Proteomic profiling : methods and protocols [Second ed.] 9781071611852, 1071611852

Table of contents :
Preface
Contents
Contributors
Chapter 1: How Modern Mass Spectrometry Can Solve Ancient Questions: A Multi-Omics Study of the Stomach Content of the Oldest ...
1 Mass Spectrometry and the Revolution in Biological Discovery
2 Archeology Meets Multi-Omics Science
3 Oetzi Under the Multi-Omics Microscope
3.1 Proteomics
3.2 Lipidomics
3.3 Metabolomics
3.4 Glycomics
3.5 Metallomics
4 Concluding Remarks
References
Chapter 2: Step-by-Step Sample Preparation of Proteins for Mass Spectrometric Analysis
1 Introduction
2 Materials
2.1 High-Throughput In-Gel Digestion of Proteins
2.1.1 Excision of Protein Spots/Bands from Gels
2.1.2 Destaining of Gel Plugs
2.1.3 Reduction, Alkylation, and Enzymatic Digestion
2.1.4 Peptide Cleaning and Extraction
2.2 Filter-Aided Sample Preparation (FASP)
2.3 One-Step Guanidine Method for Protein Lysis and Digestion
2.4 StageTip Peptide Cleaning and Desalting
2.5 TMT (Tandem Mass Tag) Labeling of Peptides
3 Methods
3.1 HTP In-Gel Digestion of Proteins
3.1.1 Excision of Protein Spots/Bands from Gels
3.1.2 Destaining of Gel Plugs
3.1.3 Reduction, Alkylation, and Enzymatic Digestion
3.1.4 Peptide Cleaning and Extraction
3.2 Filter-Aided Sample Preparation (FASP)
3.3 One-Step ``Guanidine´´ Method for Protein Lysis and Digestion
3.4 StageTip Peptide Cleaning and Desalting
3.5 Modified TMT Labeling of Peptides
4 Notes
References
Chapter 3: Lab-on-a-Filter Techniques for Economical, Effective, and Flexible Proteome Analysis
1 Introduction
2 Materials
2.1 Reagents and Equipment
2.2 Suspension Trapping (STrap) Digestion-Specific Reagents
2.3 Filter-Aided Sample Preparation (FASP) Digestion-Specific Reagents
2.4 PVDF Membrane Digestion-Specific Reagents
3 Methods
3.1 Protein Preparation from Different Sources
3.2 STrap Digestion
3.3 FASP Digestion
3.4 PVDF Membrane Digestion
3.5 Performance Analysis
4 Notes
References
Chapter 4: Released N-Glycan Analysis for Biotherapeutic Development Using Liquid Chromatography and Mass Spectrometry
1 Introduction
2 Materials
2.1 Sample Preparation
2.2 LC-FLR/MS Analysis
3 Methods
3.1 Sample Preparation
3.2 LC-FLR-MS Analysis of Released N-Glycans
3.3 Data Analysis
4 Notes
References
Chapter 5: Comprehensive Protocol to Simultaneously Study Protein Phosphorylation, Acetylation, and N-Linked Sialylated Glycos...
1 Introduction
2 Materials
2.1 Sample Lysis, Reduction, Alkylation, and Digestion
2.2 HLB Cartridge Peptide Purification
2.3 TiO2 Batch Mode Enrichment
2.4 Deglycosylation
2.5 SIMAC Enrichment
2.6 Second TiO2 Enrichment
2.7 Acetyl Lysine IP
2.8 Desalting Peptide Mixtures on R3 RP Columns
2.9 HILIC Fractionation
2.10 LC-ESI-MS/MS Analysis
3 Methods
3.1 Sample Lysis, Reduction, Alkylation, and Digestion
3.2 HLB Cartridge Peptide Purification (See Notes 5 and 6)
3.3 TiO2 Batch Mode Enrichment
3.4 Deglycosylation
3.5 SIMAC Enrichment
3.6 Second TiO2 Enrichment
3.7 Acetyl Lysine IP
3.8 Desalting Peptide Mixtures on R3 RP Columns
3.9 HILIC Fractionation
3.10 LC-ESI-MS/MS Analysis (See Note 18)
4 Notes
References
Chapter 6: Phos-Tag Fluorescent Gel Staining for the Quantitative Detection of His- and Asp-Phosphorylated Proteins
1 Introduction
2 Materials (See Note 1)
2.1 Protein Samples
2.2 Solutions for Phos-Tag Magenta Gel Staining
2.3 Gel Staining and Detection Equipment
3 Methods
4 Notes
References
Chapter 7: Polyubiquitin Profile in Down Syndrome and Alzheimer´s Disease Brain
1 Introduction
2 Materials
2.1 Sample Preparation
2.2 Isolation of Polyubiquitinated Proteins
2.3 Isolation of Total Protein
2.4 Two-Dimensional Electrophoresis
2.5 Image Analysis
2.6 Protein Digestion
2.7 Peptide Extraction and Clean-Up
3 Methods
3.1 Sample Preparation
3.2 Isolation of Polyubiquitinated Proteins
3.3 Isolation of Total Proteins
3.4 Two-Dimensional Electrophoresis
3.5 Image Analysis
3.6 Protein Digestion
3.7 Peptide Extraction
4 Notes
References
Chapter 8: Platform Methods to Characterize the Charge Heterogeneity of Three Common Protein Therapeutics by Imaged Capillary ...
1 Introduction
2 Materials
2.1 Equipment
2.2 Reagents
3 Methods
3.1 icIEF Analysis of a Trastuzumab Biosimilar
3.1.1 CpB Treatment
3.1.2 Sample Preparation and icIEF
3.2 icIEF Analysis of rhEPO
3.2.1 Sample Concentration
3.2.2 Sample Preparation and icIEF
3.3 icIEF Analysis of a Fusion Protein
3.3.1 Sample Preparation
3.3.2 icIEF Analysis
4 Notes
References
Chapter 9: A Protocol for Isolation, Purification, Characterization, and Functional Dissection of Exosomes
1 Introduction
2 Materials
2.1 Small-Scale Exosome Production
2.2 Large-Scale Exosome Production
2.3 Protein Quantitation
2.3.1 Micro BCA Assay
2.3.2 Protein Staining Densitometry
2.4 Ultracentrifugation Exosome Isolation
2.5 OptiPrep Density Gradient Exosome Isolation
2.6 Cushion-Based Separation of Exosomes
2.7 EpCAM Immunoaffinity Capture (IAC) Exosome Isolation
2.8 Western Blot Analysis
2.9 Nanoparticle Tracking Analysis (NTA)
2.10 Aldehyde/Sulfate Latex (ALS) Bead-Based Capture
2.11 Electron Microscopy (EM)
2.11.1 Transmission EM
2.11.2 Cryo EM
2.12 Proteomics: Sample Preparation
2.12.1 In-Solution Reduction, Alkylation, and Digestion
2.12.2 StageTip Sample Cleanup
2.13 Fluorometric Peptide Assay
2.14 UHPLC-MS/MS
2.15 Data Analysis
2.16 Phenotypic Reprogramming of Cells by Exosomes: Dissecting Function
2.16.1 Labeling Exosomes with Lipophilic Tracer
2.16.2 Exosome Uptake
2.16.3 Cell Activation Assay
2.16.4 Transwell-Matrigel Invasion Assay
2.16.5 Molecular Reprogramming of Cells by Exosomes
3 Methods
3.1 Small-Scale Exosome Production (2-20 μg)
3.2 Large-Scale Exosome Production
3.3 Protein Quantitation (See Note 7)
3.3.1 Micro BCA Assay
3.3.2 Protein Staining Densitometry
3.4 Ultracentrifugation Exosome Isolation
3.5 OptiPrep Density Gradient Exosome Isolation
3.6 Cushion-Based Separation of Exosomes
3.6.1 Cushion-Based Separation of Exosomes
3.6.2 Ultracentrifugation Density Gradient Separation
3.7 EpCAM Immunoaffinity Capture Exosome Isolation (See Note 15)
3.8 Western Blot Analysis (See Note 16)
3.9 Nanoparticle Tracking Analysis (NTA) (See Note 17)
3.10 Aldehyde/Sulfate Latex Bead-Based Capture
3.11 Electron Microscopy (EM) (See Note 18)
3.11.1 Transmission EM
3.11.2 CryoEM
3.12 Proteomics: Sample Preparation
3.12.1 In-Solution Reduction, Alkylation, and Digestion (See Note 19)
3.12.2 StageTip Cleanup (See Note 23)
3.13 Fluorometric Peptide Assay (See Note 25)
3.14 UHPLC-MS/MS
3.15 Data Analysis
3.16 Phenotypic Reprogramming of Cells by Exosomes: Dissecting Function
3.16.1 Labeling Exosomes with Lipophilic Tracer (See Note 27)
3.16.2 Exosome Uptake (See Note 29)
3.16.3 Cell Activation Assay
3.16.4 Invasion Assay
3.16.5 Molecular Reprogramming of Cells by Exosomes
4 Notes
References
Chapter 10: Human Plasma Extracellular Vesicle Isolation and Proteomic Characterization for the Optimization of Liquid Biopsy ...
1 Introduction
2 Materials
2.1 PBPL Isolation
2.2 Large EVs Isolation
2.3 Small EVs Isolation
2.3.1 CD63 Exo-FLOW Capture Kit
2.3.2 ExoQuick ULTRA Kit
2.3.3 exoEasy Kit
2.3.4 Purification Mini Kit
2.4 Albumin Depletion (See Note 14)
2.5 EV Lysate Preparation
2.6 Protein Quantitation (See Note 19)
2.7 Western Blot Analysis
2.8 Nanoparticle Tracking Analysis
2.9 Transmission Electron Microscopy
2.10 Proteomics: Sample Preparation
2.10.1 In-Solution Reduction, Alkylation, and Digestion
2.10.2 StageTip Sample Cleanup
2.11 Fluorometric Peptide Assay
2.12 UHPLC-MS/MS
2.13 Data Analysis
3 Methods
3.1 PBPL Isolation
3.2 Large EVs Isolation (See Note 22)
3.3 Small EVs Isolation
3.3.1 CD63 Exo-FLOW Capture Kit
3.3.2 ExoQuick ULTRA Kit
3.3.3 exoEasy Kit
3.3.4 Purification Mini Kit
3.4 Albumin Depletion (See Notes 14, 15, and 29)
3.5 EV Lysate Preparation (See Notes 16-18)
3.6 Protein Quantitation (See Note 19)
3.7 Western Blot Analysis (See Notes 32-35)
3.8 Nanoparticle Tracking Analysis (See Notes 32 and 36)
3.8.1 NanoSight NS300 System
3.8.2 ZetaView PMX-120 (See Notes 32 and 36)
3.9 Transmission Electron Microscopy (See Notes 27, 37-39)
3.10 Proteomics: Sample Preparation
3.10.1 In-Solution Reduction, Alkylation, and Digestion
3.10.2 StageTip Cleanup
3.11 Fluorometric Peptide Assay (See Note 45)
3.12 UHPLC-MS/MS
3.13 Data Analysis
4 Notes
References
Chapter 11: Isolation of Extracellular Vesicles for Proteomic Profiling
1 Introduction
2 Materials
2.1 Cell Culture with Serum-Free Culture Media
2.2 Cell Culture with EV-Depleted Serum-Containing Culture Media
2.3 Density Gradient Ultracentrifugation
2.4 Methanol/Chloroform Precipitation
2.5 In-solution Protein Digestion
2.6 Desalting Using C18 Spin Column
3 Methods
3.1 Cell Culture with Serum-Free Culture Media
3.2 Cell Culture with EV-Depleted Serum-Containing Culture Media
3.3 Density Gradient Ultracentrifugation: Serum-Free Samples
3.4 Density Gradient Ultracentrifugation: EV-Depleted Serum Samples
3.5 Determination of Fraction Density
3.6 Proteomic Profiling
3.6.1 Methanol/Chloroform Precipitation
3.6.2 In-solution Digestion of EVs
3.6.3 Desalting Using C18 Spin Column (See Note 15)
4 Notes
References
Chapter 12: Isolation of Proteins from Extracellular Vesicles (EVs) for Mass Spectrometry-Based Proteomic Analyses
1 Introduction
2 Materials
3 Methods
3.1 EV-Cell Culture
3.2 EV Isolation
3.3 EV Lysis with Different Buffers
3.3.1 Lysis with RIPA Buffer
3.3.2 Lysis with Urea-Thiourea Solution
3.3.3 Lysis with Guanidium-Hydrochloride (Gu-HCl) Solution
4 Notes
References
Chapter 13: Flow Cytometry as an Important Tool in Proteomic Profiling
1 Introduction
2 Materials
3 Methods
3.1 Panel Design
3.2 Sample Preparation
3.3 Red Cell Lysis
3.4 Antibody and Viability Staining
3.5 Data Analysis (See Fig. 4)
4 Notes
References
Chapter 14: Improved Immunoprecipitation to Mass Spectrometry Method for the Enrichment of Low-Abundant Protein Targets
1 Introduction
2 Materials
3 Methods
3.1 Biotinylation of Antibodies
3.2 Immunoprecipitation with Streptavidin Magnetic Beads
3.3 Enzyme Elution, Single-Pot Reduction Alkylation, and Mass Spectrometry Sample Preparation
3.4 Second Elution for Antibody Epitope Mapping
4 Notes
References
Chapter 15: Multiplex Fluorescent Bead-Based Immunoassay for the Detection of Cytokines, Chemokines, and Growth Factors
1 Introduction
2 Materials
3 Methods
3.1 Assay Layout
3.2 Serum Collection and Preparation
3.3 Preparation of Standard Dilution Series (See Fig. 4)
3.4 Preparation of Magnetic Beads and Samples (See Note 7)
3.5 Preparation of Biotinylated Detection Antibodies (See Note 10)
3.6 Preparation of Streptavidin- Phycoerythrin (SA-PE) (See Note 11)
3.7 Main Assay Procedure
3.8 Plate Reading
3.8.1 Hardware Setup
3.8.2 Adjustment of the Needle
3.8.3 Start-Up
3.8.4 Calibration
3.8.5 Washing
3.8.6 Protocol Setup
3.8.7 Plate Reading
3.8.8 Shutdown
3.8.9 Data Analysis
4 Notes
References
Chapter 16: Bead-Based Multiplex Immunoassays: Procedures, Tips, and Tricks
1 Introduction
2 Technical Considerations
2.1 Assay Format
2.2 Sample Preparation and Storage
2.3 Lab Materials and Equipment
2.4 Assay Workflow
2.5 Maintenance and Shutdown Procedures
2.6 Data Analysis
2.6.1 Bio-Plex Manager Software
2.6.2 Bio-Plex Data Pro Software
3 Concluding Remarks
References
Chapter 17: Immunoaffinity-Based Liquid Chromatography Mass Spectrometric Assay to Accurately Quantify the Protein Concentrati...
1 Introduction
2 Materials
2.1 Proteotypic Peptides
2.2 Proteolysis
2.3 Immunoprecipitation
2.4 LC-MS/MS and Data Analysis
3 Methods
3.1 Peptide Preparation
3.2 Proteolysis
3.3 Immunoprecipitation
3.4 LC-MS/MS-Based Peptide Quantification
3.5 Data Analysis and Analyte Quantification
4 Notes
References
Chapter 18: Recombinant Anti-idiotypic Antibodies in Ligand Binding Assays for Antibody Drug Development
1 Introduction
2 Materials
3 Methods
3.1 PK Bridging Assay
3.2 PK Assay Using Drug-Target Complex-Specific Antibodies
3.3 ADA Bridging Assay
3.4 Batch to Batch Consistency Test
3.5 Accelerated Stability Test
4 Notes
References
Chapter 19: cDNA Display-Mediated Immuno-PCR (cD-IPCR): An Ultrasensitive Immunoassay for Biomolecular Detection
1 Introduction
2 Materials
2.1 Synthesis of cnvK-rG Puromycin Linker (See Note 1) (Fig. 1a)
2.2 Synthesis of BDA Coding cDNA Display Molecules
2.2.1 In Vitro Transcription of DNA Coding for B Domain Protein A (BDA)
2.2.2 UV Photocrosslinking Between mRNA and cnvK-rG Puromycin Linker
2.2.3 In Vitro Translation (Synthesis of mRNA-BDA Fusion Molecule)
2.2.4 Purification of mRNA-BDA Fusion Molecule
2.2.5 Reverse Transcription to Form cDNA Display Molecules
2.2.6 His-tag Affinity Purification of cDNA Display Molecule
2.3 cDNA Display-Mediated Immune-PCR (cD-IPCR)
2.3.1 Immobilization of Target Protein (Immunoglobulin G)
2.3.2 Direct Type cD-IPCR
3 Methods
3.1 Synthesis of Puromycin Linker
3.1.1 Reduction of Puromycin Segment
3.1.2 EMCS Modification of Biotin cnvK Segment
3.1.3 Cross-Linking of the Puromycin Segment and Biotin cnvK Segment
3.1.4 HPLC Purification of Puromycin Linker
3.2 Construction BDA DNA Templates
3.3 Synthesis of cDNA Display Molecules
3.3.1 In Vitro Transcription of BDA DNA
3.3.2 Photocrosslinking Between BDA mRNA and cnvK-rG Puromycin Linker
3.3.3 In Vitro Translation (Synthesis of mRNA-BDA Fusion Molecule)
3.3.4 Immobilization of mRNA-BDA Fusion Molecule on Streptavidin Magnetic Beads
3.3.5 Reverse Transcription to Form cDNA Display Molecules
3.3.6 His-tag Affinity Purification of cDNA Display
3.4 Direct cD-IPCR Detection of IgG
3.4.1 Immobilization of IgG in Magnetic Beads
3.4.2 Direct Type cD-IPCR
4 Notes
References
Chapter 20: Chromatin Immunoprecipitation (ChIP) to Study DNA-Protein Interactions
1 Introduction
2 Materials
2.1 Nuclei Sample Preparation
2.2 Prep Chromatin for IP
2.3 Prep Antibody for IP
2.4 Immunoprecipitation
3 Methods
3.1 Nuclei Sample Preparation
3.2 Prep Chromatin for IP
3.3 Prep Antibody for IP
3.4 Immunoprecipitation
4 Notes
References
Chapter 21: Profiling Protein-DNA Interactions by Chromatin Immunoprecipitation in Arabidopsis
1 Introduction
2 Materials
2.1 Consumable Materials and Reagents
2.2 Equipment
3 Methods
3.1 Material Collection and Fixation
3.2 Preparation of Antibody Beads
3.3 Isolation of Nuclei, Chromatin Shearing, and ChIP
3.4 Washing, Elution, Reverse Cross-Linking, and DNA Purification
3.5 qPCR and Data Analyses
4 Notes
References
Chapter 22: Biotin Proximity Labeling for Protein-Protein Interaction Discovery: The BioID Method
1 Introduction
2 Materials
2.1 Biotin Labeling Time Course Validation
2.2 Biotinylated Protein Generation
2.3 Cell Harvesting and Lysate Generation
2.4 Biotinylated Protein Purification
3 Methods
3.1 Biotin Labeling Time Course Validation
3.2 Biotinylation and Cell Harvest
3.3 Cell Lysis
3.4 Affinity Purification with Magnetic Streptavidin Beads
3.5 Western Blot Visualization
3.6 Data Analysis
4 Notes
References
Chapter 23: Studying OTUD6B-OTUB1 Protein-Protein Interaction by Low-Throughput GFP-Trap Assays and High-Throughput AlphaScree...
1 Introduction
2 Materials
2.1 Protein Expression and Purification
2.2 Immunoprecipitation Using GFP-Traps
2.3 ``AlphaScreen´´ Homogeneous Proximity Assay
3 Methods
3.1 Protein Expression and Purification Protocols
3.1.1 Recombinant Production and Purification of His-OTUB1
3.1.2 Recombinant Production and Purification of GST-OTUD6B
3.2 GFP-Trap Immunoprecipitation
3.2.1 Seeding of Cells and Transfection
3.2.2 Harvesting the Cells
3.2.3 Bead Equilibration of GFP-Trap-Agarose
3.2.4 GFP-Trap (Binding, Washing and Elution)
3.3 ``AlphaScreen´´ Homogeneous Proximity Assay
3.3.1 AlphaScreen Matrix Titration Experiment
3.3.2 AlphaScreen PPI Assay at Defined Concentration
4 Notes
References
Chapter 24: Thermal Shift Assay for Exploring Interactions Between Fatty Acid-Binding Protein and Inhibitors
1 Introduction
2 Materials
2.1 Protein Preparation and Purification
2.2 Protein Melt Reaction Using Real-Time PCR
3 Methods
3.1 Protein Preparation and Purification
3.2 Thermal Shift Assay
3.2.1 Protein Thermal Shift Assay Mix Preparation (See Table 1)
3.2.2 Set the Real-Time PCR Program (See Table 2)
3.2.3 Data Analysis
4 Notes
References
Chapter 25: Isolation and Purification of Mitochondria from Cell Culture for Proteomic Analyses
1 Introduction
2 Materials
2.1 Isolation of Crude Mitochondria
2.2 Density Gradient
3 Methods
3.1 Isolation of Crude Mitochondria (See Note 1)
3.2 Density Gradient Centrifugation (See Note 13)
4 Notes
References
Chapter 26: Investigating the Adipose Tissue Secretome: A Protocol to Generate High-Quality Samples Appropriate for Comprehens...
1 Introduction
2 Materials
2.1 General Hardware and Consumables
2.2 Tissue Biopsies and Isolation of Preadipocytes
2.3 Differentiation of Preadipocytes
2.4 Quality Control of Differentiation by Oil Red O Protocol
2.5 Collecting Secreted Peptides/Proteins
3 Methods
3.1 Tissue Biopsies
3.2 Isolation of Preadipocytes
3.3 Differentiation of Preadipocytes
3.4 Quality Control of Differentiation by Oil Red O Staining
3.5 Collecting Secreted Peptides/Proteins
3.6 Concentration of Supernatant for Proteomic Analyses
4 Notes
References
Chapter 27: Methods for Proteomics-Based Analysis of the Human Muscle Secretome Using an In Vitro Exercise Model
1 Introduction
2 Materials
2.1 General Consumables
2.2 Cell Culture
2.3 Targeted Proteomics
2.4 Untargeted Proteomics
3 Methods
3.1 Enrichment of CD56-Positive Myoblasts
3.2 Fusion to Myotubes and Stimulation
3.3 Collection of Medium for Targeted Secretome Studies
3.4 Collection of Medium for Untargeted Proteomics
4 Notes
References
Chapter 28: Western Blotting Using In-Gel Protein Labeling as a Normalization Control: Advantages of Stain-Free Technology
1 Introduction
2 Materials
2.1 Tissue Sample Preparation
2.2 SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE)
2.3 Protein Transfer
2.4 Antibodies for Immunostaining
3 Methods
3.1 Sample Preparation
3.2 SDS-Page
3.3 Protein Transfer
3.4 Immunostaining
4 Notes
References
Chapter 29: Technical Considerations for Contemporary Western Blot Techniques
1 Introduction
2 Immunodetection
2.1 Blocking
2.2 Wash Volume and Agitation
2.3 Antibody Selection and Dilution
2.3.1 Primary Immunodetection
2.3.2 Secondary Immunodetection
2.3.3 Antibody-Specific Ligands
3 Image Acquisition
3.1 Visualization Methods
3.1.1 Chemiluminescence Detection
3.1.2 Fluorescence Detection
3.2 Stripping and Reprobing
3.3 Imaging Systems
3.3.1 Digital Imaging for Chemiluminescence Detection
3.3.2 Digital Imaging for Fluorescence
3.4 Sensitivity
3.5 Scales of Reporting
3.5.1 Log
3.5.2 Orders of Magnitude/Decades
3.5.3 Bit Depth and Bits
3.5.4 Dynamic Range
3.5.5 Resolution
4 Image Analysis
4.1 Molecular Weight (Size) Estimation
4.2 Quantitation
4.2.1 Total Protein Normalization
4.2.2 Housekeeping/Single Protein Normalization
References
Chapter 30: Simple Western: Bringing the Western Blot into the Twenty-First Century
1 Introduction
2 Materials
2.1 Simple Western Size Reagents
2.2 Simple Western Charge Reagents
3 Methods
3.1 Measuring Protein Expression with Protein Normalization
3.2 Characterizing Charge Heterogeneity
4 Notes
References
Chapter 31: Development of Peptide Ligands for Targeted Capture of Host Cell Proteins from Cell Culture Production Harvests
1 Introduction
2 Materials
2.1 Protein Mix Formulation for Library Screening
2.2 Dual Fluorescence Library Screening
2.2.1 Incubation of the OBOP Library
2.2.2 Manual Bead Sorting
2.2.3 ClonePix 2 Bead Sorting
2.3 Confirmation of HCP Binding by Selected Peptides
2.4 Proteomic Analysis of the Chromatographic Fractions
2.4.1 Filter-Aided Sample Preparation (FASP)
2.4.2 nanoLC-MS/MS Analysis
3 Methods
3.1 Protein Mix Formulation for Library Screening
3.2 Dual Fluorescence Library Screening
3.2.1 Incubation of the OBOP Library
3.2.2 Manual Bead Sorting
3.2.3 ClonePix 2 Bead Sorting
3.3 Confirmation of HCP Binding by Selected Peptides
3.4 Proteomic Analysis of the Chromatographic Fractions
3.4.1 Filter-Aided Sample Preparation (FASP)
3.4.2 nanoLC-MS/MS Analysis
3.4.3 Statistical Analysis of the DDA Proteomics Data
4 Notes
References
Chapter 32: Sample Preparation of Secreted Mammalian Host Cell Proteins and Their Characterization by Two-Dimensional Electrop...
1 Introduction
2 Materials
2.1 Sample Preparation and Protein Assay
2.2 2D Electrophoresis: First Dimension IEF
2.3 2D Electrophoresis: Equilibration and Second-Dimension SDS-PAGE
2.4 Staining and Blotting
2.5 Immunodetection and Analysis
2.6 Image Analysis Software
3 Methods
3.1 Sample Preparation and Protein Assay
3.2 2-D Electrophoresis: First Dimension IEF
3.3 2D Electrophoresis: Equilibration and Second-Dimension SDS-PAGE
3.4 Staining and Blotting
3.5 Immunodetection
3.6 Software Analysis and Generation of a Match Rate for Assessment of Antibody Coverage
4 Notes
References
Chapter 33: Quantitative Proteomic Analysis Using Formalin-Fixed, Paraffin-Embedded (FFPE) Human Cardiac Tissue
1 Introduction
2 Materials
2.1 Sample Preparation and Protein Digestion
2.2 LC-MS and Functional Analysis
3 Methods
3.1 Protein Extraction from FFPE Tissue
3.2 Protein Digestion
3.3 LC-MS/MS Analysis, Quantification, and Functional Analysis
3.4 Quantitative Proteomics Using FFPE Cardiac Tissue: An Example
4 Notes
References
Chapter 34: Chloroplast Isolation and Enrichment of Low-Abundance Proteins by Affinity Chromatography for Identification in Co...
1 Introduction
2 Materials
2.1 Chloroplast Isolation from Pea (Pisum sativum)
2.2 Chloroplast Isolation from Arabidopsis
2.3 Chloroplast Stroma Extraction
2.4 Gel Filtration
2.5 Ion Exchange Chromatography
2.6 Hydrophobic Interaction Chromatography (HIC)
2.7 ATP/Purvalanol B Affinity Chromatography
2.8 Heat Treatment of Isolated Chloroplasts and Protein Extraction
2.9 Eu3+-IDA Column Affinity Chromatography
3 Methods (See Note 5)
3.1 Chloroplast Isolation from Pea (See Note 6)
3.2 Chloroplast Isolation from Arabidopsis (See Note 6)
3.3 Estimation of the Chlorophyll Content
3.4 Chloroplast Stroma Extraction
3.5 Gel Filtration
3.6 Ion Exchange Chromatography
3.7 Hydrophobic Interaction Chromatography (HIC)
3.8 ATP/Purvalanol B Affinity Chromatography
3.9 Heat Treatment of Chloroplasts and Protein Extraction
3.10 Eu3+-IDA Column Affinity Chromatography
4 Notes
References
Chapter 35: Principles of Protein Labeling Techniques
1 Introduction
2 Materials
3 Methods
3.1 Labeling with Fluorescent Dyes Prior to Electrophoretic Separations
3.2 Protein Labeling with Stable Isotopes
4 Notes
References
Chapter 36: Mechanical/Physical Methods of Cell Disruption and Tissue Homogenization
1 Introduction
2 Bead Impact Methods: Shaking Vessel
2.1 Theory
2.2 Practical Aspects
3 Bead Impact Methods: Stirred Agitated Beads
3.1 Theory
3.2 Practical Aspects
4 Rotor-Stator Homogenizer
4.1 Theory
4.2 Practical Aspects
5 Mortar and Pestle Tissue Grinders: Shear by Mechanical Pressure
6 High Pressure Batch: Expanding Fluids
6.1 Theory
6.1.1 The French Press G-M
6.1.2 The Parr Cell Disruption Vessel
6.2 Practical Aspects
6.2.1 French Press G-M
6.2.2 PARR Cell Disruption Vessel
7 HIGH Pressure Flow: Shear Through a Valve or Tube
7.1 Theory
7.1.1 High Pressure Valve with Impingement Wall: Gaulin and Rannie
7.1.2 High Pressure Flow Narrow Tubes or Opposed Jets: Microfluidics
7.2 Practical Aspects
7.2.1 Practical Aspects Using High Pressure Valve with Impingement Wall: Gaulin and Rannie
7.2.2 Practical Aspects: High Pressure Narrow Tubes/Opposed Jets
8 Low Pressure: Shear by Droplet´s Impingement
9 Ultrasonic Processors: Shear by Collapsing Bubbles
9.1 Theory
9.2 Practical Aspects
10 Extraction Across an Electromotive Field
10.1 Theory
10.2 Practical Aspects
11 Lysis with Pulsed Electric Fields
11.1 Theory
11.2 Practical Aspects
12 Microwave-Assisted Centrifugation
12.1 Theory
12.2 Practical Aspects
References
Index

