transformed and filtered, and replicates (reps) are grouped. The data table is further filtered based on a minimum of three valid values in at least one group. Next, missing values are imputed from the normal distribution, and statistics are applied to determine significantly enriched proteins in the experimental bait versus the control eDHFR setup. The results of the statistical analysis can be visualized in a volcano plot

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8. We typically use a Q Exactive HF Orbitrap mass spectrometer (Thermo Fisher Scientific) coupled to an UltiMate 3000 RSLCnano LC system (Thermo Fisher Scientific). Ionization source: Phoenix PneuNimbus dual-column nanoESI source (MS Wil, Zurich); however, an alternative Nanospray ESI source can be used. 9. The bait construct consists of the gene of interest (GOI) genetically fused to the coding sequence of HIV-1 Gag in the pMET7-GAG vector (Table 1). To generate this construct, either classic cloning with restriction enzymes starting from the pMET7-GAG-HRAS plasmid (Addgene, plasmid no. 80604) or Gateway cloning starting from the pMET7GAG-GW (Addgene, plasmid no. 194634) plasmid can be used. 10. Each Virotrap experiment consists of a minimum of two Gag-eDHFR control samples and two Gag-bait samples, but for improved statistical power and to ensure robust data generation by having sufficient valid proteins in the final lists, we recommend performing at least three and preferentially four replicates. The HEK293T cells should always be kept at a confluence of below 70% during cultivation. Besides the 75 cm2 flasks intended for the pMET7-GAG-bait and pMET7-GAG-eDHFR (Addgene, plasmid no. 194635), we recommend including an extra 75 cm2 flask for controlling transfection efficiency prior to VLP harvest making the use of pMET7-GAG-EGFP (Addgene, plasmid no. 80605). 11. Virotrap allows the trapping of proteins under conditions of pathway activation such as by stimulation with tumor necrosis factor (TNF)-α or interferon (IFN) [4]. For the latter, we optimized the IFN concentration that does not interfere with VLP formation together with interferon-stimulated gene (ISG) expression in HEK293T cells. During Subheading 3.1, step 6 adds 10 ng/mL IFN in the DMEM supplemented with 2% (vol/vol) FBS. 7–8 h incubation (before refreshing with DMEM with 10% (vol/vol) FBS, Subheading 3.1, step 9), is sufficient to stimulate the production of ISGs without disrupting proper VLP formation (Fig. 4). 12. We strongly recommend leaving behind a residual volume (approximately 0.5 mL) when collecting culture supernatant for centrifugation and 0.45 μm filtration. Sacrificing little material will ensure that small quantities of cellular debris will not contaminate the samples during VLP purification. Of note, a new syringe and 0.45 μm filter should be used for each sample, even for replicate samples. The 14 mL round-bottom tube with filtered supernatant can be kept at room temperature during the preparation of the affinity beads. Subheading 3.2, step 3 can be omitted for WB analysis.

B. Lysate

+10ng/mL IFN 48h

-IFN

A.

VLP enrichment

George D. Moschonas et al.

+10ng/mL IFN 24h

68

GAG-bait

Actin

Actin

ISG15

ISG15

C. POI(bait).3

POI(bait).2

POI(bait).1

+IFN

POI(bait).3

IFITM1 (ISG)

POI(bait).2

POI(bait)

POI(bait).1

HIV-GAG

-IFN

Fig. 4 Virotrap under IFN stimulation. (a) HEK293T cell lysate was collected 24 and 48 h after the cells were treated with 10 ng/mL rhIFNa2a and analysed by immunoblot (IB) against ISG15 (IFN stimulation marker) and actin. (b) Cells treated with 10 ng/mL IFNa2a and the lysate and supernatant (VLP enrichment) were collected and analyzed by immunoblot (IB) against ISG15 (IFN stimulation marker), actin, and Gag. Gag-tagged bait was detected in the VPL enrichment fraction suggesting that VLPs are able to form under this condition. (c) Profile plot generated via Perseus demonstrating the LFQ intensities of Gag (VLP marker), POI (Bait), and IFITM1 interferon-induced transmembrane protein 1 (ISG marker) between three replicates that were treated or not with IFN

13. As this step results in reduced background (i.e., cellular debris) for downstream MS analysis, it is not required for the Virotrap pilot study using WB. 14. Make sure to collect all the beads from the tube wall, and resuspend them completely to avoid differences in the amount of input beads between processed samples.

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15. Take APols out of the freezer only a few minutes before use. 16. The APols-protein precipitates are colorless and hence difficult to observe at low quantities. Mind the orientation of the tubes in the centrifuge and do not touch the tube wall that was positioned on the outside of the centrifuge with pipette tips. After removal of the supernatant, the tubes with APols-protein precipitate can be kept frozen at -20 °C for several weeks. 17. Small volume incubations are always performed in an incubator to prevent evaporation. We do not recommend using thermomixers for such incubations. 18. In this step, APols is precipitated and is hence removed from the peptide sample for compatibility with LC-MS/MS analysis. Trypsin is also removed from the digest by this precipitation step. 19. Avoid bringing air bubbles in the sample, as this will interfere with injection into the mass spectrometer. The samples can be stored at -20 °C for several weeks. 20. We do not measure the peptide concentration of our Virotrap samples. We typically inject 7.5 μL or ~35% of the Virotrap sample for LC-MS. 21. As a principle, we run the control samples before the bait samples. After each sample, a blank LC run is performed to wash remaining hydrophobic peptides off the analytical column and to avoid carryover between samples. We recommend performing those blank runs in every instrument. 22. While SFINX is fast and highly accurate, LFQ implemented in MaxQuant showed higher sensitivity for designs involving direct comparisons of (three or preferentially) four repeat experiments to controls, and in case of comparable VLP production. 23. We recommend using the latest database release version (taxonomy ID 9606), downloaded from https://www.uniprot.org. 24. “Match between runs” can be enabled to increase the number of sample-specific protein identifications. 25. After filtering, about 1000–2500 proteins are identified in a typical Virotrap experiment (four-replicate experiment). From our experience, the number of identified proteins can vary substantially between different baits and additionally depends on the available MS instrument and the MS settings used. At this step of the analysis, we recommend inspection of the data using a profile plot and comparing the intensities of Gag among all the replicates of the bait and the control (Fig. 3). Note that Gag is added as a separate entry to the search database and is therefore identified as an independent protein

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rather than as bait fusion. This allows comparison of the levels of the Gag levels across different samples. In an optimally controlled Virotrap experiment, highly similar Gag (and bait) protein levels are expected between all the samples. 26. Missing values are imputed with values from the lower part of the normal distribution, representing the detection limit of the instrument. We typically impute the missing values from normal distribution in Perseus using the default parameters (Width = 0.3, Down shift = 1.8, and Mode = “replacement of missing values” should be applied to each identified protein separately). The parameters of the distribution can be altered to simulate the abundance that the missing values would have if they had been measured, assigning random numbers that resemble the valid values as much as possible. In most cases, missing values represent low-abundance measurements that represent low-abundance proteins. It is important to note that imputation from the normal distribution masks putative prey candidates with low LFQ intensities, and hence additional visual inspection of the non-imputed data is recommended. In our analysis, proteins with low values present in all replicates of the bait samples and absent in the replicates of the control samples are considered as possible interactors. It is also essential to report the used values for this imputation approach. Alternatively, the quantile regression imputation of left-censored data (QRILC) function can be used as a plugin in Perseus or using the imputeLCMD package in R to improve on the latter concern by randomly drawing values from a truncated normal distribution for imputation [19]. 27. HIV-1 budding is a process that depends on several cellular host factors [20, 21]. Specifically, the proper budding of the newly synthesized HIV-1 particles, and as an extent the Virotrap VLPs, requires proteins that belong to the ESCRT complexes (primarily ESCRT-III) and ESCRT-interacting proteins [22–24]. The ESCRT complexes are involved in a broad range of plasma membrane and endomembrane remodeling functions in eukaryotic cells. Proteins that participate in HIV-1 budding can therefore be incorporated in VLPs and exemplify typical background proteins in case of Virotrap. We recommend being critical about the validity of those proteins if they are enriched for in bait samples. 28. The default number of randomizations in Perseus is 250, but we typically use 1000 randomizations. We recommend a maximum FDR value of 0.05. We typically set the S0 value (artificial within groups variance; this value defines the relative importance of the p-value and difference between means) to 1 in the Perseus software. FDR and S0 values can be adjusted accordingly to generate a more relaxed or stringent analysis. More

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advance statistics can be applied using limma (in R Studio or as a plugin in Perseus) fitting a linear model to the data to determine significantly enriched proteins for the bait samples upon comparison with the control samples in a pairwise analysis. References 1. Titeca K, Lemmens I, Tavernier J, Eyckerman S (2019) Discovering cellular protein-protein interactions: technological strategies and opportunities. Mass Spectrom Rev 38:79–111 2. Gingras A-C, Gstaiger M, Raught B, Aebersold R (2007) Analysis of protein complexes using mass spectrometry. Nat Rev Mol Cell Biol 8: 645–654 3. Roux KJ, Kim DI, Burke B, May DG (2018) BioID: a screen for protein-protein interactions. Curr Protoc Protein Sci 91: 19.23.1–19.23.15 4. Eyckerman S, Titeca K, Van Quickelberghe E et al (2016) Trapping mammalian protein complexes in viral particles. Nat Commun 7: 11416 5. Huttlin EL, Bruckner RJ, Navarrete-Perea J et al (2021) Dual proteome-scale networks reveal cell-specific remodeling of the human interactome. Cell 184:3022–3040.e28 6. Go CD, Knight JDR, Rajasekharan A et al (2021) A proximity-dependent biotinylation map of a human cell. Nature 595:120–124 7. Masschaele D, Wauman J, Vandemoortele G et al (2018) High-confidence interactome for RNF41 built on multiple orthogonal assays. J Proteome Res 17(4):1348–1360 8. Lambert J-P, Tucholska M, Go C et al (2015) Proximity biotinylation and affinity purification are complementary approaches for the interactome mapping of chromatin-associated protein complexes. J Proteome 118:81–94 9. De Meyer M, Fijalkowski I, Jonckheere V et al (2021) Capturing salmonella SspH2 host targets in virus-like particles. Front Med 8:1–11 10. Braun P, Tasan M, Dreze M et al (2009) An experimentally derived confidence score for binary protein-protein interactions. Nat Methods 6:91–97 11. Steklov M, Pandolfi S, Baietti MF et al (2018) Mutations in LZTR1 drive human disease by dysregulating RAS ubiquitination. Science (80- ) 362:1177–1182 12. Najm P, Zhao P, Steklov M et al (2021) Lossof-function mutations in TRAF7 and KLF4 cooperatively activate RAS-like GTPase signaling and promote meningioma development. Cancer Res 81:4218–4229

13. Thery F, Martina L, Asselman C et al (2021) Ring finger protein 213 assembles into a sensor for ISGylated proteins with antimicrobial activity. Nat Commun 12:5772 14. Titeca K, Meysman P, Gevaert K et al (2016) SFINX: straightforward filtering index for affinity purification-mass spectrometry data analysis. J Proteome Res 15:332–338 15. Titeca K, Meysman P, Laukens K et al (2017) Sfinx: an R package for the elimination of false positives from affinity purification-mass spectrometry datasets. Bioinformatics 33:1902– 1904 16. Cox J, Mann M (2008) MaxQuant enables high peptide identification rates, individualized p.p.b.-range mass accuracies and proteomewide protein quantification. Nat Biotechnol 26:1367–1372 17. Tyanova S, Temu T, Sinitcyn P et al (2016) The Perseus computational platform for comprehensive analysis of (prote)omics data. Nat Methods 13:731–740 18. Lin YC, Boone M, Meuris L et al (2014) Genome dynamics of the human embryonic kidney 293 lineage in response to cell biology manipulations. Nat Commun 5:4767 19. Lazar C (2015) imputeLCMD: a collection of methods for left-censored missing data imputation. R Package version 2 20. Von Schwedler UK, Stuchell M, Mu¨ller B et al (2003) The protein network of HIV budding. Cell 114:701–713 21. Sundquist WI, Krausslich HG (2012) HIV-1 assembly, budding, and maturation. Cold Spring Harbor Perspectives in Medicine 2: a006924–a006924 22. Gupta S, Bromley J, Saffarian S (2020) Highspeed imaging of ESCRT recruitment and dynamics during HIV virus like particle budding. PLoS One 15:1–13 23. Morita E, Sandrin V, McCullough J et al (2011) ESCRT-III protein requirements for HIV-1 budding. Cell Host Microbe 9:235– 242 24. Prescher J, Baumg€artel V, Ivanchenko S et al (2015) Super-resolution imaging of ESCRTproteins at HIV-1 assembly sites. PLoS Pathog 11:e1004677

Chapter 5 Thermal Proteome Profiling for Drug Target Identification and Probing of Protein States Patricia Sauer and Marcus Bantscheff Abstract Proteins are central drivers of physiological and pathological processes in the cell. Methods evaluating protein functional states are therefore vital to fundamental research as well as drug discovery. Thermal proteome profiling (TPP) to this date constitutes the only approach that permits examining protein states in live cells, under native conditions and at a proteome-wide scale. TPP harnesses ligand/perturbationinduced changes in protein thermal stability, which are monitored by multiplexed quantitative mass spectrometry. In this chapter, we describe a modular experimental workflow for TPP experiments using live cells or crude cell extracts. We provide the tools to perform different TPP formats, i.e., temperature range experiments, TPP-TR; isothermal compound titrations, TPP-CCR; and a combination thereof, 2D-TPP. Key words Mass spectrometry, TMT-based quantitative proteomics, Cellular thermal shift assay (CETSA), Functional proteomics, Drug target engagement

1

Introduction The majority of cellular processes are orchestrated by proteins whose functions need to be tightly regulated. Knowledge of the functional state of the proteome is therefore fundamental to understanding and ultimately modulating biological systems. Aside from posttranslational modifications, mutations and changes in the global environment (e.g., phase separation), a key mechanism regulating protein function is through interaction with other molecules, be it cofactors, metabolites, drugs, nucleic acids, or other proteins. These binding events also influence the biophysical properties of proteins, most notably their temperature-dependent unfolding, or melting, behavior [1]. Traditionally, thermal shift assays are used to probe compound-binding and are performed with purified proteins [2]. In 2013, the Nordlund laboratory first described the Cellular Thermal Shift Assay (CETSA) that probes

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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compound binding directly in cells or cell extracts by heating samples to a range of temperatures before quantifying the remaining soluble fractions of select proteins via antibody-based methods [3]. Thermal proteome profiling combines the principles of CETSA with multiplexed quantitative mass spectrometry-based proteomics to enable investigations of protein functional states at a proteome-wide scale [4]. There are three commonly used TPP formats (Fig. 1). Each configuration is characterized by a particular sample multiplexing strategy and addresses different experimental questions. Temperature range TPP (TPP-TR) experiments compare discrete conditions, e.g., compound treatment at a single concentration or a gene knockout, over multiple temperatures. When paired with an appropriate control (e.g., vehicle treatment or wild-type), the resulting protein melting curves reveal target proteins as well as downstream effects in cell-based experiments. Conversely, TPP experiments performed over a range of concentrations (TPP-CCR) measure dose–response curves at a single temperature (isothermal titrations). This setup allows to measure and compare compound affinities in situ. Finally, the two-dimensional TPP (2D-TPP) approach measures dose-dependent drug effects over multiple temperatures. This approach is highly sensitive and increases the number of accessible target proteins, as changes in thermal stability are best detected near a protein’s apparent melting temperature. Depending on the experimental setup, TPP experiments may capture abundance differences in addition to liganddependent thermal stability changes, thereby indicating influences on protein solubility or turnover (Fig. 1c). Recent years have seen many developments in experimental and data analysis procedures that further expand the scope and popularity of this method (reviewed in [5–8]). Here, we present the first detailed and comprehensive TPP protocol since 2015 [9] and describe the most up-to-date experimental workflows for TPP-TR, TPP-CCR, and 2D-TPP experiments using mammalian cell lines. Experimental schemes with live cells as well as the recently introduced crude lysate protocol [10] are covered. Crude lysate denotes a type of detergent-free cell extract that provides near native conditions for studying direct effects. Because the extract is not cleared by centrifugation after mechanically disrupting the cells, most protein complexes and membrane proteins are left intact. Lysate-based assays allow for higher throughput and faster turnaround, due to the possibility of preparing lysate stocks. Additionally, protein binding of plasma membrane-impermeable molecules such as ATP can be investigated using crude lysates [10]. TPP-TR experiments were traditionally less sensitive than 2D-TPP experiments, because vehicle-treated and compoundtreated samples are analyzed in separate MS experiments [11]. However, with the advent of TMTpro 16-plex isobaric

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Fig. 1 Thermal proteome profiling (TPP) experimental design. (a) Intact cells or crude cell extracts from a variety of biological sources can be subjected to thermal proteome profiling. This chapter focuses on mammalian cells. (b) Based on the specific combination of compound concentrations and temperature ranges, the three different TPP formats serve to identify compound targets (TPP-TR), to determine the affinity of the compound to its targets (isothermal titration), or both (2D-TPP). Higher multiplexing, e.g., via TMTpro 16-plex, enables combining treatment and control in one analysis, thus increasing throughput and sensitivity of TPP-TR experiments. Isothermal titration is also called TPP-CCR for compound concentration range. (c) Full melting profiles obtained from TPP-TR experiments indicate target engagement through shifts in the apparent melting point of a protein. TPP-CCR and 2D-TPP yield dose–response curves at a single or multiple temperatures, respectively, that enable the calculation of compound affinities for each target protein

tagging reagents [12, 13], and most recently TMTpro 18-plex [14], samples of two TPP-TR experiments can be analyzed in a single MS run. This setup results in considerable gains in sensitivity, proteome coverage, and overall efficiency [15].

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Fig. 2 Workflow for thermal proteome profiling (TPP) experiments. (a) The choice of input material, live cells or crude lysate, impacts procedure details for compound treatment, heat-induced protein denaturation and recovery of the remaining soluble protein population. Subheading 3 describing the respective steps for live cell or crude lysate-based experiments are indicated. (b) Subsequent sample processing, LC-MS/MS analysis and data analysis is performed irrespective of the type of biological material used. Abbreviations: A, adherent cells; S, suspension cells

Even though diverse cellular perturbations can be studied via TPP, for clarity reasons, we will focus on compound treatment in this chapter. In short, live cells or crude extracts are treated with compound, followed by heat treatment and filter-based separation of soluble from precipitated proteins (Fig. 2a). The soluble protein fractions are subjected to bead-based MS sample processing, and the resulting peptides are labeled using isobaric tandem mass tags (TMT). The pooled multiplex sample is offline fractionated by high pH reversed-phase chromatography and finally analyzed via LC-MS/MS analysis (Fig. 2b). As a detailed description of downstream analysis of TPP data is beyond the scope of this chapter, we will provide a general overview and refer to relevant publications for specifics.

2

Materials Prepare all solutions with ultrapure water and analytical grade reagents.

2.1 Preparation of Input Material

1. Cell culture facility with humidified incubators at 37 °C and 5% CO2.

2.1.1

2. Water bath at 37 °C.

Cell Culture

3. Complete growth medium: as required for specific cell line. 4. Cell culture dishes or flasks: typically, 15 cm dishes for adherent cells and T-25 or T-75 flasks for suspension cells. 2.1.2 Crude Lysate from Cell Pellets

1. Cell pellets. 2. Dulbecco’s phosphate-buffered saline (DPBS), no calcium, no magnesium.

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3. DNA digest buffer A: DPBS, 3 mM magnesium chloride (MgCl2). 4. Benzonase® Nuclease: recombinant, purity ≥90%. 5. 2× Sample buffer: 200 mM Tris–HCl, 250 mM Tris-Base, 20% (v/v) Glycerol, 4% (w/v) SDS, 0.01% (w/v) Bromophenol Blue. The pH of the 2× sample buffer should be about 8.8. Shortly before use, add DTT to a final concentration of 50 mM. 6. Bead Mill Homogenizer equipped with a cooling unit. 7. Bead Mill Accessories: bead beating tubes of appropriate size pre-filled with 0.28 mm ceramic beads. 8. Digital thermometer. 9. Polycarbonate tubes for ultracentrifugation: various sizes. 10. Ultracentrifuge. 11. SDS-PAGE system. 12. Agarose gel electrophoresis system. 13. Assay to determine protein concentration, e.g., Bradford or BCA. 14. Kit for isolating genomic DNA. 2.2 Compound Treatment

1. Dimethyl sulfoxide (DMSO), anhydrous. 2. Compound dilution series: Prepare a 200× dilution series of the compound in DMSO (see Note 1). Prepare on day of use. 3. 1 M dithiothreitol (DTT) in water: Weigh 7.7 g DTT, add water to a final volume of 50 mL, and sterile filter (0.22 μm). Aliquot and store at -20 °C. 4. Low protein-binding 1.5 mL reaction tubes. 5. Benchtop centrifuge: refrigerated, equipped with swinging bucket rotor and buckets for microplates as well as buckets for tubes.

2.2.1

Adherent Cells

1. Treatment medium: cell line-specific growth medium supplemented with the respective concentration of compound or vehicle. 2. Cell wash buffer: DPBS supplemented with the respective concentration of compound or vehicle. 3. Trypsin solution: trypsin solution supplemented with the respective concentration of compound or vehicle. Alternative cell dissociation solutions may be used as well.

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4. 5% medium: cell line-specific growth medium including only 5% fetal bovine serum (FBS), supplemented with the respective concentration of compound or vehicle. 5. Cell counting device: hemocytometer or automated cell counter. 2.2.2

Suspension Cells

1. Cell wash buffer: DPBS supplemented with the respective concentration of compound or vehicle. 2. Conical-bottom centrifugation bottles.

2.3

Heat Treatment

1. Thermal Cycler: functionality.

96-well

block,

temperature

gradient

2. PCR 96-well plates: semi-skirted (half-skirted), polypropylene, non-sterile. 3. Adhesive aluminum foil. 2.4 Live Cell Format: Cell Lysis

1. 20% IGEPAL® CA-630: 20% (w/w) IGEPAL® CA-630 in water. Weigh 40 g IGEPAL® CA-630, add water to a final mass of 400 g, and sterile filter (0.22 μm). Store solution at 4 °C for up to 1 year. 2. Protease inhibitor mix: 500 nM aprotinin, 40 μM bestatin, 100 μM leupeptin, 10 μM phosphoramidon, and 1.46 μM pepstatin (see Note 2). Aliquot and store at -80 °C for up to 1 year. 3. Lysis buffer: DPBS, protease inhibitor mix, 1.12% (w/v) IGEPAL® CA-630, 2.1 mM MgCl2. Add the protease inhibitor mix at a ratio of 1:18 (protease inhibitor mix/lysis buffer).

2.5 Collection of Soluble Protein Fraction 2.5.1

DNA Digest

2.5.2 Removal of Aggregated Proteins

1. Microplate shaker. 2. DNA digest buffer B: 0.8% (w/v) IGEPAL® CA-630, 1.5 mM MgCl2 in DPBS. Keep on ice. 3. Benzonase® stock solution: ≥5 units/μL Benzonase® endonuclease in DNA digest buffer B. Prepare fresh. Keep on ice. 1. Thick wall polycarbonate for ultracentrifugation: 230 μL capacity. 2. 96-well filter plate with 0.45 μm hydrophilic PVDF membrane. 3. 96-well filter plate with 0.22 μm hydrophilic PVDF membrane.