Citation preview

Methods in Molecular Biology 2261

Anton Posch Editor

Proteomic Profiling Methods and Protocols Second Edition

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-bystep 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.

Proteomic Profiling Methods and Protocols Second Edition

Edited by

Anton Posch Bio-Rad Laboratories GmbH, Feldkirchen, Germany

Editor Anton Posch Bio-Rad Laboratories GmbH Feldkirchen, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1185-2 ISBN 978-1-0716-1186-9 (eBook) https://doi.org/10.1007/978-1-0716-1186-9 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved 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.

Preface This volume is a comprehensive continuation and extension of a previous edition published in 2015 and presents the latest developments of the main pillars of protein analysis, namely sample preparation, separation, and characterization. Individual technologies of each pillar combined into complementary and robust workflows render proteomic analysis of complex biological samples even more powerful and are the prerequisite to gain maximum value from biological samples in a single experiment. Various modern mass spectrometry techniques have become indispensable tools in proteome research, and their importance is very well reflected throughout this book in general and by the first chapter of this volume in particular. Rudolf Grimm provides a highly inspiring story about the first complex multi-omics study of the oldest human ice mummy, the 5300-year-old Iceman or Oetzi, and how mass spectrometry was applied. Core areas in this volume are protocols for the analysis of post-translational modifications and protein interaction partners, followed by sophisticated procedures to enrich for extracellular vesicles and organelles. In addition, several types of protein immuno-assays are described and complemented by various methods for the characterization of antibodies and host-cell protein analysis. Last but not least, a few standard sample preparation protocols and recent advances concerning immuno-chemical detection of proteins are included as well. Special thanks to the authors of the individual chapters who are well-known protein biochemists, and all of them have made the valuable choice to provide a detailed representation of their lab work and to share important tips and tricks for a successful and reproducible employment of their precious protocols in other laboratories. This book is for students of Biochemistry, Biomedicine, Biology, and Genomics and will be an invaluable source for the experienced, practicing scientist, too. Munich, Germany

Anton Posch

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

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1 How Modern Mass Spectrometry Can Solve Ancient Questions: A Multi-Omics Study of the Stomach Content of the Oldest Human Ice Mummy, the 5300-Year-Old Iceman or Oetzi . . . . . . . . . . . . . . . . . . . 1 Rudolf Grimm 2 Step-by-Step Sample Preparation of Proteins for Mass Spectrometric Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Xinping Li, Thomas Franz, Ilian Atanassov, and Thomas Colby 3 Lab-on-a-Filter Techniques for Economical, Effective, and Flexible Proteome Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 Yi-Han Lin, Rodrigo Vargas Eguez, Isha Vashee, and Yanbao Yu 4 Released N-Glycan Analysis for Biotherapeutic Development Using Liquid Chromatography and Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . 35 Ximo Zhang 5 Comprehensive Protocol to Simultaneously Study Protein Phosphorylation, Acetylation, and N-Linked Sialylated Glycosylation . . . . . . . . . 55 ˜ ez-Vea, Marcella Nunes Melo-Braga, Marı´a Iba´n Katarzyna Kulej, and Martin R. Larsen 6 Phos-Tag Fluorescent Gel Staining for the Quantitative Detection of His- and Asp-Phosphorylated Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Emiko Kinoshita-Kikuta, Eiji Kinoshita, and Tohru Koike 7 Polyubiquitin Profile in Down Syndrome and Alzheimer’s Disease Brain . . . . . . 79 Antonella Tramutola and Marzia Perluigi 8 Platform Methods to Characterize the Charge Heterogeneity of Three Common Protein Therapeutics by Imaged Capillary Isoelectric Focusing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Jiaqi Wu, Will McElroy, Charles Haitjema, ¨ ck, and Christopher Heger Carsten Lu 9 A Protocol for Isolation, Purification, Characterization, and Functional Dissection of Exosomes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Alin Rai, Haoyun Fang, Monique Fatmous, Bethany Claridge, Qi Hui Poh, Richard J. Simpson, and David W. Greening

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Human Plasma Extracellular Vesicle Isolation and Proteomic Characterization for the Optimization of Liquid Biopsy in Multiple Myeloma . . . . . . . . . . . . . . . . . Antonia Reale, Tiffany Khong, Rong Xu, Maoshan Chen, Sridurga Mithraprabhu, Nicholas Bingham, Andrew Spencer, and David W. Greening Isolation of Extracellular Vesicles for Proteomic Profiling . . . . . . . . . . . . . . . . . . . . Dongsic Choi, Janusz Rak, and Yong Song Gho Isolation of Proteins from Extracellular Vesicles (EVs) for Mass Spectrometry-Based Proteomic Analyses. . . . . . . . . . . . . . . . . . . . . . . . . . . Prabal Subedi, Michael Schneider, Michael J. Atkinson, and Soile Tapio Flow Cytometry as an Important Tool in Proteomic Profiling . . . . . . . . . . . . . . . . Michael P. Blundell, Sharon L. Sanderson, and Tracey A. Long Improved Immunoprecipitation to Mass Spectrometry Method for the Enrichment of Low-Abundant Protein Targets. . . . . . . . . . . . . . . Penny Jensen, Bhavin Patel, Suzanne Smith, Renuka Sabnis, and Barbara Kaboord Multiplex Fluorescent Bead-Based Immunoassay for the Detection of Cytokines, Chemokines, and Growth Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan M. Preuss, Ute Burret, and Sabine Vettorazzi Bead-Based Multiplex Immunoassays: Procedures, Tips, and Tricks . . . . . . . . . . . Candice Cox Immunoaffinity-Based Liquid Chromatography Mass Spectrometric Assay to Accurately Quantify the Protein Concentration of HMGB1 in EDTA Plasma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Viktoria Anselm, Andreas Steinhilber, Cornelia Sommersdorf, and Oliver Poetz Recombinant Anti-idiotypic Antibodies in Ligand Binding Assays for Antibody Drug Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stefan Harth and Christian Frisch cDNA Display-Mediated Immuno-PCR (cD-IPCR): An Ultrasensitive Immunoassay for Biomolecular Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chathuni Jayathilake and Naoto Nemoto Chromatin Immunoprecipitation (ChIP) to Study DNA–Protein Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eliza C. Small, Danielle N. Maryanski, Keli L. Rodriguez, Kevin J. Harvey, Michael-C. Keogh, and Andrea L. Johnstone Profiling Protein–DNA Interactions by Chromatin Immunoprecipitation in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hendry Susila, Zeeshan Nasim, Suhyun Jin, Geummin Youn, Hyewon Jeong, Ji-Yul Jung, and Ji Hoon Ahn

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Biotin Proximity Labeling for Protein–Protein Interaction Discovery: The BioID Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jeff E. Habel Studying OTUD6B-OTUB1 Protein–Protein Interaction by Low-Throughput GFP-Trap Assays and High-Throughput AlphaScreen Assays. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elisabeth Weber, Kenji Schorpp, and Kamyar Hadian Thermal Shift Assay for Exploring Interactions Between Fatty Acid–Binding Protein and Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jiaqing Hao Isolation and Purification of Mitochondria from Cell Culture for Proteomic Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yaschar Kabiri, Christine von Toerne, Adriana Fontes, Percy A. Knolle, and Hans Zischka Investigating the Adipose Tissue Secretome: A Protocol to Generate High-Quality Samples Appropriate for Comprehensive Proteomic Profiling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sonja Hartwig, Jorg Kotzka, and Stefan Lehr Methods for Proteomics-Based Analysis of the Human Muscle Secretome Using an In Vitro Exercise Model. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cora Weigert, Sonja Hartwig, and Stefan Lehr Western Blotting Using In-Gel Protein Labeling as a Normalization Control: Advantages of Stain-Free Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ o Silva Neris, Andrea Marie Chua Dobles, Roˆmulo Lea and Aldrin V. Gomes Technical Considerations for Contemporary Western Blot Techniques . . . . . . . . Kenneth Oh Simple Western: Bringing the Western Blot into the Twenty-First Century . . . . ¨ ck, Charles Haitjema, Carsten Lu and Christopher Heger Development of Peptide Ligands for Targeted Capture of Host Cell Proteins from Cell Culture Production Harvests. . . . . . . . . . . . . . . . . . . . . . . . R. Ashton Lavoie, Taufika Islam Williams, R. Kevin Blackburn, Ruben G. Carbonell, and Stefano Menegatti Sample Preparation of Secreted Mammalian Host Cell Proteins and Their Characterization by Two-Dimensional Electrophoresis and Western Blotting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anton Posch, Franziska Kollmann, Thomas Berkelman, and Elizabeth Dreskin Quantitative Proteomic Analysis Using Formalin-Fixed, Paraffin-Embedded (FFPE) Human Cardiac Tissue . . . . . . . . . . . . . . . . . . . . . . . . . Omid Azimzadeh, Michael J. Atkinson, and Soile Tapio Chloroplast Isolation and Enrichment of Low-Abundance Proteins by Affinity Chromatography for Identification in Complex Proteomes. . . . . . . . . Roman G. Bayer, Simon Stael, and Markus Teige

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Principles of Protein Labeling Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 Christian Obermaier, Anja Griebel, and Reiner Westermeier Mechanical/Physical Methods of Cell Disruption and Tissue Homogenization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 563 Stanley Goldberg

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

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Contributors JI HOON AHN • Department of Life Sciences, Korea University, Seoul, South Korea VIKTORIA ANSELM • Signatope GmbH, Reutlingen, Germany ILIAN ATANASSOV • Max-Planck-Institute for Biology of Ageing, Cologne, Germany MICHAEL J. ATKINSON • Institute of Radiation Biology, Helmholtz Zentrum Mu¨nchen, German Research Center for Environmental Health, Neuherberg, Germany; Department of Radiation Oncology, Klinikum rechts der Isar, Technical University of Munich, Munich, Germany OMID AZIMZADEH • Institute of Radiation Biology, Helmholtz Zentrum Mu¨nchen, German Research Center for Environmental Health, Neuherberg, Germany ROMAN G. BAYER • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria THOMAS BERKELMAN • Bio-Rad Laboratories, Inc., Hercules, CA, USA NICHOLAS BINGHAM • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia R. KEVIN BLACKBURN • Waters Corporation, Morrisville, NC, USA MICHAEL P. BLUNDELL • Bio-Rad Laboratories Inc., Kidlington, Oxfordshire, UK UTE BURRET • Institute of Comparative Molecular Endocrinology (CME), Ulm University, Ulm, Germany RUBEN G. CARBONELL • Department of Chemical and Biomolecular Engineering, North Carolina State University, Raleigh, NC, USA; Biomanufacturing Training and Education Center (BTEC), North Carolina State University, Raleigh, NC, USA MAOSHAN CHEN • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia DONGSIC CHOI • Research Institute of the McGill University Health Centre, Glen Site, McGill University, Montreal, QC, Canada BETHANY CLARIDGE • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia; Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia THOMAS COLBY • Max-Planck-Institute for Biology of Ageing, Cologne, Germany CANDICE COX • Bio-Rad Laboratories Inc., Hercules, CA, USA ANDREA MARIE CHUA DOBLES • Department of Neurobiology, Physiology, and Behavior, University of California, Davis, CA, USA ELIZABETH DRESKIN • Bio-Rad Laboratories, Inc., Hercules, CA, USA RODRIGO VARGAS EGUEZ • J. Craig Venter Institute, Rockville, MD, USA HAOYUN FANG • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia MONIQUE FATMOUS • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia; Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia ADRIANA FONTES • Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany; CNC-Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal; DCV-Department of Life Sciences, Faculty of Sciences and Technology of the University of Coimbra, Coimbra, Portugal

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THOMAS FRANZ • A&M Stabtest GmbH, Bergheim, Germany CHRISTIAN FRISCH • Bio-Rad AbD Serotec GmbH, Puchheim, Germany YONG SONG GHO • Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea STANLEY GOLDBERG • Glen Mills Inc., Clifton, NJ, USA ALDRIN V. GOMES • Department of Neurobiology, Physiology, and Behavior, University of California, Davis, CA, USA; Department of Physiology and Membrane Biology, University of California, Davis, CA, USA DAVID W. GREENING • Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia; Molecular Proteomics Group, Baker Heart and Diabetes Institute, Melbourne, VIC, Australia ANJA GRIEBEL • SERVA Electrophoresis GmbH, Heidelberg, Germany RUDOLF GRIMM • Agilent Technologies, Santa Clara, CA, USA JEFF E. HABEL • Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA KAMYAR HADIAN • Assay Development and Screening Platform, Helmholtz Zentrum Mu¨nchen, Neuherberg, Germany CHARLES HAITJEMA • ProteinSimple, a Bio-Techne Brand, San Jose, CA, USA JIAQING HAO • Department of Microbiology and Immunology, University of Louisville, Louisville, KY, USA STEFAN HARTH • Bio-Rad AbD Serotec GmbH, Puchheim, Germany SONJA HARTWIG • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Leibniz Center for Diabetes Research at Heinrich Heine University Duesseldorf, Duesseldorf, Germany; German Center for Diabetes Research (DZD), Neuherberg, Germany KEVIN J. HARVEY • Thermo Fisher Scientific, Carlsbad, CA, USA CHRISTOPHER HEGER • ProteinSimple, a Bio-Techne Brand, San Jose, CA, USA MARI´A IBA´N˜EZ-VEA • Genetics, Genomics and Microbiology Research Group, Health Science Department, Public University of Navarre, Pamplona, Navarra, Spain CHATHUNI JAYATHILAKE • Graduate School of Science and Engineering, Saitama University, Saitama, Japan PENNY JENSEN • Thermo Fisher Scientific, Rockford, IL, USA HYEWON JEONG • Department of Life Sciences, Korea University, Seoul, South Korea SUHYUN JIN • Department of Life Sciences, Korea University, Seoul, South Korea ANDREA L. JOHNSTONE • EpiCypher Inc., Durham, NC, USA JI-YUL JUNG • Department of Life Sciences, Korea University, Seoul, South Korea YASCHAR KABIRI • Institute of Toxicology and Environmental Hygiene, Technical University of Munich, School of Medicine, Munich, Germany BARBARA KABOORD • Thermo Fisher Scientific, Rockford, IL, USA MICHAEL-C. KEOGH • EpiCypher Inc., Durham, NC, USA TIFFANY KHONG • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia EIJI KINOSHITA • Department of Functional Molecular Science, Graduate School of Biomedical and Health Sciences, Hiroshima University, Hiroshima, Japan EMIKO KINOSHITA-KIKUTA • Department of Functional Molecular Science, Graduate School of Biomedical and Health Sciences, Hiroshima University, Hiroshima, Japan PERCY A. KNOLLE • Institute of Molecular Immunology and Oncology, University Hospital rechts der Isar, Technical University of Munich, Munich, Germany

Contributors

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TOHRU KOIKE • Department of Functional Molecular Science, Graduate School of Biomedical and Health Sciences, Hiroshima University, Hiroshima, Japan FRANZISKA KOLLMANN • Bio-Rad Laboratories GmbH, Munich, Germany JORG KOTZKA • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Leibniz Center for Diabetes Research at Heinrich Heine University Duesseldorf, Duesseldorf, Germany; German Center for Diabetes Research, Mu¨nchen-Neuherberg, Germany KATARZYNA KULEJ • Division of Protective Immunity and Division of Cancer Pathobiology, The Children’s Hospital of Philadelphia, Philadelphia, PA, USA; Department of Pathology and Laboratory Medicine, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA MARTIN R. LARSEN • Department of Biochemistry and Molecular Biology, University of Southern Denmark, Odense, Denmark R. ASHTON LAVOIE • Department of Chemical and Biomolecular Engineering, North Carolina State University, Raleigh, NC, USA STEFAN LEHR • Institute for Clinical Biochemistry and Pathobiochemistry, German Diabetes Center, Leibniz Center for Diabetes Research at Heinrich Heine University Duesseldorf, Duesseldorf, Germany; German Center for Diabetes Research, Mu¨nchen-Neuherberg, Germany XINPING LI • Max-Planck-Institute for Biology of Ageing, Cologne, Germany YI-HAN LIN • J. Craig Venter Institute, Rockville, MD, USA TRACEY A. LONG • Bio-Rad Laboratories Inc., Kidlington, Oxfordshire, UK CARSTEN LU¨CK • ProteinSimple, a Bio-Techne Brand, San Jose, CA, USA DANIELLE N. MARYANSKI • EpiCypher Inc., Durham, NC, USA WILL MCELROY • ProteinSimple, a Bio-Techne Brand, San Jose, CA, USA MARCELLA NUNES MELO-BRAGA • Department of Biochemistry and Immunology, Institute of Biological Science, Universidade Federal de Minas Gerais, Belo Horizonte, MG, Brazil STEFANO MENEGATTI • Department of Chemical and Biomolecular Engineering, North Carolina State University, Raleigh, NC, USA; Biomanufacturing Training and Education Center (BTEC), North Carolina State University, Raleigh, NC, USA SRIDURGA MITHRAPRABHU • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia ZEESHAN NASIM • Department of Life Sciences, Korea University, Seoul, South Korea NAOTO NEMOTO • Graduate School of Science and Engineering, Saitama University, Saitama, Japan; Epsilon Molecular Engineering, Inc., Saitama, Japan ROˆMULO LEA˜O SILVA NERIS • Department of Neurobiology, Physiology, and Behavior, University of California, Davis, CA, USA CHRISTIAN OBERMAIER • SERVA Electrophoresis GmbH, Heidelberg, Germany KENNETH OH • Bio-Rad Laboratories, Hercules, CA, USA BHAVIN PATEL • Thermo Fisher Scientific, Rockford, IL, USA MARZIA PERLUIGI • Department of Biochemical Sciences, Sapienza University of Rome, Rome, Italy OLIVER POETZ • Signatope GmbH, Reutlingen, Germany QI HUI POH • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia; Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia ANTON POSCH • Bio-Rad Laboratories GmbH, Munich, Germany

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Contributors

JONATHAN M. PREUSS • Institute of Comparative Molecular Endocrinology (CME), Ulm University, Ulm, Germany ALIN RAI • Baker Heart and Diabetes Institute, Melbourne, VIC, Australia JANUSZ RAK • Research Institute of the McGill University Health Centre, Glen Site, McGill University, Montreal, QC, Canada ANTONIA REALE • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia KELI L. RODRIGUEZ • EpiCypher Inc., Durham, NC, USA RENUKA SABNIS • Nisarga Biotech Pvt. Ltd., Satara, Maharashtra, India SHARON L. SANDERSON • Bio-Rad Laboratories Inc., Kidlington, Oxfordshire, UK MICHAEL SCHNEIDER • Institute of Radiation Biology, Helmholtz Zentrum Mu¨nchen, German Research Center for Environmental Health, Neuherberg, Germany KENJI SCHORPP • Assay Development and Screening Platform, Helmholtz Zentrum Mu¨nchen, Neuherberg, Germany RICHARD J. SIMPSON • Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia ELIZA C. SMALL • Thermo Fisher Scientific, Rockford, IL, USA SUZANNE SMITH • Thermo Fisher Scientific, Rockford, IL, USA CORNELIA SOMMERSDORF • Signatope GmbH, Reutlingen, Germany ANDREW SPENCER • Myeloma Research Group, Australian Centre for Blood Diseases, Monash University/The Alfred Hospital, Melbourne, VIC, Australia; Malignant Haematology and Stem Cell Transplantation, The Alfred Hospital, Melbourne, VIC, Australia; Department of Clinical Haematology, Monash University, Melbourne, VIC, Australia SIMON STAEL • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria; VIB Department of Plant Systems Biology, Ghent University, Ghent, Belgium ANDREAS STEINHILBER • Signatope GmbH, Reutlingen, Germany PRABAL SUBEDI • Institute of Radiation Biology, Helmholtz Zentrum Mu¨nchen, German Research Center for Environmental Health, Neuherberg, Germany HENDRY SUSILA • Department of Life Sciences, Korea University, Seoul, South Korea SOILE TAPIO • Institute of Radiation Biology, Helmholtz Zentrum Mu¨nchen, German Research Center for Environmental Health, Neuherberg, Germany MARKUS TEIGE • Department of Ecogenomics and Systems Biology, University of Vienna, Vienna, Austria; Max Perutz Labs, Department of Biochemistry and Cell Biology, University of Vienna, Vienna, Austria ANTONELLA TRAMUTOLA • Department of Biochemical Sciences, Sapienza University of Rome, Rome, Italy ISHA VASHEE • J. Craig Venter Institute, Rockville, MD, USA SABINE VETTORAZZI • Institute of Comparative Molecular Endocrinology (CME), Ulm University, Ulm, Germany CHRISTINE VON TOERNE • Research Unit Protein Science, Helmholtz Center Munich, German Research Center for Environmental Health GmbH, Munich, Germany ELISABETH WEBER • Assay Development and Screening Platform, Helmholtz Zentrum Mu¨nchen, Neuherberg, Germany CORA WEIGERT • Institute for Clinical Chemistry and Pathobiochemistry, University Hospital Tu¨bingen, Tu¨bingen, Germany; Institute for Diabetes Research and Metabolic Diseases of the Helmholtz Zentrum Mu¨nchen at the University of Tu¨bingen, Tu¨bingen, Germany; German Center for Diabetes Research (DZD), Neuherberg, Germany

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REINER WESTERMEIER • Freising, Germany TAUFIKA ISLAM WILLIAMS • Molecular Education, Technology, and Research Innovation Center (METRIC), North Carolina State University, Raleigh, NC, USA JIAQI WU • ProteinSimple, a Bio-Techne Brand, San Jose, CA, USA RONG XU • Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC, Australia GEUMMIN YOUN • Department of Life Sciences, Korea University, Seoul, South Korea YANBAO YU • J. Craig Venter Institute, Rockville, MD, USA XIMO ZHANG • Waters Corporation, Milford, MA, USA HANS ZISCHKA • Institute of Toxicology and Environmental Hygiene, Technical University of Munich, School of Medicine, Munich, Germany; Institute of Molecular Toxicology and Pharmacology, Helmholtz Center Munich, German Research Center for Environmental Health, Neuherberg, Germany

Chapter 1 How Modern Mass Spectrometry Can Solve Ancient Questions: A Multi-Omics Study of the Stomach Content of the Oldest Human Ice Mummy, the 5300-Year-Old Iceman or Oetzi Rudolf Grimm Abstract In the past 40 years, mass spectrometry has seen a stunning development regarding increased sensitivity, resolution, and accuracy, especially for biomolecule analysis. These days without any doubt mass spectrometry is the most powerful analytical tool as a standalone technique or in conjunction with separation techniques such as high-performance liquid chromatography (HPLC), gas chromatography (GC), or capillary electrophoresis (CE). It is literally used to analyze any kind of small or large molecules ranging from basic elements to metabolites, pesticides, toxins, small or large molecule drugs, oligonucleotides, peptides, proteins, and many other molecule classes. Here, various modern mass spectrometry techniques such as LC-MS, GC-MS, ICP-MS, and elemental bio-imaging are briefly described how they were used for the first complex multi-omics study of the oldest human ice mummy, the 5300-year-old Iceman or Oetzi. The study comprised of mass spectrometry–driven proteomics (protein profiling and characterization), metabolomics, lipidomics, glycomics, and metallomics. Key words Matrix-assisted laser desorption ionization time of flight (MALDI-TOF) mass spectrometry, Electrospray ionization (ESI) mass spectrometry, Inductively coupled plasma mass spectrometry (ICP-MS), Elemental bio-imaging, Proteomics, Metabolomics, Lipidomics, Glycomics, Metallomics, Oetzi, Iceman

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Mass Spectrometry and the Revolution in Biological Discovery Alongside further refinements of HPLC such as capillary-, nano-, and chip-HPLC, new developments in mass spectrometry such as matrix-assisted laser desorption ionization time of flight (MALDITOF) mass spectrometry, nano-electrospray mass spectrometry, various kinds of ion mobility mass spectrometry, and orbitrap mass spectrometry made is possible to dive much deeper in key biological application areas such as proteomics, metabolomics, lipidomics, glycomics, and other relevant omics areas.

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Nowadays, mass spectrometry is being utilized in a broad range of challenging application areas such as single-cell analysis [1], bee sperm metabolomics [2], analyzing cross-sections of first molar teeth of apes and humans including the 100,000-year-old Neanderthal [3, 4] or more than two-million-year-old Australopithecus africanus teeth samples [5] to understand their breastfeeding behavior, analyzing extraterrestrial materials from comets or meteorites [6, 7], or being currently used in the US Mars rover, just to mention a few [8]. The rise of mass spectrometry for biomolecule analysis started with the development of two new ionization techniques referred to as electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI). During the 1990s, the evolution of MALDI-TOF mass spectrometry allowed for the first time the rapid identification of large numbers of peptides and proteins separated either by 1D or 2D gel electrophoresis or by chromatographic technologies. MALDI-TOF mass spectrometry became the standard offline mass spectrometry analytical tool with key advantages such as being salt tolerant, even allowing samples in 6 M urea to be analyzed [9] as well as allowing the storage and re-analysis at a later point of time. Finally, the development of nano-electrospray mass spectrometry in combination with capillary- and nano-HPLC opened the door for deep proteomics and metabolomics analysis of samples. The rise of multi-omics or system biology studies that include complex DNA and RNA analysis and the subsequent development of very powerful new bioinformatics and pathway analytical tools gave a deeper understanding of biological processes, diseases, or drug actions at the molecular level. Another increasing use of mass spectrometry is the diagnostics area where it has started to replace more and more ELISA due to its absolute specificity, high linear range, and multiplexing capabilities using SRM assays. Mass spectrometry is also more and more widely used for the successful analysis of bacteria (microbiome) irrespective of their origin (human, animal, soil, and others). Furthermore, imaging mass spectrometry in its various versions (MALDI, DESI, SIMS, elemental bioimaging, or laser ablation ICP-MS) is contributing more and more to the understanding of all kinds of biological questions and compliments multi-omics approaches in a very powerful way.

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Archeology Meets Multi-Omics Science Complementary mass spectrometry techniques such as LC-MS, GC-MS, ICP-MS, and elemental bio-imaging were applied to the first complex multi-omics study of the oldest human ice mummy, the 5300-year-old Iceman or Oetzi. The study comprised of mass

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spectrometry–driven proteomics, metabolomics, lipidomics, glycomics, and metallomics. Non-mass spectrometric techniques such as next-gen DNA sequencing, PCR, and various types of microscopy further complemented the toolset applied [10]. This project presented a number of challenges due to the uniqueness of the historical human sample and, for that reason, was undertaken by a research consortium of world-leading scientists in the USA, Europe, Asia, and Australia [10]. Dealing with very unique and very precious ancient samples demanded the careful development of sample preparation strategies followed by applying the right set of modern mass spectrometry techniques in order to maximize the information that could be obtained. Once a unique sample is gone, it is gone forever! Other important challenges of dealing with stomach samples are the complexity of the samples as they can comprise of human, bacterial, plant, and animal materials. Furthermore, chemical compounds are likely to have degraded to a certain extent or even completely over the 5300-year period, contributing additional challenges in generating sufficient amounts of highquality data. Hence, a large focus was on the best sample preparation strategies. Typically, low milligram amounts of samples were used for the individual analyses. Exact sample preparation procedures and the analytical mass spectrometric setups are not described in detail here in this chapter as they are all available in publication number [10].

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Oetzi Under the Multi-Omics Microscope The Iceman or Oetzi is a very unique European glacier mummy as he is the only completely intact Neolithic human mummy that we currently know of. About 30 years ago, he was found by hikers in a glacier right at the border of Italy and Austria. After the recovery and some initial scientific studies of the mummy by Austrian scientists, the Iceman was transferred to the city of Bolzano, Italy where a dedicated Research Institute for Mummies and the Iceman (EURAC) was established. The institute is headed by paleopathologist Professor Albert Zink since then. Stomach samples of the Iceman were removed under sterile conditions at a temperature of 4  C several years ago by Professor Albert Zink and Dr. Eduard Egarter-Vigl (Fig. 1) and kept frozen until further usage. Numerous research projects have been undertaken since then. In this chapter, the focus will be on the complex mass spectrometric analysis of stomach samples of the Iceman. Previous radiological investigations [11] revealed that his well-preserved stomach was completely filled, allowing the identification of ancient endogenous biomolecules as well as getting an understanding of key components of a Neolithic meal, most likely the Iceman’s last meal.

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Fig. 1 Paleopathologists Prof. Albert Zink and Dr. Eduard Egarter-Vigl taking samples of the Iceman under sterile conditions at a temperature of 4  C in the Iceman’s conservation chamber at the Archeological Museum of Bolzano, Italy. (Figure is reproduced with permission of Professor Albert Zink (© EURAC/Marco Samadelli)) 3.1

Proteomics

Protein profiling and characterization studies have been performed using HPLC-Chip-QTOF mass spectrometry or Orbitrap mass spectrometry in conjunction with nano-HPLC separations of peptide samples derived either from in-gel digestion of 1D gel electrophoresis separated stomach samples dissolved in SDS-buffer or from off-gel electrophoretic separations using complete digest of the sample. As shown in Fig. 2, most of the stomach proteins were degraded and hence of very small size as expected when analyzing stomach samples. However, faint protein bands of 50 kDa and much larger could still be detected. A total of 167 unique animal and plant proteins were unambiguously identified, with 13 of them clearly assigned to the plant species of einkorn, the ancient form of wheat. Further evidence for the presence of einkorn was obtained by DNA analysis which showed that the grains belonged to either the subspecies of Triticum monococcum or Triticum urartu. The detected einkorn/wheat proteins Globulin 3A and Triticin are primarily expressed in the endosperm of wheat seeds. Oxalate oxidase 2 is typically found in the pericarp of wheat seeds. Several other identified proteins belonged to the ruminant animals of Ibex (alpine goat) and red deer, indicating that the Iceman was a meat eater. Surprisingly, several bracken (fern) proteins were identified which led to the speculation that the Iceman used fern leaves to wrap up his food. Bracken can be toxic and was most likely not part of the Iceman’s last meal. Spores might have entered his stomach unintentionally [10].

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Fig. 2 SDS-electrophoresis gel of various Iceman stomach samples

Interestingly, macro- and microscopical analysis showed that the stomach content still contained compact pieces of food such as striated muscle fibers or plant fragments identified as bran and glumes, indicating that the Iceman’s proteins derived from pieces of meat and whole seeds that he ate shortly before his death. All proteomics data were also confirmed by genomic approaches [10]. Of the proteins identified in the Iceman’s stomach, 22 had matched with proteins primarily expressed in neutrophils and with biological functions involved in inflammatory host response. The two subunits S100A8 and S100A9 of calprotectin (CP) were detected with the highest number of distinct peptide hits. This is of great interest as Helicobacter pylori (H. pylori) was detected in the Iceman’s stomach, and the entire DNA sequence of this ancient form of H. pylori was determined [12]. Inflamed gastric tissues of modern H. pylori-infected patients also show high levels of CP subunit S100A8 and S100A9 expression. Thus, the Iceman’s stomach was colonized by a cytotoxic H. pylori–type strain that triggered CP release as a result of host inflammatory immune responses. However, whether the Iceman suffered from gastric disease cannot be determined because of the poor preservation of the stomach mucosa [12]. 3.2

Lipidomics

Lipidomics studies have been performed after a modified Bligh and Dyer lipid extraction of the two samples (6.9 mg and 4.5 mg) followed by C-18 reversed phase separation and QTOF mass spectrometry. The key question addressed with this approach was whether the Iceman consumed any dairy products as it is known that domestication of cattle did take place in central Europe during

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Fig. 3 LC-ESI-MS lipid analysis of the Iceman stomach sample. TAG triacylglycerides

the Neolithic period and that milk became part of the human diet [13]. Dairy products have a very characteristic distribution of triacylglycerols (TAGs) with mostly TAGs containing between 28 and 38 carbon atoms as shown by analyzing goat milk and cheese (see Fig. 3). TAG distribution however showed predominantly TAGs with 44–56 carbon atoms similar to the profile obtained by analyzing ibex (an alpine goat) meat samples and fat attached to it. This suggests that he did not consume any dairy products in his last meal (s), but instead animal meat with a good amount of fat. Interestingly, his genome data showed that besides having brown eyes and blood group O that he was lactose intolerant [14] and most likely understood that consuming dairy products was detrimental to his health plus having a negative impact on his ability to hiking regularly over glaciers from one valley to another. Other data received from lipid analysis showed that most of the TAGs contained fully saturated fatty acid chains which was confirmed by total fatty acid analysis using GC-MS. Very interestingly, a significant amount (up to 20%) of TAGs with odd-chain fatty acids were identified. These are typically found in ruminant animals such as alpine goat or deer. Numerous phospholipids, sphingolipids, and diacylglycerols were also identified. Overall, the fatty acid composition and distribution matched very much to those of mammalian adipose tissue. Using a histology approach with Sudan III staining the presence of adipose tissue with fat droplets was verified in the Iceman’s stomach.

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3.3

Metabolomics

For metabolomics analysis, 2 mg of the stomach samples were extracted with a 4:1 methanol/water mix to allow for LC-MS and GC-MS metabolite profiling. In order to obtain comprehensive datasets LC-QTOF mass spectrometry in positive and negative ionization mode after either reversed-phase or HILIC HPLC separations were performed. In addition, GC-MS analysis of the same extracts was performed. A total of approximately 1000 metabolites of all compound classes was identified when combining the separate LC-QTOF and GC-MS data and with some of them having interesting biological function. A significant amount of cholesterol was detected which is consistent with the fact that the Iceman’s last meal and most likely in general his entire diet being very fatty. Interestingly, computed tomography scans of his arteria and aorta showed major calcifications [15]. We also identified cadaverine which is typically produced by protein hydrolysis during putrefaction of animal tissue, another indication of the Iceman’s last meal containing meat. Other interesting metabolic molecules identified were phytanic acid, indicating again the presence of ruminant fat and azelaic acid which supports the existence of whole-grain cereals in the Iceman’s diet. The identification of gamma-terpinene was interesting as it is typically found in coriander oil or essential oil made from citrus fruits, indicating that the Iceman added some spice or flavor to his meal in one way or the other. Another very interesting metabolite that got identified was diisopropylnaphthalene. This compound is formed from burning wood, suggesting that the Iceman used fire, most likely in the form of smoke, to preserve his meat. This is in great accordance with a former microscopical study of his lower intestine in 2007 that showed the presence of charcoal particles [11]. ICP-MS analysis showed no increased levels of sodium or chlorine which indicates that no salt was used for meat preservation.

3.4

Glycomics

Glycomics analysis of stomach samples was performed using HPLC-Chip QTOF mass spectrometry. Two 21 and 23 mg samples were micro-ground and delipidated. N-Glycans were isolated using enzymatic release of the glycans by PNGase F, ethanol precipitation, and solid phase extraction with porous graphite carbon (PGC). O-Glycans were chemically released by B-elimination, ethanol precipitation followed again by solid phase extraction with PGC. Both N-glycans and O-glycans were then subjected to a PGC chip that was connected to a QTOF mass spectrometer. MS/MS fragmentation was performed in order to reliably identify the structures present in the samples. Numerous N-glycan structures were detected with an abundance of core-fucosylated and core-fucosylated bi-antennary structures (Fig. 4). We also identified several high-mannose-type N-glycan structures, basic core structures, and undecorated N-glycans. The N-glycan structures that were missing and typically present in

Rudolf Grimm

Intensity (counts)

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Acquision Time (min)

Fig. 4 LC-ESI-MS analysis of the N-glycan fraction prepared from the Iceman stomach sample

fresh human samples were complex N-glycan structures, tetraantennary N-glycans, terminally fucosylated N-glycans, and sialylated N-glycans. The absence of the latter structures in the stomach samples can be explained by the acid lability of them in the stomach environment plus the abundance of exosialidases present in gut bacteria. However, the cause of absence of terminally fucosylated N-glycan structures as well as complex and tetra-antennary N-glycans remains unknown. A total of four O-glycan structures were identified with three of them belonging to typical O-glycoprotein structures while the fourth one was a typical plant cell wall O-glycan structure (data not shown) which most likely may have derived from the grain pericarp found in the Iceman’s stomach [10]. Similar N-glycan data have been obtained in a different project [16] where a small skin sample from the Iceman’s left thigh was analyzed together with tissue samples from two Siberian ice mummies, the 2400-year-old Scythian warrior and the 2400-yearold Scythian ice princess. As reference, a fresh human skin sample was analyzed together with a tissue sample of a deceased human that was mummified over a period of several years. Sample preparation in this project was of major challenge as all the ancient mummy tissue samples had become very leathery. Finally, a successful extraction protocol was optimized using beef jerky. In the mummified tissue samples, again no sialylated or terminally fucosylated N-glycans were identified opposite to fresh human samples where all potential glycan structures were found. While the rapid dephosphorylation of phosphoproteins in human tissue postmortem has

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been discussed in literature [17], there are similarly no reports on what happens to glycosylation postmortem today. What we have learned from our studies is that N-glycans can persist over thousands of years and that the size and complexity of N-glycans are decreasing over time. Core-fucosylation seems to be very robust while terminal fucosylation and sialylation of N-glycan structures seem to be very unstable. The impact of the mummification process and general environmental factors on the fate of postmortem glycan structures are not understood yet. However, it is fair to state that there is a very different postmortem stability of the two major posttranslational modifications of proteins. 3.5

Metallomics

For metallomics studies, the stomach samples were subjected to elemental bio-imaging of frozen cross-sections as well as to flowinjection ICP-MS analysis of complete digests of 28 mg and 59 mg using 1% nitric acid. Elemental bio-images showed mostly Ca and Fe ions (Fig. 5a) with some Mn, Zn, and Cu concentrations that are typically found in a stomach after consumption of red meat, grains, or dairy products, indicating the Iceman’s last meal was well balanced in terms of essential minerals required for good health. Furthermore, no toxic elements such as lead, arsenic, or cadmium were detected. The elemental bio-imaging data were confirmed by flow-injection ICP-MS (Fig. 5b). Although the analytical data did not provide any evidence of whether free or bound metal ions were present in the samples, protein profiling and characterization data however identified several Fe- and Ca-binding proteins. Among the identified Fe-binding proteins were chymotrypsin-like elastase, adult ER calcium ATPase1, calmodulin-like protein, protein S100-A3, protein

Fig. 5 (a) Elemental bioimages of a small Iceman stomach cross-section showing calcium (Ca) and iron (Fe) content. (b) Flow-injection ICP-MS analysis of a completely digested Iceman stomach sample

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S100-A8, and protein S100-A9 while among the Ca-binding proteins ferritin, hemoglobin subunit A, and hemopexin were identified.

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Concluding Remarks A set of modern powerful mass spectrometry tools such as LC-MS, GC-MS, and ICP-MS have been applied from protein profiling and characterization to a complex multi-omics approach to solve ancient questions like this: What did a Neolithic meal look like? Now we know that it contained meat, fat, and cereals. We know that fire was somehow applied to preserve the meat, and some spices have been added to the meal to make it more palatable. After all, it was a fascinating and an exceptionally inspiring research project, which has allowed us to peer to some extent into his everyday life and technologies.

Acknowledgments First of all, the author would like to thank Albert Zink and Frank Maixner from the EURAC Institute of Mummies and the Iceman for providing the Iceman samples, supporting this project and conducting together with so many other wonderful colleagues at various institutes for all the DNA analyses as well as all the microscopy work done in context of this project. The author would also like to thank the entire incredible team that was involved in all the mass spectrometry analyses of this project: Ulrike Kusebauch, Michael Hoopman, Mark Sirtain, and Robert Moritz from the Institute of Systems Biology in Seattle; Oliver Fiehn, John Meissen, Mine Palozoglu, Sureyya Ozcan, Serenus Hua, and Carlito Lebrilla from UC Davis; Amaury Cazenave Gassiot and Markus Wenk from NUS Singapore; Bum Jin Kim and Hyun-Joo An from Chungnam National University in Daejeon/South Korea; and last but not least David Bishop and Philip Doble from the University of Technology Sydney/Australia. References 1. Yin L, Zhang Z, Liu Y, Gao Y, Gu J (2019) Recent advances in single-cell analysis by mass spectrometry. Analyst 144:824–845 2. Paynter E, Millar AH, Welch M, Baer-ImhoofB, Cao D, Baer B (2017) Insights into the molecular basis of long-term storage and survival of sperm in the honeybee (Apis mellifera). Nat Sci Rep 7:40236

3. Austin C, Smith TM, Bradman A, Hinde K, Joannes-Boyau R, Bishop D, Hare DJ, Doble P, Eskenazi B, Arora M (2013) Barium distributions in teeth reveal early-life dietary transitions in primates. Nature 498:216–219 4. Smith TA, Austin A, Green DR, JoannesBoyau R, Bailey S, Dumitriu D, Fallon S, Gru¨n R, James HF, Moncel MH, Williams IS,

Modern Mass Spectrometry Solving Ancient Questions Wood R, Arora M (2018) Wintertime stress, nursing, and lead exposure in Neanderthal children. Sci Adv 4:eaau9483 5. Joannes-Boyau R, Adams JW, Austin C, Arora M, Moffat I, Herries AIR, Tonge MP, Benazzi S, Evans AR, Kullmer O, Wroe S, Dosseto A, Fiorenza L (2019) Elemental signatures of Australopithecus africanus teeth reveal seasonal dietary stress. Nature 572:112–115 6. Altwegg K, Balsiger H, Bar-Nun A, Berthelier JJ, Bieler A, Bochsler P, Briois C, Calmonte U, Combi MR, Cottin H, De Keyser J, Dhooghe F, Fiethe B, Fuselier SA, Gasc S, Gombosi TI, Hansen KC, Haessig M, J€ackel A, Kopp E, Korth A, Le Roy L, Mall U, Mousis O, Owen T, Re`me H, Rubin M, Se´mon T, Tzou CY, Hunter Waite J, Wurz P (2016) Prebiotic chemicalsamino acid and phosphorus-in the coma of comet 67P/Churyumov-Gerasimenko. Sci Adv 2:e1600285 7. Popova OP, Jenniskens P, Emel’yanenko V, Kartashova A, Biryukov E, Khaibrakhmanov S, Shuvalov V, Rybnov Y, Dudorov A, Grokhovsky VI, Badyukov DD, Yin QZ, Gural PS, Albers J, Granvik M, Evers LG, Kuiper J, Kharlamov V, Solovyov A, Rusakov YS, Korotkiy S, Serdyuk I, Korochantsev AV, Larionov MY, Glazachev D, Mayer AE, Gisler G, Gladkovsky SV, Wimpenny J, Sanborn ME, Yamakawa A, Verosub KL, Rowland DJ, Roeske S, Botto NW, Friedrich JM, Zolensky ME, Le L, Ross D, Ziegler K, Nakamura T, Ahn I, Lee JI, Zhou Q, Li XH, Li QL, Liu Y, Tang GQ, Hiroi T, Sears D, Weinstein IA, Vokhmintsev AS, Ishchenko AV, Schmitt-Kopplin P, Hertkorn N, Nagao K, Haba MK, Komatsu M, Mikouchi T (2013) Chelyabinsk airburst, damage assessment, meteorite recovery, and characterization. Science 342:1069–1073 8. Williams AJ, Eigenbrode J, Floyd M, Wilhelm MB, O’Reilly S, Johnson SS, Craft KL, Knudson CA, Andrejkovicˇova´ S, Lewis JMT, Buch A, Glavin DP, Freissinet C, Williams RH, Szopa C, Millan M, Summons RE, McAdam A, Benison K, Navarro-Gonza´lez R, Malespin C, Mahaffy PR (2019) Recovery of fatty acids from mineralogic mars analogs by TMAH thermochemolysis for the sample analysis at Mars wet chemistry experiment on the curiosity rover. Astrobiology 19:522–546 9. Grimm R, Huber R, Neumeier T, Seidl A, Haslbeck M, Seibert F (2004) A rapid method for analysing recombinant protein inclusion bodies by mass spectrometry. Anal Biochem 330(1):140–144