2.6 Mass Spectrometry-Based Proteomic Analysis 2.6.1 Sample Preparation and TMTpro Labeling

1. Bead mix: 5 μg/μL hydrophilic and 5 μg/μL hydrophobic carboxylate-modified magnetic particles in water (10 μg/μL total particle concentration, e.g., Sera-Mag SpeedBead Carboxylate-Modified Magnetic Particles, Cytiva). First, prepare a 1:1 mixture of hydrophilic and hydrophobic carboxylatemodified magnetic particles in a 1.5 mL tube. Place tube into a magnetic rack, and allow beads to settle for 30 s. Aspirate and

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discard supernatant. To wash the beads, take the tube off the magnetic rack, add 200 μL water, and mix by pipetting up and down three times. Place the tube back into the magnetic rack, and allow beads to settle for 30 s. Aspirate and discard supernatant. Repeat the wash step twice. Finally, add 200 μL water and keep bead mix on ice. Prepare fresh on day of use. 2. Protein cleanup solution: 0.5 μg/μL bead mix, 75% (v/v) ethanol, 3.75% (v/v) formic acid in water. Prepare on day of use. Keep at 4 °C. 3. 70% ethanol in water. Store at 4 °C. 4. 0.1 M Tris(2-carboxyethyl)phosphine hydrochloride (TCEP): 0.1 M TCEP in 1 M HEPES pH 8.5. Weigh 64.5 mg TCEP, and add 1 M HEPES pH 8.5 to a final volume of 2250 μL. Check pH. It should be about pH 7.5. Aliquot and store at 20 °C, protected from light, for up to 3 months (see Note 3). 5. 0.1 M 2-chloroacetamide (CAA): 0.1 M CAA in 0.1 M HEPES pH 8.5. Weigh 132.3 mg CAA, and add 14.1 mL 0.1 M HEPES pH 8.5. Check pH. It should be about pH 8.0. Aliquot and store at -20 °C. 6. Lysyl endopeptidase (LysC) solution: 0.1 μg/μL LysC in 0.1 M HEPES pH 8.5 (see Note 4). Prepare shortly before use. Keep on ice. 7. Trypsin solution: 0.1 μg/μL trypsin in 0.1 M HEPES pH 8.5 (see Note 4). Prepare shortly before use. Keep on ice. 8. Digest solution: 111.25 mM HEPES pH 8.0, 6.25 ng/μL LysC, 6.25 ng/μL trypsin, 5 mM CAA, 1.25 mM TCEP. Prepare shortly before use. Keep on ice. 9. Tandem mass tag (TMT) reagents: TMT10™ or TMTpro™ 16-plex label reagents (Thermo Scientific). Prepare 8.6 μg/μL stock solution in acetonitrile. TMT reagents are sensitive to moisture, so equilibrate to room temperature before opening to avoid condensation. Store solutions at -20 °C for up to 1 month. 10. Stop solution: 2.5% (w/w) hydroxylamine (NH2OH) in water. Prepare shortly before use. 11. Wash solution: 60% (v/v) acetonitrile in water. Store at 4 °C. 12. Saturated potassium chloride (KCl) solution: about 5 M KCl in water. Weigh 11.2 g potassium chloride, add water to a final volume of 30 mL, and mix by vortex for 3 min. The salt will not dissolve completely. Store at room temperature. 13. Activation solution: 80% (v/v) acetonitrile and 0.5% (v/v) trifluoroacetic acid (TFA) in water. Store at 4 °C. 14. 4% TFA in water. Store at 4 °C.

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15. Reversed-phase sorbent for solid-phase extraction cleanup, e.g., 96-well HLB μElution plate, 2 mg sorbent per well, 30 μm (Oasis). 16. Vacuum centrifuge. 17. 96-well polypropylene plate. 18. Polyolefin microtiter plate foil: sealing foil for microtiter plates using glue with high solvent resistance, e.g., polyolefin foil with glue in pressure-activated microcaps (HJ Bioanalytik). 19. Polyethylene microtiter plate foil, e.g., MS suitable polyethylene film with 96 adhesive-free zones. 2.6.2 Offline Reverse Phase Fractionation at Basic pH

1. Mobile phase A: 1.25% (v/v) ammonia in water. Prepare fresh. 2. Mobile phase B: 70% (v/v) acetontrile, 1.25% (v/v) ammonia in water. Prepare fresh. 3. UPLC system suitable for basic pH. Required components: biocompatible dual gradient pump, solvent rack, well plate autosampler, column compartment, variable wavelength detector (see Note 5). 4. Reversed-phase C18 trap column, e.g., XBridge BEH C18 Sentry Guard Cartridge, 130 Å, 5 μm, 2.1 mm × 10 mm (Waters). 5. Reversed-phase C18 analytical column, e.g., XBridge BEH C18 Column, 130 Å, 3.5 μm, 1 mm × 150 mm (Waters).

2.6.3 LC-MS/MS Analysis and Data Analysis

1. NanoLC mobile phase A: 3.5% (v/v) DMSO, 0.1% (v/v) formic acid in water. Prepare fresh. 2. NanoLC mobile phase B: 3.5% (v/v) DMSO, 0.1% (v/v) formic acid in acetonitrile. Prepare fresh. 3. NanoLC mobile phase C: 0.05% (v/v) trifluoroacetic acid in water. Prepare fresh. 4. Nano-flow ultra-performance liquid chromatography system, e.g., UltiMate 3000 RSLCnano LC or Vanquish Neo UHPLC systems (Thermo Scientific). 5. Reversed-phase C18 trap column, e.g., Acclaim™ PepMap™ 100 C18, 100 Å, 2 μm, 1 mm × 50 mm (Thermo Scientific). 6. Reversed-phase C18 analytical column: we use a custom-made C18 column, 120 Å, 1.9 μm, 0.1 × 500 mm. 7. High-resolution Orbitrap tandem mass spectrometer, e.g., Q Exactive Plus (Thermo Scientific). 8. Nano-electrospray ion source. 9. Column oven. 10. Search engine of your choice for protein identification, e.g., Mascot Server (www.matrixscience.com).

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11. Download isobarQuant (https://github.com/protcode/isob. git). 12. Download the R package TPP from Bioconductor (https:// doi.org/10.18129/B9.bioc.TPP). 13. Download the R package TPP2D from Bioconductor (https:// doi.org/10.18129/B9.bioc.TPP2D). 14. Download the R package RTSA (https://doi.org/10.5281/ zenodo.4274152).

3

Methods Perform TPP experiments at least as biological duplicates (see Fig. 2).

3.1 Preparation of Input Material 3.1.1

Cell Culture

3.1.2 Crude Lysate from Cell Pellets

Culture cells as appropriate for the specific cell line. Only work with authenticated cell lines. Follow standard cell culture practice and adhere to aseptic techniques to avoid cross-contamination. Keep an eye on morphology and viability of the cells over time. Always check cultures for visual signs of microbial and fungal infections. Test cells periodically for mycoplasma contamination. Use ice-cold reagents, and carry out all steps in a cold room or on ice. We recommend using tight fitting cooling racks to insure efficient cooling. Lysate-based TPP experiments should be performed in the absence of protease inhibitors; therefore, lysate quality critically depends on keeping samples cold continuously (see Note 6). 1. Partially thaw frozen cell pellets in a room temperature water bath until only small clumps of frozen cells remain, and then transfer tubes to ice. 2. Once cell pellets have thawed completely, estimate the pellet volume (see Note 7). 3. Add 1 volume of ice-cold DPBS to the pellet. 4. Incubate samples at 4 °C with overhead rotation at intermediate speed for 10 min until a homogenous suspension has formed. 5. Distribute the cell suspension to bead beating tubes of appropriate size that are pre-filled with 0.28 mm ceramic beads. 6. Set the temperature of the Bead Mill’s processing chamber to 5 °C or colder, if possible (see Note 8). 7. Monitor the temperature of the samples before and after homogenization, using a digital thermometer with stainless steel stem. Temperatures typically range from 1 °C before homogenization to 5–8 °C after homogenization.

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8. Lyse cells using the following settings: speed 4 m/s, time 0.10 s, number of cycles 1, dwell between runs 0 (see Note 9). 9. Immediately after the run, place tubes back on ice. 10. Pool the lysate in a precooled reaction tube. 11. Add 1 volume of DNA digest buffer A to the lysate. Mix by inverting the tube. 12. Take a 25 μL aliquot of the crude lysate for DNA digest quality control (see Subheading 3.1.3, step 1). Store at -20 °C if the test is not performed the same day. 13. Add Benzonase® nuclease to the crude lysate reaching a final concentration of ≥0.25 units/μL. Incubate at 4 °C with overhead rotation at intermediate speed for 1 h (see Note 10). 14. Following incubation, take another 25 μL aliquot of the Benzonase® nuclease-digested crude lysate for DNA digest QC (see Subheading 3.1.3, step 1). Store at -20 °C if the test cannot be performed the same day. 15. In addition, take a 50 μL aliquot of the crude lysate to assess protein integrity (see Subheading 3.1.3, step 2). If the aliquot is not processed the same day, snap-freeze in liquid nitrogen, and store at -80 °C. 16. The protein concentration used to calculate input amounts for the TPP assay is measured from cleared lysate (see Note 11). Transfer 50 μL of crude lysate to a thick wall polycarbonate tube for ultracentrifugation, and centrifuge for 20 min at 100,000 × g at 4 °C using an ultracentrifuge. 17. Afterward, carefully transfer the supernatant to a precooled reaction tube and keep on ice. Discard the pellet. 18. Measure the protein concentration via the assay of your choice, e.g., Bradford or BCA. 19. Aliquot the crude lysate into precooled cryotubes. Snap-freeze in liquid nitrogen, and store at -80 °C. 3.1.3 Quality Control of Crude Lysates

1. DNA digests quality control: isolate the genomic DNA from the lysate aliquots, e.g., using the commercial kit of your choice. Monitor the genomic DNA content by agarose gel electrophoresis. Use 0.8–1.0% agarose gel and a wide range marker for double-stranded DNA (e.g., 100–15,000 bp). 2. Protein integrity quality control: separate and visualize the proteins in the crude lysate via SDS-PAGE and Coomassie staining. Defined protein bands spread across the entire length of the gel indicate that most proteins are intact in the lysate. In contrast, the presence of an intense, diffuse band at the lower edge of the gel indicates poor quality lysates that contain high amounts of degraded proteins.

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3.2.1 Compound Treatment of Live Adherent Cells

83

Perform compound treatment as described on the subsection that is relevant to your biological system, i.e., live adherent cells (see Subheading 3.2.1), live suspension cells (see Subheading 3.2.2), or crude lysate (Subheading 3.2.3). 1. Prepare an adequate number of 15 cm cell culture dishes, at least one dish per compound concentration (see Note 12). Plate cells in advance so that they reach at least 70% confluency on the day of the experiment. We recommend using cells only up to passage number 15. 2. Pre-warm all media and buffers to 37 °C unless stated otherwise. 3. For the compound treatment, aspirate growth medium without disturbing the cells. 4. Add 15 mL treatment medium, starting with the lowest compound concentration (see Note 13). 5. Incubate cells for 90 min at 37 °C and 5% CO2 in a humidified incubator. 6. Aspirate treatment medium without disturbing the cells. 7. Carefully wash cells once using 10 mL cell wash buffer to remove FCS. Gently swirl the buffer on the plate, and then aspirate. 8. Add 4 mL trypsin solution, and incubate cells at 37 °C and 5% CO2 in a humidified incubator to detach cells. 9. Inactivate trypsin by adding 11 mL of 5% medium. 10. Transfer the cell suspensions to 15 mL tubes using serological 10 mL pipettes. Rinse each plate with a few milliliters of the respective cell suspension to transfer as many cells as possible. 11. Sediment cells by centrifugation for 3 min at 216 g at room temperature. Aspirate supernatant. 12. Add 10 mL DPBS containing the corresponding compound concentration or DMSO to the cell pellets. The solution should be at room temperature. Resuspend cells by pipetting up and down using a serological pipette until a homogenous single cell suspension is achieved. Yet, be careful not to damage cells during this step. 13. Sediment cells by centrifugation for 3 min at 216 × g at room temperature. Aspirate supernatant (see Note 14). 14. Repeat the wash step once but take a 50 μL aliquot of each homogenous cell suspension before centrifugation. Use these aliquots to assess cell viability post compound treatment (see Note 15). 15. Meanwhile, preheat thermocyclers to the respective temperature(s).

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16. After the second wash step, aspirate the supernatant completely without losing cells. 17. Add 1.3 mL room temperature DPBS containing the corresponding compound concentration or DMSO to the cells. From now on, it is no longer necessary to work under sterile conditions. 3.2.2 Compound Treatment of Live Suspension Cells

1. Expand suspension cells to a cell density of about 1.4 × 106 cells/mL. Approximately three to ten million cells are required per data point (see Note 12). We recommend using cells only up to passage number 15. 2. Pre-warm all media and buffers to 37 °C unless stated otherwise. 3. Determine cell density of the suspension cell culture via your preferred method. 4. Transfer the required amount of cells for the entire experiment to conical-bottom centrifugation bottles, and centrifuge for 7 min at 216 × g at room temperature. If the supernatant is still turbid afterward, extend the centrifugation time (see Note 14). 5. Carefully aspirate growth medium without disturbing the cell pellet. 6. Add fresh complete medium to the pellet. Adjust the cell density such that 15 mL of cell suspension contains the required number of cells per compound condition. 7. Resuspend cells by pipetting up and down using serological pipettes until a homogenous single cell suspension is achieved. Yet, be careful not to damage cells during this step. 8. Distribute 15 mL of the cell suspension to labeled T75 cell culture flasks. 9. Add 75 μL of the respective compound stock or DMSO (vehicle control), starting with the lowest compound concentration (see Note 13). 10. Incubate cells for 90 min at 37 °C and 5% CO2 in a humidified incubator with orbital shaking. 11. Transfer cells from T75 flasks to 15 mL conical tubes. 12. Wash cells, determine cell viability, and prepare cells for heat treatment as described in Subheading 3.2.1, steps 11–16.

3.2.3 Compound Treatment of Crude Lysate

1. For crude lysate TPP experiments, 30 μg protein is required per data point. For instance, each compound condition in a TPP-TR experiment comprising ten different temperatures requires a minimum of 300 μg protein. Calculate the necessary lysate volume in excess.

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2. If working with frozen lysate stocks, thaw crude lysate in a water bath at 25 °C. 3. Adjust the protein concentration of the crude lysate to 3 mg/ mL using ice-cold DPBS. 4. Distribute the respective volume of the compound solution or vehicle to 1.5 mL protein low-bind reaction tubes. 5. Using wide orifice pipet tips, briefly mix the diluted crude lysate to ensure a homogenous suspension, and transfer respective volumes to the prepared reaction tubes. 6. Thoroughly mix lysate and treatment solutions by inverting the tubes several times. 7. Incubate the lysate-treatment solution mixtures at 25 °C with orbital shaking at 800 rpm for 15 min. 8. Meanwhile, preheat thermocyclers to the respective temperature(s). 9. Proceed with heat treatment immediately after the compound incubation. 3.3

Heat Treatment

What constitutes a suitable temperature range depends on the TPP format, multiplexing strategy, and proteins of interest. Here are a few guidelines to help select the temperature or temperature gradient best suited to your application: Most proteins (>90%) heat denature, or “melt,” somewhere between 37 and 67 °C [4]. Studying particularly heat-stable proteins, such as quinone reductase NQO2, may require an extended or shifted temperature range between 60 and 80 °C [16]. When designing the temperature range, also take into account the particular capabilities of available thermocyclers as there will be a limit as to which temperature gradients are technically feasible. For standard TPP-TR experiments, we recommend using ten heating temperatures, with 37 °C as the lowest and 67 °C as the highest temperature. When combining both compound-treated and vehicle control samples in one TMTpro 16-plex experiment, we suggest a seven-point temperature gradient (e.g., 44–66 °C, ~3.5 °C steps) plus 37 °C as reference temperature. Heat treatment for TPP-CCR experiments should be performed at a temperature just above the melting point of the protein(s) of interest [17]. If the protein is stabilized by the compound, select the temperature based on the melting behavior in vehicle-treated samples. This way most of the protein is precipitated and not detectable without compound, but easily detectable upon compound addition. Conversely, for ligand-destabilized proteins, draw on to the melting point observed in the presence of saturation exceeding amounts of compound to achieve the maximum abundance window. TPP-CCR experiments performed under such conditions may even reveal thermal stability effects that cannot be detected as significant shift in a TPP-TR.

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Finally, for 2D-TPP experiments, we typically heat samples to 12 different temperatures ranging from 42 to 64 °C (equidistant steps). Activate the heated lid function on the thermocycler for heat treatment. 1. Thoroughly resuspend cells by pipetting up and down using wide orifice pipet tips (live cells) or invert tubes several times to insure a homogenous suspension of cellular fragments (crude lysate). A homogenous suspension of cells or crude lysate is crucial at this step. 2. Distribute 100 μL aliquots of each compound condition to PCR plates using wide orifice pipette tips. Mix samples in between aliquoting steps to avoid sedimentation or precipitation (see Note 16). 3. Live cells only: centrifuge PCR plates for 2 min at 256 × g at room temperature to sediment the cells. Then, aspirate 80 μL of the supernatant (see Note 17). 4. Crude lysate only: briefly spin down PCR plates (20 s) at room temperature to remove any air bubbles trapped at the bottom of wells. 5. Tightly seal PCR plates using adhesive aluminum foil. 6. Double-check that the thermocyclers are still active and set to the correct temperatures, as some instruments automatically switch off after some time. Only place samples in the thermocycler when designated temperatures have been reached. 7. Place PCR plates into preheated thermocyclers, and incubate plates for 3 min. 8. Remove plates from the thermocycler, and keep them at room temperature for 3 min (see Note 18). 9. Briefly spin down PCR plates at room temperature to collect condensate. 10. Crude lysate only: Place samples on ice, and continue with Subheading 3.5. 3.4 Live Cell Format: Cell Lysis

1. Add 50 μL lysis buffer to each sample. 2. Pipet up and down 25 times using regular pipet tips to disrupt the cells (see Note 17). Check for residual cell chunks. Especially for samples treated at higher temperatures, it may be necessary to add additional pipetting rounds. 3. Place PCR plates on ice.

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DNA Digest

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Carry out all steps at 4 °C or on ice unless stated otherwise. We recommend using an automated 96-channel liquid handling system, e.g., VIAFLO 96/384 (Integra), for all subsequent steps up to MS sample processing. 1. Crude Lysate only: to improve coverage of membrane proteins, add 4 μL of a 20% IGEPAL® CA-630 solution to each sample (final concentration of 0.8% detergent). 2. To remove DNA and reduce viscosity of the lysates, add an appropriate volume of the ≥5 units/μL Benzonase® nuclease stock solution to reach a final concentration of ≥0.25 units/μL Benzonase® nuclease. 3. Thoroughly mix each sample by pipetting up and down ten times. Make sure that all pellets are sufficiently dissolved, and lysates appear homogenous. 4. Seal PCR plates using adhesive aluminum foil. 5. Incubate samples for 1 h on a microplate shaker with 700 rpm at 4 °C.

3.5.2 Removal of Aggregated Proteins (Live Cell Format)

1. Pre-wet a 96-well filter plate with 0.45 μm pore size using 50 μL DNA digest buffer B. Place the filter plate on a 96-well collection plate, and centrifuge for 2 min at 400 × g at 4 °C. Discard flow-through, and then place the filter plate on a fresh 96-well collection plate. 2. Centrifuge PCR plates containing the DNA-digested samples for 3 min at 400 × g at 4 °C. 3. Transfer 55 μL supernatant from the PCR plate to the pre-wet filter plate. 4. Centrifuge plates at 500 × g at 4 °C for 5 min. Repeat centrifugation once if residual lysate remains in the filter plate. 5. Distribute 40 μL of 2× sample buffer to a new 96-well collection plate (see Note 19). 6. Transfer 40 μL filtered sample to the prepared sample buffer plate. 7. Incubate samples at room temperature with orbital shaking at 450 rpm for 30 min. 8. Use the remaining 10 μL of filtered lysate to determine the protein concentration. Measure for each compound concentration the lowest two temperatures as well as the highest temperature (see Note 20). 9. Dilute samples 1:10 using water. The final IGEPAL® CA-630 concentration should be 0.08%. 10. Protein standard dilutions should be prepared such that they contain the same amount of detergent as the TPP samples.

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Therefore, first dilute DNA digest buffer B 1:5 using water, and then mix it in a 1:1 ratio with protein standard solutions. 11. Determine the protein concentration using a DTT-compatible assay of your choice, e.g., Bradford or BCA. 12. Harmonize the protein concentration across samples. Separate for each compound condition, average the protein concentration of the samples heated to the two lowest temperatures. Based on this average, calculate the volume required to adjust the protein concentration to 1.25 mg/mL. Dilute samples accordingly with sample buffer, using the same volume of dilution buffer for all samples treated with the respective compound concentration. 13. Proceed directly with Subheading 2.6.1. Alternatively, samples can be stored at -20 °C. 3.5.3 Removal of Aggregated Proteins (Crude Lysate Format)

1. During DNA digest, label an appropriate number of ultracentrifugation tubes. Precool tubes on ice or at 4 °C. 2. Following DNA digest, centrifuge PCR plates for 1 min at 311 × g at 4 °C. 3. Remove aluminum foil, and transfer samples to labeled, precooled ultracentrifuge tubes. 4. Clear lysates by ultracentrifugation for 30 min at 100,000 × g at 4 °C (see Note 21). 5. Meanwhile, pre-wet a 96-well filter plate with 0.2 μm pore size as described in Subheading 3.5.2, step 1. 6. Carefully remove ultracentrifuge tubes from the rotor, optionally with the help of tweezers, and place them into a precooled cooling rack. Immediately continue with the next step. 7. Transfer 55 μL supernatant from the ultracentrifuge tubes to the pre-wet filter plate. Take great care not to disturb or touch any pellets when recovering cleared crude lysate from ultracentrifuge tubes. Note that pellets may not be clearly visible, so avoid touching the inner wall of the tubes as well (see Note 17). 8. Centrifuge plates at 500 × g at 4 °C for 5 min. Repeat centrifugation once if residual lysate remains in the filter plate. 9. Distribute 45 μL of 2× sample buffer to a new 96-well collection plate (see Note 19). 10. Transfer 45 μL filtered sample to the prepared sample buffer plate. 11. Incubate samples at room temperature with orbital shaking at 450 rpm for 30 min. 12. Proceed directly with Subheading 2.6.1. Alternatively, samples can be stored at -20 °C.

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3.6 Mass Spectrometry-Based Proteomics Analysis

All steps are performed at room temperature unless specified otherwise.

3.6.1 Sample Cleanup and Proteolytic Digest

The MS sample processing workflow we use is a modified version of the single-pot solid-phase-enhanced sample preparation (SP3) approach [18, 19]. Proteins are trapped on paramagnetic beads in the presence of 50% ethanol that acts as binding solvent. After removing contaminants, proteins are simultaneously alkylated, reduced, and digested using both LysC and trypsin, each at protease-to-protein ratio of 1:100. The 96-well plate format allows for parallelization and make easier/convenient facilitates wash steps as well as peptide recovery. 1. If starting with frozen samples, thaw samples for 10 min at 55 ° C to insure that SDS is completely dissolved. 2. Spin down plates for 1 min at 300 × g. 3. Mix protein cleanup solution by briefly vortexing, and distribute 40 μL per sample into a 96-well filter plate with 0.45 μm pore size. Take care not to mechanically damage the membrane while pipetting. 4. Promptly add 20 μL sample (i.e., 25 μg) to the protein cleanup solution (see Note 22). The ratio of paramagnetic beads to protein should be between 0.8:1 and 1:1 (w/w). 5. Cover the filter plate using a lid, and incubate samples for 15 min with orbital shaking at medium speed to insure complete protein binding. 6. Remove the unbound fraction by centrifuging for 1 min at 1000 × g. 7. Place filter plate onto a fresh waste collection plate. 8. To remove contaminants, wash beads four times using 200 μL of a 70% ethanol solution, and centrifuge for 2 min at 1000 × g. 9. Place the filter plate on top of a new polypropylene 96-well collection plate. 10. Add 40 μL digest solution to the bead-bound sample proteins. 11. Tightly seal the filter plate. We use pressure-sensitive polyolefin microtiter plate foil. 12. Incubate samples for at least 16 h with 500 rpm orbital shaking.