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10. Maixner F, Turaev D, Cazenave-Gassiot A, Janko M, Krause-Kyora B, Hoopmann MR, Kusebauch U, Sartain M, Guerriero G, O’Sullivan N, Teasdale M, Cipollini G, Paladin A, Mattiangeli V, Samadelli M, Tecchiati U, Putzer A, Palazoglu M, Meissen J, Lo¨sch S, Rausch P, Baines JF, Kim BJ, An HJ, Gostner P, Egarter-Vigl E, Malfertheiner P, Keller A, Stark RW, Wenk M, Bishop D, Bradley DG, Fiehn O, Engstrand L, Moritz RL, Doble P, Franke A, Nebel A, Oeggl K, Rattei T, Grimm R, Zink A (2018) The Iceman’s last meal consisted of fat, wild meat, and cereals. Curr Biol 28:2348–2355 11. Oeggl K, Kofler W, Schmidt A, Dickson JH, Egarter-Vigl E, Gaber O (2007) The recon¨ tzi”, the struction of the last itinerary of “O Neolithic Iceman, by pollen analyses from sequentially sampled gut extracts. Quat Sci Rev 26:853–861 12. Maixner F, Krause-Kyora B, Turaev D, Herbig A, Hoopmann MR, Hallows JL, Kusebauch U, Egarter Vigl E, Malfertheiner P, Megraud F, O’Sullivan N, Cipollini G, Coia V, Samadelli M, Engstrand L, Linz B, Moritz R, Grimm R, Krause J, Nebel A, Moodley Y, Rattei T, Zink A (2016) The 5,300-year-old Helicobacter pylori genome of the Iceman. Science 351:162–165 13. Evershed RP, Payne S, Sherratt AG, Copley MS, Coolidge J, Urem-Kotsu D, Kotsakis K, Ozdog˘an M, Ozdog˘an AE, Nieuwenhuyse O, Akkermans PM, Bailey D, Andeescu RR, Campbell S, Farid S, Hodder I, Yalman N, Ozbas¸aran M, Bic¸akci E, Garfinkel Y, Levy T, Burton MM (2008) Earliest date for milk use in the Near East and southeastern Europe linked to cattle herding. Nature 455:528–531 14. Keller A, Graefen A, Ball M, Matzas M, Boisguerin V, Maixner F, Leidinger P, Backes C, Khairat R, Forster M, Stade B, Franke A, Mayer J, Spangler J, McLaughlin S, Shah M, Lee C, Harkins TT, Sartori A, Moreno-Estrada A, Henn B, Sikora M, Semino O, Chiaroni J, Rootsi S, Myres NM, Cabrera VM, Underhill PA, Bustamante CD, Vigl EE, Samadelli M, Cipollini G, Haas J, Katus H, O’Connor BD, Carlson MR, Meder B, Blin N, Meese E, Pusch CM, Zink A (2012) New insights into the Tyrolean Iceman’s origin and phenotype as inferred by whole-genome sequencing. Nat Commun 3:698 15. Murphy WA Jr, Dz N, Gostner P, Knapp R, Recheis W, Seidler H (2003) The iceman: discovery and imaging. Radiology 226:614–629

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16. Ozcan S, Kim BJ, Ro G, Kim JH, Bereuter T, Reiter C, Dimapasoc L, Garrido D, Mills DA, Grimm R, Lebrilla CB, An HY (2014) Glycosylated proteins preserved over millennia: N-glycan analysis of Tyrolean Iceman, Scythian Princess and Warrior. Nat Sci Rep 4:4963

17. Li J, Gould TD, Yuan P, Manji HK, Chen G (2003) Post-mortem interval effects on the phosphorylation of signaling proteins. Neuropsychopharmacology 28:1017–1025

Chapter 2 Step-by-Step Sample Preparation of Proteins for Mass Spectrometric Analysis Xinping Li, Thomas Franz, Ilian Atanassov, and Thomas Colby Abstract Nowadays identification and quantification of proteins from biological samples by mass spectrometry are widely used. For the identification of proteins, there are two scenarios. Proteins are either pre-fractionated in some way, e.g., by gel electrophoresis or chromatography, or analyzed as complex mixture (shotgun). Because of technological developments of mass spectrometry, the identification of several thousand proteins from complex biological matrix becomes possible. However, in many cases, it is still useful to separate proteins first in a gel. For quantifying proteins, label-free, isotopic labeling, and data-independent acquisition (DIA) library are widely used. Not only mass spectrometry technology made progress. This is also true for the sample preparation. Protocols and techniques developed recently not only make the analysis of starting material in the low microgram range possible but also simplify the whole procedure. Here, we will describe some detailed protocols of preparing samples for mass spectrometry-based protein identification and protein quantification, as in-gel digestion, in-solution digestion, peptide cleaning, and TMT labeling. This will allow also inexperienced beginners to get good results. Key words Sample preparation, Proteomics, In-solution digest, In-gel digest, FASP , StageTip, OASIS, TMT labeling, Mass spectrometry

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Introduction Mass spectrometry has become a valuable and indispensable tool in biological analysis and characterization of proteins. During the recent years, the sensitivity of the instruments improved dramatically, and this development is still ongoing. In addition, protein sample preparation for mass spectrometry has made very much progress toward handling of small samples and easiness. However, multi-step procedures are usually not suited for starting material in the low microgram range. The loss of material during sample preparation is substantial. Therefore, one-step procedures are usually preferred. Inexperienced scientific staff is facing the problem of old protocols in the literature, incomplete handling procedures, or protocols that are scattered throughout several publications,

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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making it not easy to get a comprehensive working protocol. The lack of important information or little “tricks” is mainly the reason for irreproducible experiments and results. In many relevant publications, the experimental part is mostly shortened in a way that it is impossible to reproduce the experimental procedure without prior knowledge. Very often, the protocols are written by people using the procedures on a daily basis, and handling of samples and techniques used are so obvious for them that little but important information is simply forgotten or not well communicated. In this chapter, we will disclose all details about high-throughput in-gel digestion [1], filter-aided sample preparation (FASP) [2, 3], one-step in-solution digestion [4], peptide cleaning [5], and TMT labeling [6]. We are using these protocols our self for daily work, but also in training courses for scientists and scientific staff.

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Materials Prepare all solutions and buffers using ultrapure water of 18 MΩ cm at 25
C, use LC-MS grade solvent and reagents of highest purity. Follow all waste disposal regulations when disposing waste materials.

2.1 High-Throughput In-Gel Digestion of Proteins 2.1.1 Excision of Protein Spots/Bands from Gels

1. OASIS®HLB μElution Plate, 30 μm for high-throughput (HTP) analysis (Waters). 2. Positive pressure-96 stand (PPS). 3. Scalpel or spot cutting tool, e.g., disposable pipet tip with 1.5 mm in diameter. 4. Acetonitrile (ACN). 5. Deep-well MTP waste plate, 24  10 mL.

2.1.2 Destaining of Gel Plugs

1. 100 mM ammonium bicarbonate (ABC), pH 8.0, in water. 2. Acetonitrile (ACN, HPLC grade). 3. 30 mM potassium ferricyanide K3[Fe(CN)6] in water. 4. 100 mM sodium thiosulfate in water.

2.1.3 Reduction, Alkylation, and Enzymatic Digestion

1. Reduction solution: 10 mM dithiothreitol (DTT) in 100 mM ABC, pH 8.0. 2. Alkylation solution: 55 mM iodoacetamide (IAA) in 100 mM ABC, pH 8.0. 3. Digestion solution: 7.4 ng/μL trypsin (MS grade) in 50 mM ABC, pH 8.0. 4. Digestion buffer: 50 mM ABC, pH 8.0, in water. 5. Incubator.

Sample Preparation for Mass Spectrometry 2.1.4 Peptide Cleaning and Extraction

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1. 0.1% formic acid (FA) for LC-MS sample preparation. 2. 0.1% trifluoroacetic acid (TFA) for MALDI-MS sample preparation. 3. Extraction solution for MALDI-MS: acetonitrile:water:trifluoroacetic acid (TFA) ¼ 49.95:49.95:0.1% (v/v/v). 4. Extraction solution for FA ¼ 47.5:47.5:5% (v/v/v).

LC-MS:

acetonitrile:water:

5. Vacuum centrifuge. 2.2 Filter-Aided Sample Preparation (FASP)

1. Lysis buffer: 2% SDS, 100 mM DTT in 100 mM Tris–HCl, pH 7.6. Add DDT always just prior use. 2. Buffer B: 8 M urea, 100 mM Tris–HCl, pH 8.0. 3. Buffer C: Add 10 mg of iodoacetamide to 1 mL of buffer B. 4. Buffer D: 20 mM Tris–HCl, pH 8.5, 10% ACN. 5. 1 μg/μL (w/v) Trypsin (MS grade) in 50 mM acetic acid. 6. Diagenode Bioruptor® plus connected to a water cooler. 7. Nanosep® Centrifugal Devices with Omega™ Membrane 30K. 8. Electric mortar and pestle grinder. 9. High-speed centrifuge for 1.5 mL reaction tubes. 10. Incubator. 11. Vacuum centrifuge. 12. NanoDrop 2000 (Thermo Scientific).

2.3 One-Step Guanidine Method for Protein Lysis and Digestion

1. Lysis buffer: 6 M guanidine hydrochloride (GdmCL), 10 mM Tris-(2-carboxyethyl)phosphine hydrochloride (TCEP), 40 mM 2-chloroaceteamide (CAA), 100 mM Tris–HCl, pH 8.5 (see Note 1). 2. Thermomixer. 3. High-speed centrifuge. 4. Diagenode Bioruptor® plus connected to a water cooler. 5. Sample dilution buffer: 20 mM Tris–HCl, pH 8.5. 6. 1 μg/μL (w/v) Trypsin (MS grade) in 50 mM acetic acid. 7. NanoDrop 2000 (Thermo Scientific). 8. Incubator. 9. Stop solution: 50% (v/v) formic acid.

2.4 StageTip Peptide Cleaning and Desalting

1. Solution A: 100% MeOH. 2. Solution B: 0.1% (v/v) FA in 60% (v/v) ACN. 3. Solution C: 0.1% (v/v) FA.

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Fig. 1 Spring-loaded syringes for the preparation of StageTips with different binding capacities

Fig. 2 Centrifuge adapters for 200 μL tips and its assembly with a 2 mL tube

4. Empore C18-SD solid phase extraction disk for StageTip production (see steps 1 and 2 in Subheading 3.4). 5. Syringes and syringe needles with inner diameter of 1.0, 1.5, and 2.0 mm (Fig. 1). 6. Tommy benchtop centrifuge or pipette tip adaptors (Fig. 2). 7. NanoDrop 2000 (Thermo Scientific). 8. High-speed centrifuge. 9. Vacuum centrifuge.

Sample Preparation for Mass Spectrometry

2.5 TMT (Tandem Mass Tag) Labeling of Peptides

17

1. TMT10plex™ Isobaric Label Reagent Set, 1  0.8 mg. 2. LC-MS grade water. 3. LC-MS grade anhydrous acetonitrile. 4. 100 mM triethylammonium bicarbonate (TEAB). 5. 5% (v/v) hydroxylamine. 6. 0.1% (v/v) FA. 7. Vortex mixer. 8. Vacuum centrifuge. 9. Tabletop centrifuge.

3

Methods

3.1 HTP In-Gel Digestion of Proteins [1] 3.1.1 Excision of Protein Spots/Bands from Gels

1. With a scalpel, excise a protein band of interest from the stained gel, place it at the upper rim of an OASIS® MTP well (see Note 2) and cut it in 1 mm3 pieces. Then push them down into the well by a pipette tip. For a 2D-gel, use a spot cutter to excise the gel plugs and transfer directly into the OASIS® MTP well. 2. Remove the liquid from the excision process into the deep-well waste plate using the positive pressure-96 stand (PPS). 3. Add 70 μL acetonitrile (ACN) and push the gel pieces down into the solution if necessary. 4. Wait for 10–15 min until they have dehydrated, they will become small and white (see Note 3). 5. Use the PPS to remove all liquid into waste plate.

3.1.2 Destaining of Gel Plugs

For destaining of Coomassie gel plugs, follow steps 1–4. The destaining of silver-stained gel plugs is described in steps 5–13. 1. Rehydrate Coomassie stained gel pieces in 100 μL of 100 mM ABC solution. 2. After 10 min, add an equal volume of ACN and wait for 10–15 min until the gel pieces have destained (see Note 4). 3. Remove all liquid using the PPS and add 100 μL ACN for 10 min to dehydrate the gel pieces. 4. Remove all liquid using the PPS. 5. Rehydrate silver-stained gel pieces with 50 μL potassium ferricyanide solution and incubate for 10 min at 37
C. 6. Add 50 μL sodium thiosulfate solution and incubate for 20 min at 37
C. 7. After 15 min, remove the liquid. 8. Add 50 μL water and after 10 min, 50 μL ACN.

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9. Remove all the liquid after 15 min. 10. Add 70 μL ABC solution and incubate for 10 min. 11. Remove all the liquid after 15 min. 12. Add 100 μL ACN for 10 min to dehydrate the gel pieces (see Note 3). 13. Remove ACN by PPS. 3.1.3 Reduction, Alkylation, and Enzymatic Digestion

1. Rehydrate gel pieces in 100 μL reduction solution and incubate for 1 h at room temperature to reduce the cysteine bridges in the protein (see Note 5). 2. Remove excess liquid using the PPS. 3. Incubate the gel pieces with 100 μL ACN and wait for 10–15 min until the gel pieces have dehydrated (see Note 3). 4. Remove all liquid with the PPS. 5. Swell gel pieces with 100 μL alkylation solution and incubate for 20 min at room temperature in the dark. 6. Remove alkylation solution with the PPS and wash gel pieces with 100 μL ABC solution for 15 min. 7. Remove ABC solution with the PPS, add 100 μL ACN, and wait for 10 min for the gel pieces to dehydrate and shrink. 8. Remove all liquid with the PPS and then rehydrate the gel pieces in 33 μL digestion solution at room temperature. 9. After 20 min, remove the remaining buffer with the PPS. 10. Add 50 μL of digestion buffer (without trypsin) to cover the gel pieces and keep them wet during enzymatic digestion. 11. Leave samples covered at room temperature overnight or if available in an incubator at 37
C.

3.1.4 Peptide Cleaning and Extraction

1. With the PPS slowly remove the digestion solution into the waste plate (peptides will bind to the HLB sorbent in the OASIS plate) (see Note 6). 2. Add 70 μL of 0.1% FA to the OASIS® HLB column and slowly remove the liquid (see Note 6). 3. Exchange the waste plate for the collection plate (0.5 mL/well) (see Note 7). 4. Add 50 μL of extraction solution (for LC-MS) to each well of the OASIS plate and wait for 20 min, then slowly transfer the extraction solution into a collection plate with the PPS. 5. Add 50 μL of acetonitrile to each well and wait for 10 min, then slowly transfer the extraction solution into collection plate with the PPS.

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6. Evaporate the eluted peptide solution in the collection plate using a vacuum centrifuge at 45
C for 60 min. 7. Store the dried peptide samples at 20 or 80
C (for storage >1 month). 3.2 Filter-Aided Sample Preparation (FASP) [2, 3]

FASP is useful if the sample lysis buffer contains SDS. The total protein amount of the starting material should be above 50 μg. Particular attention is drawn to possible filter leakage of the membrane and the total loss of sample during the procedure (see Note 8). 1. 1.5 mL Eppendorf tube containing crude sample (cell pellet, tissue, or similar) containing about 150 μg protein. 2. Heat 1 mL of lysis buffer to 95
C (2 min) and then add 100 μL to the sample. 3. With an electric mortar, homogenize the sample for 2 min. 4. Then heat for 3 min at 95
C. 5. For further homogenization at 4
C, use the Diagenode Bioruptor®. Set 15 cycles each 30 s sonication; 30 s break; position: high performance. 6. Centrifuge the sample at 16,000  g for 10 min. 7. Add 200 μL of buffer B to a Nanosep 30k Omega Centrifugal Device followed by 50 μL (about 75 μg protein) of the supernatant of the homogenized sample. 8. Centrifuge at 12,000  g for 15 min. 9. Collect the flow-through (in case filter was leaking) and add 250 μL of buffer B to the residue in the filter. 10. Centrifuge again at 12,000  g for 15 min. 11. Add 250 μL of buffer B and centrifuge again at 12,000  g for 15 min. 12. Add 300 μL of buffer C, mix with a pipette carefully, and wait for 20 min at RT in the dark (see Note 9). 13. Centrifuge at 12,000  g for 15 min. 14. Add 250 μL of buffer B and centrifuge again (12,000  g for 15 min). 15. Add 200 μL of buffer D and centrifuge (12,000  g for 15 min). Repeat this step once more. 16. Add 50 μL of buffer D to the sample followed by 3 μg of trypsin and mix carefully by pipetting up and down. 17. Close the lid of the centrifugal device, place the complete device in a floating foam tube rack. Fill a box with water, put the floating rack in the box and close the box. Put the box into an incubator at 37
C overnight.

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18. Following the digest, add 200 μL buffer D and centrifuge at 10,000  g for 15 min. 19. Place the tube with the flow-through in a vacuum centrifuge for 30 min at 45
C and concentrate the solution to a final volume of 100 μL. 20. Take 2 μL of the digested sample for a NanoDrop OD 260/280 measurement with water as reference (see Note 10). 21. Perform StageTip cleaning as described in Subheading 3.4. 3.3 One-Step “Guanidine” Method for Protein Lysis and Digestion

1. Add lysis buffer to a sample pellet, e.g., a protein or cell pellet. The volume of the lysis buffer should exceed the volume of the sample pellet by a factor of 2. 2. Incubate the sample solution in a Thermomixer at 95
C for 10 min at 600 rpm. 3. Homogenize sample with Bioruptor at 4
C: 30 s sonication, 30 s off, 10 cycles, position: high performance. 4. If necessary, repeat steps 2 and 3. 5. Centrifuge the sample solution at 20,000  g for 20 min at room temperature. 6. Keep the supernatant. 7. Take 2 μL of the supernatant, dilute ten times with sample dilution buffer. 8. Measure the concentration of the protein solution with NanoDrop 2000 [7]. 9. According to the measured concentration, take some μL of the supernatant corresponding to 100–300 μg of protein, dilute ten times with sample dilution buffer to reduce the concentration of Guanidine to 0.6 M, add trypsin, and digest at 37
C overnight (see Note 11). Use a trypsin/protein (w/w) ratio of 1/200 (see Note 12). 10. Acidify the protein digest by the addition of stop solution to a final concentration of 1% FA to stop the digestion process. 11. Centrifuge the digest at 15,000  g for 10 min and retain the supernatant. 12. Perform StageTip cleaning as described in Subheading 3.4.

3.4 StageTip Peptide Cleaning and Desalting

1. Overlay three pieces of EmporeTM C18-SD solid phase extraction disk for StageTip production. 2. Punch out a plug with a syringe needle and press the plug into the narrow end of a 200 μL pipette tip (see Note 13 and Fig. 3). Make sure that the plug is stuck.

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21

Fig. 3 The preparation of StageTips

3. Wash the tips with 200 μL of solution A, centrifuge at 3000  g for 1–2 min to remove the liquid. Leave 1–2 mm of the liquid on the top. 4. Wet the tips with 200 μL of solution B, centrifuge at 3000  g for 1–2 min to remove the liquid. Leave 1–2 mm of the liquid on the top. 5. Equilibrate the tips with 200 μL of solution C, centrifuge at 3000  g for 2 min to remove the liquid. Leave 1–2 mm of the liquid on the top. 6. Load the protein digest to the bottom of the StageTips. Centrifuge for 2–2.5 min at 3000  g. 7. Apply 200 μL of solution C to the StageTips and centrifuge at 3000  g for 2–2.5 min until the liquid goes through. 8. Repeat step 7. 9. Apply 80–100 μL of solution B and elute the peptides by centrifugation at 1500  g for 4–8 min. Collect the eluate (see Note 14). 10. Transfer the eluates to 500 μL tubes. 11. Dry the digest completely in a vacuum centrifuge at 45
C for 40–45 min. Store sample at 20
C. 3.5 Modified TMT Labeling of Peptides [6]

With the TMT-10plex isobaric mass tag labeling set, ten peptide samples can be labeled and analyzed simultaneously. Additionally, due to the co-isolation and fragmentation of the isobaric precursor ions, there are much less missing values compared to label-free quantification leave techniques [8]. 1. Adjust TMT vials (stored at 20
C) at room temperature for 10–15 min.

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2. Add 70 μL of anhydrous acetonitrile to the TMT vials and leave at room temperature for 10 min with occasional vortexing. 3. Dissolve 4 μg of dried and cleaned peptides (after StageTiping) in 9 μL of 100 mM TEAB solution, vortex and briefly spin down. 4. Aliquot 7 μL of each TMT reagent to each peptide tube, vortex, and briefly spin down. 5. After 60 min, stop the reaction by adding 2 μL of 5% hydroxylamine solution. Quenching takes about 15 min. 6. Pool all peptide samples in a new tube and dry in a vacuumcentrifuge for 1 h at 45
C. 7. Resuspend the dried peptides with 200 μL of 0.1% formic acid solution. 8. Split the 200 μL of sample into two equal parts. Using a StageTip of 45 μg binding capacity, clean each part as described in Subheading 3.4. 9. Dry the cleaned peptide and store at 20
C. One part is for further mass spectrometric analysis, the other is for backup.

4

Notes 1. It is important to prepare the Guanidine/TCEP/CAA/Tris lysis buffer always fresh, because the components react with each other over time. 2. Put the excised gel spots/bands directly into the OASIS® HLB μElution Plate. The OASIS platform allows the use of an eightchannel pipette to add solutions, and with the help of the positive pressure-96 stand, the solutions are then simultaneously removed. Thus, one can make a manual digest of 96 samples in the same time and of the same quality that one could typically digest only 12 samples manually using standard 0.5 mL tubes. The in-gel digestion protocol for protein spots contains six steps: excision of spots/bands, destaining, reducing, alkylation, digestion, and peptide extraction. 3. It happens that acetonitrile runs through the HLB column without pressure. In this case, just add acetonitrile again after 5 min. 4. If gel pieces are not destained, repeat the steps. Do not destain more than twice as it will not improve the results. 5. The reaction can be shortened to 30 min by incubating at 37
C. 6. First use 2 psi of nitrogen pressure and wait for 1 min. Then increase to 9 psi to remove the digestion solution completely.

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7. The PPS stand requires a certain height of the collection plate. Use suitable 96-well collection plates as spacers. 8. Some filters may even leak from beginning. To test this, add 200 μL of buffer B to a couple of filters and centrifuge at 12,000  g for 5 min. Compare the amount of liquid left in the filter. Discard the filters with no or significantly less liquid. It is not unusual that during the procedure the filter starts leaking too. 9. DTT for reduction is already in the lysis buffer. 10. The peptide concentration is used to calculate how much one needs to take for StageTip desalting and cleaning for MS analysis. 11. Dilute at least 1:10, otherwise trypsin digest will not work. 12. For SILAC samples with only lysine label, use LysC. 13. StageTips can be produced with different capacities. To produce a 10 μg binding capacity tip, three layers of 3M Empore™ C18-SD solid phase extraction disk and 1 mm inner diameter syringe are needed. For an inner diameter of 1.5 or 2.0 mm, the binding capacities are about 30, 45 μg, respectively (Fig. 1). For larger quantities of protein up to 150 μg, commercial 4 mm (1 mL) Empore™ C18-SD solid phase extraction cartridges from 3M are available. 14. For centrifuging the StageTips, a 2 mL reaction tube with a custom made adapter (Fig. 2) was used to hold the tip. Alternatively, a TOMY “StageTip” centrifuge (Sonation GmbH, Germany) can be used. References 1. Franz T, Li X (2012) The OASIS® HLB μElution plate as a one-step platform for manual high-throughput in-gel digestion of proteins and peptide desalting. Proteomics 12:2487–2492 2. Manza LL, Stamer SL, Ham AJ, Codreanu SG et al (2005) Sample preparation and digestion for proteomic analyses using spin filters. Proteomics 5:1742–1745 3. Wisniewski RJ, Zougman A, Nagaraj N, Mann M (2009) Universal sample preparation method for proteome analysis. Nat Methods 6:359–362 4. Kulak NA, Pichler G, Paron I, Nagaraj N, Mann M (2014) Minimal, encapsulated proteomicsample processing applied to copy-number estimation in eukaryotic cells. Nat Methods 11:319–324 5. Rappsilber J, Ishihama Y, Mann M (2003) Stop and go extraction tips for matrix-assisted laser

desorption/ionization, nanoelectrospray, and LC/MS sample pretreatment in proteomics. Anal Chem 75:663–670 6. Thompson A, Schafer J, Kuhn K, Kienle S, Schwarz J, Schmidt G, Neumann T, Johnstone R, Mohammed AK, Hamon C (2003) Tandem mass tags: a novel quantification strategy for comparative analysis of complex protein mixtures by MS/MS. Anal Chem 75:1895–1904 7. Desjardins P, Hansen JB, Allen M (2009) Microvolume protein concentration determination using the NanoDrop 2000c spectrophotometer. J Vis Exp 33:1610 8. O’Connell JD, Paulo JA, O’Brien JJ, Gygi SP (2018) Proteome-wide evaluation of two common protein quantification methods. J Proteome Res 17:1934–1942

Chapter 3 Lab-on-a-Filter Techniques for Economical, Effective, and Flexible Proteome Analysis Yi-Han Lin, Rodrigo Vargas Eguez, Isha Vashee, and Yanbao Yu Abstract Effective and reliable protease digestion of biological samples is critical to the success in bottom-up proteomics analysis. Various filter-based approaches using different types of membranes have been developed in the past several years and largely implemented in sample preparations for modern proteomics. However, these approaches rely heavily on commercial filter products, which are not only costly but also limited in membrane options. Here, we present a plug-and-play device for filter assembly and protease digestion. The device can accommodate a variety of membrane types, can be packed in-house with minimal difficulty, and is extremely cost-effective and reliable. Our protocol offers a versatile platform for general proteome analyses and clinical mass spectrometry. Key words Filter-aided sample preparation (FASP), Suspension trapping (STrap), Polyvinylidene fluoride (PVDF), Polyethersulfone (PES), Glass fiber, Membrane, Shotgun proteomics, Urine, Saliva, Tissue

1

Introduction Substantial investments have been made in the past decades aiming to develop robust and easy-to-use technologies for mass spectrometry (MS)-based proteomic analysis of biospecimens, tissues, and in vitro cell cultures. One of the methods commonly used in protease digestion in modern proteomics is the filter-based sample processing technique. Including the classical filter-aided sample preparation (FASP) approach using cutoff membranes and derivatives such as the use of PVDF membranes and glass fiber [1–4], filter-based sample processing techniques have been greatly implemented in proteomic researches and biomarker discoveries. However, although largely applied by the proteomic community, the available products in the market not only are expensive but also have limited flexibility to adjust for the development of novel technologies [1, 2, 5–7].