3.6.2 Peptide Labeling Using Tandem Mass Tags

1. Briefly spin down overnight digested samples to collect any condensate (1 min, 1000 × g). 2. Remove the cover, and repeat the centrifugation step to elute peptides.

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3. Add 10 μL water to the beads, and elute into the same collection plate. This step serves to insure complete peptide elution. 4. Vacuum centrifuge eluates until dry. This will take approximately about 1 h. 5. Resuspend dried peptides by adding 10 μL water and incubating the samples for 5 min with 500 rpm orbital shaking. 6. Add 10 μL of the respective TMT or TMTpro labeling reagent to reach a final concentration 4.3 μg/μL of the reagent (see Note 23). 7. Tightly seal the plate using a polyolefin microtiter plate foil, and incubate the labeling reaction for 1 h with 500 rpm orbital shaking. 8. Shortly before the end of the hour, prepare the stop solution. 9. Quench non-reacted TMT reagent by adding 5 μL stop solution per sample. 10. Tightly seal the plate using polyolefin microtiter plate foil, and incubate for 15 min with 500 rpm orbital shaking. 11. Pool samples belonging to one multiplexed experiment in a low-bind reaction tube. 12. Rinse each well using 10 μL wash solution. Combine the wash solution with the respective sample pool. 13. Vacuum centrifuge samples until dry. This will take about 3–5 h. Dried samples can be stored at -20 °C. 3.6.3 Final Peptide Purification

These final purification steps are warranted by the fact that, at this stage, samples still contain considerable amounts of contaminants incompatible with MS analysis. First, SDS is precipitated using potassium ions. The associated centrifugation simultaneously sediments any bead particles. Finally, peptides are enriched and desalted via reversed-phase solid-phase extraction. 1. Pre-warm a saturated KCl solution for 10 min in a 38 °C water bath. 2. Resolubilize dried peptide samples using 400 μL of the saturated KCl solution. Briefly mix by vortex. 3. Centrifuge samples for 5 min at 16,000 × g to precipitate any potassium dodecyl sulfate and bead particles. 4. Meanwhile, activate the SPE matrix using 100 μL activation solution. Centrifuge for 1 min at 240 × g. 5. Equilibrate the SPE matrix using 200 μL 4% TFA. Centrifuge for 1 min at 240 × g. 6. Carefully aspirate the supernatant of the KCl precipitated samples without disturbing precipitates. Load the supernatant onto the equilibrated SPE matrix. Centrifuge for 5 min at 140 × g. The reduced speed improves peptide-binding efficiency.

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7. Wash bound peptides using 100 μL 4% TFA. Centrifuge for 1 min at 240 × g. 8. Place the SPE plate onto a fresh 96-well polypropylene collection plate. Elute peptides using 200 μL activation solution. Centrifuge for 1 min at 240 × g, and collect eluate. 9. Vacuum centrifuge eluates until dry. Dried samples can be stored at -20 °C. 3.6.4 Offline Fractionation by High pH Reversed-Phase Chromatography

1. Add 30 μL of mobile phase A to the dried samples. 2. Seal the plate using polyethylene microtiter plate foil with adhesive-free zones. 3. Thoroughly dissolve samples via incubation for 15 min with orbital shaking at 500 rpm at room temperature. 4. Place the sample plate as well as new 96-well plates for fraction collection into the autosampler of the UPLC system. 5. Recommended settings for fractionation: samples are loaded completely at a flow of 50 μL/min onto the trap column. Samples are then separated on the analytical column using a gradient of 3–57% mobile phase B over 70 min followed by a step to 90% B that is held for 5 min. Separation is performed at a flow of 50 μL/min, and the separation column is heated to 35 °C. Depending on the final number, fractions are collected for 60 s (n = 24 fractions), 80 s (n = 16 fractions), or 90 s (n = 11 fractions) (see Note 24). Once the respective maximum number of fractions is reached (n), collection starts over at the first position. Specifically, fraction n + 1 is eluted into the first fraction; fraction n + 2 is eluted into the second fraction, etc. This way three to four elution steps are combined into one final fraction, resulting in similar peptide amounts across fractions as well as ensuring optimal use of instrument time for subsequent LC-MS/MS analysis. 6. Following fractionation, vacuum centrifuge samples until dry. Dried samples can be stored at -20 °C for several weeks.

3.6.5 LC-MS/MS Analysis

1. Add 10 μL of NanoLC mobile phase C to dried fractionated samples. 2. Seal the plate using polyethylene microtiter plate foil. 3. Incubate samples for 15 min with orbital shaking at 500 rpm at room temperature. 4. Place the plate on the autosampler of the online nanoLC-MS/ MS system. 5. Inject 5 μL of the first sample (i.e., 50% of the sample).

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6. Apply a linear gradient of 7–30% NanoLC mobile phase B at a flow rate of 350 nL/min over 50 min. Heat the column oven to 60 °C. 7. Different mass spectrometry equipment can be used and may be available at institutions and contract research organizations. The following is intended to provide guidance for commonly used Q Exactive Plus mass spectrometers. Acquire the survey scan with an AGC target value of 3 × 106 ions in a 375–1200 m/z range with 250 ms maximum injection time at a resolution of 70,000. Isolate the top ten most intense precursor ions, and apply an isolation window of 0.4 m/z. Fragment precursor ions by higher-energy C-trap dissociation (HCD) with normalized collision energies set to 33 eV. For the data-dependent MS/MS scans, apply an AGC target value of 2 × 105, an intensity threshold of 1.7 × 103, a loop count of 10 and a maximum injection time of 120 ms at a resolution of 35,000. Set peptide match to be preferred. Enable dynamic exclusion for 60 s. Use a fixed first mass of 100 m/z. Exclude ions having unassigned charge states as well as those with charge states of one, seven, and above. Record both full MS and fragment spectra in the profile mode. 8. Measure fractions of one multiplexed sample consecutively (see Note 24). 9. Wash the system with one wash run before loading the first fraction of the next experiment. 3.7 Peptide and Protein Identification and Quantification

1. Analyze MS data using the search engine of your choice, e.g., MASCOT 2.5 (Matrix Science). 2. (Optional) Perform a first-pass database search using a 30 ppm precursor peptide tolerance (monoisotopic mass) and a 30 mDa (HCD) fragment ion tolerance, and use the result as basis for recalibration via the “software locked mass” strategy by Cox et al. [20]. 3. Peptide and protein identification: search the data using a 10 ppm precursor peptide tolerance and 20 mDa (HCD) fragment ion tolerance. Set carbamidomethyl cysteine and TMT or TMTpro modification of lysine as fixed modifications. Set methionine oxidation, N-terminal acetylation, and TMT or TMTpro modification of peptide N-termini as variable modifications. Specify trypsin as cleavage enzyme. Allow a maximum of three missed cleavages and peptide charges of 2+ and 3+. Use a decoy database to control the false discovery rate (FDR). Filter peptide and protein identifications using appropriate false discovery rate (FDR) cutoffs. Additional filter may be applied such as a signal-to-background value of the precursor ion of >4 and a signal-to-interference >0.5. Refer to [21] for more details.

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4. Relative quantification: quantify TMT or TMTpro reporter ions as described in Savitski et al. [22]. Calculate fold changes using a sum-based bootstrap algorithm of peptides unique for identified proteins [23]. For higher quantification accuracy, correct fold changes both for isotope purity and for interference caused by co-eluting peptides as described in [21]. 3.8

Data Analysis

Use the analysis strategy best suited to the respective TPP format. Software packages for each referenced analysis procedure can be freely downloaded. All of these packages perform the necessary steps to determine proteins with altered thermal stability profiles, from data normalization and protein fold change calculations to curve fitting. A great resource for further information on available analysis strategies for classical TPP-TR, TPP-CCR, and 2D-TPP experiments is the review by Mateus et al. [7]. 1. For most robust results, only consider proteins quantified with at least two unique peptides for downstream analysis. 2. Classical TPP-TR, TPP-CCR: for classical TPP-TR experiments and isothermal dose–response experiments (TPP-CCR), use the analysis approach described in Franken et al. [9]. You will need the two software packages “isobarQuant,” written in Python, and “TPP”, written in R. 3. 2D-TPP: to detect ligand-protein interactions from 2D thermal profiles, use the R package “TPP2D” [24] (see Note 25). 4. Combined TPP-TR: for temperature range TPP experiments that combine both treatment and control samples in one TMT experiment, we recommend applying the recently introduced abundance ratio-based thermal shift assay analysis (RTSA) approach [25]. This analysis procedure enables robust and sensitive detection of both changes in thermal stability as well as protein abundance.

4

Notes 1. Small molecules are commonly solubilized using DMSO. However, DMSO is not biologically inert, and its concentration should be kept as low as possible in the cell culture medium to avoid severe cytotoxicity. A final concentration of 0.5% (v/v) DMSO is generally a good starting point. Some cell lines even tolerate up to 1% (v/v), while other cell lines as well as primary cells are much more sensitive to DMSO. We therefore recommend performing a pilot experiment to assess the DMSO sensitivity of your cell line of choice.

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2. Dissolve all protease inhibitors separately at first. Add ice-cold water to prepare stock solutions of 250 μM aprotinin, 1 mM bestatin, 10 mM leupeptin, and 5 mM phosphoramidon. Use ice-cold ethanol to prepare a stock solution of 1 mM pepstatin. Finally, mix these stock solutions at a ratio of 1:20:5:1:0.73 (aprotinin:bestatin:leupeptin:phosphoramidon:pepstatin). 3. Note that TCEP is dissolved in 1 M HEPES pH 8.0 buffer. The comparatively high buffer concentration is necessary to compensate for the very acidic nature of TCEP solutions. We use TCEP as reducing agent because it is more resistant to oxidation by air than DTT or β-mercaptoethanol. This makes TCEP particularly suitable for our one-step protocol for protein reduction, alkylation, and digest protocol overnight. 4. One 20 μg vial of trypsin and LysC each suffice to digest 80 samples according to this protocol. Use freshly reconstituted protease for optimal performance. Still, HEPES protease solutions may be stored at -20 °C for up to 4 weeks. Freezethaw aliquots no more than three times. 5. Use PEEK™ tubing for all connections, because fused silica is not resistant to high pH in the long term. 6. Protease inhibitors can introduce artifacts and should be avoided when generating crude cell extracts for TPP experiments. If this is not possible, perform pilot experiments to evaluate the extent to which they interfere with target proteins or the biological system under study. 7. We find it helpful to add about 0.5 volumes of cold DPBS while thawing cell pellets. Pellets often skew to the side walls, and gradation marks on reaction tubes can be imprecise. We therefore use pipets to determine the volume of cell pellets, and the presence of buffer facilitates the pipetting. Do remember to subtract volume of added buffer from the total volume measured to determine the correct pellet volume. 8. We set the temperature of the processing chamber to -20 °C. The temperature in the processing chamber can fluctuate drastically, especially when processing many samples or samples requiring multiple cycles. Take appropriate measures to ensure samples do not heat up to more than 15 °C, e.g., by adjusting dwell time. 9. These settings are sufficient to efficiently lyse most cell lines. Still, when generating crude lysate from a cell type for the first time, we recommend checking the resulting lysis efficiency. To this end, collect 10 μL aliquots of the sample before and after lysis. Keep aliquots on ice until further processing. Measure the number of intact cells using an automated cell counter, and then determine the degree of cell disruption. Be sure to use the measurement parameters that apply to the specific cell line.

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10. The Benzonase® nuclease digest step serves to remove DNA and, importantly, reduce the sample viscosity to facilitate liquid handling. 11. Crude lysate is highly viscous and contains supramolecular structures and cellular debris. This makes it difficult to measure protein concentrations in crude lysates. Moreover, it is the amount of soluble protein in the reference sample (i.e., vehicle-treated and subjected to lowest temperature) after the removing aggregated proteins and residual cell debris that counts. Hence, protein concentration is not measured directly in crude lysate. 12. The number of cells and thus cell culture plates or flasks required per compound concentration varies between cell types. The cell material needs to yield 25–50 μg of protein per data point when extracted using the described lysis buffer. For adherent cells, a good starting point is to use one 15 cm cell culture dish per compound concentration. At 70–90% confluency, the protein yield from one 15 cm dish of, e.g., HepG2 or HeLa cells, is sufficient for up to 12 data points. For suspension cells, the required cell number per data point may range from 3 × 106 cells (e.g., K562 cells) or 6.5 × 106 cells (e.g., THP-1 cells) to 1 × 107 cells (e.g., Jurkat cells). Before using a cell type for the first time, determine how many cells are needed to get 25–50 μg of protein when applying lysis conditions of the live TPP workflow (see Subheadings 3.4, 3.5.1, and 3.5.2). 13. The compound concentration is kept constant for all subsequent cell handling steps up to and including heat treatment. Adhere to the following order for handling samples during all steps that involve compound or vehicle: start with the lowest compound concentration, continue with the next higher concentration up to the maximum, and treat the vehicle control last. 14. The centrifugal force may need to be adjusted for certain cell types as they may be more sensitive to mechanical damage or, conversely, may be more difficult to pellet. Always make sure that the supernatant appears clear before aspirating to avoid losing cells. 15. The treatment compound may be toxic at the concentrations tested. It is therefore important to determine the number of viable cells in compound-treated samples relative the vehicletreated control sample. If cell viability levels fall below 80%, repeat the experiment using revised treatment conditions. We recommend using the CellTiter-Glo® Luminescent Cell Viability Assay, as the luminescent signal both develops rapidly and remains stable for several hours, providing flexibility and allowing for easy integration into the TPP workflow.

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16. Homogenous crude lysate aliquots can also be distributed by multi-dispenser pipette when using wide orifice tips. This method is rapid enough to make repeated mixing unnecessary. 17. We recommend using a 96-channel liquid handling system, either a manual device (e.g., Rainin Liquidator™ 96, Mettler Toledo) or an electronic platform (e.g., VIAFLO 96, Integra). In addition to increased throughput and continuous temperature control, these systems importantly enable a degree of robustness which is difficult to achieve by hand. Precise immersion depths insure supernatant transfer without disturbing pelleted material, and fixed pipetting speeds insure consistent lysis conditions. 18. To achieve efficient protein aggregation, it is crucial to incubate samples for no less than 3 min at room temperature following heat treatment. Samples can be left at room temperature for up to 10 min. 19. In our hands, the presence of 2% (v/v) SDS in the sample positively impacts the performance of the subsequent sample preparation steps. 20. Measure the concentration of samples treated at the highest temperature as temperature gradient control. For example, cells treated at 64 °C should yield a protein concentration of less than 0.5 mg/mL. 21. Preclearing crude lysate samples via ultracentrifugation prevents clogged wells during filtration later on. 22. Proteins bind to the surface of hydrophilic carboxylatemodified beads via ethanol-driven solvation capture. This process is most efficient at a solvent concentration of at least 50% (v/v). But, prolonged exposure to high ethanol concentrations may damage the filter matrix. We found incubating samples for 15 min in 50% (v/v) ethanol to be a good compromise. 23. TMT reagents are amine reactive. Efficient peptide labeling crucially depends on completely removing any aminecontaining contaminants. 24. Adjust the number of fractions per experiment that are analyzed depending on what analytical depth is required. Typically, TPP-TR and TPP-CCR experiments are fractioned into 24 samples of which either 12 or all 24 fractions are measured. A full 2D-TPP experiment consists of six multiplexed experiments, each of which constitutes a pool of samples heated to two consecutive temperatures. The protein content will therefore vary across the six multiplexed samples. As a rule of thumb, separate pools of samples treated up to ~56 °C into 24 fractions, pools of samples heated between 57 and 60 °C

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into 16 fractions, and pools of samples heated above 60 °C into 13 fractions. Then measure 13/24, 11/13, and 8/11 fractions. 25. TPP-TR experiments of vehicle-treated cells provide useful reference data sets of protein melting curves. These meltomes can help to identify true positives among putative hits from 2D-TPP experiments. Stabilization effects should be most prominent at temperatures slightly above the melting point in the reference data set. Note that this rule may not apply for shallow curves. Ligand-dependent destabilization should be most pronounced at the reference melting point. Measure meltomes as biological replicates for each cell type, applying a temperature gradient encompassing 37, 40, 44, 47, 50, 53, 56, 59, 62, and 66 °C. References 1. Hanash S (2003) Disease proteomics. Nature 422:226–232 2. Pantoliano MW, Petrella EC, Kwasnoski JD et al (2001) High-density miniaturized thermal shift assays as a general strategy for drug discovery. J Biomol Screen 6:429–440 3. Martinez Molina D, Jafari R, Ignatushchenko M et al (2013) Monitoring drug target engagement in cells and tissues using the cellular thermal shift assay. Science 341:84–87 4. Savitski MM, Reinhard FB, Franken H et al (2014) Tracking cancer drugs in living cells by thermal profiling of the proteome. Science 346:1255784 5. Mateus A, Kurzawa N, Perrin J et al (2021) Drug target identification in tissues by thermal proteome profiling. Annu Rev Pharmacol Toxicol 62:465 6. Sridharan S, Gu¨nthner I, Becher I et al (2019) Target discovery using thermal proteome profiling. In: Mass spectrometry-based chemical proteomics. Wiley, Hoboken, pp 267–291 7. Mateus A, Kurzawa N, Becher I et al (2020) Thermal proteome profiling for interrogating protein interactions. Mol Syst Biol 16:e9232 8. Dai L, Prabhu N, Yu LY et al (2019) Horizontal cell biology: monitoring global changes of protein interaction states with the proteomewide cellular thermal shift assay (CETSA). Annu Rev Biochem 88:383–408 9. Franken H, Mathieson T, Childs D et al (2015) Thermal proteome profiling for unbiased identification of direct and indirect drug targets using multiplexed quantitative mass spectrometry. Nat Protoc 10:1567–1593

10. Sridharan S, Kurzawa N, Werner T et al (2019) Proteome-wide solubility and thermal stability profiling reveals distinct regulatory roles for ATP. Nat Commun 10:1155 11. Becher I, Werner T, Doce C et al (2016) Thermal profiling reveals phenylalanine hydroxylase as an off-target of panobinostat. Nat Chem Biol 12:908–910 12. Thompson A, Wolmer N, Koncarevic S et al (2019) TMTpro: design, synthesis, and initial evaluation of a proline-based isobaric 16-plex tandem mass tag reagent set. Anal Chem 91: 15941–15950 13. Li J, Van Vranken JG, Pontano Vaites L et al (2020) TMTpro reagents: a set of isobaric labeling mass tags enables simultaneous proteome-wide measurements across 16 samples. Nat Methods 17:399–404 14. Li J, Cai Z, Bomgarden RD et al (2021) TMTpro-18plex: the expanded and complete set of TMTpro reagents for sample multiplexing. J Proteome Res 20:2964–2972 15. Zinn N, Werner T, Doce C et al (2021) Improved proteomics-based drug mechanismof-action studies using 16-plex isobaric mass tags. J Proteome Res 20:1792–1801 16. Miettinen TP, Bjorklund M (2014) NQO2 is a reactive oxygen species generating off-target for acetaminophen. Mol Pharm 11:4395–4404 17. Jafari R, Almqvist H, Axelsson H et al (2014) The cellular thermal shift assay for evaluating drug target interactions in cells. Nat Protoc 9: 2100–2122 18. Hughes CS, Moggridge S, Muller T et al (2019) Single-pot, solid-phase-enhanced

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sample preparation for proteomics experiments. Nat Protoc 14:68–85 19. Moggridge S, Sorensen PH, Morin GB et al (2018) Extending the compatibility of the SP3 paramagnetic bead processing approach for proteomics. J Proteome Res 17:1730–1740 20. Cox J, Michalski A, Mann M (2011) Software lock mass by two-dimensional minimization of peptide mass errors. J Am Soc Mass Spectrom 22:1373–1380 21. Savitski MM, Mathieson T, Zinn N et al (2013) Measuring and managing ratio compression for accurate iTRAQ/TMT quantification. J Proteome Res 12:3586–3598 22. Savitski MM, Zinn N, Faelth-Savitski M et al (2018) Multiplexed proteome dynamics

profiling reveals mechanisms controlling protein homeostasis. Cell 173:260–274.e25 23. Savitski MM, Sweetman G, Askenazi M et al (2011) Delayed fragmentation and optimized isolation width settings for improvement of protein identification and accuracy of isobaric mass tag quantification on Orbitrap-type mass spectrometers. Anal Chem 83:8959–8967 24. Kurzawa N, Becher I, Sridharan S et al (2020) A computational method for detection of ligand-binding proteins from dose range thermal proteome profiles. Nat Commun 11:5783 25. Kalxdorf M, Gunthner I, Becher I et al (2021) Cell surface thermal proteome profiling tracks perturbations and drug targets on the plasma membrane. Nat Methods 18:84–91

Chapter 6 Improved Coverage of the N-Terminome by Combining ChaFRADIC with Alternative Proteases Xuehui Jiang, Ying Lao, Victor Spicer, and Rene´ P. Zahedi Abstract Many proteolytic cleavage events cannot be covered with conventional trypsin-based N-terminomics workflows. These typically involve the derivatization of protein N-termini and Lys residues as an initial step, such that trypsin will cleave C-terminal of arginine but not lysine residues (ArgC-like cleavage). From 20,422 reviewed human protein sequences in Uniprot, 3597 have known N-terminal signal peptides. An in silico ArgC-like digestion of the corresponding 3597 mature protein sequences reveals that—even for these well-known and well-studied proteolytic events—trypsin-based N-terminomics workflows may miss up to 50% of signaling cleavage events as the corresponding neo-N-terminal peptides will have an unfavorable length of 30 (911 peptides) amino acids. In this chapter, we provide a protocol that can be applied to all kinds of samples to improve access to this “inaccessible” N-terminome, by making use of the alternative, broad-specificity protease subtilisin for fast and reproducible digestion of proteins. Key words N-terminal peptides, N-terminomics, Alternative proteases, ChaFRADIC

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Introduction The human genome encodes for nearly 600 proteases [1]. Proteases play a role in many important physiological and pathophysiological processes, such as protein maturation, degradation, and translocation, as well as cell migration, coagulation, viral infection, and tumor invasion. Although proteases have been studied for more than 100 years, the substrates and specificities of many proteases are still unknown. Mass spectrometry has greatly expanded our knowledge on protease substrates and cleavage specificity, in particular with the advent of “N-terminomics” methods. These methods enable the specific enrichment of N-terminal peptides that represent endogenous protein N-termini, including neo-N-termini that have been generated upon endogenous protease activity. N-terminomics methods use different strategies to separate, after proteolytic digestion of samples, N-terminal peptides from

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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the large majority of internal (non-N-terminal) peptides. This allows not only to identify which proteins are cleaved by a specific protease but also where these proteins are cleaved to ultimately deduce cleavage motifs. The first method that allowed such a dedicated and in-depth analysis of N-terminal peptides was combined fractional diagonal chromatography (COFRADIC) developed by Gevaert et al. in 2003 [2], while later terminal amine isotopic labeling of substrates (TAILS) was introduced by the Overall lab in 2010 [3]. We have developed a modified version of COFRADIC which we termed charge-based FRADIC (ChaFRADIC) [4]. Instead of inducing hydrophobic retention time shifts between two reversed chromatography (RP) runs in order to separate internal peptides from N-terminal ones, as done in COFRADIC, we used the same concept to induce more defined retention time shifts based on charge reduction of internal peptides between strong cation exchange chromatography (SCX) runs. We later transferred the concept from high-performance liquid chromatography (HPLC) systems to pipette tips (ChaFRAtip) [5]. The advantage of these FRADIC strategies is their versatility, as different chemistries as well as enzymatic approaches can be used to enrich for specific subsets of peptides [6–9]. A general limitation of N-terminomics workflows, however, is that many N-terminal and neo-N-terminal peptides are not wellsuited for trypsin-based LC-MS analysis. After blocking of primary amines (protein N-termini and Lys residues) and subsequent tryptic digestion, the generated peptides have ArgC-like features (i.e., no cleavage C-terminal to blocked Lys) and are often either too short (30 amino acids) to be compatible with conventional bottom-up proteomics approaches. Querying the UniProt database shows that from 20,422 reviewed human proteins, 3597 have known N-terminal signal peptides. An in silico digestion of the corresponding 3597 mature protein sequences without these signal peptides reveals that—even for these well-known and well-studied proteolytic events—trypsinbased N-terminomics workflows with ArgC-like specificity may miss up to 50% of these cleavage events because of unfavorable length of the respective neo-N-terminal peptides, i.e., generating neo-N-terminal peptides that are 30 (911 peptides) amino acids long. Here, we provide a protocol that can be applied to all kinds of samples to improve the coverage of this “inaccessible” N-terminome by making use of the alternative, broad-specificity protease subtilisin [10].

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Materials

2.1 Cell Lysis, Reduction, and Alkylation

1. Cell pellets, stored at -80 °C. 2. Lysis buffer: 2% w/v sodium dodecyl sulfate (SDS), 100 mM Tris–HCl pH 8.0. 3. Eppendorf Thermomixer. 4. Thermo Sonic Dismembrator or equivalent sonication probe. 5. Benchtop centrifuge capable of centrifuging microcentrifuge tubes up to 20,000 × g. 6. Reduction buffer: 0.5 M tris(2-carboxyethyl)phosphine (TCEP). 7. Alkylation buffer: 0.5 M iodoacetamide (IAA) in 100 mM Tris–HCl pH 8.0. Prepare fresh before use. IAA needs to be protected from light (see Note 1).

2.2 Protein Concentration Determination

1. Bicinchoninic acid assay (BCA). 2. 15 mL Falcon tubes. 3. Eppendorf Thermomixer. 4. Microplate compatible spectrophotometer.

2.3 Labeling of Primary Amines on Protein Level Using iTRAQ 8-Plex

1. 100% ethanol (EtOH), stored at -40 °C. 2. Ice cold acetone. 3. 10% SDS. 4. iTRAQ 8-plex reagents. 5. iTRAQ labeling buffer: 0.5 M triethylammonium bicarbonate in 20% isopropanol, pH 8.5. 6. Isopropanol. 7. Quenching solution: 600 mM glycine. 8. Hydroxylamine solution: 1.3 M hydroxylamine.

2.4 Proteolytic Digestion and Peptide Cleanup

1. S-TRAP Mini cartridge (Protifi LLC, Huntington NY) (see Note 2). 2. 50% phosphoric acid diluted in water. 3. S-TRAP loading buffer: 9:1 methanol:water in 100 mM Tris–HCl (pH 8.0). 4. Benchtop centrifuge. 5. Subtilisin (Sigma Aldrich). 6. Eppendorf Thermomixer. 7. Digestion buffer/S-TRAP elution buffer 1: 100 mM Tris–HCl pH 8.0, freshly prepared. 8. S-TRAP elution buffer 2: 0.2% formic acid (FA).

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9. S-TRAP elution buffer 3: 50% acetonitrile (ACN). 10. Centrifugal vacuum concentrator. 2.5

Desalting

1. Solid phase extraction materials—SOLA HRP 30 mg/3 mL cartridge. 2. Syringe or positive pressure manifold for SPE. 3. Activation buffer: 100% ACN. 4. SPE wash buffer: 0.1% FA (FA; see Note 3). 5. SPE elution buffer: 60% ACN, 0.1% FA (see Note 3).

2.6 Enrichment of NTerminal Peptides Using ChaFRAtip

1. Whatman glass microfiber filter (Sigma Aldrich). 2. 2–200 μL pipette tips. 3. SCX beads: Polysulfoethyl A, 5 μm particle size, 200 Å pore size (PolyLC Inc.). 4. SCX buffer A: 10 mM KH2PO4, 20% ACN, pH 2.7. 5. SCX buffer B: 250 mM KCl, 10 KH2PO4, 20% ACN, pH 2.7 (see Note 4). 6. Mix SCX buffers A and B to obtain: 15% B, 20% B, 30% B, 40% B. 7. NHS-trideutero acetate. 8. 10% FA. 9. Benchtop centrifuge. 10. Protein LoBind tubes. 11. C18 SPE: Oligo R3 beads. 12. C18 Empore Disc (3 M). 13. 70% ACN, 0.1% FA. 14. 20–200 μL pipette tips. 15. Gel loader tip. 16. Air-filled syringe. 17. 100% ACN. 18. 0.1% FA. 19. 60% ACN, 0.1% FA.

2.7 Nano-LC-MS/MS Analysis (See Note 5)

1. Easy nanoLC 1200 (Thermo Scientific). 2. Mobile Phase A: 0.1% FA. 3. Mobile Phase B: 84% ACN, 0.1% FA. 4. Precolumn, Acclaim™ PepMap™ 100 C18, 2 cm × 100 μm. 5. Analytical column, 25 cm × 75 μm.

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6. Q-Exactive Plus mass spectrometer (Thermo Scientific).

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1. Proteome Discoverer 3.0 (Thermo Scientific). 2. Data analysis tools.

Method

3.1 Cell Lysis, Reduction, and Alkylation

1. Resolubilize cells pellet in 200 μL of lysis buffer (see Note 6). 2. Heat samples to 85 °C in a thermomixer for 10 min, gently shaking (see Note 7). Let block cool down before removing tubes. 3. Use probe-based sonication to support the solubilization and shear DNA, e.g., using a Thermo Sonic Dismembrator at 25% amplitude for 3 cycles × 5 s (see Note 8). 4. Pellet the remaining debris by centrifugation at 4 °C and 20,000 × g for 5 min. 5. From each sample, remove a 10 μL aliquot, and dilute to a final volume of 100 μL water (tenfold dilution) such that the final concentration of SDS is 50% and protein:bead aggregates are immobilized on a magnetic rack. Harsh washing and subsequent on-bead digestion result in clean peptide samples for analysis by LC–MS

Ensure all measures for appropriate ethical votes and patient consent forms. Store all biological samples at -80 °C until the first steps. All used chemicals or pre-made solvents are prepared at LC– MS grade. Only use ultrapure LC–MS grade water. Not all materials described below are needed for every sample type and input amount. 2.1 Specimen Lysis, DNA Shearing, and Protein Extraction

1. Appropriate type and format of tubes depending on the anticipated processing method (see below): (a) 1.5 mL Protein LoBind tubes, Eppendorf. (b) 2 mL Protein LoBind tubes, Eppendorf. (c) PCR 8-stripes. (d) 8 AFA-TUBE TPX Strip, Covaris, Inc. (Woburn, USA). (e) 96-well SuperPlate, skirted. (f) 96 AFA-TUBE TPX Plate, Covaris, Inc. (g) 8 AFA-TUBE TPX Strip Caps, Covaris, Inc. 2. Tubes, stripes, or plates need to be closed/covered during most processing or incubation steps: (a) Covaris Plate holder. (b) Covaris 8-Strip holder. (c) PCR Foil Seal. (d) X-Pierce Film. 3. Phosphate buffered saline (PBS). 4. Lysis buffer A: 1% SDS, 100 mM ammonium bicarbonate (NH4HCO3), used for resuspending, diluting, or transferring samples (see Note 1): 5. A device capable of solubilizing tissue to extract proteins and properly shear DNA (see Note 2). For example, but not limited to the following (see Note 3):

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6. A benchtop centrifuge capable of holding tubes, stripes, or plates. The centrifuges indicated below with the corresponding rotors allow processing of either tubes, stripes, or plates: (a) Centrifuge 5424, Eppendorf. (b) Centrifuge 5424 Rotor F-45-32-5-PCR, Eppendorf. (c) Centrifuge Eppendorf.

5424

(d) Centrifuge 5430R Eppendorf.

Rotor Rotor

FA-45-24-11

(Tubes),

FA-45-48-11

(Tubes),

(e) Centrifuge 5430R Rotor A-2-MTP (Plates), Eppendorf. 2.2 Protein Quantification

It is recommended to quantify the amount of protein extracted from a given specimen. Use a suitable method that is compatible with the utilized lysis buffer. For example: 1. Pierce BCA protein assay. 2. Nanodrop spectral photometer. 3. Qubit protein BR assay. 4. Qubit protein assay. This does not hold for FFPE samples, as they will contain paraffin, which interferes with most quantification methods (see Note 4).

2.3 Protein Reduction and Alkylation

Denaturation, reduction, and alkylation of proteins are performed in a manual procedure of choice off-deck or as integral part of the autoSP3 protocol. The reaction may be performed with different reagents in either a single reaction step or a two-step procedure (see Note 5). The following components are used by default: 1. Lysis buffer B: 4% SDS, 360 mM NH4HCO3, 160 mM CAA, 40 mM TCEP, 1× PIC (see Notes 1, 6, and 7): 2. A thermomixer capable of holding tubes, stripes, or plates and temperatures up to 95 °C with appropriate lid heating to avoid evaporation (see Note 8).

2.4 AutoSP3 Processing and Protein Digestion

The following section describes the required hardware, solvents, reagents, and consumables for running the autoSP3 protocol performed on a Bravo liquid handling system (Agilent Technologies). The protocol can be adapted to other liquid handling systems, which in turn, may require other accessory equipment/ consumables.

2.4.1 Hardware Equipment and Accessories

The following section summarizes the required hardware when using the Bravo liquid handling system:

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1. Bravo liquid handling system. 2. Heating station. 3. 96-well ring magnet, Alpaqua Engineering (Beverly, USA, Magnum FLX) (see Note 9) 4. Orbital shaking station. 5. Bravo 96 channel disposable tip head LT. 6. Inheco Single TEC controller. 7. Peltier thermal station. 8. PCR plate insert. 9. A benchtop thermomixer that allows heating to 95 °C with lid heating to avoid evaporation (see Note 8). 2.4.2

Consumables

The following section summarizes the required consumables to run autoSP3 using the default settings on a Bravo liquid handling system: 1. 2 mL Protein LoBind tubes, Eppendorf (see Note 10). 2. Magnet for 1.5–2 mL tubes. For example, Dyna Mag 2, Life Technologies (Carlsbad, USA) (see Note 10). 3. 250 μL LT head tips boxes, Agilent Technologies. 4. SuperPlate PCR Plate, 96-well (see Note 11). 5. Reagent plate, Reservoir 6 columns, Agilent Technologies (see Note 12). 6. Waste plate, Agilent Reservoir 8 rows, Agilent Technologies (see Note 12). 7. 96-well format sealing foil (see Note 13).

2.4.3 Solvents and Reagents

The following section summarizes the required solvents and reagents to run autoSP3 using the default settings on a Bravo liquid handling system: 1. Ethanol (EtOH) (see Note 14). 2. Isopropanol (see Note 14). 3. Acetonitrile (ACN) (see Notes 14 and 15). 4. Lysis buffer B: 4% SDS, 360 mM NH4HCO3, 160 mM CAA, 40 mM TCEP, 1× PIC (see Subheading 2.3) (see Note 5–7). 5. SP3 beads (see Notes 17–19): 6. Digestion buffer: e.g.,100 mM NH4HCO3. 7. Sequencing grade-modified trypsin (see Note 20).

2.5

Peptide Recovery

It is recommended to perform the digestion of proteins in a lid-heated thermomixer. For short digestion periods, this can be done on-deck of the liquid handling system. For the acidification and recovery of peptides to a new sample plate, the 96-well plate is placed back on the deck.

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1. SuperPlate PCR Plate, 96-well, Thermo Fisher Scientific (see Note 21). 2. 250 μL LT head tips boxes, Agilent Technologies 3. Trifluoroacetic acid (TFA). 4. Formic acid (FA). 5. PCR Foil Seal (see Note 13). 6. X-Pierce Film (see Note 13). 7. A suitable -20 °C or preferably -80 °C freezer for sample storage until LC–MS measurement. 8. Pierce Quantitative Fluorometric Peptide Assay. 9. Pierce Quantitative Colorimetric Peptide Assay. 2.6 LC–MS Data Acquisition and Processing

3

The analysis of peptide samples requires a suitable LC–MS setup. Several combinations of nanoLCs (e.g., Thermo Dionex, Vanquish, nanoEasy 1200, Bruker nanoElute 2, and Evosep) and mass spectrometers (e.g., Thermo Orbitrap or Bruker timsTOF instruments) are commonly used in the proteomic field. Since all are compatible with autoSP3, they will not be described in detail here. The same is true for a wide range of search algorithms and downstream software packages for data analysis, which should be chosen dependent on the utilized acquisition mode [10]. For example, data-dependent acquisition (DDA) data can be processed with Bruker ProteoScape, Proteome Discoverer, PEAKS, MaxQuant, Mascot, or MS Fragger. Data-independent acquisition (DIA) data can be processed using Bruker ProteoScape, Spectronaut, DIA-NN, or MaxQuant (version 2 or higher). Follow the manufacturer’s instructions for data acquisition, processing, and subsequent analyses.

Methods The following section assumes that a Bravo liquid handling system with the necessary accessories is available in the laboratory (see Subheading 3.3.2). The processing of liquid samples, cells, freshfrozen tissue, and FFPE requires different parameters. Specific guidelines for the individual sample types are noted throughout the protocol (see Subheading 3.1), which holds mostly for the protein extraction and optional emulsification and paraffin removal. Not all reagents scale linearly with the protein input amount and can be limited by the geometry of the 96-well format plates. The excess amount of SP3 beads in this protocol assumes the processing of 10–20 μg protein material. For higher amounts of protein, the bead:substrate ratio may require optimization, while carefully considering the magnet strength and well geometry (see Notes 18 and 19).

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3.1 Specimen Lysis, DNA Shearing, and Protein Extraction

The protocol provides guidelines and limitations for ranges of input material. The most important parameters that need to be considered are the number of samples and the expected protein content per sample. The general aim is the extraction of proteins from a given sample material, while additionally shearing DNA. Downstream processing with autoSP3 is compatible with other lysis and protein extraction procedures as long as DNA shearing is sufficient (see Note 2). The amount of protein to be expected from a given specimen can be estimated based on rules of thumb provided in Table 1. Estimates will vary for different cell types and organs.

3.1.1 Selection of a Suitable Sample Lysis Method

The following section serves as a guideline to select a suitable method for protein extraction: 1. Based on the amount of starting material (e.g., cell pellet or tissue), select the volume of lysis buffer A (see Note 1), aiming to achieve ~1–2 μg/μL protein concentration (Table 1): 2. Select the lysis method dependent on the volume per sample and the required throughput. As a selection guideline, use the conditions low (high) volume and low (high) throughput (Fig. 2). The availability of instrumentation and equipment limits the number options. (a) Low volume (50% organic (see Note 15). This is achieved by column-wise adding 21 μL 100% acetonitrile from column 4 in the 6-column reservoir at position 5 to each well of the sample plate at position 9. The protein binding to the beads is enhanced by consecutive fast (30 s at 1200 rpm) and slow (90 s at 400 rpm) shaking in the orbital shaking accessory installed at position 9 for a total of 18 min. 4. The sample plate is transferred from position 9 to the 96-well format ring magnet at position 7 to allow trapping of beadbound proteins. The beads are forced to the inner well wall in a ring shape at a defined height to allow bead-free pipetting. The supernatant is aspirated and dispensed to the waste reservoir at deck position 2 (see Note 9). 5. The washing tasks in autoSP3 are performed from column one to three of the 6-column reservoir at position 5. The combination of wash solvents and number of cycles can be adapted by the user considering the maximum capacity of 45 mL per reservoir column. Each wash solvent 1, 2, and 3 corresponds to the 6-column reservoir columns 1, 2, and 3, respectively (see Notes 14 and 34). The default method uses 80% EtOH in two wash cycles of 200 μL in a column-wise manner. In each wash cycle, solvent is added from the 6-column reservoir at position 5 to each well of the sample plate at position 9 followed by brief agitation. The wash solvents are discarded from the 96-well ring magnet at position 7 to the waste reservoir at position 2. For efficient removal of paraffin from emulsified FFPE samples (see Note 29), it is recommended to use two wash cycles of 200 μL 80% EtOH, followed by four wash cycles with 150 μL 100% Isopropanol and heating for 10 min at 50 °C between cycles (see Note 23) (see settings for wash Solv 2 and 3 in Fig. 4a). The heat incubation steps are performed in a lid-heated PCR cycler off-deck. The autoSP3 protocol is paused by clicking the “Pause all” button in VWorks. The resulting pause prompt is used to continue autoSP3 when the sample plate is placed on-deck after the incubation. 6. The final wash cycle is performed with 180 μL of 100% acetonitrile from column 4 of the 6-column reservoir at position 5 as described for the previous washes (see Note 14). The solvent is removed and additionally air-dried before the addition of digestion buffer in the next step. 7. Digestion buffer is added to each well in the sample plate at position 9 from the 6-column reservoir column 5 at position

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5 (see Note 16). By default, autoSP3 transfers 35 μL digestion buffer per sample to immerse the bead-bound proteins for an efficient reaction. 8. Finally, autoSP3 transfers 5 μL of the protease solution in column-wise manner from column 3 of the reagent plate at position 8 to each well of the sample plate at position 9 (see Note 20). 9. At this point, the first part of autoSP3 is finished, and the sample plate is incubated automatically at 37 °C for 30 min, allowing the user to collect and seal the plate (see Note 37). Take the plate from the Bravo deck at position 4, seal it properly (see Note 13), and transfer it to a suitable lid-heated PCR device for prolonged incubation (1–16 h) at 37 °C (see Note 37). 3.4 Peptide Recovery After Digestion

After digestion, the sample plate is manually transferred back to the Bravo system at position 9, and the deck setup is updated as described in the following section (Fig. 5). The aim of the described procedure here is to quench protease activity and to transfer clean peptides to a new sample plate.

3.4.1 Peptide Recovery Deck Setup and Supplying Reagents

1. Position 3: The tip box may require a refill depending on the number of previously used tasks. Two columns of clean tips are required for peptide recovery located in the first two columns of the tip box. The tip box setup is specified in the user interface (Fig. 4c).

Fig. 5 Schematic illustration of the peptide recovery part of autoSP3, including the deck setup with the required hardware accessories, and the default reagents with their respective on-deck reservoir position. The required tips and their position are highlighted in positions 3 and 6. The acid solvent used to quench the digest reaction is supplied at position 5. The plate to retrieve clean peptides and the digest plate are placed at positions 8 and 9, respectively, just prior to starting the protocol

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2. Position 5: The 6-column deep-well reservoir is supplied with an acidic solvent in column six for peptide recovery (e.g., 5% TFA or FA) (see Note 38) (Fig. 5). The required volume is indicated by the user interface depending on the user-supplied settings (Fig. 4a, d). 3. Position 6: Replace the tip box with clean tips (Fig. 5). 4. Position 8: Place a new 96-well PCR SuperPlate AB2800 to receive the acidified peptides (Fig. 5). 5. Position 9: The sample plate holding the digested protein samples is replaced on the Bravo deck. The plate will be automatically transferred between position 9 and the magnetic rack at position 7 during the procedure (Fig. 5). 3.4.2 Running Peptide Recovery

Upon preparation of the Bravo deck hardware, the following section describes the individual steps of the autoSP3 protocol for acidification and recovery of peptides (see Note 38). Here, it is assumed that the digest was carried out at the previously described default settings for 96 samples in a total volume of 40 μL per sample. The Bravo system first requires initialization upon starting of the VWorks environment (see Note 36). 1. By default, autoSP3 transfers 5 μL of 5% TFA (or e.g., 5% FA) column-wise from column 6 at position 5 to each well of the sample plate at position 9 (see Note 38). The supernatant is automatically transferred to the new PCR 96-well plate at position 8 (see Note 21). The transfer is performed in several consecutive steps to reduce residual volume. The total transfer volume can be adapted in the user interface (Fig. 4a). 2. The 96-well plate with acidified peptides at position 8 can be removed, covered with a suitable lid for short- (-20 °C) or long-term storage (-80 °C) (see Note 13). 3. The sample plate can be directly transferred to an autosampler for LC–MS, to inject a suitable volume for proteome analysis. Alternatively, samples can be (manually) transferred to another vial system that is compatible with the used autosampler or with other downstream procedures (see Subheading 3.5) (see Note 21). 4. Discard the waste volumes in the reservoir at position 2 at the end of running autoSP3.

3.4.3 Peptide Quantification for FFPE Material

Different methods are available to determine the quantity of peptides that are present in a processed sample after autoSP3, such as a Pierce peptide quantification assay. The quantification of peptides is not required after autoSP3 except for FFPE samples in which the true amount of peptides, and the relative difference between samples is yet unknown (see Note 4). The following steps are required:

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1. Prepare a peptide standard dilution from 2 to 0.125 μg/μL as described by the instructions of the specific quantification kit that is used (see Subheading 2.5). 2. Transfer 20 μL of the peptide standard curve to a 96-well flat bottom plate in triplicates. 3. Transfer a defined fraction of each peptide sample (e.g., 5–10%, approximately matching to the peptide standard dilution to achieve the highest assay accuracy) to the 96-well flat bottom plate containing the peptide standard curve to perform the quantification assay (Table 1). 4. Store the remaining peptide sample on ice for short-term processing on the same day or at -20 °C for processing on the next day. 5. Supplement the volume of each well with ddh2O to match the peptide standard curve. 6. Prepare the working reagent (WR) according to the manufacturer’s instructions. 7. Use the formula to determine the necessary volume of WR: μL WR = ð#of standards þ #of samplesÞ  180 μL ð50 parts reagent A þ 48 parts reagent B þ 2 parts reagent CÞ þ 10% 8. Add 180 μL WR to each well containing either standard or peptide sample and incubate for 30 min at 23 °C. Subsequently, use a spectrophotometer device to read out the absorbance at wavelength 480 nM. 9. Calculate a standard curve by plotting the average peptide standard measurements against the respective μg/μL concentrations. Generate a linear trendline to calculate the unknown concentration of each peptide sample. 3.5 Compatibility with Downstream Applications

After protein cleanup and digestion by autoSP3, recovered peptide samples are ready for LC–MS without further processing. Alternatively, peptides are compatible with a wide range of downstream applications prior to LC–MS [10, 17]. For example, isobaric labeling with TMT reagents shall be used for sample multiplexing during LC–MS data acquisition (see Note 16). Therefore, the digestion buffer of autoSP3 is required to be free of amines (e.g., TEAB) that could interfere with the labeling reaction and acidification in the final step of autoSP3 is omitted. Other methods, such as high-pH peptide fractionation, can be readily applied with the default autoSP3 method. An offline HPLC system equipped with a suitable fractionation column is required. Traditionally applied C18-based sample desalting only remains necessary when a buffer is used for proteolysis that is not compatible with MS (e.g., Tris-

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based) [7, 17]. For the above and other downstream applications, refer to the manufacturer’s instructions or literature and the appropriate equipment. 3.6 Quality Control of Acquired MS Data

Peptide samples generated by autoSP3 can be analyzed by LC–MS with a data acquisition method of choice (see Subheading 2.6). The resulting peptides and proteins that are identified and quantified can be used as direct quality control of the autoSP3 processing. The following points serve as a guideline. Other metrics can be used to judge the performance: 1. The observed total ion chromatogram can indicate if the overall intensity corresponds to the amount of injected protein to reflect proper protein recovery from SP3 beads, or if hydrophilic or hydrophobic peptides are lost, potentially indicating incomplete protease digestion. Although autoSP3 efficiently removes detergents, residual contamination of detergents, salts, or PEG would be readily visible from the chromatogram. Detergent CHAPS is observed at a m/z of 615.4043, while PEG results in a Gaussian peak envelope with a characteristic m/z 44 difference. Detergent SDS causes ion suppression, while salts can precipitate and clog LC columns or emitter tips. 2. The degree of missed cleavages is a direct readout for digestion efficiency. By default, two missed cleavages are allowed in most database search algorithms. If they occur at a high frequency within identified peptide sequences, the digestion was likely incomplete. This could be due to a low protease to protein ratio, insufficient volume to cover the protein-binding beads to enable digestion, or too high residual organic solvent, or protease activity incompatible pH. 3. High charge states (>7–8) can be indicative for undigested protein. In case of FFPE samples, a high charge state distribution can indicate insufficient de-crosslinking of peptides. 4. Undigested proteins are typically visible by a Gaussian peak pattern at >1000 Th with high charge states (>7–8 z or higher). Undigested proteins accumulate over time on the analytical column leading to suppression of its retaining capacity. 5. It is recommended to establish a reference sample from which these and other metrics are known. For example, an aliquot of pelleted cells that can be run with every autoSP3 processing as a standard reference sample. This allows to immediately pinpoint whether the autoSP3 run was hampered by, e.g., supplying or accidentally omitting required solvents, buffers, and reagents.