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Design of the lab-on-a-filter platform. With a self-packable filter device, membranes on the user’s own choice can be purchased separately in bulk, cut, and packed. A variety of membranes with different functionalities can be selected, such as glass microfiber, molecular weight cutoff (e.g., regenerated cellulose, polyethersulfone), binding-based (e.g., polyvinylidene fluoride, PVDF), or mixed-mode (e.g., glass fiber and C18, PVDF, and cutoff). This results in a versatile reactor for general proteomic sample preparation

We have previously demonstrated the usage of an in-house packed glass fiber filter for the analysis of global HeLa cell proteome and salivary biomarker discovery [8]. We benchmarked the device’s high reliability, reproducibility, and cost-effectiveness. In an effort to lower the barriers for applying filter-based techniques to a wide range of biomedical problems, here in this chapter, we attempt to further expand the capacity and flexibility of this method by packing a variety of membranes with different chemistries and functionalities (e.g., cutoff, binding, enrichment, and fractionation), thus resulting in a lab-on-a-filter platform that will provide unprecedent convenience for proteomic analysis (Fig. 1). This platform can accommodate membranes of one’s choice and can be applied to various biospecimen sample processing (e.g., urine, saliva, tissue, cells). The capability of packing one’s own filter drastically reduces the cost, especially when large number of samples need to be processed, as in the case of clinical proteomics. In addition, it offers an opportunity to the design or employment of unique filter types currently unavailable in the market and the emerging techniques. The high adaptability of our method also leverages the existing filter-based technologies and opens a new window for proteomic applications.

2

Materials

2.1 Reagents and Equipment

1. PBS buffer: Phosphate-buffered saline with protease inhibitor included (see Note 1). 2. SDS buffer: 4% SDS, 10 mM EDTA, 0.05% Tween-20, 100 mM Tris–HCl, pH 8.0. Add fresh DTT (20 mM) before use.

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3. Tris buffer: 1 M Tris–HCl, pH 8.0. 4. Amicon Ultra-15 Centrifugal Filter Units (10,000 molecular weight cutoff, MWCO). 5. Alkylation reagent: 0.5 M iodoacetamide. 6. Sequencing Grade Modified Trypsin at a stock concentration of 0.1 μg/μL. 7. Bead beater. 8. Bead tube with 2.8 mm ceramic beads. 9. Benchtop centrifuge. 10. Temperature-controlled shaking incubator. 11. Sonicator. 12. Filter device: From Microcon (Millipore) or Vivacon (Sartorius) filters, disassemble the plastic components including a tube, an O-ring, and a base (Fig. 2a). New collection tubes can be purchased separately. Alternatively, an “O-ring free” filter can be used by disassembling Thomson filter vials (Fig. 2b). 13. Membranes for packing: (a) Glass fiber membrane (e.g., 0.7, 1.6 μm). (b) Regenerated cellulose (RC) membrane, 30,000 MWCO. (c) Polyethersulfone (PES) 300,000 MWCO.

membrane,

30,000

and

Fig. 2 Components of self-packable filter device. (a) Three major components to assemble a filter include a base, an O-ring, and a tube. These can be reused after simple washes. New collection tubes can be purchased separately. Membrane discs can be cut from a large membrane sheet using a 10/3200 hole puncher. (b) A plugand-play filter without O-ring. 2-mL microtubes can be used as collection tubes and 9/3200 hole puncher is used to cut membrane discs

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(d) Polyvinylidene fluoride (PVDF) membrane, 0.2 or 0.45 μm. (e) Mixed mode membranes: PVDF and 30,000 MWCO; glass fiber and C18. 14. Hole puncher, 10/3200 or 9/3200 . 15. Collection tubes. 16. SpeedVac. 2.2 Suspension Trapping (STrap) Digestion-Specific Reagents

1. SDS lysis solution: 10% SDS in 100 mM triethylammonium bicarbonate (TEAB), pH 7.5; add 25 mM DTT before use. 2. Binding solution: 90% methanol in 100 mM TEAB, pH 7.1 (see Note 2). 3. Neutralization solution: 12% phosphoric acid. 4. Elution solution I: 50 mM TEAB, pH 8. 5. Elution solution II: 0.2% formic acid in H2O. 6. Elution solution III: 0.2% formic acid in 50% acetonitrile/50% H2O.

2.3 Filter-Aided Sample Preparation (FASP) DigestionSpecific Reagents

1. Prewash solution for PVDF mix-mode membrane: 50% ethanol. 2. Urea lysis solution: 8 M urea in 50 mM Tris–HCl, pH 8; prepare freshly. 3. Digestion solution: 50 mM ammonium bicarbonate (ABC) in H2O. 4. Elution solution for PVDF mix-mode membrane: 0.1% formic acid in 40% acetonitrile.

2.4 PVDF Membrane Digestion-Specific Reagents

1. Activation solution: 100% methanol. 2. Prewash solution: 70% ethanol. 3. Urea lysis solution: 8 M urea in 50 mM Tris–HCl, pH 8; prepare freshly. 4. Digestion solution: 50 mM ABC in H2O. 5. Elution solution: 0.1% formic acid in 40% acetonitrile.

3

Methods

3.1 Protein Preparation from Different Sources

1. From cell culture: Resuspend cell pellet in PBS buffer. Add SDS buffer at a 1:1 ratio and sonicate the cell solution for 3 min (pulse: 30 s on, 30 s off). After sonication, incubate the sample at 95  C for 10 min. Cool the sample to room temperature. Obtain clear cell lysate by centrifuging at 16,100  g for 20–30 min.

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2. From whole tissue: Dissect tissue and place in the bead tube with 2.8 mm ceramic beads (see Note 3). Add PBS buffer and place the tube in the bead beater (homogenizing speed: 3450 oscillations/min) for a beating cycle of 15–30 s depending on the size of the tissue. Centrifuge the tube at 6000  g at 4  C for 10 min, collect the supernatant (PBS lysate). Resuspend the pellet with the same amount of SDS buffer and sonicate for 3 min (pulse: 30 s on, 30 s off). Centrifuge at 16,100  g for 20–30 min and collect the supernatant (SDS lysate). Combine the PBS lysate and the SDS lysate at a 1:1 ratio and incubate at 95  C for 5–10 min. Cool the sample to room temperature and spin down the liquid condensed on the cap. 3. From urine: Urine specimens are added with Tris buffer to a final concentration of 50 mM to neutralize the specimens. Sediment can be removed by centrifuging the specimens at 1350  g for 15 min. Use Amicon Ultra-15 Centrifugal Filter Units (10,000 MWCO) to concentrate soluble urinary proteins. 4. From saliva: Centrifuge raw saliva at 2600  g for 20 min at 4  C to remove residual food and cell debris. Collect the supernatant and add protease inhibitor to prevent protein degradation. 5. For STrap digestion of urine and saliva samples, add SDS lysis solution at a 1:1 ratio and incubate at 95  C for 10 min. Cool the sample to room temperature and spin down the liquid condensed on the cap. 6. Determine protein concentration in cell lysate, tissue homogenate, or body fluids using SDS-PAGE as described previously [9]. 7. Take the desired amount of protein (see Note 4), alkylate free cysteines by adding alkylation reagent to 50 mM, and incubate in dark for 20–30 min. 3.2

STrap Digestion

1. Use the 10/3200 hole puncher to cut discs from the glass microfiber membrane sheet or disc. The analysis in Fig. 3 was accomplished using Whatman GF/F membrane (pore size 0.7 μm). Use a tweezer to pack one or two layers of glass fiber discs in the filter base, assemble the O-ring, and snap on the sample tube (see Note 5). 2. Transfer the filter device onto a collection tube and flush the filter disc with 300 μL binding solution by centrifugation at 400  g for 1 min, discard the flow-through. 3. Add neutralization solution to 1.2% to the protein sample from step 7 in Subheading 3.1, mix well, and incubate for 1–2 min at room temperature (see Note 6).

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Fig. 3 Comparison of performance for different types of membrane evaluated in this chapter. (a) Time duration for sample preparation using different membranes. (b) Comparison of the number of proteins identified. GF/F, Whatman glass microfiber membrane (pore size, 0.7 μm). YM-30, regenerated cellulose membrane (Millipore). PES-30K and 300K, polyethersulfone (Synder Filtration), PVDF-30K (Synder Filtration). PVDF, 0.2 and 0.45 μm (Millipore). (c) Comparison of peptide identification. PSMs peptide spectrum matches. (d) Overlapping analysis of protein-level identifications. Three replicate experiments were performed for each membrane

4. Add six volumes of binding solution, mix well, and incubate for 1–2 min (see Note 7). 5. Load the sample onto the STrap filter device, centrifuge at 400  g for 2 min or until all the liquid has been spun down to the collection tube (see Note 8). Discard flow-through. 6. Wash the protein particulate on the filter by 200 μL binding solution and centrifuge at 400  g for 2 min or until all the liquid has been spun down, discard the flow-through. Repeat the wash for two more times. 7. Transfer the filter device to a clean collection tube. Add trypsin at a 1:30–1:100 (wt/wt) ratio prepared in 150 μL 50 mM TEAB, pH 8. Quick spin for 1–2 s to ensure the filter has been immersed with the trypsin solution. Transfer the liquid in the collection tube back to the top of the filter.

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8. Incubate the tube at 37  C overnight (see Note 9). 9. Next day, without opening the cap, centrifuge the tube at 400  g for 2 min to collect digested peptides in the collection tube, add 200 μL Elution buffer I and centrifuge, repeat one more time, and transfer the eluent to a clean tube. Add 200 μL Elution buffer II and centrifuge. Add 200 μL Elution buffer III and centrifuge. Pool all eluents together. 10. Dry the combined eluent solution on a SpeedVac. 11. Desalt peptides using C18 StageTip as described previously [9], dry and store in 80  C until further use (or send directly to LC-MS/MS analysis). 3.3

FASP Digestion

1. Similar to step 1 in Subheading 3.2, use the hole puncher to cut discs from RC, PES, or PVDF mix-mode membranes. Pack one layer of the membrane disc in the filter device. Place the filter device onto a collection tube. 2. For the RC and PES membrane, wash the membrane by adding 200 μL H2O and centrifuge at 10,000  g for 15 min or until the remaining solution on top is less than 20 μL (see Note 10). Repeat H2O wash one more time. Add 200 μL urea lysis solution and centrifuge, repeat one more time. Discard all flow-throughs (see Note 11). 3. For the PVDF mix-mode membrane (30,000 MWCO), add 200 μL prewash solution and centrifuge at 10,000  g for 15 min or until the remaining solution on top is less than 20 μL. Repeat wash for two more times, followed by the H2O and urea lysis solution wash as described in step 2. Discard all flow-throughs. 4. Load protein aliquots from step 7 in Subheading 3.1 to the membrane device, centrifuge at 10,000  g for 15 min or until the remaining solution on top is less than 20 μL. Discard the flow-through. Add 200 μL urea lysis solution to the top and pipet up and down for thorough mixing. Centrifuge at 10,000  g for 15 min or until the remaining solution on top is less than 20 μL. Repeat the addition of 200 μL urea lysis solution and centrifugation one more time. Discard the flowthrough. 5. Add 200 μL digestion solution and centrifuge, repeat one more time. 6. Transfer the filter device to a clean collection tube. Add trypsin at a 1:30–1:100 (wt/wt) ratio prepared in 150 μL digestion solution. Incubate the filter device overnight at 37  C with 650 rpm shaking. 7. Next day, centrifuge the tube at 10,000  g for 10 min to collect digested peptides. For the RC and PES membrane, add

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200 μL digestion solution and centrifuge until the remaining solution on top is less than 20 μL, repeat for a total of three times and collect eluents in a clean tube. For the PVDF mix-mode membrane, perform elution for two times with 200 μL digestion solution, followed by elution with 200 μL elution solution for two times. Combine eluents in a clean tube and dry the peptides on a SpeedVac. 8. Desalt peptides using C18 StageTip as described previously [9]. 3.4 PVDF Membrane Digestion

1. Pack one layer of PVDF membrane in the filter device as described in step 1 in Subheading 3.2. Transfer the filter device onto a collection tube. 2. Activate the PVDF membrane by adding 200 μL activation solution and centrifuge at 2000  g for 1–2 min. Add 200 μL prewash solution and centrifuge. Add 200 μL urea lysis solution and centrifuge, repeat one more time. Transfer the filter device to a new collection tube. 3. Mix the protein aliquot from step 7 in Subheading 3.1 with 200 μL urea lysis solution. Load the solution onto the PVDF filter and centrifuge at 2000  g for 1–2 min. Take the flowthrough and reload onto the filter. Repeat the loading for a total of three times. Discard the final flow-through. 4. Add 200 μL urea lysis solution and centrifuge at 2000  g for 1–2 min. Repeat one more time. 5. Add 200 μL digestion solution and centrifuge. Repeat one more time. 6. Transfer the filter device to a new collection tube. Add trypsin at a 1:30–1:100 (wt/wt) ratio prepared in 150 μL digestion solution. Incubate the filter device overnight at 37  C with gentle shaking. 7. Next day, centrifuge the tube at 2000  g for 1–2 min to collect digested peptides in the collection tube. Add 200 μL digestion solution and centrifuge, repeat one more time and collect eluents in a clean tube. Add 200 μL elution solution to the filter and centrifuge, repeat one more time. Combine eluents and dry the peptides on a SpeedVac. Desalt peptides as described previously [9].

3.5 Performance Analysis

1. Figure 3 illustrates the time effectiveness and identification performance of each membrane tested in this chapter. Glass fiber-based digestion (i.e., STrap) is the method with the least centrifugation time (Fig. 3a). PVDF membrane–based digestion is also faster than cutoff membranes; however, the protein and peptide identification rates are relatively low (Fig. 3b, c).

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2. Interestingly, complementary identification of proteins can clearly be seen between different types of membrane (Fig. 3d), suggesting that more than one method can be employed if in-depth proteome coverage is desired. Overall, our method implicates the value using the lab-on-a-filter platform for flexible proteomics analysis.

4

Notes 1. Protease inhibitor needs to be included in the PBS buffer used for cell or tissue resuspension. If phosphopeptide enrichment will be performed downstream, phosphatase inhibitor will also need to be included. 2. To prepare 100 mL binding solution, add 90 mL methanol, 10 mL 1 M TEAB, and adjust the pH by adding ~471 μL 37% HCl. 3. Select the material and size of bead according to manufacturer’s suggestions. In this protocol, 2.8 mm ceramic beads were used to homogenize mouse lung, kidney, and liver tissues. 4. In our experience, input protein between 30 and 100 μg yields a decent protein coverage for global proteome analysis. 5. Make sure the O-ring is positioned all around the filter base, and visually inspect if the O-ring has been evenly sandwiched between the tube and the filter disc after snap-on. If the O-ring is distorted or out of position, the device needs to be disassembled and readjusted to prevent sample leakage during centrifugation. 6. Vortex the tube or pipet the solution up and down to ensure thorough mixing of phosphoric acid to the solution. 7. Solution should turn cloudy after adding the binding solution. Vortex the tube or pipet the solution up and down to ensure thorough mixing and to avoid the formation of large clumps of protein particulate. 8. In our experience, two layers of GF/F glass fiber in the STrap device can accommodate up to 500 μg of proteins without sample loss. 9. Shaking is not required for this step, but if large clump of protein is present on the filter, shaking (at 650 rpm) is suggested for efficient protease digestion. 10. Centrifugation time needed for PES 300,000 MWCO membrane is shorter than 30,000 MWCO (Fig. 3a). Adjust centrifugation time according to the amount of solution remaining on top (98%. 3. Working reagent (WR): 100 μL for each standard/sample. Mix 25 parts Reagent A, 24 parts Reagent B, and 1 part Reagent C. 4. 1% (w/v) SDS in ultrapure water. 5. Exosome or cell lysate sample for measurement. 6. 1.5 mL microcentrifuge tubes. 7. Flat-bottom, clear 96-well plates. 8. Incubator (37  C). 9. Microplate reader with 562 nm filter.

2.3.2 Protein Staining Densitometry

1. 2
SDS sample buffer: 4% (w/v) SDS, 20% (v/v) glycerol, 0.01% (v/v) bromophenol blue, 125 mM Tris–HCl, pH 6.8. If reducing conditions are required, supplement 2
SDS sample buffer with dithiothreitol (DTT) to a final concentration of 2% (w/v). 2. Exosome or cell lysate sample for measurement. 3. NuPAGE Bis-Tris Precast gels: 1 mm thickness, 10- or 12-well, 4–12% gradient. 4. 1
MES running buffer: 50 mM MES, 50 mM Tris base, 0.1% SDS, 1 mM EDTA, pH 7.3. Do not use acid or base to adjust the pH. 5. XCell Surelock™ gel tank, with compatible power supply (e.g., Bio-Rad Laboratories, Hoefer, Thermo Fisher Scientific). 6. BenchMark™ Protein Ladder standard of known protein concentration: 1.7 μg/μL. 7. SYPRO® Ruby staining solution. 8. SYPRO® Ruby fixation solution: 40% (v/v) methanol, 10% (v/v) acetic acid in water.

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9. SYPRO® Ruby destaining solution: 10% (v/v) methanol, 6% (v/v) acetic acid in water. 10. Orbital shaker. 11. Typhoon 9410 variable mode imager with green (532 nm) excitation laser and 610BP30 emission filter. 12. ImageQuant software or suitable densitometry-based analysis software. 2.4 Ultracentrifugation Exosome Isolation

1. Conditioned medium. 2. Sterile/filtered PBS. 3. Optima XPN Ultracentrifuge and matched rotor (Table 1). 4. SW28/SW32 Ti swinging-bucket (large-scale spins) with 38.5 mL Open-Top Thinwall Ultra-Clear Tube, 25
89 mm. 5. TLA-55 fixed angle (small scale/washing) with 1.5 mL Polypropylene Tube with Snap-on Cap, 9.5
38 mm.

2.5 OptiPrep™ Density Gradient Exosome Isolation

1. Conditioned medium. 2. Optima XPN Ultracentrifuge and matched rotor (Table 1). 3. SW 41 Ti swinging-bucket (large scale) with 13.2 mL Thinwall Polypropylene Tubes, 14
89 mm. 4. TLA-55 fixed angle (small scale/washing) with 1.5 mL Polypropylene Tube with Snap-on Cap, 9.5
38 mm. 5. OptiPrep™ stock solution: 60% (w/v) aqueous iodixanol. 6. 0.25 M sucrose in 10 mM Tris–HCl, pH 7.5. 7. Sterile/filtered PBS.

2.6 Cushion-Based Separation of Exosomes

1. Conditioned medium. 2. Optima XPN Ultracentrifuge and matched rotor (Table 1). 3. SW28/SW32 Ti swinging-bucket (large scale spins) with 38.5 mL Open-Top Thinwall Ultra-Clear Tube, 25
89 mm. 4. SW 41 Ti swinging-bucket (large scale) with 13.2 mL Thinwall Polypropylene Tubes, 14
89 mm. 5. TLA-55 fixed angle (small scale/washing) with 1.5 mL Polypropylene Tube with Snap-on Cap, 9.5
38 mm. 6. OptiPrep™ stock solution: 60% (w/v) aqueous iodixanol. 7. Sterile/filtered PBS.

2.7 EpCAM Immunoaffinity Capture (IAC) Exosome Isolation

1. Conditioned medium. 2. EpCAM (CD326) magnetic microbeads (Miltenyi Biotec, Auburn, CA).

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3. 3 mL LS Microcolumn for the gentle isolation of MicroBeadlabeled cells. 4. Solid support magnet (SSM). 5. IAC Rinsing Solution: MACS® BSA Stock Solution diluted 1:20 with autoMACS® Rinsing Solution. 6. Optima XPN Ultracentrifuge and matched rotor (Table 1). 7. TLA-55 fixed angle (small scale/washing) with 1.5 mL Polypropylene Tube with Snap-on Cap, 9.5
38 mm. 8. Sterile/filtered PBS. 9. IAC Elution buffer: 0.2 M glycine, Tris–HCl, pH 2.8. 10. 2
SDS sample buffer (see item 1 in Subheading 2.3.2). 2.8 Western Blot Analysis

1. Exosome, cell lysate preparations (~10–20 μg protein). 2. 2
SDS sample buffer (see item 1 in Subheading 2.3.2). 3. NuPAGE Bis-Tris Precast gel (see Subheading 2.3.2). 4. 1
MES running buffer (see Subheading 2.3.2). 5. XCell Surelock™ gel tank, with compatible power supply (see Subheading 2.3.2). 6. SeeBlue Plus 2. 7. Pre-stained protein standard (e.g., SeeBlue™ Plus2, or Dual Color Standard). 8. iBlot™ dry blotting system and nitrocellulose transfer membranes. 9. TTBS solution: 0.05% Tween® 20 in Tris-buffered saline (TBS). 10. Blocking buffer: 5% (w/v) skim milk powder in TTBS. 11. Primary antibody Mouse anti-TSG101 (#612696; BD Biosciences): 1:500 in TTBS. 12. Primary antibody Mouse anti-Alix (#2171, Cell Signaling Technology): 1:1000 in TTBS. 13. Secondary antibody IRDye 800 goat anti-mouse IgG: 1:15,000 in TTBS. 14. Orbital shaker. 15. Odyssey Infrared Imaging System, v3.0.

2.9 Nanoparticle Tracking Analysis (NTA)

1. Exosome preparation (~1–2 μg protein). 2. NanoSight NS300 system. 3. Ultrapure water. 4. Disposable 1 mL syringe for sample loading.

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1. Exosome preparations (~2–5 μg protein). 2. 1.5 mL Polypropylene Tube with Snap-on Cap. 3. 4% w/v Aldehyde/Sulfate Latex Beads, 4 μm. 4. Blocking buffer: 0.2% (v/v) Triton X-100, 2% (w/v) BSA in PBS. 5. Wash buffer: 0.2% (v/v) Triton X-100 in PBS. 6. 1 M glycine in PBS. 7. Primary antibody Mouse anti-Alix (#2171; Cell Signaling Technology): 1:1000 in blocking buffer. 8. Secondary antibody IRDye800 1:15,000 in blocking buffer.

goat

anti-mouse

IgG:

9. Benchtop centrifuge. 10. Microscope slide. 11. Zeiss AxioObserver Z1 microscope. 2.11 Electron Microscopy (EM)

1. Exosome preparations (~2 μg protein).

2.11.1

3. Fixing solution: 1% (v/v) glutaraldehyde.

Transmission EM

2. Sterile/filtered PBS. 4. Formvar coated 200 mesh copper grids. 5. 1% (w/v) aqueous uranyl acetate. 6. Gatan UltraScan 1000 (2k
2k) CCD camera coupled to a Tecnai F30 electron microscope.

2.11.2

Cryo EM

1. Exosome preparations (~2 μg protein). 2. Aurion Protein-G gold 10 nm. 3. Sterile/filtered PBS. 4. Glow-discharged C-flat holey carbon grids. 5. Vitrobot or automated sample preparation device. 6. Liquid ethane. 7. Liquid nitrogen. 8. Gatan cryoholder. 9. Tecnai G2 F30 electron microscope.

2.12 Proteomics: Sample Preparation 2.12.1 In-Solution Reduction, Alkylation, and Digestion

1. Exosome and cell lysate preparations (~5–10 μg protein). 2. 1.5 mL Protein LoBind Tubes or Protein LoBind deep 96-well Plates (1000 μL). 3. 1 M tetraethylammonium bromide (TEAB) stock solution in LC-MS grade water. 4. 50 mM TEAB, pH 8.0, in LC-MS grade water. For pH adjustment use 1 M hydrochloride in LC-MS grade water.

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5. 100 mM TEAB, pH 8.0, in LC-MS grade water. For pH adjustment use 1 M hydrochloride in LC-MS grade water. 6. 100% (v/v) acetone (LC grade). Stored at 20  C. 7. 90% (v/v) acetone in water (LC grade). Stored at 20  C. 8. Lysis buffer: 1% (w/v) SDS, 50 mM TEAB, pH 8.0. 9. 500 mM DTT stock solution in 100 mM TEAB, pH 8.0. Prepare fresh before the reduction step. 10. 1 M iodoacetamide (IAA) stock solution in 100 mM TEAB, pH 8.0. Prepare fresh before the alkylation step. 11. Trypsin, sequencing grade. Store lyophilized or frozen at 80  C. 12. Formic acid (LC-MS grade), prepare 1.5% (v/v) stock. 13. Water (LC-MS grade). 14. ThermoMixer with 1.5 mL microfuge tube capacity. 15. Thermostat oven or incubator at 37  C. 16. pH strip, range 1–14. 17. Minsox S-4000 600W Sonicator with microtip (or related sonicator). 2.12.2 StageTip Sample Cleanup

1. Empore SDB-RPS (Styrene Divinyl Benzene-Reversed Phase Sulfonate) solid-phase extraction disks for StageTip preparation. 2. SDB-RPS StageTip loading buffer: 1% (v/v) trifluoroacetic acid (TFA) in acetonitrile (ACN). 3. SDB-RPS StageTip wash buffer 1: 1% (v/v) TFA in ACN. 4. SDB-RPS StageTip wash buffer 2: 0.2% (v/v) TFA in 5% (v/v) ACN. 5. SDB-RPS StageTip elution buffer: 20 μL of NH4OH in 4 mL of 60% (v/v) ACN. Elution buffer must be prepared fresh (within 1 h of use) because the pH will begin to increase due to its high volatility, thereby reducing its elution strength. 6. Water (LC-MS grade). 7. Vacuum centrifuge (lyophilizer). 8. MS loading buffer: 0.07% (v/v) TFA in LC-MS water. This buffer is stable for >6 months at RT.

2.13 Fluorometric Peptide Assay

1. Quantitative Fluorometric Peptide Assay kit (Thermo Scientific #23290). 2. MS loading buffer (see item 9 in Subheading 2.12.2). 3. Peptide samples for analysis.

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4. Fluorescence compatible 96-well microplate (black). 5. Fluorescent plate reader compatible with Ex 390 nm/Em 475 nm. 2.14

UHPLC-MS/MS

1. MS loading buffer: 0.07% (v/v) TFA in LC-MS water. 2. Transparent MS sample vials, 300 μL: snap ring vial with PP insert. 3. UHPLC System coupled to a Q Exactive HF-X Hybrid Quadrupole-Orbitrap MS System. 4. Load column (Acclaim PepMap100 C18 5 μm beads with ˚ pore-size). 100 A 5. Analytical/separation column (50-cm fused-silica emitter reversed-phase PepMapRSLC C18, 75 μm inner diameter, ˚ pore-size). 2 μm resin with 100 A 6. Mobile Phase A: 0.1% formic acid. 7. Mobile Phase B: 0.1% formic acid in acetonitrile.

2.15

Data Analysis

1. MaxQuant software. 2. Microsoft Office Excel and Perseus software [45] (Max-Planck Institute of Biochemistry, Munich). 3. Online web-based bioinformatics resources such as gProfiler (https://biit.cs.ut.ee/gprofiler/gost), STRING (https:// string-db.org/), and Database for Annotation, Visualization and Integrated Discovery (DAVID) (https://david.ncifcrf. gov/).

2.16 Phenotypic Reprogramming of Cells by Exosomes: Dissecting Function

1. 1–2.5 mg/mL Dil (1,10 -dioctadecyl-3,3,30 ,30 -tetramethylindocarbocyanine Perchlorate) stock solution in DMSO.

2.16.1 Labeling Exosomes with Lipophilic Tracer

3. OptiPrep™ density gradient reagents (see Subheading 2.5).

2.16.2

1. Dil stained exosomes.

Exosome Uptake

2. Dil staining solution: 1 μM in PBS (10 mL). 4. 96-Well plate (black). 5. Fluorescence plate reader (Ex 549 nm/Em 565 nm).

2. Recipient cells of choice (~70% confluency). 3. 1 μM DiO or DiR staining solution in PBS. 4. Six-well plate or eight-well microscopy cover slides. 5. 10 μg/mL Hoechst nuclei stain in PBS. 6. PBS. 7. Phenol-red free media.

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8. Formaldehyde. 9. Dako fluorescence mounting media. 10. Fluorescence microscope. 2.16.3 Assay

Cell Activation

1. Recipient cells of choice (e.g., fibroblasts (5
104)). 2. 96-Well plate. 3. Serum-free culture media. 4. Purified exosomes. 5. PBS.

2.16.4 TranswellMatrigel™ Invasion Assay

1. Recipient cells of choice (e.g., fibroblasts (5
104)). 2. Purified exosomes. 3. PBS. 4. Growth factor-reduced Matrigel™ matrix. 5. Transwell inserts, 8 μm pore size. 6. 24-Well plate. 7. Incubator at 37  C. 8. 4% (v/v) formaldehyde. 9. 10 μg/μL Hoechst nuclei stain in PBS (or other nuclei stain). 10. Cotton swab. 11. Fluorescence microscope. 12. ImageJ image analysis tool.

2.16.5 Molecular Reprogramming of Cells by Exosomes

1. 70–80% confluent cells. 2. Ice-cold PBS. 3. Cell scraper. 4. 2% (w/v) SDS sample buffer. 5. 1.5 mL, Polypropylene Tube with Snap-on Cap. 6. Benchtop Centrifuge (up to 20,000
g spin). 7. Heat block (up to 95  C). 8. Minsox S-4000 600 W Sonicator with microtip (or related sonicator) (optional for ultracentrifugation approach) – 1.5 ml ultracentrifuge tubes, polypropylene (#41121703, Beckman Coulter). – TLA-55 rotor (Beckman Coulter). – Ultracentrifuge (Optima MAX-MP Tabletop Ultracentrifuge, #393315, Beckman Coulter).

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Methods See Table 2.

3.1 Small-Scale Exosome Production (2–20 μg)

Typically, cells grown to 80% confluency on 15 cm2 tissue culture plate yield ~1–2 μg (24 h incubation) of exosomes. This yield is cell type-dependent and should be determined for each cell type. 1. Culture cells to 60–70% confluency on 15 cm2 tissue culture plate in suitable culture media with supplements as required: 5% (v/v) FCS, 1% (v/v) Pen/Strep) at 37  C with 5% CO2 (see Note 1). 2. Gently wash cells three times with 10 mL (pre-warmed at 37  C) of PBS. 3. Culture for 24–48 h in cell culture medium (see Note 2) containing either 5% (v/v) EV-depleted FCS or 0.6–1% (v/v) ITS. 4. After 24–48 h incubation, collect conditioned medium (CM) into 50 mL polypropylene tubes and centrifuge:

Table 2 Overview of commonly used exosome isolation methods Isolation method

Mechanism

Advantage(s)

– Low/medium recovery Differential Sedimentation velocity yield ultracentrifugation (size, volume, density). – Scalability (DC) Typically used to isolate crude EVs from conditioned media and various biological fluids [92, 93] with the potential to purify with wash steps and DGC [6] Stepwise DC approach includes: initial 500
g/2000
g centrifugation (remove cells, membrane debris, apoptotic bodies), membrane filtration including 0.1 μm [48] or 0.22 μm membrane filtration [60], 10–14,000
g to isolate crude sMVs [18, 94, 95], 100,000
g to isolate crude exosomes [6]

Disadvantage(s) – High heterogeneity/ low purity – Co-purification with non-EV components – Yield dependent on sample viscosity and concentration [96] – Reproducibility may be influenced by rotor type and G-force [97] – Additional wash steps and handling may decrease yield [98, 99]

(continued)

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Table 2 (continued) Isolation method

Mechanism

Advantage(s)

Density-gradient centrifugation (DGC)

Buoyant density (density, – High purity – Potential for EV subtype size) isolation Used for further – Applicable to clinical purification of EV settings [104, 105] populations by a discontinuous gradient of sucrose (or lessviscous iodixanol, OptiPrep™ [6, 86]) [44, 70, 81] Iodixanol gradients are readily measured by refractive index [46], less toxic than sucrose in downstream functional cell assays [100], form iso-osmotic solutions at all densities (preserves vesicle size) [101], and allow non-vesicular components to be differentially fractionated [71]. Typically, use of DC (ultracentrifugation) at 100,000
g to establish gradients. Different variations of DGC include float-down Use to separate subpopulations of EVs, including exosomes of low (1.12–1.19 g/mL), high density (1.26–1.29 g/mL) [102, 103], sMVs of low (1.09 g/mL), and high density (1.12 g/mL) [102]

Sedimentation velocity and – Preservation of physical Density-cushioned integrity and biological buoyant density ultracentrifugation activity Layer of 40% iodixanol (DCGC) – Avoids aggregates below CM, EVs, and aggregates are concentrated in this cushion during ultracentrifugation Can be followed by DGC for purification

Disadvantage(s) – Time consuming – Unable to separate different EVs based on density – Low recovery (sample loss due to additional handling) – Can co-purify with different density lipoproteins [106, 107]

– Requires concentrated CM

(continued)

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Table 2 (continued) Isolation method

Mechanism

Advantage(s)

Affinity isolation

Surface marker selectivity – High purity – Potential to purify (protein or peptide different EV epitope target) (sub)populations The tag may be a – Ability of coupling with biospecific surface other methods of protein, such as a characterization (i.e., monoclonal antibody flow cytometry, western (mAb), that targets an blotting and rt-PCR) EV-surface antigen [108] (e.g., mAb 763.74-specific CSPG4 epitope uniquely expressed on melanoma cells), biospecific peptide (e.g., designer synthetic peptides with high affinity for HSPs [109]), or proteoglycan affinity reagent (e.g., heparin [110–112]). mAbs that have been successfully employed include those directed against A33 [74], EpCAM [48, 113], MHC-II antigens [114, 115], CD45 [77, 116], CD63 [117, 118], CD81 [118], CD9/CD1b/ CD1a/CD14 [119], CD9 [120], HER2 [76], and L1CAMb [121]. Heparin affinitybased affinity capture [112] is generally applicable for EV isolation from cell culture media and biofluids, given it overcomes limitations with availability of suitable mAbs directed to specific EV-surface antigens

Disadvantage(s) – Expensive (if antibody based) – EV elution might damage surface proteins and functionality – Typically, dependent on availability of suitable mAbs directed to specific EV-surface antigens – Low scalability – Low yield (binding capacity)

(continued)

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Table 2 (continued) Isolation method

Mechanism

Advantage(s)

– High scalability Size exclusion and gel Size, molecular weight This approach has been permeation widely applied for chromatography isolating EVs from plasma samples [122] and adapted (using commercially available columns) for highthroughput clinical samples [123]. Gel permeation chromatography overcomes many of the problems associated with EV isolation from plasma/serum using DC/DGC—e.g., co-isolation of EVs with large-Mr protein aggregates and lipoproteins [122–124] Precipitation

Disadvantage(s) – Dilution in elution buffer

– Low purity – Applicable for large Salting out using a – PEG chain might volumes polyethylene glycol/salt envelope the EVs, – Recent advances in solution possibly interfering sequential This approach provides with their precipitation/ rapid, but impure EV functionality absorption have preparations, and is indicated potential for therefore unsuitable for select types of EVs to be detailed biophysical/ differentially isolated functional assay purposes. However, the method acts as an isolation/concentration step for crude EV preparation for diagnostic assays of known EV-associated biomarkers. Recent developments using sequential polyethylene glycol precipitation and adsorption to immobilized lectin concanavalin A [125] have demonstrated both exosomes and sMVs can be selectively enriched (continued)

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Table 2 (continued) Isolation method

Mechanism

Advantage(s)

Sequential filtration

Membrane filtration (size, – Medium scalability – Time of separation molecular weight) of (efficiency) [129] proteins and other – No direct requirement macromolecules for ultracentrifugation Nanomembrane (i.e., preserves integrity ultrafiltration spin of exosomes) devices equipped with low protein binding membranes (e.g., polyether sulfone or hydrophilic polyvinylidene difluoride (PVDF)) can be used for EV isolation [126–128]. In combination with DC and DGC, nanomembrane ultrafiltration has enabled fractionation of EV subpopulations; sMVs and exosomes from the same cancer cell origin [43]

Disadvantage(s) – EV can clog the membrane filter – Sample loss (yield) – Does not discriminate between specific subtypes of EVs which are similar in size

480
g for 5 min followed by 2000
g for 10 min at 4  C to remove intact cells and cell debris. Carefully transfer the supernatant into a new tube and subject to desired exosome isolation protocol or store at 20  C until further use. This supernatant contains both soluble and vesicle components (see Note 3). 5. Cell viability should be assessed in cell culture medium (i.e., 24 or 48 h incubation) (see Note 4). 3.2 Large-Scale Exosome Production

Bioreactor flasks contain a lower cell-cultivation chamber separated from an upper nutrient supply chamber by a semipermeable membrane with a molecular weight cutoff of 10 kDa (see Note 5). This membrane allows continuous diffusion of nutrients from the upper to the lower chamber and waste elimination from the lower to the upper chamber. Exosomes released by cells are retained in the lower chamber. For detailed notes on large-scale exosome production (see Note 6). 1. Add 500 mL of prewarmed (37  C) DMEM with 5% (v/v) FCS, 1% (v/v) Pen/Strip) to the nutrient-supply chamber.

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2. Prepare cells (15–30
106 cells) in 15 mL of DMEM medium with 5% (v/v) FCS and 1% (v/v) Pen/Strep. 3. Transfer cells to the cultivation chamber and allow to seed/ expand for 3–4 days. 4. Remove the cultivation chamber medium (the cells have now adhered to cultivation chamber). 5. Wash the cultivation chamber three times with serum-free DMEM. 6. Add 15 mL of DMEM with 0.6–1% (v/v) ITS, 1% (v/v) Pen/Strep to the cultivation chamber. 7. Replace media in the nutrient chamber every 4–7 days (cell type dependent). 8. Replace media in the cultivation chamber every 1–2 days (cell type dependent). 9. Subject the conditioned medium from the cultivation chamber to exosome isolation protocol (see Subheadings 3.4–3.7). 10. Collect CM into 50 mL polypropylene tubes and centrifuge at 480
g for 5 min followed by 2000
g for 10 min at 4  C to remove intact cells and cell debris (see Note 3). 11. Carefully transfer the supernatant into a new tube and subject to desired exosome isolation protocol or store at 20  C until further use. This supernatant contains both soluble secreted and EV components. 3.3 Protein Quantitation (See Note 7) 3.3.1 Micro BCA Assay

1. Prepare a set of standard dilutions with BSA, starting with 200 μg/mL, performing dilutions in 1% SDS solution to 0.5 μg/mL and place them at RT (1% SDS solution can crystallize on ice). 2. Thaw protein sample (CM, exosomes, or cell lysates samples) and place on ice. 3. In a flat-bottom 96-well plate, load 100 μL of each standard BSA dilution/blank. The blank is dependent on what stock/ lysis solution exosomes are prepared in. 4. Load 1–4 μL of CM, exosomes, or cell lysate samples into wells, making up the total volume to 100 μL using 1% SDS solution. 5. Prepare working reagent (WR) by mixing reagent A, B, and C (as provided in the commercial kit) with the ratio of 25:24:1. 6. Add 100 μL of WR to each sample/standard. 7. Incubate at 37  C for 1.5–2 h. 8. Cool plate to RT. 9. Measure absorbance at 562 nm.

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10. Prepare standard curve, subtract the blank (abs) from the sample (abs), and determine the concentration of starting sample. 3.3.2 Protein Staining Densitometry

1. Solubilize samples (2–5 μL of CM, exosomes, or cell lysates) in reducing 2
SDS sample buffer (10 μL), and heat at 95  C for 5 min (closed cap) (optional but recommended to ultracentrifuge cell lysate at 400,000
g for 30 mins (TLA-100.2 rotor) to remove genomic DNA). 2. Briefly centrifuge samples using benchtop centrifuge and resolve on NuPAGE™ 4–12% (w/v) Bis-Tris Precast gels. 3. Load BenchMark™ Protein Ladder (2–5 μL, same volume as sample; 1.7 μg/μL) for quantitation. An empty lane will serve as background control. 4. Perform electrophoresis at 150 V for 1 h in 1
MES running buffer. 5. Remove the gel (and plastic casing) and place it in 50 mL fixing solution for 30 min on an orbital shaker and stain with 30 mL SYPRO® Ruby for 40 min, followed by destaining in SYPRO® Ruby destaining solution for 1 h. 6. Image gel on a Typhoon 9410 variable mode imager, using a green (532 nm) excitation laser and a 610BP30 emission filter at 100 μm resolution. 7. Perform densitometry quantitation using ImageQuant software to determine protein concentration relative to a BenchMark™ Protein Ladder standard of known protein concentration (1.7 μg/μL). Normalize by subtracting background signal from empty lane.

3.4 Ultracentrifugation Exosome Isolation

1. Centrifuge CM (~30 mL) at 10,000
g (SW28/ SW32 Ti, swinging-bucket) for 30 min at 4  C to pellet shed microvesicles. Retain the supernatant which contains exosomes. 2. Centrifuge the supernatant at 100,000
g (SW28/ SW32 Ti, swinging-bucket) for 1 h at 4  C to pellet exosomes (see Notes 8–11). 3. Resuspend exosome pellet in 1 mL sterile/filtered PBS and re-centrifuge at 100,000
g (TLA 55 rotor) for 1 h to obtain crude exosomes. 4. Resuspend crude exosomes in 50 μL PBS and either use immediately or store at 80  C.

3.5 OptiPrep™ Density Gradient Exosome Isolation

1. Prepare OptiPrep™ density gradient by diluting a stock solution of 60% (w/v) OptiPrep™ with 0.25 M sucrose/10 mM Tris–HCl, pH 7.5, to obtain 40% (w/v), 20% (w/v), 10% (w/v), and 5% (w/v) solutions of iodixanol. These solutions

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are made immediately prior to centrifugation and allow up to 30 min mixing time. The gradient is formed by adding 3 mL of 40% iodixanol solution, followed by careful layering of 3 mL each of 20% and 10% solutions, and 2 mL of the 5% solution (see Note 11). 2. Overlay crude exosomes (from Subheading 3.4) (500 μL) on top of the discontinuous iodixanol gradient. 3. Overlay 500 μL of 0.25 M sucrose/10 mM Tris–HCl, pH 7.5, on top of the discontinuous iodixanol gradient (control experiment). 4. Centrifuge at 100,000
g (SW41Ti swinging-bucket) for 18 h at 4  C (Table 1). 5. After centrifugation, collect 12 individual 1 mL gradient fractions (with increasing density—top-bottom collection) using a 1 mL pipette. Isolation from the meniscus is imperative for this careful procedure (see Note 12). 6. Dilute fractions with 2 mL PBS and centrifuge at 100,000
g for 1 h at 4  C followed by washing with 1 mL PBS (repeat 100,000
g for 1 h at 4  C), and resuspend in 50 μL PBS. 7. Determine density of each fraction of the control OptiPrep™ gradient run in parallel (see step 3). Collect 12 fractions from the control gradient, serially dilute 1:10,000 with distilled water, and determine the iodixanol concentration based on absorbance at 244 nm using a molar extinction coefficient of 320 L/g/cm [46] (see Note 12). 8. Use exosomes immediately or store at 80  C (see Note 13). 3.6 Cushion-Based Separation of Exosomes

Exosome pellet obtained following ultracentrifugation potentially contains protein aggregates sticking to the surface of exosomes [47]. Thus, interpretation of down-stream biochemical, biophysical, and functional characterization may be compromised. Although IP-based isolation of exosomes can overcome these limitations, it only enables enrichment of a specific subset of exosomes (see Note 14). Here, we outline ultracentrifugation coupled to cushion-based separation of exosomes to gently obtain exosomes and prevent their pelleting/aggregation. We highly recommend using CM from CELLine adhere/classic bioreactor flasks, as the concentration of exosomes in the CM is high.

3.6.1 Cushion-Based Separation of Exosomes

1. Add 1 mL of 40% OptiPrep™ solution (prepared as described in Subheading 3.5) to Open-Top Thinwall Ultra-Clear Tube (38.5 mL). 2. Gently overlay with 30 mL conditioned media (post 10,000
g spin) (see Subheading 3.4, step 1).

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3. Centrifuge at 100,000
g for 1 h at 4  C (SW28/ SW32 Ti swinging-bucket). 4. Exosomes float at CM and OptiPrep™ solution interphase as a band. 5. Aspirate out the CM, being careful not to disturb the exosome band. 6. Collect the exosome band. 7. Dilute exosomes in 30 mL PBS and repeat ultracentrifugationcushion separation or subject to density-based separation. 8. Use exosomes immediately or stored at 80  C. 3.6.2 Ultracentrifugation Density Gradient Separation

1. Prepare 4 mL 5–40% OptiPrep™ gradient: 1 mL, 5%; 1 mL, 10%; 1 mL, 20%; 1 mL, 40%) in an Open-Top Thinwall UltraClear tube (38.5 mL). 2. Gently overlay with 8 mL conditioned media (post 10,000
g spin) (see Subheading 3.4, step 1). 3. Centrifuge at 100,000
g for 18 h at 4  C (SW28/ SW32 Ti swinging-bucket). 4. Just as in regular OptiPrep™ density gradient exosome isolation (see Subheading 3.5), exosomes enter the gradient, float at ~1.07–1.09 g/mL, and visible often as a band if the yield is high. 5. Aspirate out the CM, being careful not to disturb the OptiPrep™ gradient. 6. Carefully collect twelve 330 μL fractions. 7. Dilute 330 μL fraction with 1 mL PBS. Ultracentrifuge 10% of the sample at 100,000
g for 1 h and subject the pellets to Western blotting (ALIX and TSG101 primary antibodies) to determine exosome-containing fraction(s) (see Subheading 3.8). 8. Pool and dilute exosome-containing fraction(s) in 10 mL PBS and overlay on 13.2 mL tube containing 500 μL 40% OptiPrep™ solution. Fraction pooling is based on density and exosome marker presence. 9. Ultracentrifuge at 100,000
g for 1 h (SW41Ti swingingbucket). 10. Exosomes float atop the OptiPrep™-solution as a band. 11. Aspirate out the supernatant, being careful not to disturb the exosome band. 12. Collect the exosome band. 13. Use exosomes immediately or store at 80  C.

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3.7 EpCAM Immunoaffinity Capture Exosome Isolation (See Note 15)

1. Incubate concentrated CM with 100 μL EpCAM magnetic microbeads for 4 h at 4  C [18, 48]. 2. Place 3 mL LS Microcolumn in a solid support magnet (SSM) and rinse three times with Rinsing Solution. 3. Pipette exosome-bound microbeads into the LS column and wash three times with 1 mL Rinsing Solution. 4. Remove LS column from the SSM and recover exosomebound microbeads by rinsing the LS column at room temperature (RT) with 3
1 mL Rinsing Solution. 5. Wash exosome-bound microbeads twice with 1 mL PBS, place in ultracentrifuge microfuge vials, and centrifuge at 100,000
g for 1 h at 4  C (Table 1). 6. Remove the supernatant and elute IAC-Exosomes (yield ~195 μg) from the microbeads with either 100 μL of IAC elution buffer for EM imaging, or lyse with 100 μL of 2
SDS sample buffer (without DTT).

3.8 Western Blot Analysis (See Note 16)

1. Lyse the samples (~10–15 μg protein) in 2
SDS sample buffer with or without DTT (depending on the primary antibody reducing/non-reducing compatibility, respectively) and heat for 5 min at 95  C (closed cap, although cap can be opened for evaporation to reduce load volume). 2. Perform electrophoresis on lysed samples at 150 V for 1 h. 3. Following electrophoresis, electro-transfer proteins onto nitrocellulose membranes using iBlot™ Dry Blotting System. 4. Incubate membranes with blocking buffer for 1 h at RT. Care should be taken to not touch and disrupt the membrane (only the immediate corners/edges). 5. Probe membranes with primary antibody solution for 1 h in TTBS followed by incubation with secondary antibody solution for 1 h in dark. Carry out antibody incubations using gentle orbital shaking at RT. 6. Wash Western blots three times in TTBS for 10 min after each antibody incubation step and subsequently visualize at 800 nm using the Odyssey Infrared Imaging System.

3.9 Nanoparticle Tracking Analysis (NTA) (See Note 17)

1. Dilute sample (exosome or CM) to a final volume of 1 mL (final concentration 1 μg/μL in ultrapure water (1:10,000 dilution)). 2. Load sample onto NS300 flow-cell top plate using syringe pump. 3. Set analysis settings as per [6]: detection threshold: 10; flow rate: 100; temperature: 25  C. 4. Adjust camera level and screen gain until particles are visible.