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Notes 1. The extraction of proteins can vary between different sample types (e.g., different organs) and different protein localizations (e.g., membrane proteins). SP3 and autoSP3 are compatible with a wide range of detergents and other contaminants at a high dynamic range [7, 10], thus making it an ideal procedure for universal processing with different buffer components (e.g., UREA, CHAPS, Triton X-100, or others). In case of specific lysis buffers that are not mentioned, we recommend performing pilot experiments before running biological samples. High detergent concentrations can lead to foaming and accompanied sample loss. For example, 1–5 mg of wet weight fresh-frozen tissue can be resuspended in 100 μL lysis buffer A or in 3 times the tissue volume. On the other hand, pellets of 100.000 cells correspond to ~10 μg protein and can be resuspended in 10 μL or 3 times the pellet volume. 2. DNA shearing efficiency may be monitored by running agarose gels. 3. The solubilization of tissue and extraction of proteins can be achieved through various methods. The selection of a suitable method largely depends on the sample amount and cohort size. Here, we provide a selection of suitable equipment that are available to us for different scenarios. Other equipment may be applied if extraction efficiencies, throughput, and nucleic acid shearing are achieved sufficiently. Exemplary solutions include: (a) Bioruptor Pico, Diagenode SA (Seraing, Belgium), e.g., 30 s on/off with 15–20 cycles depending on the nucleic acid shearing (check with agarose gels). (b) Probe sonicator, Branson Digital Sonifier, Branson Ultrasonic Corporation (USA), two cycles of 10–15 s at 10% frequency depending on the nucleic acid shearing (check with agarose gels). (c) LE220R-plus Focused-ultrasonicator, Covaris, Inc. (Woburn, USA). Optimal settings are described in the Subheading 3. 4. The protein concentration in emulsified FFPE samples cannot be quantified without prior paraffin removal. This can be achieved efficiently with SP3 and autoSP3 (see Subheading 3.1.5), for which, however, it is recommended to process (approximately) equal amounts of protein. A rough estimate can be achieved by using literature references (e.g., similar tissue types and amount of material) or according to the rule of thumb that 10% of the tissue weight corresponds to protein (Table 1). The true protein concentration will depend on the

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tissue composition (e.g., tumor cell content, microenvironment, etc.). It is recommended to perform a peptide quantification assay after autoSP3 before LC–MS (see Subheading 3.4.3). 5. Reduction and alkylation within the autoSP3 protocol are optional and can also be performed manually off-deck prior to autoSP3. Alternatively, the user has the choice to perform a single-step or two-step reduction and alkylation with the desired reagents and incubation conditions. In a two-step reaction, the reducing reagent is prepared in column 1 of the reagent plate at position 8, while the alkylation reagent is prepared in column 2. The time and temperature settings can be selected in the user interface. Pilot experiments are recommended to verify efficiency of reduction and alkylation. 6. We recommend chloroacetamide instead of commonly used iodoacetamide, as the latter can lead to off-site modifications and double carbamidomethylation that mimics the same mass as a ubiquitination [23]. 7. We recommend TCEP because it allows a single-step reaction together with chloroacetamide. The pH should be monitored carefully as TCEP is acidic [9]. 8. Heating to 95 °C on the Bravo deck takes considerable amount of time (approximately 45 min). Alternatively, the protocol could be paused for off-deck incubation at higher temperatures to speed-up the protocol. Here, lid heating is an essential measure to avoid evaporation of the sample. For example, a PCR cycler with lid heating, e.g., CHB-T2-D ThermoQ, Hangzhou BIOER Technologies (Binjiang, China) or ThermoMixer C, Eppendorf (Hamburg, Germany) may be used. 9. Not every available 96-well format magnet is suitable for autoSP3. The compatibility of the magnet to autoSP3 is determined by the magnetic strength as well as the position of the magnetic field. The latter will determine the position of immobilized beads in the sample plate. For example, pillar magnets center the beads in all compass directions at the well-bottom and are thus incompatible with 96-well pipetting. In case that beads are observed in the pipette tips during discarding of the wash solvents, it is recommended to check the concentration of beads and its compatibility to the utilized magnet. Alternatively, downscaling the bead concentration should be considered. The beads are added in excess, and lower concentrations are sufficient to capture the same amount of protein (Notes 17 and 18).

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10. SP3 beads are provided in a PEG-containing solution. Therefore, the manual preparation of the bead stock should be done with caution as described by Hughes et al. [17]. Washing is an essential part of the protocol. Other beads may be used according to the manufacturer’s instructions and with considering appropriate pilot experiments. 11. Other 96-well plates that follow the American National Standards Institute (ANSI): Society for Biomolecular Screening (SBS) (https://www.slas.org/education/ansi-slas-microplatestandards/) standards for microplates can be used. This includes the Covaris 96 AFA-TUBE TPX plates, which are designed in accordance with the SBS-plate format. Thus, plates can be transferred from the Covaris instrument to the Bravo for direct autoSP3. 12. Other reservoirs following the American National Standards Institute (ANSI): Society for Biomolecular Screening (SBS) (https://www.slas.org/education/ansi-slas-microplatestandards/) standards for microplates can be used as long as the requirements for solvent and waste volumes are met. The utilized plates require specification in the labware editor of VWorks and the autoSP3 user interface (Fig. 4b). 13. Foil or sealing mats other than those indicated can work as well, but it is highly recommended to use seals without glue at the position of the wells as it may result in contaminating the injection needle or sample. It should be tested whether the utilized sealing can effectively prevent evaporation and is temperature stable at up to 95 °C or during storage at -20 °C. Suitable solutions are, e.g., PCR Foil Seal, 4titude Ltd. (Berlin, Germany) and X-Pierce Film, Sigma-Aldrich (Steinheim, Germany). 14. The wash solvents in autoSP3 can be replaced by other organic solvents. It is recommended to consider pilot experiments prior to using biological samples. Further, it is recommended to use 100% ACN in the last wash cycle, as its residual moisture may remain behind on the bead surface, will not interfere with the digestion reaction. Instead, low concentrations of organic up to 10% have the potential to promote the digestion efficiency [24]. To avoid a residual organic concentration higher than 10%, the processes are paused for a 2 min air-drying incubation. 15. Protein-bead binding is induced by implementing a >50% (v/v) organic concentration in each sample. The required volume can be specified in the user interface (Fig. 4a) and depends on the starting volume of the sample, the reduction and alkylation reagent volumes, as well as the added bead volume. The volume can be modified to reach higher than 50% (v/v) of

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organic within the maximum capacity of the sample well as the limiting factor (approximately 200 μL for a typical 96-well plate). 16. It is recommended to use LC–MS-compatible buffers to maintain the pH and corresponding protease activity during digestion to avoid the need for additional cleanup steps, such as C18 desalting. For example, low concentrations of NH4HCO3 (>25–30 mM) are LC–MS compatible, which can be achieved by diluting a sample digested in 100 mM NH4HCO3 with 0.1% formic acid (FA) before injection. It is recommended to use 100 mM NH4HCO3 to compensate for the 50 mM acetic acid used to store trypsin on the Bravo deck (see Note 20). Other buffers can be used with or because of the appropriate downstream workflows. For example, isobaric labeling of peptides with tandem mass tags (TMT) requires an amine-free digestion buffer, such as triethyl-ammonium bicarbonate (TEAB, Sigma-Aldrich, Steinheim). 17. SP3 beads or other beads might come at different stock concentrations by different distributors. Take care to use the recommended amount of 250 μg beads in 5 μL for binding up to 50 μg protein. For other bead types, the optimal concentration requires pilot experiments (e.g., [7]). In this framework Sera-Mag A (Sera-Mag Speed Beads A, Fisher Scientific, Slurry at 50 μg/μL) and Sera-Mag B (Sera-Mag Speed Beads B, Fisher Scientific, Slurry at 50 μg/μL), beads were used. 18. The amount of SP3 beads added to each sample does not scale linearly with the protein input amount. By default, an excess concentration of beads is added to each sample, which is sufficient to bind up to 50 μg protein. 19. The hardware of the Bravo accessories (e.g., the ring magnet) and its consumables set limits to the maximum amount of SP3 beads that can be used. For example, the magnetic strength should be capable of complete immobilization of the beads. If beads are left unbound (i.e., remain in suspension), clean pipetting from the mid-well positions is no longer possible. In addition, too high bead concentrations may interfere with homogeneous bead pipetting. 20. It is recommended to use trypsin stored in 50 mM acetic acid to avoid autolysis during the autoSP3 run. In a 96-well scenario, it will take ~2–2.5 h (depending on number of wash cycles) until the protease is added to each sample. Other proteases can be used and may require different storage conditions to ensure maximum efficiency during the digestion process [25]. 21. Despite the autoSP3 compatibility with the standard microplate format (ANSI:SBS) ((https://www.slas.org/education/

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ansi-slas-microplate-standards/), it is recommended to use the listed SuperPate PCR plate in case that direct transfer to a nanoLCs autosampler is anticipated. 22. Covaris, Inc. offers a range of different sonication instruments including the LE220Rsc as the successors of the LE220R+. Both and other Covaris instrumentation are available to deliver highly focused AFA energetics to the samples in either single vial, stripe, or plate format. More details can be obtained from the manufacturer. 23. The processing of FFPE requires deparaffinization and de-crosslinking. This can be achieved by protocols that are considered routine practice in pathology [22], e.g., comprising xylol- and ethanol washes or alternatively, using the AFA sonication procedure developed by Covaris, Inc. in combination with autoSP3. In the latter case, paraffin is removed by autoSP3, which requires additional wash cycles. Specifically, 4 additional cycles of 150 μL 100% isopropanol with intermediate 10 min at 50 °C (off-deck) are needed in a lid-heated device (see Subheading 3.3.4). 24. AutoSP3 can process extracted proteins from any sample type whenever the buffer is compatible (see Note 1) [18]. We recommend using autoSP3 preferably for low protein amounts (3 mg), it is recommended to use cryo-pulverization to homogenize each individual sample. Aliquots of the powder can be processed as described for 1 mg wet tissue (see Subheading 3.1.4). 29. The FFPE material is emulsified and consequently turned into an opaque white solution from the paraffin. The latter is subsequently removed from each sample during autoSP3, producing a transparent peptide solution. 30. The Pierce BCA protein quantification assay is compatible with a range of buffer components as detailed in the manufacturer’s instructions. Other lysis buffers may be used (see Note 1) considering their compatibility to the protein quantification. 31. Importing the autoSP3 default profile might overwrite existing profiles of other protocols. It is recommended to generate backups of other protocols prior to starting the import wizard. 32. The Bravo liquid handling robot offers a range of hardware accessories and compatibility to consumable or inserts. For each protocol and custom deck setup, all deck positions require appropriate teach points, defining the exact positioning (X and Y directions) and height (Z direction) of a tip, to reach the hardware. The teach points are defined in the VWorks device diagnostics at the Jog/Teach table A single tip is moved manually to the mark at each position to save the exact position. 33. The columns for each used reagent in the reagent plate at position 8 are specified in the user interface (Fig. 4e). Thus, allowing to adjust the protocol for a single- or two-step reduction and alkylation. It further enables the re-use of the same 96-well plate for multiple consecutive autoSP3 runs to reduce the consumable waste. The autoSP3 sanity checks will not allow multiple selections of the same column in a single run, although a corresponding process might be deactivated. 34. The number of wash cycles and the volume per cycle can be modified. Up to 6 cycles per column of the 6-column reservoir at position 5 are possible. The reservoir capacity and the actual number of samples (columns) at position 9 limit the number (max. 6) of wash cycles with a defined volume. The autoSP3 sanity check will prompt an error message if the user input is not possible. 35. Parameters in autoSP3 that may require optimization include increased starting volume, reduction of speed for orbital shaking for sample volumes >150 μL, or dispensing heights into the sample plate at position 9. 36. Every time that the VWorks environment is started or has been idle for extended periods, the Bravo requires initialization to confirm the ability to move accurately. Upon loading of the

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autoSP3 protocol, the user will be asked to initialize. For later initialization, the user must enter the device diagnostics profile tab. 37. The incubation time for protein digestion depends on the used protease and the protease to substrate ratio. The incubation times typically range from 1 to ~16 h overnight digestion. 38. The acidification in autoSP3 stops the digestion process. Omitting the quenching can lead to highly abundant peptide species originating from autolysis of the protease. This can lead to suppression of other peptide features. Other acids can be used. The supplied stock concentration of acidic solvent and the anticipated final concentration per sample are specified in the user interface (Fig. 4a).

Acknowledgments This work was supported by the German Ministry of Education and Research (BMBF), as part of the National Research Node “Mass spectrometry in Systems Medicine” (MSCoreSys), under grant agreement 161L0212A. Conflict of Interest M.A.C. is an employer of Agilent Technologies

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6. Messner CB, Demichev V, Wendisch D et al (2020) Ultra-high-throughput clinical proteomics reveals classifiers of COVID-19 infection. Cell Syst 11:11–24.e4. https://doi.org/10. 1016/j.cels.2020.05.012 7. Hughes CS, Moggridge S, Mu¨ller T et al (2019) Single-pot, solid-phase-enhanced sample preparation for proteomics experiments. Nat Protoc 14:68–85. https://doi.org/10. 1038/s41596-018-0082-x 8. Wis´niewski JR (2018) Filter-aided sample preparation for proteome analysis. Methods Mol Biol 1841:3–10. https://doi.org/10. 1007/978-1-4939-8695-8_1 9. Doellinger J, Schneider A, Hoeller M, Lasch P (2020) Sample preparation by easy extraction and digestion (SPEED) – a universal, rapid, and detergent-free protocol for proteomics based on acid extraction. Mol Cell Proteomics 19:209–222. https://doi.org/10.1074/mcp. TIR119.001616 10. Mu¨ller T, Kalxdorf M, Longuespe´e R et al (2020) Automated sample preparation with

Robust Processing of Large Sample Cohorts for Proteome Profiling SP3 for low-input clinical proteomics. Mol Syst Biol 16:e9111. https://doi.org/10.15252/ msb.20199111 11. Leutert M, Rodrı´guez-Mias RA, Fukuda NK, Ville´n J (2019) R2-P2 rapid-robotic phosphoproteomics enables multidimensional cell signaling studies. Mol Syst Biol 15:e9021. https://doi.org/10.15252/msb.20199021 12. Zhu Y, Weiss T, Zhang Q et al (2019) Highthroughput proteomic analysis of FFPE tissue samples facilitates tumor stratification. Mol Oncol 13:2305–2328. https://doi.org/10. 1002/1878-0261.12570 13. Buczak K, Ori A, Kirkpatrick JM et al (2018) Spatial tissue proteomics quantifies inter- and intratumor heterogeneity in hepatocellular carcinoma (HCC). Mol Cell Proteomics 17:810– 825. https://doi.org/10.1074/mcp.RA117. 000189 14. Marchione DM, Ilieva I, Devins K et al (2020) HYPERsol: high-quality data from archival FFPE tissue for clinical proteomics. J Proteome Res 19:973–983. https://doi.org/10.1021/ acs.jproteome.9b00686 15. Kuljanin M, Dieters-Castator DZ, Hess DA et al (2017) Comparison of sample preparation techniques for large-scale proteomics. Proteomics 17. https://doi.org/10.1002/pmic. 201600337 16. Feist P, Hummon AB (2015) Proteomic challenges: sample preparation techniques for microgram-quantity protein analysis from biological samples. Int J Mol Sci 16:3537– 3 5 6 3 . h t t p s : // d o i . o r g / 1 0 . 3 3 9 0 / ijms16023537 17. Hughes CS, Sorensen PH, Morin GB (2019) A standardized and reproducible proteomics protocol for bottom-up quantitative analysis of protein samples using SP3 and mass spectrometry. Methods Mol Biol 1959:65–87. https:// doi.org/10.1007/978-1-4939-9164-8_5

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Chapter 12 Proteomics-Based Analysis and Diagnosis of Formalin-Fixed Paraffin-Embedded Amyloidosis Samples Delphi Van Haver, Ame´lie Dendooven, and Francis Impens Abstract Amyloidosis is a group of rare pathologies characterized by abnormal folding and deposition of susceptible proteins in tissues and organs. Diagnosis of amyloidosis often relies on immunohistochemistry of formalinfixed paraffin-embedded (FFPE) patient samples; however, dependency on antibodies for protein staining is one of the major pitfalls of this approach, especially for the detection of rare amyloidosis types. In recent years, mass spectrometry-based proteomics has emerged as a promising alternative for adequate detection and amyloid typing, despite the fact that preparing FFPE samples for proteomics remains a challenging task. Major hurdles are removal of formalin-induced protein cross-links and water-insoluble paraffin prior to mass spectrometry analysis. With the recent development of the suspension trapping protocol, enabling the use of high concentrations of SDS, these obstacles can be overcome. In this chapter, we describe the implementation of suspension trapping for FFPE sample processing and its application to analyze human amyloidosis samples, comparing a standard procedure with probe sonication with a more advanced workflow based on ultrasonication. Key words Clinical proteomics, FFPE , HYPERsol, S-Trap, Mass spectrometry

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Introduction Formalin fixation in combination with paraffin embedding (FFPE) has been the go-to method to preserve clinical tissue samples for decades [1]. During the FFPE process, tissues are fixed with formalin—a solution of formaldehyde in water—to halt all biological processes and prevent autolysis of cells [2]. Formaldehyde is a reactive electrophilic species that induces cross-links between biomolecules in two steps, as shown in Fig. 1. When applied to proteins, first primary amines (on protein N-termini and Lys) and thiol groups (on Cys) are altered to a methylol modification, which converts to a Schiff base when dehydration occurs. Next, the Schiff base can react with primary amides (on Gln and Asn), guanidine groups (on Arg, guanine base), thiols (on Cys), imidazole (on His), indole (on Trp), or Tyr ring carbons

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 Protein cross-linking by formaldehyde. During formalin fixation protein cross-linking occurs in two steps. (a) First, primary amines (on protein N-termini and Lys residues) are altered to a methylol modification, which converts to a Schiff base when dehydration occurs. (b) Next, the Schiff base can react with another nucleophilic group (Nuc, e.g., amides on Gln and Asn residues), resulting in two cross-linked proteins. (Created with BioRender.com)

to form methylene bridges [2–4]. After the cross-linking step, water is removed from the tissues with ascending ethanol concentrations, xylene is used to remove ethanol, and replaced by molten paraffin. Finally, the paraffinized tissue is embedded in a paraffin block, giving it the ability to be stored safely at room temperature for long periods of time [1, 5–7]. Due to the contained costs and applicability to FFPE samples, (immuno)histochemistry (IHC) staining of FFPE sections is still the most widely used technique in histopathology for diagnosis, prognosis, and biomarker identification [7]. With immunohistochemistry, specific antigens are detected by interaction with monospecific antibodies that are linked to horseradish peroxidase or alkaline phosphatase. When a chromophore is added, oxidation occurs and development of a red or brown color can be detected by light microscopy [8]. Although IHC has been the most popular method for pathological diagnostics since the 1980s, some issues arose: the demanding lab procedure requires expensive monospecific antibodies, preservation is not equal for all antigens, and antigen detection is not possible when antibodies are not available [7–9]. The latter complicates the diagnosis of amyloidosis, pathologies characterized by abnormal folding of susceptible proteins. The abnormal folding leads to a conformational change with formation of an insoluble beta-pleated sheet secondary structure. Extracellular deposition and accumulation of these beta-pleated amyloid proteins result in tissue and organ dysfunction. There are specific histologic characteristics shared by the different forms of amyloidosis, for example, positivity on a histochemical Congo red stain with (green) birefringence, positivity on thioflavin immunofluorescence, or an ultrastructure made of 7–12 nm thin microfibrils on

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electron microscopy [10]. However, the cause and the nature of the precursor protein differ between patients, with around 30 different types of amyloid described. Determination of protein type is mandatory for implementation of specific therapy. Although common forms of amyloid make up for more than 90% of cases [11], it is difficult to type and detect rare forms due to uncommon precursor proteins. Immunohistochemical stains for these rare forms are often not available in routine practice. Additionally, sensitivity and specificity of immunohistochemistry or immunofluorescence, when available, are imperfect [12]. With current mass spectrometers allowing detection of analytes in the attomolar range (10-18) [13], mass spectrometry (MS)-based proteome analysis can be superior to IHC as a diagnostic tool. Moreover, MS has proven very helpful for adequate detection and typing of amyloid [12–14] as it is highly specific and it allows detection of previously unsuspected amyloid types because there is no reliance on a preselected antibody panel [10]. Small deposits with a low abundance of amyloid may, however, remain difficult to pick-up. Additionally, preanalytical factors need to be taken care of [14, 15]. In standard bottom-up proteomics, extracted proteins are enzymatically digested into peptides typically using trypsin, followed by liquid-chromatography tandem mass spectrometry (LC– MS/MS) analysis of the resulting peptide mixture. Searching the recorded peptide mass spectral data against protein sequence databases using specialized software algorithms then allows confident identification and quantification of the peptides and their parent proteins [16]. Despite the fact that FFPE preservation of tissues has been customary for decades, development of a standard protocol for MS-based proteome analysis of FFPE samples has not been straightforward. The water-insoluble paraffin used to embed samples and formaldehyde-induced cross-links between proteins and other biomolecules are the major problems when performing standard protein extraction protocols preceding proteomics analysis [9, 17]. Some successful protein extraction methods, using heat to cleave methylene bridges [5, 18, 19], have been developed, but often involve a long deparaffinization process with harmful solvents (heptane or xylene) and several washes with decreasing ethanol concentration. After this laborious process, proteins are extracted and dissolved in aqueous buffers containing chaotropes such as urea or guanidine hydrochloride or detergents like sodium dodecyl sulfate (SDS) or Triton X-100 [5, 9, 17, 20]. Although lysis buffers containing anionic detergents like SDS often result in the best lysis and solubilization of proteins [21], detergents are known to have a negative effect on enzymatic digestion and LC–MS/MS analysis, even at concentrations as low as 0.01% [22, 23]. Hence, detergent removal before enzymatic digestion is essential to perform successful proteomics experiments [24, 25]. Several methods for elimination of SDS have been described including protein precipitation