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5. For video capture, set conditions as follows: number of capture: 3 (for triplicate); capture duration: 60 s. 6. For acquisition, change base file name to preferred destination folder. 7. Create and run script to begin capture and analysis with NTA software. Ensure that detection threshold is set to include as many particles as possible with the restriction that 10–100 red crosses were counted while only 500–1000 mL) are often utilized in order to generate exosome yields for characterization/functional studies. This introduces issues with the capacity of centrifugation, availability (rotors, instrumentation), and time. If researchers are limited by large-scale ultracentrifugation, then the CM preparation approach, with IAC, or commercial kits to isolate EVs would be suggested. 9. Requirement to isolate shed microvesicles for analysis (using the CM preparation vesicles >0.1 μm are not retained and therefore restricting their subsequent analysis). Shed microvesicles are heterogeneous ~150–~1000 nm diameter vesicles, distinct in their biogenesis and marker expression composition to exosomes [68]. If researchers are interested in the isolation of shed microvesicles, then this fraction can be easily isolated from culture medium in the process of obtaining the crude exosome pellet. For the isolation of shed microvesicles, a direct ultracentrifugation approach is employed. Following 10,000
g centrifugation, the shed microvesicle pellet is resuspended in 1 mL of PBS and 10,000
g centrifugation performed to obtain the washed shed microvesicle fraction (resuspend in 50–100 μL PBS). Further use of density-based separation can purify this EV type [6]. The isolation and proteomic characterization of shed microvesicles has been described previously [6, 18, 43, 69]. 10. For ultracentrifugation, mark each ultracentrifuge tube and orient the tube in the rotor with the mark facing up. The mark should be used as a reference for the location of a pellet following centrifugation. For swinging-bucket rotors, the

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pellet is at the bottom of the tube. For fixed-angle rotors, the pellet is on the side of the tube near the base. 11. The ultracentrifuge tubes are thin walled and will collapse if improperly filled. All tubes should be at least 80% capacity to ensure the tube is not flexible (add PBS if required). Tube specifications should be checked with the manufacturer prior to use. Ensure that the tubes are sealed (no condensation which can directly affect vacuum) and that the rotor is properly seated within the centrifuge. 12. Exosomes float at densities ranging from 1.09 to 1.15 g/mL on continuous sucrose or iodixanol density gradients following centrifugation [18, 48, 69, 70]. OptiPrep™ density gradients have low viscosity, isosmotic gradients that provide rapid and efficient separation of extracellular vesicles. OptiPrep™ density gradients have been shown to efficiently separate exosomes from HIV-1 particles [71]. Following OptiPrep™ density gradient, isolated protein fractions (in this case 12 fractions) can be assessed for density (e.g., range 1.01–1.30 g/mL), and protein yield, and assessing expression of exosomal markers using immunoblot analysis. As an example, OptiPrep™ density gradient fractions (10 μg per fraction) revealed exosomal marker Alix detected/enriched in density fraction 1.09 g/mL (fraction 7) [7, 69, 72]. 13. Exosome stability during different storage conditions has been examined (i.e., at 20, 4, and 37  C) [73]. The size of exosomes decreases at 4  C (3–4 days) and 37  C (from 2 days), indicating a possible structural change or degradation. Multiple freezing to 20  C and thawing does not affect exosome size based on nanoparticle tracking analysis (NTA) [73]. In our studies, exosome degradation has been monitored at 4  C (within 72 h) and 37  C (within 24 h) (data not shown). We recommend storage of exosomes at 4  C on ice for short-term use (within 3 days), and for long periods at either 20  C/ 80  C or lyophilization (over 12 months at 4  C). Exosome samples should be stored in small (50–100 μL) aliquots to avoid repeated freezing and thawing. 14. The availability and suitability of exosome markers for IAC is dependent on their target specificity and application [21, 42]. Currently, several markers for exosomes have been used for IAC including Glycoprotein A33+ exosomes derived from human epithelial colon cancer cells [74], EpCAM+ exosomes derived from human epithelial colon cancer cells [48], sequential Glycoprotein A33+ and EpCAM+ exosomes from human epithelial colon cancer cells to reveal specific subpopulations of exosomes [18], MHC II+ exosomes derived from dendritic cells [75], HER2+ exosomes derived from BT-474

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breast cancer cells [76], and CD45+ exosomes derived from Jurkat and SupT1/CCR5 cells [77]. Commercially, Dynabeads®-based CD63-specific reagent (#10606D, Life Technologies), and streptavidin reagent with choice of a biotinylated antibody to purify a specific vesicle population based on a surface antigen (#10608D, Life Technologies) are available. The use of density-based separation provides significant advantages for exosome isolation when the use of IAC is limited (due to availability/suitability of exosome markers and their localization on exosomes, i.e., surface capture). 15. CD326 (EpCAM)+ exosomes are bound with EpCAM MicroBeads, with the CM loaded onto a MACS® Column which is placed in the magnetic field of a MACS Separator (SSM). The magnetically labeled EpCAM+ exosomes are retained within the column. The EpCAM vesicles are passed through; this fraction is depleted of EpCAM+ exosomes. After removing the column from the magnetic field, the magnetically retained EpCAM+ exosomes are eluted as the positively selected exosome fraction. For this application, 100 μL of CD326 (EpCAM) MicroBeads were used for 500 mL CM (1.5 mg CCM) generated from 2
109 cells. Working on ice may require increased incubation times. Higher temperatures and/or longer incubation times may lead to nonspecific vesicle binding. 16. To characterize extracellular vesicles as exosomes, it is important to demonstrate the expression of common exosomal proteins using immunoblotting. The commonly used markers may include Alix (PDCD6IP, programmed cell death 6 interacting protein), TSG101 (tumor susceptibility gene 101), CD63 (tetraspanin CD63), and CD81 (tetraspanin CD81). As recommended by MISEV guidelines [42], Western blotting should be performed by loading side-by-side extracellular vesicles as exosomes and source material lysates either in specified protein amount or in cell-equivalent amounts to determine if the analyzed proteins are enriched in extracellular vesicles as exosomes as compared with their producing cells. Further, to specify small EV subtype (including exosomes), further protein identification should be performed in/on intracellular compartments of eukaryotic secreting cells other than the plasma membrane and endosomes (i.e., components of the nucleus, mitochondria, endoplasmic reticulum, Golgi apparatus, autophagosomes, peroxisomes) not enriched in the smaller EVs of plasma membrane or endosomal origin [42]. 17. Nanosight analysis settings and sample dilution may be adjusted to obtain ideal particle per frame value (20–100 particle/frame). Dilution should be made in ultrapure water free from particle contamination (i.e., dust).

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18. We provide two methods (TEM and cryoEM) for electron microscopy exosome sample preparation. For TEM analysis, exosomes are fixed using glutaraldehyde and negatively stained using uranyl acetate. This approach typically has a rapid sample preparation approach (100 days [88, 89]. 29. Optimal concentration and time length must be determined for different recipient cells. From our experience using live fluorescence imaging, exosome uptake (10–30 μg/mL) can start within the first 15 min [90, 91]. 30. It is recommended to label the plasma membrane of recipient cells with DiO or DiR or to stain intracellular structures (such as organelle probes; e.g., https://www.thermofisher.com/au/ en/home/references/molecular-probes-the-handbook/ tables/molecular-probes-organelle-selective-probes.html).

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31. Important: Do not subject the samples to detergent permeabilization as the Dil dye labeling the exosomes will be washed away with the lipids resulting in loss of signal. 32. Resource for rotor speed, conversion, tubes, and adapters can be found at https://www.beckmancoulter.com/wsrportal/ wsr/research-and-discovery/products-and-services/centrifu gation/rotors/index.htm?t¼3 and https://www. beckmancoulter.com/wsrportal/wsr/research-and-discov ery/products-and-services/centrifugation/tubes-andadapters/index.htm.

Acknowledgments This work was funded by NHMRC project grants (#1057741 and #1139489; D.W.G.), Helen Amelia Hains Fellowship (D.W.G.), Australian Government Research Training Program Scholarship (M.F., B.C.), BrightSparks Scholarship (B.C.), and La Trobe University-Baker Institute Postgraduate Scholarship (Q.P.). References 1. van Niel G, D’Angelo G, Raposo G (2018) Shedding light on the cell biology of extracellular vesicles. Nat Rev Mol Cell Biol 19:213–228. https://doi.org/10.1038/ nrm.2017.125 2. Mathieu M, Martin-Jaular L, Lavieu G, The´ry C (2019) Specificities of secretion and uptake of exosomes and other extracellular vesicles for cell-to-cell communication. Nat Cell Biol 21:9–17. https://doi.org/10.1038/s41556018-0250-9 3. Bidarimath M, Khalaj K, Kridli RT, Kan FWK, Koti M, Tayade C (2017) Extracellular vesicle mediated intercellular communication at the porcine maternal-fetal interface: a new paradigm for conceptus-endometrial cross-talk. Sci Rep 7:40476–40476. https://doi.org/ 10.1038/srep40476 4. Zhang X, Hubal MJ, Kraus VB (2020) Immune cell extracellular vesicles and their mitochondrial content decline with ageing. Immun Ageing 17(1):1. https://doi.org/10. 1186/s12979-019-0172-9 5. Evans J, Rai A, Nguyen HPT, Poh QH, Elglass K, Simpson RJ, Salamonsen LA, Greening DW (2019) Human endometrial extracellular vesicles functionally prepare human trophectoderm model for implantation: understanding bidirectional maternalembryo communication. Proteomics. https://doi.org/10.1002/pmic.201800423

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Chapter 10 Human Plasma Extracellular Vesicle Isolation and Proteomic Characterization for the Optimization of Liquid Biopsy in Multiple Myeloma Antonia Reale, Tiffany Khong, Rong Xu, Maoshan Chen, Sridurga Mithraprabhu, Nicholas Bingham, Andrew Spencer, and David W. Greening Abstract Cancer cells secrete membranous extracellular vesicles (EVs) which contain specific oncogenic molecular cargo (including oncoproteins, oncopeptides, and RNA) into their microenvironment and the circulation. As such, EVs including exosomes (small EVs) and microvesicles (large EVs) represent important circulating biomarkers for various diseases, including cancer and its progression. These circulating biomarkers offer a potentially minimally invasive and repeatable targets for analysis (liquid biopsy) that could aid in the diagnosis, risk stratification, and monitoring of cancer. Although their potential as cancer biomarkers has been promising, the identification and quantification of EVs in clinical samples remain challenging. Like EVs, other types of circulating biomarkers (including cell-free nucleic acids, cf-NAs; or circulating tumor cells, CTCs) may represent a complementary or alternative approach to cancer diagnosis. In the context of multiple myeloma (MM), a systemic cancer type that causes cancer cells to accumulate in the bone marrow, the specific role for EVs as biomarkers for diagnosis and monitoring remains undefined. Tumor heterogeneity along with the various subtypes of MM (such as non-secretory MM) that cannot be monitored using conventional testing (e.g. sequential serological testing and bone marrow biopsies) render liquid biopsy and circulating tumor-derived EVs a promising approach. In this protocol, we describe the isolation and purification of EVs from peripheral blood plasma (PBPL) collected from healthy donors and patients with MM for a biomarker discovery strategy. Our results demonstrate detection of circulating EVs from as little as 1 mL of MM patients’ PBPL. High-resolution mass spectrometry (MS)-based proteomics promises to provide new avenues in identifying novel markers for detection, monitoring, and therapeutic intervention of disease. We describe biophysical characterization and quantitative proteomic profiling of disease-specific circulating EVs which may provide important implications for the development of cancer diagnostics in MM. Key words Extracellular vesicles, Exosomes, Liquid biopsy, Multiple myeloma, Blood

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Introduction Multiple myeloma (MM) is a blood cancer that originates from the clonal expansion of plasma cells in the bone marrow (BM) [1– 3]. MM remains incurable despite advances in its treatment due to high rates of relapse and drug resistance [4, 5]. Risk stratification remains a major challenge in plasma cell dyscrasias where current recommendations for therapy are observation for asymptomatic patients (smoldering MM; premalignant condition Monoclonal Gammopathy of Undetermined Significance, MGUS), with the initiation of therapy only at the time of the emergence of symptomatic disease [6–9]. Therefore, a better understanding of the molecular characteristics that define the risk of progression to symptomatic MM would provide a framework for early initiation of systemic treatment [10–15]. Liquid biopsies represent less-invasive diagnostic alternatives or additions to single-site tissue biopsies being able to capture the spatial and temporal tumor heterogeneity—a major limitation of tissue biopsies [16–20]. The latter are often invasive or not feasible due to patient compliance/capacity (e.g., comorbidities, logistics) or tumor (e.g., location, size, type/subtype) characteristics [21– 26]. Conversely, liquid biopsies are innovative tools in precision medicine and cancer diagnostics with the ability to detect, monitor, and characterize tumors in a minimally invasive and repeatable way [16–20, 27–31]. It is well established that tumor cell-derived proteins, nucleic acids and extracellular vesicles (EVs), enter the circulation and reach distant sites where they establish a favorable microenvironment for tumor expansion [32–35]. These circulating factors represent useful biomarkers for cancer diagnosis with studies highlighting their prognostic and predictive significance with important clinical implications [36–40]. Liquid biopsies have the potential to detect low-abundant biomarkers from complex biofluids like blood, making them exceptional candidates for early detection and monitoring of cancers (diagnosis and/or residual disease) or for risk stratification [16, 17, 41–44]. Highly sensitive and selective omics (including genomics and proteomics) technologies and strategies have been developed to overcome the inherent challenge posed by the low-abundance of tumor-derived circulating factors [19, 28, 31, 45–48]. Importantly, next-generation sequencing (NGS) technologies and mass spectrometry (MS)based proteomics, with the aid of advanced bioinformatic tools, have been successfully utilized for cancer biomarker discovery [43, 46, 49–57]. Circulating cf-NA (cf-DNA/RNA) together with CTCs are the most developed biomarkers detected by liquid biopsy, with increasing evidence that combined analyses (i.e., combination of cf-NA and CTC), rather than single-source strategies (cf-NA or CTC),

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represent the key for increasing specificity of cancer detection [16, 18–20, 27, 58]. The interest for EVs as liquid biopsies is also increasingly expanding through preclinical and clinical evaluation in both solid tumors and hematological malignancies including MM [42, 47, 59–68]. EVs are particles delimited by a lipid bilayer and cannot replicate as they do not contain a functional nucleus [69, 70]. EVs are secreted by almost all cell types including cancer cells and have been shown to play an important role in cell-to-cell communication by horizontal transfer of their contents (lipids, proteins, nucleic acids) [71–75]. They are able to influence pathological as well as normal homeostatic cellular processes by reprogramming signaling pathways to modulate the function and activity of target cells [76– 79]. EVs have the capacity to signal at distant sites and reprogram organs conducive toward a metastatic microenvironment [80– 82]. Moreover, EVs share common surface markers with their cell/tissue of origin, designating them as a promising source for biomarker discovery and diagnostics [40, 83, 84]. EV subtypes including large (~50–1300 nm) and small (30–150 nm) originate from different cellular compartments, plasma membrane- and endosome-derived, respectively [69, 70]. Omic technologies represent key strategies for the study of EV cargo [46, 55, 76, 85, 86] with several recent data suggesting a critical role for EVs in the context of cancer liquid biopsy [41, 43, 47, 63]. A role for large EVs as predictive and prognostic biomarkers has also been suggested in MM [59, 60], while data on small EVs are still exploratory and require further investigation [62, 64–68]. In a rapidly growing field, a major challenge is related to the isolation, purification, and characterization of EVs. Several position statements from the International Society for Extracellular Vesicles (ISEV) and the Extracellular RNA Communication Consortium (ERCC) have been issued, highlighting the importance of standardization of sample collection and EV isolation and characterization approaches and reporting metrics [70, 87]. It is important to consider the starting material, the downstream application, and the end use, as a guide to choose the most suitable isolation method wherein balance between purity, cost, and time must be achieved [70, 88]. The isolation of small EVs from blood represents one of the most demanding applications. Blood is a complex biofluid that contains highly abundant proteins (HAP; e.g., albumin) which are often co-isolated and impair the enrichment of low-abundant particles such as small EVs [89–91]. While several strategies have been used for EV isolation [92–98], commercially available kits represent valuable tools for blood-derived EVs with readily translational implications when compared to time-consuming methods such as ultracentrifugation which are more suitable for in vitro studies. Commercially available kits are based on different isolation modalities such as immunoaffinity capture, precipitation, membrane-

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a

Myeloma cell clones

small EVs

Secreteome Bone marrow microenvironment

small EVs characterisation Characterization

b

small EVs isolation

Plasma preparation

6,000 rpm 1 min, RT

Blood PPP

PFP

1,800 x g 10 min

2,000 x g 15 min

4 °C

4 °C

1 ml PBPL + 3 ml H2O + Resin

-Vortex 10 sec -Incubate 5 min -Vortex 10 sec -2,000 rpm, 2 min, RT

-Vortex 10 sec -Incubate 5 min -Vortex 10 sec -500 rpm 2 min RT

-Discard supernatant - Add ExoR buffer

-Transfer the supernatant to a Mini Filter Spin column

Depletion

Western Blotting Nanoparticle Tracking Analysis Electron Microscopy

HAP depletion

Proteomic profiling

Discard spin column

Protein

Peptide STRECK RNA tube 10 ml

EVs

CTC

nLC and high resolution MS HAP Platelets cf-DNA cf-RNA

Informatic analyses

Fig. 1 Protocol for isolation and characterization of PBPL-derived EVs. (a) Multiple myeloma cell clones secrete various factors into the local tumor microenvironment and circulation, including EVs, CTCs, and cf-DNA/RNA. Therefore, liquid biopsies are able to capture the spatial and temporal heterogeneity of tumors which is often underestimated by single-site tissue biopsy. Blood, a complex bio-fluid, comprises HAP (e.g., albumin) and platelet-derived EVs which represent major challenges in the pre-analytical and purification steps prior to omic approaches to define the composition of EV cargo. (b) Strategy to isolate and characterize small EVs from blood is provided. Isolation of small EVs from 1 mL of PBPL utilizing a resin-based approach is shown (refer to Subheadings 3.1–3.4). Biochemical and biophysical characterization of EVs can include western blotting (EV marker proteins), nanoparticle tracking analyses (particle detection, particle size distribution) and transmission electron microscopy (EV size, morphology), proteomic-based profiling of EVs cargo, with analysis, and informatic approaches depending on the question and strategy. It is important to deplete albumin or other highly abundant proteins which may be co-purified during EV isolation due to MS identification issues. [EVs extracellular vesicles, CTC circulating tumor cells, cf-DNA/RNA circulating cell free DNA/RNA, HAP highly abundant proteins, PPP platelet-poor plasma, PFP platelet-free plasma, PBPL peripheral blood plasma, MS mass spectrometry, nLC nanoscale liquid chromatography, RT room temperature]

based affinity, and resin [92, 93, 95–97]. Here, we provide a detailed isolation and purification protocol together with methods for EV characterization which are employed to optimize the pre-analytical and isolation/purification steps prior to MS-proteomics to define the composition of EV cargo (Fig. 1). Methodologies including immunoblotting, nanoparticle tracking analyses (NTA), and electron microscopy are of critical importance for EV characterization [70, 94, 98]. These techniques show that small EVs derived from 1 mL of fresh PBPL and isolated utilizing a commercial kit (resin-based) are homogeneous in terms of size and morphology and enrichment of EV markers (Alix, TSG101, and tetraspanins CD63 and CD81) in comparison to whole blood.

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Of critical importance in the study of blood-derived EVs is the demonstration of the source of EVs of interest. Platelet-derived vesicles (microparticles, MPs, and small EVs) represent a significant proportion of the source of EVs in blood [99–101]. The levels of protein expression of platelet EV markers (e.g., CD41, CD62) may indicate the amount of “contamination” of the isolated EV sample by platelets EVs. Specific protocols for blood collection and PBPL preparation are tailored for minimizing the activation of platelets with subsequent release of platelet EVs in the sample of interest [102–104]. Immunocapture has also been successfully utilized to exclude this population from EVs of interest (e.g., CD41
) [105]. In vitro/ex vivo [106, 107] models utilizing cell lines (human myeloma cell lines, HMCL) and primary (MM and stromal) cells are also suggested not only for EV functional studies but also to complement the validation of the PBPL-EV findings. Sideby-side comparison of EVs with source material (cells/tissue) [70] utilizing omic strategies may provide important insights into the specific EV cargo enrichment. The use of blood collected from healthy controls as a comparator is strategic as it implements the strength of MM-derived data when a normal background (reference) is defined.

2 2.1

Materials PBPL Isolation

1. Blood obtained from healthy donors, MGUS, and MM patients utilizing 2  10 mL STRECK RNA Complete BCT™ tubes (see Notes 1–4). 2. 1.5 mL centrifuge low-protein binding tubes (see Note 5). 3. Sterile pipette tips with filters. 4. Pipettes, stripettor, and pipettors. 5. Allegra X-15R refrigerated benchtop centrifuge with SX4750 swinging bucket rotor for large-scale preparation. 6. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL) for smallscale preparation.

2.2 Large EVs Isolation

1. Fresh/frozen PBPL (see Note 6). 2. Sterile/filtered phosphate-buffered saline (PBS). 3. 1.5 mL centrifuge low-protein binding tubes (see Note 5). 4. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL).

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2.3 Small EVs Isolation 2.3.1 CD63 Exo-FLOW Capture Kit

1. CD63 Exo-FLOW capture kit (# EXOFLOW300A-1) from System Biosciences (SBI) (see Notes 7–9) containing streptavidin magnetic beads, biotinylated capture antibody (CD63), bead wash buffer, exosomes stain buffer, Exo-FITC universal exosome stain, and exosome elution buffer. 2. ExoQuick™ precipitant (see Subheadings 2.3.2 and 3.3.2). 3. 100–200 μg EV protein. 4. 1.5 mL centrifuge tubes. 5. Vortex mixer. 6. Rotating shaker. 7. Magnetic stand. 8. Canto II flow cytometer. 9. FACSAria flow sorter (see Note 8).

2.3.2 ExoQuick ULTRA Kit

1. ExoQuick ULTRA kit (#EQULTRA-20A-1) from SBI containing proprietary precipitant and purification columns, collection tubes, and buffer A and B (see Note 10). 2. Thrombin Plasma prep for Exosome precipitation (see Note 11). 3. 1.5 mL centrifuge tubes. 4. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL). 5. Rotating shaker. 6. Syringe filters for excluding particles larger than EVs of interest (see Note 10).

2.3.3 exoEasy Kit

1. exoEasy kit (#76064) from Qiagen containing proprietary membrane columns and buffers (see Note 12). 2. Conical tubes. 3. Syringe filters for excluding particles larger than EVs of interest (see Note 12). 4. Allegra X-15R refrigerated benchtop centrifuge with SX4750 swinging bucket rotor for large-scale preparation.

2.3.4 Purification Mini Kit

1. Purification Mini Kit (#57400) from Norgen Biotek Corporation containing proprietary resin, mini filter spin columns, elution tubes, and buffers (see Note 13). 2. 15 mL conical tubes. 3. Nuclease-free water. 4. Sterile pipette tips with filters. 5. Allegra X-15R refrigerated benchtop centrifuge with SX4750 swinging bucket rotor for large-scale preparation.

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6. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL) for smallscale preparation. 7. Vortex mixer. 2.4 Albumin Depletion (See Note 14)

1. Pierce™ Albumin Depletion Kit (see Note 15) containing resin, buffers, and spin columns. 2. Sterile pipette tips with filters. 3. 1.5 mL centrifuge low-protein binding tubes. 4. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL).

2.5 EV Lysate Preparation

1. 10 RIPA buffer: 0.22% (w/v) beta glycerophosphate, 10% (v/v) 4-nonylphenol (branched, ethoxylated), 0.18% (w/v) sodium orthovanadate, 5% (w/v) sodium deoxycholate, 0.38% (w/v) EGTA, 1% (v/v) sodium lauryl sulfate, 6.1% (w/v) Tris-base, 0.29% (v/v) EDTA, 8.8% (w/v) sodium chloride, and 1.12% (w/v) sodium pyrophosphate decahydrate, pH 7.5 (see Notes 16 and 17). 2. 25 protease inhibitors (PI) cocktail (see Note 18). 3. 1.5 mL centrifuge tubes. 4. Eppendorf 5424R refrigerated benchtop centrifuge with FA45-24-11 fixed angle rotor (24  1.5/2.0 mL).

2.6 Protein Quantitation (See Note 19)

1. Mirco BCA Protein assay kit. 2. Bovine serum albumin (BSA) as standard protein. BSA purity >98%. 3. Working reagent (WR): Mix 25 parts Reagent MA, 24 parts Reagent MB, and one part Reagent MC. 150 μL for each standard/sample. 4. 1% (w/v) sodium dodecyl sulfate (SDS) in ultrapure water. 5. EV lysate sample for measurement. 6. 1.5 mL centrifuge tubes. 7. Flat-bottom, clear 96-well plates. 8. Incubator (37  C). 9. Microplate reader with 562 nm filter.

2.7 Western Blot Analysis

1. EV lysate preparations: ~7–20 μg protein. 2. 4–15% Mini-PROTEAN TGX Precast Gels; 4–15% MiniPROTEAN TGX Stain-Free Protein Gels. 3. TTBS solution: Tris-buffered saline (TBS) with 0.05% (v/v) Tween-20. 4. Blocking solution: 5% (w/v) skim milk powder in TTBS.