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and gel-based or filter-aided sample preparation (FASP), but often these resulted in poor peptide recovery or proved to be time consuming [26, 27]. The development of the suspension trapping (S-Trap) method has been a game-changer for bottom-up proteomics sample preparation [26]. S-Trap enables removal of up to 15% SDS before protein digestion and LC–MS/MS analysis. In brief, proteins are extracted from the sample with a buffer containing at least 5% SDS and cysteine residues are reduced and alkylated to completely unfold and solubilize proteins. Then, phosphoric acid is added to lower the pH to 1, and the mixture is combined with methanol at neutral pH, resulting in the formation of a protein particulate suspension. With self-made or commercially available S-Trap columns, proteins are then captured on a quartz filter with a C18 plug, while methanol-soluble SDS molecules and other contaminants are washed away with a methanol-containing buffer. Next, proteins are digested on the filter with a protease such as trypsin. Finally, hydrophilic peptides are eluted from the column with triethylammonium bicarbonate (TEAB) and formic acid (FA), while hydrophobic peptides, trapped by the carbon-18 chains, are recovered from the S-Trap column with a buffer containing acetonitrile (ACN). The eluted peptides are pooled, dried, and reconstituted in the appropriate buffer to obtain MS-ready samples [26–28]. In 2020, Marchione et al. [29] introduced the HYPERsol (high-yield protein extraction and recovery by direct solubilization) and DPS (direct solubilization in SDS with probe sonication) protocols, new methods applying S-Trap to efficiently process FFPE tissue samples for proteomics. Both methods start with incubation of diced FFPE cores in 5% SDS lysis buffer at 50 °C to soften the paraffin wax. Because of its amphiphilic structure, SDS does not only facilitate solubilization of proteins but also of the hydrophobic paraffin [21]. Then, homogenization of the samples is achieved either with a micropestle and passing of the sample through small-gauge needles in combination with probe sonication (DPS) or with a single adaptive-focused acoustics (AFA) ultrasonication step (HYPERsol). After cleavage of the heat-labile methylene crosslinks at 80 °C, sonication is repeated and cysteine disulfide bridges are broken by reduction and alkylation, resulting in fully dissolved proteins. The protein concentration of the centrifuged sample can be determined with a bicinchoninic acid (BCA) assay and up to 100 μg protein in 5% SDS is processed with the S-Trap columns as described above. The HYPERsol and the DPS protocols were shown to outperform the traditional, lengthy xylene/ethanol method, and improve proteome coverage when processing FFPE samples. Additionally, when comparing results of FFPE samples processed by the HYPERsol or DPS protocol with results from flash frozen tissue samples prepared by the S-Trap procedure,

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similar protein identifications and quantifications could be achieved [29]. To enable clinical diagnosis of FFPE amyloidosis samples, we recently implemented the DPS and HYPERsol protocols. Even though HYPERsol was shown to yield the best results, ultrasonicators are expensive devices that are not available in every lab. Therefore, we describe here our implementation of the DPS and HYPERsol protocol comparing laborious but low-cost probe sonication with simple but cost-intensive ultrasonication. A visual representation of the Probe (~DPS) and PIXUL (~HYPERsol) protocols is shown in Fig. 2. Our implementation generally follows the published DPS and HYPERsol protocols, except that samples are processed in 96-well format using S-Trap plates and a PIXUL Multi-Sample sonicator along with extra methanol/chloroform washes of the proteins on the S-Trap columns to ensure efficient removal of all wax before trypsin digestion. With these refinements, we have established a method that is easy to apply and to scale up to process large sample quantities. To illustrate the performance of both protocols, we here applied the Probe and PIXUL methods to four equal biopsy punches of the same FFPE myocardial tissue sample from a patient with senile systemic transthyretin (TTR) amyloidosis. IHC on slides from the same sample resulted in identification of TTR as the accumulated protein thus leading to the diagnosis of TTR amyloidosis as shown in Fig. 3. In our comparison, we examined the amount of extracted proteins, number of peptide, and protein identifications as well as the presence of several protein modifications indicated in Table 1. We were able to extract at least 50 μg per sample, largely sufficient for LC–MS/MS analysis. Strikingly, with the PIXUL sonicator, we could extract on average about seven times more protein from the same amount of FFPE starting material. Nevertheless, after injecting a similar amount of peptides for LC–MS/MS analysis, more than 2800 peptides and 600 proteins could be identified in each sample, with even slightly more peptides and proteins identified in the samples processed with the Probe protocol. Minor differences in protein modifications were also observed between the two protocols, most notably a higher amount of oxidized methionine residues observed with the Probe protocol. In Table 2, we listed the top ten most abundant proteins identified in each sample as determined with intensity-based absolute quantification (iBAQ) that corrects the raw protein intensity for theoretical detectability [30]. In all samples, TTR was identified as most abundant protein, confirming the diagnosis of senile systemic TTR amyloidosis established by IHC. Thus, we successfully implemented both DPS and HYPERsol protocols to detect amyloid proteins and to determine the type of amyloidosis in FFPE tissue samples.

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Fig. 2 Schematic outline of the Probe and PIXUL protocols. (a) Formalin-fixed paraffin-embedded (FFPE) biopsy cores, in an Eppendorf tube or PIXUL plate, were submerged with a buffer containing 5% sodium dodecyl sulfate (SDS) and heated to 50 °C. (b) With mechanical friction (b-1), sonication, and incubation at 80 °C (b-1/ 2), protein cross-links are broken. (c) Cysteine residues are reduced with dithiothreitol (DTT) and alkylated with iodoacetamide (IAA). Phosphoric acid (PA) and methanol (MeOH) are added, and the protein extract is loaded on an S-trap plate. After several washes with MeOH and chloroform (CHCl3), trypsin is added to digest proteins. Triethylammonium bicarbonate (TEAB), formic acid (FA), and acetonitrile (ACN) are used for elution of peptides, which are ready to analyze with liquid-chromatography tandem mass spectrometry (LC–MS/MS). (Created with BioRender.com)

2 Materials All solutions should be prepared using ultrapure water (with a maximum resistance of 18.2 MΩ-cm at 25 °C, purified from deionized water) and high-quality LC–MS grade reagents. Stock solutions should be stored at room temperature (RT) unless stated otherwise. When disposing waste material, meticulously follow waste disposal regulations. 2.1 Protein Extraction and Solubilization from FFPE Samples

1. FFPE tissue blocks, stored at RT. 2. Disposable 3 mm biopsy punch. 3. Sterile scalpel. 4. Micropestle. 5. 18-gauge needle 6. 21-gauge needle. 7. 1 mL Luer syringes.

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Fig. 3 Light microscopy images of formalin-fixed paraffin-embedded myocardial tissue from a patient with senile systemic transthyretin amyloidosis. Histology characteristics of the disease are shown. (a) Hematoxylin Eosin stain of myocardium showing pink amorphous material between the cardiomyocytes. (b) Congo red stain showing orange pink staining of the amorphous material. (c) Immunohistochemistry for transthyretin proves the amorphous Congo red-positive material as transthyretin amyloidosis. (Created with BioRender. com) Table 1 List of most important result parameters used to compare the Probe and PIXUL protocols Probe

PIXUL

Replicate 1

Replicate 2

Replicate 1

Replicate 2

Amount of extracted proteins (μg/100 μL)

122.4

62.7

799.8

619.0

Number of identified protein groups

803

701

630

663

Number of identified peptides

3407

3052

2896

2902

% Carbamidomethyl (C)

93.03

92.08

93.05

90.34

% Acetylation (protein N-term)

1.64

1.77

1.07

1.09

% Oxidation (M)

33.04

27.42

16.00

18.85

% Methylol (K)

3.21

4.86

2.95

3.89

% Methyl (K)

13.79

14.02

15.73

16.56

% Methyl (R)

3.19

3.45

2.06

3.66

Table 2 List of the ten most abundant proteins identified in each sample Protein ID

Gene name

iBAQ

Relative abundance (%)

PIXUL P02766 Transthyretin replicate 1 P68032 Actin, alpha cardiac muscle 1 P17661 Desmin P06576 ATP synthase subunit beta, mitochondrial P02511 Alpha-crystallin B chain P08590 Myosin light chain 3 P12883 Myosin-7 P25705 ATP synthase subunit alpha, mitochondrial P09493 Tropomyosin alpha-1 chain P02743 Serum amyloid P-component

TTR ACTC1 DES ATP5B

837,140,000 260,950,000 175,010,000 138,430,000

20 6 4 3

CRYAB MYL3 MYH7 ATP5A1

135,520,000 120,190,000 98,109,000 91,070,000

3 3 2 2

TPM1 APCS

91,020,000 89,735,000

2 2

P02766 Transthyretin PIXUL replicate 2 P68032 Actin, alpha cardiac muscle 1 P17661 Desmin P68871 Hemoglobin subunit beta P02511 Alpha-crystallin B chain P06576 ATP synthase subunit beta, mitochondrial P02743 Serum amyloid P-component P08590 Myosin light chain 3 P69905 Hemoglobin subunit alpha P25705 ATP synthase subunit alpha, mitochondrial

TTR ACTC1 DES HBB CRYAB ATP5B

1,545,100,000 313,350,000 245,030,000 238,400,000 193,100,000 193,010,000

24 5 4 4 3 3

APCS MYL3 HBA1 ATP5A1

158,890,000 147,770,000 146,200,000 128,360,000

2 2 2 2

Probe P02766 Transthyretin replicate 1 P68032 Actin, alpha cardiac muscle 1 P06576 ATP synthase subunit beta, mitochondrial P25705 ATP synthase subunit alpha, mitochondrial P02511 Alpha-crystallin B chain P00761 Trypsin P17661 Desmin O14558 Heat shock protein beta-6 P08590 Myosin light chain 3 P68871 Hemoglobin subunit beta

TTR 835,640,000 ACTC1 333,940,000 ATP5B 250,760,000

16 6 5

ATP5A1 211,640,000

4

CRYAB 164,100,000 150,670,000 DES 130,310,000 HSPB6 122,080,000 MYL3 119,680,000 HBB 91,247,000

3 3 2 2 2 2

P02766 Transthyretin Probe replicate 2 P68133 Actin, alpha skeletal muscle P06576 ATP synthase subunit beta, mitochondrial P00761 Trypsin P25705 ATP synthase subunit alpha, mitochondrial P08590 Myosin light chain 3 P02511 Alpha-crystallin B chain P02743 Serum amyloid P-component O14558 Heat shock protein beta-6 P26678 Cardiac phospholamban

TTR ACTA1 ATP5B

18 7 4

Sample

Protein name

447,220,000 182,770,000 109,790,000

108,200,000 ATP5A1 87,477,000

4 4

MYL3 CRYAB APCS HSPB6 PLN

3 3 2 2 2

86,189,000 82,912,000 59,033,000 53,903,000 52,276,000

The relative abundance was determined based on the intensity-based absolute quantification (iBAQ) value reported by MaxQuant

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8. Lysis buffer: 5% SDS, 50 mM TEAB in water, pH 8.5 (see Note 1). 9. PIXUL 96-well plate with sealer (Active Motif). 10. PIXUL coupling liquid (Active Motif). 11. Polypropylene 96-well plate or Eppendorf tubes. 12. Probe sonicator, equipped with microtip (e.g., VCX130 with a 3 mm probe, Sonics and Materials). 13. PIXUL sonicator (Active Motif). 14. Benchtop centrifuge with adaptors for Eppendorf tubes and 96-well plates. 15. Incubator. 2.2 Protein Concentration Determination and Reduction/Alkylation of Proteins

1. BCA assay kit (Thermo Scientific). 2. Bovine serum albumin standards (BSA, VWR) at concentrations 0, 0.125, 0.25, 0.5, 0.75, 1, 1.5 and 2 mg/mL in lysis buffer (see Note 2). 3. Clear UV-star 96-well plate (Greiner Bio-one). 4. Clear 96-well plate seal. 5. Lysis buffer: 5% SDS, 50 mM TEAB in water, pH 8.5. 6. 0.5 M dithiothreitol (DTT) in water. Store at -20 °C (see Note 3). 7. 0.5 M iodoacetamide (IAA) in water (see Note 4). 8. Microplate reader. 9. Incubator.

2.3 S-Trap, Digestion of Proteins and Peptide Purification

1. 96-well S-Trap plate (Protifi). 2. Polypropylene 96-well plates, 1.2 mL (Agilent). 3. 12% phosphoric acid in water (see Note 5). 4. Binding buffer: 100 mM TEAB in 90% methanol, pH 7.55 (see Note 6). 5. Wash buffer: 50% chloroform, 50% methanol (see Note 7). 6. Digestion buffer: 8 ng/μL sequencing grade modified trypsin (Promega) in 50 mM TEAB (see Note 8). 7. Elution buffer 1: 50 mM TEAB in water (see Note 9). 8. Elution buffer 2: 0.2% formic acid (FA) in water (see Note 10). 9. Elution buffer 3: 0.2% FA in water/acetonitrile (ACN, 50:50, v/v) (see Note 11). 10. Polypropylene HPLC vial. 11. Benchtop centrifuge with adaptors for deep well plates (see Note 12).

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12. Incubator. 13. Vacuum concentrator with adaptors for HPLC vials. 2.4 LC–MS/MS Analysis

1. Lunatic chips (Unchained labs). 2. Loading solvent A: 0.1% trifluoroacetic acid (TFA) in water/ ACN (98:2, v/v). 3. MS solvent A: 0.1% FA in water. 4. MS solvent B: 0.1% FA in water/ACN (2:8, v/v). 5. 20 mm trapping column (made in-house, 100 μm internal diameter (ID), 5 μm beads, C18 Reprosil-HD, Dr. Maisch). 6. 200 cm μPAC column with C18-endcapped functionality (PharmaFluidics). 7. Fused silica PicoTip emitter (10 μm ID) (New Objective). 8. Lunatic UV-Vis spectrophotometer (Unchained labs). 9. LC-MS/MS system, for example, Ultimate 3000 RSLC nanoLC in line connected with an Orbitrap Elite mass spectrometer (Thermo Fisher Scientific) equipped with a pneuNimbus dual ion source (Phoenix S&T) (see Note 13).

3

Methods

3.1 Protein Extraction and Solubilization from FFPE Samples

3.1.1 Protein Extraction from FFPE Samples Using a Probe Sonicator

As a result of the FFPE process, proteins are cross-linked and samples are embedded in water-insoluble paraffin. To extract and dissolve proteins in an aqueous buffer, samples are homogenized by mechanical friction, heating, and sonication in a buffer containing SDS. Additionally, DNA present in the samples is sheared during the sonication step as full-length DNA can clog S-Trap columns. In this section, we describe two protein extraction methods using different sonication equipment. 1. Select the area of interest in the tissue block and obtain FFPE cores with a Biopsy punch (see Note 14). 2. Trim away excess paraffin with a sterile scalpel and weigh the trimmed sample in a weighing dish (see Note 15). 3. Dice sample in small pieces and transfer to a 1.5 mL Eppendorf tube (see Note 16). 4. Add 20× volume/weight lysis buffer to the sample pieces. 5. Incubate the sample overnight at 50 °C (see Note 17). 6. Homogenize the FFPE pieces with a micropestle. 7. Pass the sample 10× through an 18-gauge needle connected to a 1 mL syringe (see Note 18). 8. Repeat step 7 with a 21-gauge needle.

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9. Sonicate the sample with a probe sonicator for 3 × 30 s pulses, 20% power, and 50% duty ratio (see Note 19). 10. Place the sample in an incubator at 80 °C for 1 h. 11. Repeat steps 9 and 10. 12. Centrifuge the homogenized lysate at 16,000 × g for 15 min. 13. Transfer the soluble fraction to a new 96-well plate or Eppendorf tube. 3.1.2 Protein Extraction from FFPE Samples Using a PIXUL Ultrasonicator

1. Select the area of interest in the tissue block and obtain FFPE cores with a Biopsy punch (see Note 14). 2. Trim away excess paraffin with a sterile scalpel and weigh the trimmed sample in a weighing dish (see Note 15). 3. Dice sample in small pieces and transfer to a PIXUL 96-well plate. 4. Resuspend sample pieces in 10–20× volume/weight lysis buffer. The maximum volume is 100 μL (see Note 20). Seal the plate well. 5. Incubate the sample overnight at 50 °C (see Note 17). 6. Check if the plate is still sealed completely. If necessary, fix seal better or replace the seal. 7. Ultrasonicate the sample with a PIXUL sonicator for 5 min or until pieces have been dissolved completely with the following settings: 50 N pulse, 1 kHz, 20 Hz burst rate (see Note 21). 8. Spin down shortly (see Note 22) and place the sample in an incubator at 80 °C for 1 h. 9. Repeat ultrasonication of step 7 for 6 min. 10. Centrifuge 96-well plate containing the homogenized lysate at 16,000 × g for 15 min (see Note 22). 11. Transfer the soluble fraction to a new 96-well plate or Eppendorf tube.

3.2 Protein Concentration Determination and Reduction/Alkylation of Proteins

To determine the protein concentration in samples containing high amounts of SDS, we use the detergent-compatible BCA assay. In this assay, the reduction of Cu2+ to Cu+ by proteins in an alkaline environment causes the chromophore bicinchoninic acid to change color resulting in a measurable difference in absorbance. As the color formation is directly related to the amount of Cu2+ that is reduced to Cu+, reducing agents interfere with correct absorbance measurement at 562 nm [31, 32]. Therefore, protein concentration measurement should be performed before reduction and alkylation of the protein extract.

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1. Determine the protein concentration of the soluble fraction with a BCA assay, using a BSA standard curve and microplate reader. 2. Calculate the amount of working reagent (WR) necessary for the protein concentration measurement of the unknown samples using the following equation: Total volume WR required = ð#standards þ #unknownsÞ × replicates × 200 μL: 3. Mix 50 parts of BCA reagent A with 1 part of BCA reagent B to get the appropriate amount of WR. 4. Pipette 10 μL of each standard (see Note 23) and sample (see Note 24) in duplicate into a clear UV-star 96-well plate. 5. Add 200 μL of the WR to all wells (see Note 25) containing standard or sample and cover with a plate seal. 6. Mix the contents of the plate using a plate shaker for 30 s. 7. Incubate the plate in a nonshaking incubator at 37 °C for 30 min (see Note 26). 8. Allow the plate to cool down to RT and measure the absorbance at or near 562 nM (see Note 27) with a plate reader. 9. Calculate the average absorbance measurement of the blank standards and subtract this average from all other absorbance measurements. 10. Per standard, average the corrected absorbance measurements and plot the averaged absorbance measurements of the standard vs. the concentrations in μg/mL. 11. Fit a curve to the plotted standard measurements and calculate the protein concentration of all samples using the equation of the curve and the averaged sample measurements. 12. Isolate 100 μg protein extract (see Note 28) and adjust volume to 100 μL with lysis buffer (see Note 29). 13. Reduce the protein extract by adding 3 μL 0.5 M DTT for a final concentration (f.c.) of 15 mM and incubate in a shaking incubator for 30 min at 55 °C. 14. Cool the sample to RT by putting it on ice. 15. Pipette 1 μL of a sample on a pH strip to check that the pH is approximately 8. If the pH is too low, add TEAB to increase the pH. 16. Add 6 μL 0.5 M IAA (30 mM f.c.) and incubate in a shaking incubator in the dark for 15 min at RT (see Note 30).

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225

After completely unfolding all proteins with SDS, DTT, and IAA, these reagents need to be removed to allow efficient digestion of proteins into peptides for LC–MS/MS analysis. In the following part, we describe an adapted version of the S-Trap 96-well plate protocol [33]. 1. Add 10 μL 12% aqueous phosphoric acid to the reduced and alkylated lysate (1.2% f.c.) and mix well. 2. Check the pH of a sample by spotting 1 μL on a pH strip and add more 12% phosphoric acid if the pH is higher than 1. 3. Add 700 μL binding buffer to the acidified lysate and mix well to form the colloidal protein particulate (see Note 31). 4. Place the S-Trap plate on top of a 96-well plate with a well volume of at least 1 mL (see Note 32). 5. Transfer the colloidal suspension to the S-Trap plate in parts of 400 μL, seal the plate well and centrifuge for 2 min at 1500 × g or until all solution has passed through (see Note 33). Discard the flow-through. 6. Wash the S-Trap plate containing the protein particulates once with 200 μL binding buffer and centrifuge for 2 min at 1500 × g. Visually check that all buffer has passed through. The flowthrough should be discarded. 7. Repeat step 6 three times with wash buffer (see Note 34) and twice with binding buffer to remove all wax. Ensure that all buffer has passed through at each wash step. Discard the flowthrough. 8. Transfer the S-Trap plate to a new 96-well plate with a well volume of at least 1 mL (see Note 32). 9. Pipette 125 μL digestion buffer containing 1 μg trypsin (see Note 35) (trypsin/protein ratio of 1/100 (w/w)) directly on top of the protein capture matrix. Ensure the protease solution covers the complete matrix (see Note 36). 10. Cover the S-Trap plate loosely with the plastic lid supplied with the S-Trap plate and incubate overnight at 37 °C in a nonshaking incubator (see Note 37). 11. Elute the peptides by adding 80 μL elution buffer 1 to each well of the S-Trap plate, seal the plate well, and centrifuge for 2 min at 1500 × g. Do not centrifuge before adding elution buffer 1. 12. Repeat elution step 11 once with 80 μL elution buffer 2 and once with 80 μL elution buffer 3. 13. Transfer the combined eluates to a HPLC vial and dry with a vacuum concentrator.

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3.4 LC–MS/MS Analysis

In theory, any LC–MS/MS system can be used to analyze FFPE samples prepared with this protocol. In order to inject similar amounts of each sample on the LC–MS/MS system, we advise to measure the peptide concentration with a microfluidic UV-Vis Lunatic spectrophotometer [34]. Below we provide a description of our setup and how the LC–MS/MS analysis of the amyloidosis samples is performed in our laboratory. 1. Dissolve peptides in 20 μL loading solvent A, cap the HPLC vial, vortex vigorously, and spin down shortly to gather all solution at the bottom of the HPLC vial. 2. Pipette 2 μL of loading solvent A and each sample on the Lunatic chip (see Note 38). 3. Insert the chip in the Lunatic spectrophotometer and measure the peptide concentration of the sample. 4. Inject the volume corresponding to 1.5 μg peptide material on the LC–MS/MS system (see Note 39). To analyze the samples described here we used an Ultimate 3000 RSLC nanoLC in-line connected to an Orbitrap Elite mass spectrometer equipped with a pneu-Nimbus dual ion source. Trapping of the peptide material was performed at 10 μL/min for 4 min in loading solvent A on a 20 mm trapping column and the sample was loaded on a 200 cm μPAC column mounted in the Ultimate 3000s column oven at 50 °C. For proper ionization, a fused silica PicoTip emitter (10 μm ID) was connected to the μPAC outlet union and a grounded connection was provided to this union. Peptides were eluted by a nonlinear increase from 1% to 55% MS solvent B over 80 min, first at a flow rate of 750 nL/min, then at 300 nL/min, followed by a 10 min wash reaching 99% MS solvent B and re-equilibration with MS solvent A. The mass spectrometer was operated in data dependent, positive ionization mode, automatically switching between MS and MS/MS acquisition for the 20 most abundant peaks in a given MS spectrum. The source voltage was 3.3 kV, and the capillary temperature was set at 275 °C. In the Orbitrap Elite, full scan MS spectra were acquired in the Orbitrap (m/z 300–2000, AGC target 3 × 106 ions, and maximum ion injection time 100 ms) with a resolution of 60,000 (at 400 m/z). The 20 most intense ions fulfilling predefined selection criteria (AGC target 5 × 103 ions, maximum ion injection time 20 ms, and spectrum data type: centroid, exclusion of unassigned and 1+ charged precursors, dynamic exclusion time 20 s) were then isolated in the linear ion trap and fragmented in the high-pressure cell of the ion trap. The CID collision energy was set to 35 V and the polydimethylcyclosiloxane background ion at 445.120028 Da was used for internal calibration (lock mass). QCloud was used to control instrument longitudinal performance during the project [35].

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3.5

Data Analysis

227

Different search engines and data analysis packages can be used to analyze the LC–MS/MS spectral data. The data analysis of the LC– MS/MS runs reported here was performed with MaxQuant [36] (version 1.6.17.0) using the Andromeda search engine with default search settings including a false discovery rate set at 1% on peptide and protein level. The mass tolerance for precursor and fragment ions is set to 4.5 ppm and 0.5 Da, respectively, during the main search. A minimum ratio count of two unique or razor peptides is required for quantification. LC–MS/MS runs of the four amyloidosis samples were searched separately and the specific search settings are listed below. 1. After loading the raw file and setting the experiment name, go to the group-specific parameters tab, select the “specific” digestion mode, choose “Trypsin/P” (see Note 40) as the enzyme, and set the maximum number of missed cleavages at 2 (see Notes 41 and 42). 2. Set the following modifications as variable: Oxidation(M), Acetyl(Protein N-term), Methyl(KR), Methylol(K). Set Carbamidomethyl(C) as a fixed modification (see Note 43). 3. In the global parameters tab, add the correct database(s) in FASTA file format. We used the human Uniprot Reference Proteome (database release version of June 2020, downloaded from http://www.uniprot.org) containing 20,621 protein sequences. 4. Check the iBAQ box in the label-free quantification section and uncheck the “Use .NET Core” box in the advanced section of the global parameters tab. 5. Set the number of threads to a number that does not exceed the number of logical cores available on the used computer and start the database search. 6. When the database search is finished, load the proteingroups. txt, peptides.txt and evidence.txt output files in Excel. 7. Remove all reverse database hits in all files, filter out protein groups with a Q-value above 0.01 and peptides with a PEP score above 0.01. The filtered matrices are ready to use for determining number of identified proteins, peptides, and the percentage of peptides that are modified.

4

Notes 1. For 100 mL lysis buffer, weigh 5 g SDS in a weighing dish, and transfer to a 100 mL graduated cylinder containing a magnetic stir bar. Add 5 mL 1 M TEAB solution (pH 8.5) and add water

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Table 3 Preparation of diluted BSA standards Volume of BSA stock solution Tube (μL)

Volume of lysis buffer (μL)

Final BSA concentration (mg/mL)

1

2000

0

2

2

1500

500

1.5

3

1000

1000

1

4

750

1250

0.75

5

500

1500

0.5

6

250

1750

0.25

7

125

1875

0.125

8

0

2000

0 (Blank)

to a total volume of 100 mL. Mix on a magnetic stirrer until all SDS is dissolved. To avoid foam production, limit the stirring speed when mixing the lysis buffer. The lysis buffer can be stored at RT up to 1 year. Never store the lysis buffer at 4 °C, as SDS precipitates at this temperature. If precipitation of SDS would occur during sample preparation, dissolve the precipitate by mixing and heating at 37 °C. 2. Weigh 20 mg BSA (stored at 4 °C) in a weighing dish and transfer to a conical 15 mL tube. Add lysis buffer up to 10 mL and vortex to dissolve all BSA. Dilute the 2 mg/mL stock solution with lysis buffer in 2 mL Eppendorf tubes using the volumes shown in Table 3. Vortex well, transfer 50 μL of each standard to PCR tube strips, and use one PCR tube strip for each experiment. Store the residual aliquots at -20 °C. 3. Weigh 77 mg DTT in an Eppendorf tube and dissolve in 1 mL water. Transfer 50 μL aliquots to new (PCR) tubes and store at -20 °C for up to 1 year. 4. Weigh 92 mg IAA in an Eppendorf tube in a fume hood, wrap with tinfoil and keep at RT. Only add 1 mL water immediately before use. Always prepare fresh and protect from light. 5. Add 141 μL 85% phosphoric acid stock solution to 859 μL water to obtain a 12% solution that can be stored up to 1 year at RT when kept sealed. 6. Due to the high percentage of organic solvent, adjusting the pH is not possible after mixing TEAB and methanol. To circumvent this problem, first adjust the pH of 1 M TEAB with phosphoric acid to pH 7.55. Freeze 10 mL aliquots at -20 °C for future use. For 100 mL binding buffer, add 10 mL 1 M TEAB pH 7.55 to 90 mL methanol in the fume hood. Store up to 1 year at 4 °C.

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7. Add 50 mL methanol to 50 mL chloroform in the fume hood. 8. Dissolve 20 μg lyophilized trypsin in 2.5 mL 50 mM TEAB under the fume hood for 8 ng/μL final concentration. Store aliquots at -20 °C. 9. For 100 mL, dilute 5 mL 1 M TEAB to 95 mL water. Store up to 1 year at 4 °C. 10. For 100 mL 0.2% FA, add 200 μL FA to 99.8 mL water under the fume hood. Store up to 1 year at 4 °C. 11. For 100 mL 50% ACN, 0.2% FA, mix 50 mL ACN, 200 μL FA and 49.8 mL water in the fume hood. Store up to 1 year at 4 °C. 12. Make sure the centrifuge is equipped with adaptors for deep well plates with a height of at least 50 mm. If the maximum height is lower than 50 mm, the S-Trap combined with a deep well plate will get stuck and buffers will not flow through correctly. 13. Any modern mass spectrometer and LC-MS/MS method will do to analyze the purified peptides. 14. To remove the FFPE core from the biopsy punch, use a long, thin object (e.g., inoculation needle) to push the core out. Alternatively, microtome-cut sections of FFPE blocks can also be used as starting material. 15. We advise to use maximum 10 mg of trimmed material and to use only part of the sample if this limit is exceeded. Use clean tweezers to hold the FFPE core still during trimming. 16. It is essential to use 1.5 mL Eppendorf tubes and not 2 mL tubes to ensure good homogenization with a micropestle. 17. The original HYPERsol protocol describes rehydration for only 10 min at 50 °C. We changed this to an overnight step as, in our experience, samples are too rigid for homogenization with a micropestle after 10 min. 18. Try to push out as much sample as possible from the needle and syringe. Use a micropipette to recover leftover droplets. If the volume is less than 200 μL, add more lysis buffer before starting probe sonication. Be careful when using needles, for your own safety do not recap needles after use and dispose them in the right waste container. 19. During sonication, continuously check that the probe is submerged sufficiently to prevent excessive foaming and ensure good circulation of the sample. Do not touch the plastic tube while sonicating to avoid introduction of polymers in your sample.

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20. If bigger samples need to be processed, cut the selected FFPE material in 10 mg pieces and divide over several wells. 21. In our experience, some samples completely dissolve after 5 min, others need up to 30 min. 22. After sonication, some coupling liquid might stick to the bottom of the PIXUL 96-well Plate. To avoid pollution of the centrifuge, remove this liquid before centrifugation. Tapping the plate on a tissue and removing the liquid with a vacuum aspiration pump work best. 23. To use diluted BSA standards that were stored at -20 °C, thaw a PCR tube strip from the freezer, vortex well and spin down before use. Use a multichannel pipette to dispense 10 μL of each standard to the 96-well plate. 24. When the protein concentration in samples exceeds 2 mg/mL, the samples should be diluted before performing the BCA assay. To get an idea about the protein concentration, prepare dilutions of one sample (e.g., 5× and 10× dilution) and add 1 μL of the undiluted and diluted sample to separate Eppendorf tubes containing 20 μL WR. Also prepare 2 tubes containing 20 μL WR and 1 μL blank or 1 μL 2 mg/mL BSA standard. Incubate all tubes at 37 °C for 15 min and compare the color difference. The dilution with a color between the blank (green) and 2 mg/mL BSA standard (purple) should be used for actual protein concentration measurement. With this approach repetitive protein concentration measurements of several diluted samples and excessive use of reagents can be avoided. 25. Transfer the WR to a multichannel pipette reservoir and use a multichannel pipette to easily dispense WR to all wells as fast as possible. 26. When the protein concentration is low, it is possible that no color difference can be observed after 30 min. In this case, increase the incubation time up to 2 h. 27. If the plate reader is not able to measure the absorbance at 562 nm, wavelengths between 540 and 590 nm can be used. 28. If the protein concentration is too low, increase the adjusted volume to continue with 100 μg and adjust all volumes in the following steps for the correct concentrations and ratios. More protein can be processed but do not continue with more than 300 μg protein as this is the maximum amount that can be processed with the S-Trap 96-well plate. 29. Freeze the leftover at -20 °C. The protocol can be paused at this point by storing the 100 μg aliquots at -20 °C. When continuing the protocol at a later point, vortex the sample when thawing to ensure all SDS is re-dissolved. Spin down shortly to gather solution.

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30. Efficient alkylation of sulfhydryl groups by IAA requires pH 8 and a dark environment. Wrapping the plate or rack with Eppendorf tubes in tinfoil works fine. Longer incubation times must be avoided to prevent over-alkylation of other chemical groups [37]. 31. Do not centrifuge samples at this point! 32. Push the S-Trap plate on the receiver plate until you hear a click to yield a stable setup. 33. Transfer the complete sample, including any insoluble parts, to the S-Trap columns. Pipetting up and down may be necessary to enable good transfer. As an alternative to centrifugation, positive (e.g., with a TECAN Resolvex A200) or negative pressure (via a vacuum manifold) can be applied to the S-Trap plate. 34. As chloroform is highly toxic, perform all steps involving the wash buffer in the fume hood. 35. Do not use less than 1 μg trypsin as this might result in incomplete digestion. Other proteases (e.g., Tryp-N, endoproteinase Lys-C, endoproteinase Arg-C,) can be used to digest proteins to peptides, but might need different digestion buffer and temperature. MS-compatible detergents like Rapigest can also be used to aid digestion. 36. Do not damage the matrix with the pipette tip when transferring protease solution to the S-Trap columns. 37. Do not seal the S-Trap plate with an adhesive seal as pressure will build up and this might cause the solution to flow through prematurely. 38. Use reverse pipetting to avoid introduction of air in the microfluidic channels. Air in the channels might impair correct peptide concentration measurement. 39. The injection amount depends on the LC-MS/MS system and columns used. Some systems require less than 0.5 μg (e.g., Evosep One); others require more than 1 μg for optimal results. 40. The ‘Trypsin/P’ setting allows detection of peptides resulting from cleavage after Arg or Lys, also if these amino acids are followed by a Pro. 41. Change this setting according to the protease that was used for digestion of the proteins. The number of missed cleavages can be increased to 3 or more, but this will expand the search space and significantly lengthen the database search time.

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42. When several raw files need to be searched with the same parameters, load the mqpar.xml file that was created with the first search and change the raw file and experiment name before starting the next database search. 43. To evaluate if reduction and alkylation of cysteine residues is complete, the carbamidomethyl(C) modification can be set as a variable modification.

Acknowledgments F.I. and D.V.H. acknowledge support from VIB. F.I. is supported by Ghent University Concerted Research Action grant BOF21/ GOA/033. References 1. Bronsert P, Weißer J, Biniossek ML et al (2014) Impact of routinely employed procedures for tissue processing on the proteomic analysis of formalin-fixed paraffin-embedded tissue. Proteom Clin Appl 8:796–804. https://doi.org/ 10.1002/prca.201300082 2. Fox CH, Johnson FB, Whiting J, Roller PP (1985) Formaldehyde fixation. J Histochem Cytochem 33:845–853. https://doi.org/10. 1177/33.8.3894502 3. Feldman MY (1973) Reactions of nucleic acids and nucleoproteins with formaldehyde. Prog Nucleic Acid Res Mol Biol 13:1–49 4. Metz B, Kersten GFA, Hoogerhout P et al (2004) Identification of formaldehydeinduced modifications in proteins. J Biol Chem 279:6235–6243. https://doi.org/10. 1074/jbc.M310752200 5. Gustafsson OJR, Arentz G, Hoffmann P (2015) Proteomic developments in the analysis of formalin-fixed tissue. Biochim Biophys Acta Proteins Proteom 1854:559–580. https://doi. org/10.1016/j.bbapap.2014.10.003 6. Kokkat TJ, Patel MS, McGarvey D et al (2013) Archived formalin-fixed paraffin-embedded (FFPE) blocks: a valuable underexploited resource for extraction of DNA, RNA, and protein. Biopreserv Biobank 11:101–106. https://doi.org/10.1089/bio.2012.0052 7. Grillo F, Bruzzone M, Pigozzi S et al (2017) Immunohistochemistry on old archival paraffin blocks: is there an expiry date? J Clin Pathol 70: 9 8 8 – 9 9 3 . h t t p s : // d o i . o r g / 1 0 . 1 1 3 6 / jclinpath-2017-204387

8. Schacht V, Kern JS (2015) Basics of Immunohistochemistry. J Investig Dermatol 135:1–4. https://doi.org/10.1038/jid.2014.541 9. Crockett DK, Lin Z, Vaughn CP et al (2005) Identification of proteins from formalin-fixed paraffin-embedded cells by LC-MS/MS. Lab Investig 85:1405–1415. https://doi.org/10. 1038/labinvest.3700343 10. Picken MM (2021) Diagnosis of amyloid beyond Congo red. Curr Opin Nephrol Hypertens 30:303–309. https://doi.org/10. 1097/MNH.0000000000000695 11. Machleder H (1988) Vaso-occlusive disorders of the upper extremity. Curr Probl Surg 25:1– 67. https://doi.org/10.1016/0011-3840 (88)90030-5 12. Gonzalez Suarez ML, Zhang P, Nasr SH et al (2019) The sensitivity and specificity of the routine kidney biopsy immunofluorescence panel are inferior to diagnosing renal immunoglobulin-derived amyloidosis by mass spectrometry. Kidney Int 96:1005–1009. https://doi.org/10.1016/j.kint.2019.05.027 13. Overview of Mass Spectrometry – BE. www. thermofisher.com/uk/en/home/life-sci ence/protein-biology/protein-biologylearning-center/protein-biology-resourcelibrary/pierce-protein-methods/overviewmass-spectrometry.html. Accessed 23 Nov 2021 14. Canetti D, Rendell NB, Gilbertson JA et al (2020) Diagnostic amyloid proteomics: experience of the UK National Amyloidosis Centre. Clin Chem Lab Med 58:948–957. https://doi. org/10.1515/cclm-2019-1007

Proteome Analysis of FFPE Amyloidosis Samples 15. Lavatelli F, Mazzini G, Ricagno S et al (2020) Mass spectrometry characterization of light chain fragmentation sites in cardiac AL amyloidosis: insights into the timing of proteolysis. J Biol Chem 295:16572–16584. https://doi. org/10.1074/jbc.RA120.013461 16. Aebersold R, Mann M (2016) Massspectrometric exploration of proteome structure and function. Nature 537:347–355. https://doi.org/10.1038/nature19949 17. Addis MF, Tanca A, Pagnozzi D et al (2009) Generation of high-quality protein extracts from formalin-fixed, paraffin-embedded tissues. Proteomics 9:3815–3823. https://doi. org/10.1002/pmic.200800971 18. Shi S-R, Cote RJ, Taylor CR (1997) Antigen retrieval immunohistochemistry: past, present, and future. J Histochem Cytochem 45:327– 3 4 3 . h t t p s : // d o i . o r g / 1 0 . 1 1 7 7 / 002215549704500301 19. Yamashita S (2007) Heat-induced antigen retrieval: mechanisms and application to histochemistry. Prog Histochem Cytochem 41: 141–200. https://doi.org/10.1016/j.proghi. 2006.09.001 20. Palmer-Toy DE, Krastins B, Sarracino DA et al (2005) Efficient method for the proteomic analysis of fixed and embedded tissues. J Proteome Res 4:2404–2411. https://doi.org/10. 1021/pr050208p 21. Reynolds JA, Tanford C (1970) The gross conformation of protein-sodium dodecyl sulfate complexes. J Biol Chem 245:5161–5165. https://doi.org/10.1016/S0021-9258(18) 62831-5 22. Loo RRO, Dales N, Andrews PC (1994) Surfactant effects on protein structure examined by electrospray ionization mass spectrometry. Protein Sci 3:1975–1983. https://doi.org/10. 1002/pro.5560031109 23. Elinger D, Gabashvili A, Levin Y (2019) Suspension trapping (S-Trap) is compatible with typical protein extraction buffers and detergents for bottom-up proteomics. J Proteome Res 18:1441–1445. https://doi.org/10. 1021/acs.jproteome.8b00891 24. Bereman MS, Egertson JD, MacCoss MJ (2011) Comparison between procedures using SDS for shotgun proteomic analyses of complex samples. Proteomics 11:2931–2935. https://doi.org/10.1002/pmic.201100045 25. Wis´niewski JR, Zougman A, Nagaraj N, Mann M (2009) Universal sample preparation method for proteome analysis. Nat Methods 6:359–362. https://doi.org/10.1038/ nmeth.1322

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Chapter 13 A Robust and Clinically Applicable Sample Preparation Protocol for Urinary Extracellular Vesicle Isolation Suitable for Mass Spectrometry-Based Proteomics Leyla A. Erozenci, Irene V. Bijnsdorp, Sander R. Piersma, and Connie R. Jimenez Abstract Urinary extracellular vesicles (uEVs) are a rich source of noninvasive protein biomarkers. However, for translation to clinical applications, an easy-to-use uEV isolation protocol is needed that is compatible with proteomics. Here, we provide a detailed description of a quick and clinical applicable uEV isolation protocol. We focus on the isolation procedure and subsequent in-depth proteome characterization using LC–MS/MS-based proteomics. As an example, we show how differential analyses can be performed using urine samples obtained from prostate cancer patients, compared to urine from controls. Key words Urine, Extracellular vesicles, Isolation of urinary extracellular vesicles, Clinically applicable, Proteomics, Mass spectrometry

1

Introduction Urine is frequently used in clinical diagnostics. Urine is advantageous over blood because it can easily be collected at multiple times a day in large volumes and, importantly, in a noninvasive manner [1]. Urine contains a large number of extracellular vesicles (EVs) [2]. EVs are small vesicles that are secreted by most, if not all, cells of the body and can be present in biofluids [3]. The content of EVs is in partial overlap with that of the cell of origin, and protein changes once the cells become diseased are frequently represented within the secreted EVs [4]. Recent technological advances in mass spectrometry (MS)-based proteomics have allowed for in-depth analysis of proteins in clinical samples, including urinary EVs (uEVs). The uEV proteome has been shown to be stable both at a short- [5, 6] and long-time [7] period within and between individuals, demonstrating its suitability for biomarker applications.

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Therefore, proteomics measurements of the uEVs cargo resulted in the discovery of several potential uEV-based biomarkers for several diseases [8]. While most studies in the literature focus on the diseases of proximal organs to urine (i.e., organs of the urogenital tract such as bladder, kidney, and prostate), recent studies reveal that protein signatures of distant diseases can also be detected in uEVs [6, 9–11]. To date, several methods to isolate and study uEVs proteomes have been developed [8]. However, their translation to the clinic remained limited mainly by the lack of an easy-to-use uEV isolation protocol. While ultracentrifugation is currently still the gold standard method to purify uEVs, it can process a limited number of samples (6 per run) in a relatively long time period (~6 h) using highly specialized equipment that is not necessarily available, especially in clinical laboratories. Recently, we and others employed the VN96-peptide [12], which isolates EVs in urine by complexing the heat shock proteins located on the outer surface of the EVs, allowing for high-throughput processing (>50 samples/day) and fast recovery of uEV pellets (~2 h), compatible with downstream single-shot MS-based proteomics [13, 14]. Using this method, high-depth protein identification (>2000 proteins) was previously achieved in uEVs using both data-dependent-acquisition (DDA) [14, 15] and data-independent-acquisition (DIA)-based MS [7, 16]. Here, we describe a detailed sample preparation protocol for the investigation of uEV proteomes using VN96-peptide, a simple, robust, and clinically applicable method to isolate uEVs (Fig. 1). Furthermore, we detail our downstream MS-based proteomics workflow, either in DDA- or DIA-mode (Fig. 2). Additionally, we show a proof-of-concept differential analyses of uEV proteome profiles of patients that are diagnosed with prostate cancer, compared to controls (Fig. 3).

2 2.1

Materials Urine Collection

1. Urine collection cups. 2. 15 mL or 50 mL tubes (depending on the volume of urine to be used). 3. Refrigerated centrifuge.

2.2 Urine Concentration

1. 100 kDa cutoff filters (15 mL). 2. Milli-Q water (MQ). 3. Refrigerated centrifuge.

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Fig. 1 Schematic representation of urinary EV isolation and MS-based proteomics workflow. Urine pre-cleared by centrifugation at 500 × g and 2000 × g is subsequently concentrated to 1 mL using 100 kDa molecular weight (MW) cutoff filters. Extracellular vesicles (EVs) are pelleted from 1 mL concentrated urine by capture with VN96-peptide [12] after 1 h incubation on rotating wheel. After centrifugation, the EV pellet is separately recovered from the soluble secretome via centrifugation. Subsequently, the EV pellet can be lysed in sample buffer to obtain protein extracts. Sample preparation for mass spectrometry (MS) can be performed via in-gel digestion (*described in this chapter, see Fig. 2 for schematic workflow), or any other suitable workflow that allows to remove the VN96-peptide-complex prior to peptide extraction. Label-free MS quantification can be performed in data-dependent- (DDA) or data-independent-acquisition (DIA) mode, followed by protein identification, quantification and comprehensive data analyses 2.3 Urinary Extracellular Vesicle Isolation

1. 1.5 mL or 2 mL Eppendorf tubes. 2. Phosphate-buffered saline (PBS). 3. Protease Inhibitor Cocktail (PIC) solution: Dissolve 1 tablet of PIC in 525 μL PBS (see Note 1).

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Fig. 2 Schematic representation of the in-gel-digestion (IGD) workflow of urinary EVs. A urinary extracellular vesicle (EV) pellet isolated using VN96-peptide [12] is lysed in sample buffer. Subsequently, urinary EV proteins are fractionated on SDS-PAGE and the EV-VN96-complex (located at the bottom of the gel; 1HZH.pdb). 2. Create the alignment file HERCAB_1hzh.ali for the Modeller: >P1;1hzh structureX:1HZH.pdb:1:H:+1331:M:::-1.00:-1.00 QVQLVQSGAEVKKPGASVKVSCQASGYRFSNFVIHWVRQAPGQRFEWMGWINPYNGNKEF SAKFQDRVTFTADTSANTAYMELRSLRSADTAVYYCARVGPYSWDDSPQDNYY-MDVWGK GTTVIVSSASTKGPSVFPLAPSSKSTSGGTAALGCLVKDYFPEPVTVSWNSGALTSGVHT FPAVLQSSGLYSLSSVVTVPSSSLGTQTYICNVNHKPSNTKVDKKAEPKSCDKTHTCPPC PAPELLGGPSVFLFPPKPKDTLMISRTPEVTCVVVDVSHEDPEVKFNWYVDGVEVHNAKT KPREEQYNSTYRVVSVLTVLHQDWLNGKEYKCKVSNKALPAPIEKTISKAKGQPREPQVY

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Zuzana Kalaninova´ et al. TLPPSRDELTKNQVSLTCLVKGFYPSDIAVEWESNGQPENNYKTTPPVLDSDGSFFLYSK LTVDKSRWQQGNVFSCSVMHEALHNHYTQKSLSLSPGK/ QVQLVQSGAEVKKPGASVKVSCQASGYRFSNFVIHWVRQAPGQRFEWMGWINPYNGNKEF SAKFQDRVTFTADTSANTAYMELRSLRSADTAVYYCARVGPYSWDDSPQDNYY-MDVWGK GTTVIVSSASTKGPSVFPLAPS-------TAALGCLVKDYFPEPVTVSWNSGALTSGVHT FPAVLQSSGLYSLSSVVTVPSSSLGTQTYICNVNHKPSNTKVDKKAEPKSCDK---CPPC PAPELLGGPSVFLFPPKPKDTLMISRTPEVTCVVVDVSHEDPEVKFNWYVDGVEVHNAKT KPREEQYNSTYRVVSVLTVLHQDWLNGKEYKCKVSNKALPAPIEKTISKAKGQPREPQVY TLPPSRDELTKNQVSLTCLVKGFYPSDIAVEWESNGQPENNYKTTPPVLDSDGSFFLYSK LTVDKSRWQQGNVFSCSVMHEALHNHYTQKSLSLS---/ EIVLTQSPGTLSLSPGERATFSCRSSHSIRSRRVAWYQHKPGQAPRLVIHGVSNRASGIS DRFSGSGSGTDFTLTITRVEPEDFALYYCQVYGASSYTFGQGTKLERKRTVAAPSVFIFP PSDEQLKSGTASVVCLLNNFYPREAKVQWKVDNALQSGNSQESVTEQDSKDSTYSLSSTL TLSKADYEKHKVYACEVTHQGLRSPVTKSFNRGEC/ EIVLTQSPGTLSLSPGERATFSCRSSHSIRSRRVAWYQHKPGQAPRLVIHGVSNRASGIS DRFSGSGSGTDFTLTITRVEPEDFALYYCQVYGASSYTFGQGTKLERKRTVAAPSVFIFP PSDEQLKSGTASVVCLLNNFYPREAKVQWKVDNALQSGNSQESVTEQDSKDSTYSLSSTL TLSKADYEKHKVYACEVTHQGLRSPVTKSFNRGEC* >P1;HERCAB sequence:HERCAB:1: :: ::: 0.00: 0.00 EVQLVESGGGLVQPGGSLRLSCAASGFNIKDTYIHWVRQAPGKGLEWVARIYPTNGYTRY ADSVKGRFTISADTSKNTAYLQMNSLRAEDTAVYYCSRWG--------GDGFYAMDYWGQ GTLVTVSSASTKGPSVFPLAPSSKSTSGGTAALGCLVKDYFPEPVTVSWNSGALTSGVHT FPAVLQSSGLYSLSSVVTVPSSSLGTQTYICNVNHKPSNTKVDKKVEPKSCDKTHTCPPC PAPELLGGPSVFLFPPKPKDTLMISRTPEVTCVVVDVSHEDPEVKFNWYVDGVEVHNAKT KPREEQYNSTYRVVSVLTVLHQDWLNGKEYKCKVSNKALPAPIEKTISKAKGQPREPQVY TLPPSREEMTKNQVSLTCLVKGFYPSDIAVEWESNGQPENNYKTTPPVLDSDGSFFLYSK LTVDKSRWQQGNVFSCSVMHEALHNHYTQKSLSLSPGK/ EVQLVESGGGLVQPGGSLRLSCAASGFNIKDTYIHWVRQAPGKGLEWVARIYPTNGYTRY ADSVKGRFTISADTSKNTAYLQMNSLRAEDTAVYYCSRWG------–GDGFYAMDYWGQ GTLVTVSSASTKGPSVFPLAPSSKSTSGGTAALGCLVKDYFPEPVTVSWNSGALTSGVHT FPAVLQSSGLYSLSSVVTVPSSSLGTQTYICNVNHKPSNTKVDKKVEPKSCDKTHTCPPC PAPELLGGPSVFLFPPKPKDTLMISRTPEVTCVVVDVSHEDPEVKFNWYVDGVEVHNAKT KPREEQYNSTYRVVSVLTVLHQDWLNGKEYKCKVSNKALPAPIEKTISKAKGQPREPQVY TLPPSREEMTKNQVSLTCLVKGFYPSDIAVEWESNGQPENNYKTTPPVLDSDGSFFLYSK LTVDKSRWQQGNVFSCSVMHEALHNHYTQKSLSLSPGK/ DIQMTQSPSSLSASVGDRVTITCRASQDVNT-AVAWYQQKPGKAPKLLIYSASFLYSGVP SRFSGSRSGTDFTLTISSLQPEDFATYYCQQHYTTPPTFGQGTKVEIKRTVAAPSVFIFP PSDEQLKSGTASVVCLLNNFYPREAKVQWKVDNALQSGNSQESVTEQDSKDSTYSLSSTL TLSKADYEKHKVYACEVTHQGLSSPVTKSFNRGEC/ DIQMTQSPSSLSASVGDRVTITCRASQDVNT-AVAWYQQKPGKAPKLLIYSASFLYSGVP SRFSGSRSGTDFTLTISSLQPEDFATYYCQQHYTTPPTFGQGTKVEIKRTVAAPSVFIFP PSDEQLKSGTASVVCLLNNFYPREAKVQWKVDNALQSGNSQESVTEQDSKDSTYSLSSTL TLSKADYEKHKVYACEVTHQGLSSPVTKSFNRGEC*

3. Create the python script model-single.py for the model building:

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from modeller import * from modeller.automodel import * env = Environ() a = AutoModel(env, alnfile=’HERCAB_1hzh.ali’, knowns=’1hzh’, sequence=’HERCAB’, assess_methods=(assess.DOPE, assess.GA341)) a.starting_model = 1 a.ending_model = 10 a.make()

4. Run the script: python model-single.py > model-single.log Check the log file for the errors. Look for the output structure filename with the lowest DOPE score at the end of the log file. 5. Rename chain names (chain A to chain H, chain B to chain K, chain C to chain L, and chain D to chain M) in the build model to the original naming using the pdb_tools: pdb_rplchain -A:H lowest_DOPE_score.pdb | pdb_rplchain -B:K | pdb_rplchain -C:L | pdb_rplchain -D:M > final_structure.pdb

Where lowest_DOPE_score.pdb is the structure with the lowest DOPE score in the log file (step 4).

4

Notes 1. HDX-MS uses reversed-phase chromatography as a first analytical step. Here, the reversed-phase trap cartridge is used to desalt the peptides generated upon digestion. Any buffer, salt, or other additives (glycerol, sucrose, etc.) used to stabilize the protein (antibody) in solution that are not retained on a RP resin can thereby be efficiently removed. Special attention must be paid to detergents or polymers whose removal may require specific conditions [64]. The ability of HDX-MS to run experiments at different pH and temperature values is not strictly straightforward, as the exchange rate is dependent on these variables. Therefore, correction must be applied if an antibody is studied at different pH or temperature. Compensation factor for pH difference is 10exp(pHhigh - pHlow), which for instance means that exchange is 100× faster at pH 8 than at pH 6 [65]. The temperature effect is corrected based on an Arrhenius equation, for example, factor compensating temperature effect between 25 and 37 °C is 3.035. The Excel file for these calculations can be found here: http://peterslab.org/ downloads/SW/HDXtools/HDX-T-correction.zip.

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2. Antibodies are large molecules, the class IgG has a molecular weight of approximately 150 kDa, and is composed of two light (24 kDa) and two heavy chains (50 kDa). This should be considered for concentration calculations, whether it is expressed with respect to the entire intact antibody (150 kDa) or to the sum of the two amino acid sequences (74 kDa). Other factors to consider are the dilution factor (typically 10× in HDX), quench ratio, and the sensitivity of the instrument used. However, typically tens-hundreds of pmols are used per experiment and thus around 100 μg is needed for triplicated analysis of 5 time points. This also needs to be multiplied by the number of different experimental conditions that are being investigated. 3. Different buffers, for example, 20 mM HEPES and addition of salts (50–150 mM NaCl), are also possible. Other additives can be also included, but their compatibility with LC–MS workflow must be considered. This mostly means avoiding the majority of detergents, polymers, and other compounds, which cannot be removed during desalting on the reversed-phase trap column. 4. There are several recommendations on how to report pD value. Calculation using correction factor as described by Glasoe [66] is one way; pD = pHread + 0.41. This can be eventually corrected to D2O content using the equation pD = pHread + Dfrac × 0.41 as described recently [67]. Recommendations provided by the community-wide publication suggest using direct pHread as the unambiguous value not affected by eventual mistakes in calculation though isotope correction. 5. The quench solution serves two purposes. First, it lowers the pH to a point where the exchange (and thus deuterium loss) is minimal. Second, it aids in sample denaturation and facilitates digestion. In the case of disulfide bond containing proteins (such as antibodies), the disulfide bond reduction is also achieved during the quenching step which means lowering the pH to 2.5. The low pH conditions disfavor reduction by common agents such as beta-mercaptoethanol or dithiothreitol as they are not functional at low pH. Luckily, phosphine-based reagents (e.g., tris-carboxyethyl phosphine, TCEP) are capable of disulfide bond reduction even at acidic pH. However, its effectivity is drastically reduced [44], and thus large molar excess is required. Common practice also includes post-quench incubation for couple of minutes on ice. An alternative solution is to perform the incubation/reduction before the sample injection. It is advisable to prepare several different quench buffers, for example, without denaturing agents, then with 6 M guanidinium chloride or 8 M urea or mixture of 6 M urea and 3 M thiourea. With these concentrations, 1:1 quench

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ratio reaches the highest advisable denaturant (final) concentration. Dissolving urea and thiourea might be slow but slight heating (40 °C) and sonication helps. Keep in mind that TCEP is provided as hydrochloride and thus it will introduce HCl into the solution. That is why adjustment of glycine to glycine hydrochloride should be done only after the full dissolution of all buffer components. TCEP is expensive and not very stable. 6. The most typical setup nowadays consists of a robotic station capable of sample preparation and injection and a Peltiercooled compartment that houses the LC setup. Another option is to place the entire UPLC system into a temperaturecontrolled cabinet. Alternatively, the LC setup comprising injection and switching valves, protease, trap, and analytical columns can be mounted onto a metallic plate and placed at the top of the box filled with water-ice. This also ensures highly stable temperature (0 °C) for several hours. 7. All tubing should have the shortest possible length to minimize dead volumes. Exception applies to the line that delivers loading/desalting solvent at a high flow rate (100–200 μL/min). This solvent is pumped isocratically and thus the tubing length is insignificant. However, to ensure sufficient cooling, additional tens of centimeters should be added, spirally coiled and immersed in the bath or closed in the thermo-controlled compartment. 8. The column setup may include individual columns or their combinations, coupled either in series (the specific order of the proteases matters, as their different serial combinations provide slightly different results) or in parallel. Alternatively, coimmobilized enzymes can be employed as well. Except for pepsin, all other proteases mentioned in this chapter withstand a pH above 5, but they all have very low tolerance toward organic solvents. Protease columns were shown to be the biggest source of carryover and need to be washed, but the cleaning procedures must be designed according to their stability in various solvents. 9. A peptide length distribution is easily obtained if the peptide list also contains the from–to (or start–end) peptide limits. Redundancy distribution can be calculated by an Excel macro [24] available here: http://peterslab.org/downloads.php under the HDX tools section. Upon calculation, the second list shows the overall digestion metrics, adjacent column lists residue resolved redundancy numbers. 10. TCEP is also an ionic compound and not completely removed during the desalting step. Therefore, its signal is found in the LC–MS data. Therefore, it is advised to use the lowest possible

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concentration that provides reduction. Our previous work showed that there was no difference between the digestion pattern done with 0.5 M or 0.1 M TCEP [43], but the background signal of TCEP was significantly lower at 0.1 M concentration. 11. The stock solution of myoglobin (1 mg/mL (59 μM) in water) can be stored at -20 °C. Take 1 μL and add 294 μL of glycine buffer to make 0.2 μM acidic solution for quality control runs. When making this solution, always add buffer to the concentrated protein and not protein into the buffer. 12. If the subsequent workflow includes freezing samples in liquid nitrogen, include the same step in the optimization phase as well. It is important to assess the effect of the freeze/thaw cycle as it may lead to sample precipitation. 13. To calculate the time required for efficient digestion and desalting, volumes of the injection loop, protease and trap columns, and all tubing connecting these elements must be added together to get the total volume. Based on that and the employed flow, sufficient digestion time can be calculated. Typically, 3 min are enough for the setup and flows described here. It should be extended to 4 min if guanidine is used in the quench buffer as it requires longer desalting. 14. Duration of the gradient and its slope depend on the column and the LC system used. In general, the gradient should be fast, lasting just a few minutes, but efficient elution still needs to be achieved. Gradients running from 5% or 10% acetonitrile (solvent B) and ending at 35–45% acetonitrile are generally recommended. In specific cases, the upper acetonitrile percentage can be raised to higher percentages. 15. The timsTOF mass spectrometer used here is very sensitive and able to efficiently fragment/sequence even peptides of very low intensity. Therefore, it is not rare to obtain hundreds of identified peptides, even from relatively small proteins. However, a substantial number of these identified peptides will not be useful in HDX-MS due to very low intensity and thus need to be removed from the list of HDX target peptides (as discussed in the main text). Also, comparisons of different digestion conditions cannot be easily performed without a thresholding set and one should focus on relatively high intensity levels (50–100 k) when doing so. However, the list of peptides for subsequent interpretation of HDX-MS data should be taken from 10 k level data to maximize the HDX coverage and resolution. 16. For difficult proteins such as antibodies, it is also advisable to prepare a fully reduced sample (reduction at neutral/slightly basic pH) to see the optimal digestion when full disulfide bond

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reduction was achieved. In addition to a fully reduced sample, prepare also a deglycosylated protein (e.g., by PNGase F treatment) to cover the glycosylation site(s). For the deglycosylated sample, deamidation of Asn (Asn to Asp conversion is a product of glycan hydrolysis) must be considered in the search. Finally, search the native, glycosylated LC–MS/MS data for the glycopeptide signals using oxonium ions. Optimally, the N-glycosylation site(s) should be covered, the type of the most prominent glycan structure identified, and the glycopeptide signals should be included in the HDX-MS processing. 17. If the reaction is prepared by one person, the shortest reasonably reproducible time point is 10 s. Shorter times, even 2 s are possible but require specific setup and two coordinated users, one performing the pipetting with both hands and the other monitoring the time and instructing the other accordingly. 18. The acid solution for quenching must be titrated with proteinfree solutions in advance. The variables to test are the volume ratios and the concentration of the acid. Good mixing is achieved using 1:1 quench and volumes of approx. 50–100 μL. Quenching using buffers, preferably very strong ones (e.g., 0.5 M glycine HCl, pH 2.3), does not need this initial step because strong buffer will always override the buffer used during the exchange step. 19. Sample analysis should be done without unnecessary delays, optimally within hours or maximally days following sample collection. Storage at -20 °C can be problematic as highly denaturing quench conditions will lead to partial melting. That is why optimal storage is only achieved at -80 °C or in liquid nitrogen. 20. There are several ways how fully deuterated proteins can be prepared. A first one is based on acid-aided denaturation followed by deuteration at elevated temperatures. This can, however, lead to precipitation and most likely, acid hydrolysis will occur, mostly via the cleavage after Asp. A second way is to denature the protein using denaturing agents. Here, special attention must be paid to the total deuteration level of the reagents prior to their use. For instance, 6 M urea delivers a lot of hydrogens into the solution and thus the overall deuterium level will be lower than in the original experimental deuterated buffer. Another alternative is to use predenaturation using organic solvents [68] or to predigest the protein and perform deuteration on the peptide level, which is an option that is described later. 21. This step can eventually be omitted and performed just before the analysis. This means thawing the sample 5 min before

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injection and placing it on ice. Both options were tested without any significant difference. 22. We will use 5 μL of this solution per aliquot and thus 25 μL are enough for five time points. The additional 2 μL are reserve, ensuring that even the last aliquot will be taken in full volume. Each sample contains 100 pmol of Ab, but it is possible to use 50 or 20 pmol or even less. However, very low concentration of Ab will cause worsening of signal-to-noise ratio and might result in not detecting low intensity peptides. At the same time, one has to be careful with very high signal intensities as well because ion trap type of mass analyzers can be overloaded, resulting in isotope beating due to space charge issues [69]. 23. The tubes are not completely gastight and thus may acquire some nitrogen which then tends to expand (explode) upon thawing of the sample. It is advisable to open/close the tubes and let the nitrogen evaporate. 24. Such preparation of FD control does not require deuterated denaturing agents, does not suffer from protein acid hydrolysis and is suitable for less stable proteins. But there is one potential risk involved: if the protein contains cysteines and the quench solution does not contain TCEP, Cys-containing peptides will likely form disulfides during the deuteration step. For antibody, one uses quench with TCEP anyway and performs reduction on ice. But even for other proteins, it is advisable to add 1 mM TCEP to the deuterated buffer while making sure that pH is not affected. 25. Soft conditions are mostly determined by lower desolvation temperature, lower gas flows, and minimization of ion activation during the transfer. Unfortunately, lowering these parameters will also result in a significant drop in sensitivity. Low scrambling can be validated using ETD/UVPD with an HDX peptide probe [22]. If these fragmentation techniques are not available on the instrument used for HDX, parameters developed on a different MS system with identical or highly similar ESI source design can be used. For timsTOF, an instrument with a similar design is the maXis q-TOF instrument as previously tested [70]. 26. The protease column was identified as the biggest source of carryover and specific washes were designed to minimize this effect [71]. 27. Using this sample order, one ensures that there is no carryover effect of the ND sample in the FD or partially deuterated samples.

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INDEX A

F

Acetylation.............................. 2, 9, 18, 64, 92, 109, 111, 112, 130, 138, 144, 147, 148, 219, 279 Affinity purification ............................................ 23, 44, 53 Alternative proteases ...............................................99–109 Antibody ........................................ 12, 30, 39–41, 47, 48, 54, 58, 62, 63, 65, 147, 168, 214, 215, 287, 291, 293, 300, 304, 306, 308–311, 313, 315–317, 319, 321, 324, 326, 328, 330 Automation ................................................. 182, 257, 293 AutoSP3...............................................182, 183, 185–210

Fibroblast......................................................................... 25 Formalin fixation in combination with paraffin embedding (FFPE) ..................182, 185, 187, 189, 191, 195, 197, 199, 201, 203, 204, 208, 209, 213–218, 222–223, 226, 229, 230, 272, 275–277, 279, 281 Furan-biotin ....................................................... 12–16, 19

B BioID ................................................................. 24–32, 37, 39–44, 49, 53, 54 Biopsy ................................................ 182, 189, 217, 218, 222, 223, 229, 286, 287, 289, 290 Biotin .................... 12–14, 17, 24, 27, 29, 38–40, 45, 47

C Cell culture ................ 28, 29, 37, 45, 54, 56, 57, 61, 76, 81, 83, 84, 93, 95, 190, 208, 255, 256, 258, 259 Cell surface ................................................................11–19 Cellular dynamics ......................................................23–49 Cellular thermal shift assay (CETSA) ......................73, 74 Charge-based fractional diagonal chromatography (ChaFRADIC)................................. 100, 109, 112 Clinical proteomics ...........................................v, 181–210

G Glycopeptides .................... 153, 155, 157–160, 162, 329 Glycoproteomics .................................................. 153, 157 Glycosylation ......................................... 11, 152–154, 329

H Heterologous expression .............................................. 137 Histone post-translational modifications....................271, 272, 275, 277, 278, 280 Hydrogen/deuterium exchange (HDX) ........... 304–306, 308–310, 313, 315, 319–321, 323, 326–328, 330 HYPERsol ................................................... 216, 217, 229

I Interaction network ...................................................... 240 In vivo phosphorylation ...................................... 167–178

K Kinase assay ................................................................... 168

D

L

Degradomics ................................................................. 112 Density gradient ................ 254, 257, 260–263, 265–267 Drug-target engagement ................................... 12, 73–97

Laser microdissection (LMD) .....................272–276, 281 Lentivirus ............................................... 26–29, 32, 37, 49 Liquid chromatography (LC).........................3, 6, 67, 69, 79, 116, 121, 122, 132, 158, 273, 278, 281 Liquid chromatography tandem mass spectrometry (LC-MS/MS) ......................................1, 2, 4, 6, 9, 12, 13, 19, 27, 43, 44, 56, 59, 62, 64, 66, 69, 76, 79, 91, 117, 122, 123, 129, 132, 139, 143, 155, 157–159, 168, 175, 222, 229, 231, 310

E Epigenetics .................................................................... 271 Escherichia coli ..................................................24, 59, 138 Extracellular vesicle (EV)........................... 235, 237–243, 248, 253–255, 258, 259, 262–268

Kris Gevaert (ed.), Mass Spectrometry-Based Proteomics, Methods in Molecular Biology, vol. 2718, https://doi.org/10.1007/978-1-0716-3457-8, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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336 Index M

S

Mass spectrometry (MS)................... 4, 6, 12, 23, 53, 58, 92, 125, 138, 143, 181–210, 285, 286, 303–331 Multi-omics ................................................................... 255

N-acetyltransferase (NAT).................................. 137, 138, 140, 141, 145–147 N-terminal peptides...................................... 1, 2, 6, 9, 64, 92, 99, 100, 102, 105, 108, 109, 112, 138 post-translational modifications ............................. 112 N-terminomics ............................................. 99, 100, 108, 112, 113, 137, 138

Sample preparation ........................... 4, 5, 39, 56, 58, 62, 78, 89, 96, 112, 115, 121, 149, 154–156, 169, 175, 182, 216, 228, 236, 237, 241–245, 327 Single sample ........................................................ 144, 293 Size-exclusion chromatography ......................... 254, 256, 258, 260, 263 SP3 ...............................................89, 156, 162, 183, 184, 186, 187, 193–197, 203, 204, 206, 207 S-Trap ................................................. 2–5, 7–9, 101, 102, 104, 105, 109, 216–219, 222, 225, 229–231 Streptavidin..................12, 24, 27, 30, 31, 40–42, 47, 65 Substrate specificity .............................................. 130, 138 Surfaceome .................................................. 11, 12, 14, 16

P

T

N

Phosphoproteomics ............................................. 286, 291 Plants .......................................... 167–170, 172, 177, 178 Plasma ...................................................3, 6, 7, 11, 13, 19, 24, 55, 70, 74, 153, 189, 190, 254 Plasma membrane proteins................................ 11, 13, 19 Protease substrate discovery .................................................. 129 Protein complex...........................................13, 23, 24, 53, 54, 74, 112, 121, 152, 183, 190, 246, 249, 323 dynamics ....................................................11, 24, 152, 167, 168, 190, 204, 305, 319 fusion ........................................................... 31, 32, 54, 64, 70, 138, 169, 304 structure................................................ 23, 31, 59, 95, 153, 214, 216, 305, 306, 319, 323, 329 Protein kinase ....................................................... 167–169 Protein-protein interaction (PPI) ..................... 23, 54, 55 Proteolysis.......................................................49, 202, 317 Proximity-dependent biotinylation (PDB).............................. 24–26, 31, 47, 321, 323 Proximity labeling ........................................................... 38

Terminal amine isotopic labeling of substrates (TAILS)........................................... 100, 113–115, 118, 121, 123, 125, 127–129, 132, 133 TMT-based quantitative proteomics.............................. 74 Transient protein expression ............................... 168, 169 Tryp-N ..........................................................1–4, 6–9, 231 Trypsin ....................................... 1–4, 6, 7, 15, 18, 24, 31, 39, 42, 43, 45, 46, 49, 54, 58, 62, 64, 65, 69, 77, 79, 83, 89, 92, 94, 116, 119, 131, 139, 142, 154, 156, 157, 159, 171, 174, 175, 186, 198, 207, 215–220, 225, 227, 229, 231, 241, 244, 247, 249, 272, 273, 277, 278, 286–288, 291, 293

U Urine......................... 189, 235–237, 242, 247, 248, 254

V Viral-like particles............................................................ 54 Virotrap............................................ 53–57, 59–61, 65–70