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5. 4 SDS-PAGE sample buffer: 0.2 M Tris-HCl, pH 6.8, 0.4 M DTT, 8.0% (w/v) SDS, 6 mM bromophenol blue, 40% (w/v) glycerol. 6. Heat block. 7. Precision Plus Dual Color Protein Standard. 8. SDS-PAGE electrophoresis buffer: 0.1% (v/v) SDS, 25 mM Tris-base, 190 mM glycine, pH 8.3. Do not titrate pH. 9. Transfer buffer: 25 mM Tris-base, 190 mM glycine, 20% (v/v) methanol. 10. Immobilon-P PVDF blotting membrane (0.45 μm pore size). 11. 100% methanol for activating PVDF membranes. 12. Whatman® cellulose blotting papers (3 mm grade). 13. Frozen cooling unit. 14. Mini-PROTEAN Tetra cell system. 15. Mini Trans-Blot Cell. 16. Ready-to-use Ponceau Red Staining solution. 17. Primary antibodies in TTBS: 1:500 (mouse anti-TSG101, BD Biosciences, #612697); 1:1000 (mouse anti-Alix, Cell Signaling Technology, #2171S; mouse anti-CD81, Santa Cruz Biotechnology, #7637; rabbit anti-CD63, Abcam, #ab134045; rabbit anti-Integrin alpha 2b, Cell Signaling Technology, #13807S; mouse anti-P-selectin, Santa Cruz Biotechnology, #8419; rabbit anti-Albumin, Abcam, #ab207327; rabbit antiGAPDH, Cell Signaling Technology, #8884). 18. Secondary antibodies, HRP conjugated, in TTBS: 1:20,000 (anti-rabbit, Dako, #P0217); 1:15,000 (anti-mouse, Dako, #P0447). 19. SuperSignal™ West Pico PLUS ECL reagent. 20. Orbital shaker. 21. Imager for chemiluminescence, e.g., ChemiDoc™ Touch Imager and software for data analysis. 2.8 Nanoparticle Tracking Analysis

1. NanoSight NS300 system or ZetaView PMX-120 system. 2. EV preparation (~1–2 μg protein). 3. Ultrapure water. 4. Disposable 1 mL syringe for sample loading.

2.9 Transmission Electron Microscopy

1. EV preparation (~1–2 μg protein). 2. Sterile/filtered PBS. 3. Fixing solution: 1% (v/v) glutaraldehyde. 4. Carbon-coated copper 400 mesh grids (#GSCU400CC) from ProSciTech.

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5. 2% (w/v) aqueous uranyl acetate. 6. Gatan CCD camera coupled to a Jeol JEM-2100 electron microscope (80 kV). 2.10 Proteomics: Sample Preparation

1. EV lysate preparations (+/
depletion with albumin, ~5–10 μg protein).

2.10.1 In-Solution Reduction, Alkylation, and Digestion

2. 1.5 mL protein LoBind tubes or protein LoBind deep 96-well plates (1000 μL). 3. 1 M tetraethylammonium bromide (TEAB) stock solution in LC-MS grade water. 4. 50 mM TEAB, pH 8.0, in LC-MS grade water. For pH adjustment use 1 M hydrochloride in LC-MS grade water. 5. 100 mM TEAB, pH 8.0, in LC-MS grade water. For pH adjustment use 1 M hydrochloride in LC-MS grade water. 6. 100% (v/v) acetone (LC grade). Stored at
20  C. 7. 90% (v/v) acetone in water (LC grade). Stored at
20  C. 8. Lysis buffer: 1% (w/v) SDS, 50 mM TEAB, pH 8.0. 9. 500 mM DTT stock solution in 100 mM TEAB, pH 8.0. Prepare fresh before the reduction step. 10. 1 M iodoacetamide (IAA) stock solution in 100 mM TEAB, pH 8.0. Prepare fresh before the alkylation step. 11. Trypsin, sequencing grade. Store lyophilized or frozen at
80  C. 12. 1.5% (v/v) formic acid (LC-MS grade). 13. Water (LC-MS grade). 14. ThermoMixer with 1.5 mL microfuge tube capacity. 15. Thermostat oven or incubator at 37  C. 16. pH strip, range 1–14. 17. Sonicator with microtip, e.g., Minsox S-4000, 600 W.

2.10.2 StageTip Sample Cleanup

1. Empore SDB-RPS (Styrenedivinylbenzene–reversed phase sulfonate) solid-phase extraction disks for StageTip preparation. 2. SDB-RPS StageTip loading buffer: 1% (v/v) trifluoroacetic acid (TFA) in acetonitrile (ACN). 3. SDB-RPS StageTip wash buffer 1: 1% (v/v) TFA in ACN. 4. SDB-RPS StageTip wash buffer 2: 0.2% (v/v) TFA in 5% (v/v) ACN. 5. SDB-RPS StageTip elution buffer: 20 μL of NH4OH in 4 mL of 60% (v/v) ACN. Elution buffer must be prepared fresh (within 1 h of use) because the pH will begin to increase due to its high volatility, thereby reducing its elution strength.

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6. Water (LC-MS grade). 7. Vacuum centrifuge (lyophilizer). 8. MS loading buffer: 0.07% (v/v) TFA in LC-MS water. This buffer is stable for >6 months at RT. 2.11 Fluorometric Peptide Assay

1. Quantitative Fluorometric Peptide Assay kit. 2. MS loading buffer (see item 8 in Subheading 2.10.2). 3. Peptide samples for analysis. 4. Fluorescence compatible 96-well microplate (black). 5. Fluorescent plate reader compatible with Ex 390 nm/Em 475 nm.

2.12

UHPLC-MS/MS

1. MS loading buffer: 0.07% (v/v) TFA in LC-MS water. 2. Transparent MS sample vials: 300 μL capacity and snap ring vial with PP insert. 3. UHPLC System coupled to a Q Exactive HF-X Hybrid Quadrupole-Orbitrap MS System. 4. Load column: Acclaim PepMap100 C18, 5 μm beads with ˚ pore size. 100 A 5. Analytical/separation column: 50-cm fused-silica emitter reversed-phase PepMap RSLC C18, 75 μm inner diameter, ˚ pore size. 2 μm resin with 100 A 6. Mobile phase A: 0.1% formic acid. 7. Mobile phase B: 0.1% formic acid in acetonitrile.

2.13

Data Analysis

1. MaxQuant software. 2. Microsoft Office Excel. 3. Perseus software (see Note 46) from Max Planck Institute of Biochemistry, Munich, Germany.

3 3.1

Methods PBPL Isolation

1. Whole blood, collected using STRECK RNA tubes, is immediately transferred to the laboratory (same site) avoiding agitation and allowed to sit at room temperature for 30 min (see Notes 1–4). 2. Centrifuge at 1800  g for 10 min at 4  C to separate plasma. 3. Carefully transfer the upper plasma phase to low-protein binding tubes (see Note 5), without disturbing the intermediate buffy coat layer which contains white blood cells and platelets. Normally up to 4–5 mL plasma can be recovered from 10 mL whole blood.

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4. Centrifuge at 2000  g for 15 min at 4  C (see Notes 20 and 21). 5. Carefully transfer the cleared supernatant to new low-protein binding tubes without disturbing the pellet (see Note 5). 6. Store aliquots in low-protein binding tubes at
80  C. 7. Depending on EV isolation method, 0.5–4 mL aliquots (fresh or frozen) are utilized for EV isolation (see Note 6). 3.2 Large EVs Isolation (See Note 22)

1. Centrifuge plasma at 10,000  g for 30 min at 4  C to pellet large EVs. 2. Resuspend the pellet in 1 mL of sterile/filtered PBS. 3. Centrifuge at 10,000  g for 30 min at 4  C to obtain the washed large EVs pellet. 4. The pellet can be resuspended in either SDS sample buffer for PAGE analysis (i.e., Subheadings 3.6 and 3.7) or sterile PBS for other downstream applications (i.e., Subheadings 3.8–3.10).

3.3 Small EVs Isolation

1. Briefly vortex the bead slurry and then load 40 μL of bead slurry solution into a 1.5 mL tube per sample.

3.3.1 CD63 Exo-FLOW Capture Kit

2. Place tubes on magnetic stand for 2 min. 3. Carefully remove the supernatant making sure to not disturb the magnetic bead pellets. 4. Remove the samples from magnetic stand and add 500 μL of Bead Wash buffer. 5. Invert the tubes a few times, place samples on magnetic stand for 2 min, and discard the buffer. Wash steps are repeated one more time and all liquid removed. Beads are on the side of the tube. 6. After removing tubes from magnetic stand, add 10 μL of CD63 biotinylated capture antibody, using the pipette tip to move the beads to the bottom of the tube, and mix by pipetting up and down three times. 7. Place tubes on ice for 2 h, flicking the tubes every 30 min to gently mix. 8. Add 200 μL of Bead Wash buffer, mix by flicking, and place samples on magnetic stand for 2 min. 9. Carefully remove the supernatant making sure not to disturb the magnetic bead pellets. 10. Add 500 μL of Bead Wash buffer after removing the samples from magnetic stand. Beads are washed by inverting 2–3 times and flicking the tubes a few times.

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11. Place samples on magnetic stand for 2 min and discard the buffer. Repeat wash steps for a total of three washes and remove all liquid. 12. Resuspend capture antibody beads with 400 μL of Bead Wash buffer per sample. 13. Add 100 μL (100–200 μg of protein) of ExoQuick™ precipitated EVs (see Subheading 3.3.2) to each bead sample for a total volume of 500 μL. 14. Incubate on a rotating rack at 4  C overnight for capture. 15. Place samples on magnetic stand for 2 min. 16. Carefully remove the supernatant making sure to not disturb the magnetic bead pellets. 17. Add 500 μL of Bead Wash buffer after removing the samples from magnetic stand. Beads are washed by inverting 2–3 times and flicking the tubes a few times. 18. Place samples on magnetic stand for 2 min and discard the buffer. Repeat wash steps for a total of two washes and remove all liquid. 19. Add 240 μL of Exosome Stain Buffer and 10 μL of Exo-FITC exosome stain for a final volume of 250 μL per sample, and place tubes on ice for 2 h (flicking the tubes every 30 min to gently mix). 20. Place samples on magnetic stand for 2 min. 21. Carefully remove the supernatant making sure to not disturb the magnetic bead pellets. 22. Add 500 μL of Bead Wash buffer after removing the samples from magnetic stand. Beads are washed by inverting 2–3 times and flicking the tubes a few times. 23. Place samples on magnetic stand for 2 min and discard the buffer. Repeat wash steps for a total of three washes and remove all liquid. 24. Resuspend samples in 300 μL of Bead Wash buffer for flow cytometry/sorting (avoid vortexing prior to loading into FACS instrument). 25. For EV Stain removal and elution (if desired), place samples on the magnetic stand for 2 min and remove buffer. 26. Add 300 μL of Exosome Elution Buffer and invert samples a few times. Vortexing is not recommended. Mix by flicking. 27. Incubate on a rotating rack or shaker at 25  C for 2 h. 28. Place samples on magnetic stand for 2 min. 29. Carefully transfer the supernatant containing eluted EVs to a fresh tube, making sure to not disturb the magnetic bead pellets. Discard the beads after use.

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1. Add 5 μL thrombin to 500 μL of plasma (see Notes 10, 11, and 23) followed by gentle mixing (flick the tube). 2. Incubate at RT for 5 min. 3. Centrifuge at 9400  g for 5 min. Transfer the supernatant to a new tube. 4. Filter the supernatant (see Note 10) to exclude particles larger than 0.2 μm (for small EVs isolation). 5. Add appropriate volume of ExoQuick™ (i.e., 67 μL for each 250 μL of plasma) and mix by inverting or flicking the tube. 6. Incubate on ice for 30 min and then centrifuge at 3000  g for 10 min (RT or 4  C) to pellet the EVs from the solution. 7. Aspirate and discard the supernatant, making sure not to disturb the pellet containing EVs. Resuspend the pellet in 200 μL of provided Buffer B, proceeding to protein quantitation (see Subheadings 2.5, 2.6, 3.5, 3.6 and Note 24). 8. Add 200 μL of provided Buffer A. 9. Loosen the screw cap, snap off the bottom closure, and place the purification column into a collection tube (save the bottom closure for step 11). Centrifuge the column at 1000  g for 30 s to remove storage buffer. 10. Discard the flow through and replace back the column into the collection tube and wash twice by applying 500 μL of Buffer B on top of the resin. 11. Centrifuge at 1000  g for 30 s. Discard the flow through. 12. Prime the column by applying 100 μL of Buffer B on top of the resin. The entire content from steps 6 to 7 (400 μL or up to 4 mg of total protein content) is then added (plug the bottom of the column with the bottom closure, see step 8) and mixed at RT on a rotating shaker for 5 min. 13. Loosen the screw cap, remove the bottom closure, and immediately transfer the column to a 2 mL Eppendorf tube. 14. Centrifuge at 1000  g for 30 s to obtain purified EVs. 15. EVs are aliquoted depending on intended use, and either used immediately or stored at
80  C.

3.3.3 exoEasy Kit

1. All steps should be performed at RT. 2. Filter 2–4 mL of plasma (see Notes 12, 23, and 25) to exclude particles larger than 0.2 μm (for small EVs isolation). 3. Add one volume of buffer XBP to one volume of sample and mix well by gently inverting the tube five times. 4. Load mixture onto the exoEasy spin column and centrifuge at 500  g for 1 min.

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5. Discard flow through and place column back into the same collection tube. Add 10 mL of buffer XWP, and residual buffer from the column is removed by centrifuging at 5000  g for 5 min. Discard the flow-through and the collection tube. 6. Place the spin column into a fresh collection tube. 7. Add 400 μL of buffer XE to the membrane, incubate for 1 min, and centrifuge at 500  g for 5 min. Collect the eluate containing EVs. 8. Reapply the EV containing eluate to the spin column membrane, incubate for 1 min, and centrifuge at 5000  g for 5 min. 9. Aliquot depending on intended use (see Notes 26 and 27), and either use immediately or store at
80  C. 3.3.4 Purification Mini Kit

1. All steps should be performed at RT. 2. Add 3 mL of nuclease-free water to 1 mL of plasma (see Notes 13, 23, and 28). 3. Add 100 μL of ExoC buffer. 4. Add 200 μL of Slurry E (mix well Slurry E prior to use). 5. Vortex sample for 10 s and incubate for 5 min. 6. Vortex for 10 s and centrifuge at 930  g (2000 rpm) for 2 min. Discard the supernatant. 7. Add 200 μL of ExoR buffer, mix well by vortexing for 10 s, and incubate at RT for 5 min. 8. Vortex the sample for 10 s, centrifuge at 500 rpm for 2 min, and transfer the supernatant to a Mini Filter Spin column assembled with an elution tube. 9. Centrifuge at 3400  g (6000 rpm) for 1 min. 10. EV containing eluate is collected and aliquoted depending on intended use, and either used immediately or stored at
80  C.

3.4 Albumin Depletion (See Notes 14, 15, and 29)

1. Resuspend well the resin by shaking the resin bottle. 2. Transfer 400 μL of the slurry (corresponding to 200 μL settled resin volume) into a spin column (loosely cap the column). 3. Twist off the bottom closure of the spin column and place the spin column into a collection tube. 4. Centrifuge at 12,000  g for 1 min to remove excess liquid. Discard the flow-through and place the spin column back into the same collection tube. 5. Add 200 μL of Binding/Wash Buffer into the spin column. Centrifuge at 12,000  g for 1 min and discard the flowthrough. Place the spin column into a new collection tube.

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6. Load (maximum) 50 μL of albumin-containing EV sample onto the resin and incubate for 1–2 min at RT. Centrifuge at 12,000  g for 1 min. Collect the flow-through. 7. Reapply the flow-through to the spin column, and incubate for 1–2 min at RT to ensure maximal albumin binding. Thus, centrifuge at 12,000  g for 1 min, retaining the flow-through. 8. Add 50 μL of Binding/Wash Buffer for each 200 μL of resin used and centrifuge at 12,000  g for 1 min to wash the resin (to release unbound proteins). Retain the flow-through. Place spin column into a new collection tube. 9. Step 8 is repeated one more time and the three EV albumindepleted fractions are combined (see Notes 30 and 31). 3.5 EV Lysate Preparation (See Notes 16–18)

1. Add RIPA buffer and PI cocktail (final concentrations 1) to EV samples. 2. Vortex sample for 15 s and incubate for 1 h at 4  C (vortexing every 10 min). 3. Centrifuge at 16,000  g for 15 min at 4  C to remove insoluble debris. 4. Collect the supernatants and store at
30  C (up to 6 months) or proceed to protein quantitation (see Subheading 3.6).

3.6 Protein Quantitation (See Note 19)

1. A set of standard dilutions with BSA is prepared starting with 200 μg/mL, performing dilutions in 1% SDS solution to 0.5 μg/mL, and placed at RT (1% SDS solution can crystallize on ice). 2. EV lysates are thawed on ice. 3. Load in a clear flat-bottom 96-well plate 150 μL of EV eluent/ lysis solution (blank), 150 μL of each standard BSA dilution, and 150 μL of diluted EV lysate sample (2 μL in 148 μL of 1% SDS solution). 4. Add 150 μL of the WR to each well and mix the plate (sealed with adhesive film) thoroughly on a plate shaker for 30 s. Plate is covered, incubated at 37  C for 2 h and then cooled to RT. 5. Absorbance is measured at 562 nm on a plate reader. 6. Determine the protein concentration of each EV sample using the BSA standard curve (plotting the average Blank-corrected 562 reading for each BSA standard vs its concentration in μg/ mL). Multiply by the diluting factor.

3.7 Western Blot Analysis (See Notes 32–35)

1. Mix EV samples (7–20 μg protein) with SDS-PAGE sample buffer at a final concentration of 1, heat for 5 min at 95–100  C, and cool on ice before proceeding to electrophoresis which is performed at constant 140–180 V for 45–60 min (until the dye reaches the reference line).

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2. Activate PVDF membranes: Soak in 100% methanol for 30 s and then in ice-cold transfer buffer. 3. Following electrophoresis, perform a tank (wet) blotting procedure: Sandwich gel and membrane between sponge and blotting paper (sponge/2 blotting paper/gel/membrane/2 blotting paper/sponge) and clamp tightly after ensuring no air bubbles have formed between the gel and membrane using a roller. 4. Insert the cassette/sandwich into the tank (the black side of the cassette should face the black side of the central core) together with the frozen cooling unit. Complete transfer at 90–100 V for 60–90 min (protein transfer is confirmed by the presence of the protein standard on the membrane and further confirmed by Ponceau Red staining). 5. Block membranes with blocking buffer for 1 h at 4  C. Care should be taken not to touch and disrupt the membrane (only the immediate corners/edges). 6. Probe membranes with primary antibodies overnight in TTBS. 7. Incubate with appropriate secondary antibody for 1 h. 8. All antibody incubations are carried out using gentle orbital shaking at RT. 9. Western blots are washed six times in TTBS for 5 min after each incubation step and subsequently visualized using ECL reagents, developed using ChemiDoc™ Touch Imager, and analyzed with the aid of Image Lab software. 3.8 Nanoparticle Tracking Analysis (See Notes 32 and 36)

1. Dilute EV sample to a final volume of 1 mL (final concentration 1 μg/mL in ultra-pure water, for example, 1:1000 dilution, depending on the concentration of EVs).

3.8.1 NanoSight NS300 System

2. Load sample onto NS300 flow-cell top plate using syringe pump. Avoid the introduction of air bubbles into the chamber. 3. Operation settings: Detection threshold ¼ 10; Flow rate ¼ 50; Temperature ¼ 25  C (standard measurement). 4. Camera level and screen gain need adjustment until particles are visible. 5. Conditions for video capture are as follows: Number of capture ¼ minimum 3 (for triplicate); Capture duration ¼ 60 s. 6. For acquisition, change base file name to preferred destination folder. 7. Click “Create” and “Run script” to begin capture and analysis with NTA software, ensuring that detection threshold is set to include as many particles as possible with the restriction that 20–100 red crosses are counted while only 50 μg based on protein amount) for proteomics and other assays, it is necessary to use as many cells as possible (i.e., a minimum of 50
150 mm culture plates, corresponding to ~1
109 cells). In addition, it is better to purify EVs from large volumes of conditioned medium, because the loss of EVs is decreased in large starting volumes.

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2. Serum contains ample extracellular proteins at high concentrations, as well as serum-derived EVs, which contaminate cultured cells and the culture plate. Without careful washing, these components are co-purified during the isolation of EVs. To entirely remove these contaminants, use PBS corresponding to 1.25-fold the volume of culture medium to wash the cells (e.g., 25 mL PBS and 20 mL culture medium in a 150 mm culture plate). 3. Cell culture with serum-containing media can cause the co-purification of serum-abundant extracellular proteins and serum-derived EVs, which can lead to the identification of serum-derived components at the expense of vesicular proteins because of the increased dynamic range of the sample. To overcome this problem, EV-depleted serum-containing media or serum-free media are used for proteomics analysis of EVs. Despite the use of EV-depleted serum-containing media, however, considerable amounts of serum-abundant extracellular proteins would remain non-specifically bound to the purified EVs without a selective enrichment step, such as density gradient ultracentrifugation. In serum-free cell culture, we recommend to check the viability of cells using an appropriate assay (e.g., annexin V staining or the PARP cleavage apoptosis assay). Cell viability of 95% is generally acceptable for proteomic analysis of EVs. Cell culture in serum-free medium for >24 h can cause cell death and thus generate apoptotic vesicles, membrane fragments, and other cellular debris. 4. Low-speed centrifugation will pellet dead cells, large membrane fragments, and cellular debris. It is recommended to use fixed-angle rotors for these steps. If swinging bucket rotors are used, the supernatant should be transferred to other centrifuge tubes using a pipette, leaving about 0.5 cm supernatant above the pellet. 5. After conditioned medium is collected, it is strongly recommended that purification of EVs is conducted as soon as possible. Alternatively, it is possible to store conditioned medium at 80  C after quick-freezing using liquid nitrogen, but this likely leads to breakdown of the EVs. 6. Although there is an advantage of serum-free culture to isolate the high-purity EVs in proteomics (see Note 3), it starves the cells and may cause the change of physiological release rate and molecular composition of EVs. Moreover, long-time serumfree culture condition can induce the cellular death. To avoid these concerns, EV-depleted fetal bovine serum is widely used. In this study, EV-depleted serum culture for 3 days in U373 and U373vIII showed the similar growth rate with about 97% viability (see Fig. 3a).

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7. About 200–250 μL is the minimum concentrated volume from starting volume of 15 mL in a Filter Unit. From the concentration by four Filter Units, the total volume is approximately 1 mL. After checking the exact volume by 1 mL pipet, additional HBS is added up to final 1.92 mL. Careful pipetting should be necessary not to generate bubble because concentrated medium is viscous. 8. To determine the density of iodixanol density gradient fractions, it is necessary that control SW41 Ti ultracentrifuge tubes (containing the same iodixanol gradient without sample) are co-centrifuged with the sample. Iodixanol can form selfgenerating gradients during ultracentrifugation, and thus, the densities of iodixanol density gradient fractions are changed after ultracentrifugation (see Fig. 1a). 9. The EV-enriched fraction is determined by Western blotting for vesicular marker proteins, such as CD63 and CD81. Generally, EVs are localized in the third fraction (density, ~1.096 g/mL) from the top (see Fig. 1a). To further determine the enrichment of EVs, the enriched fraction should be analyzed by electron microscopy, which is the gold standard for visualizing EVs (see Fig. 1b). Moreover, dynamic light scattering analysis and nanoparticle tracking analysis (NTA) are used to determine the size distribution and particle concentration of EVs (see Fig. 2a, b). Dynamic light scattering analysis and NTA have determined average diameters of SW480-derived EVs of 176.70  7.35 nm and 172.41  16.42 nm, respectively. NTA also shows that the particle concentration is 7.56  0.49
109 particles/μg of total protein.

a

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Number (%)

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Fig. 2 Size distribution and particle concentration of purified SW480-derived EVs. (a) The size distribution of EVs was measured by dynamic light scattering and indicated that the average diameter is 176.70  7.35 nm. (b) NTA showed that the average diameter of purified EVs is 172.41  16.42 nm, with a particle concentration of 7.56  0.49
109 particles/μg

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10. EVs can be stored in the density gradient fraction at 80  C as 100 μL aliquots after quick-freezing using liquid nitrogen. Avoid repeated freezing and thawing. 11. The EV-enriched fraction was determined by NTA and Western blotting for vesicular marker proteins, including CD81, TSG101, syntenin-1, representing that the third fraction (density, ~1.096 g/mL) from the top was enriched with EVs (see Fig. 3b, c). Moreover, electron microscopy images showed the vesicular morphological features (see Fig. 3d) and NTA showed the average diameters of U373- and U373vIII-derived EVs of 215.9  1.1 nm and 237.6  2.0 nm with differential release rates, respectively (see Fig. 3e, f).

b

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Fig. 3 Purification of U373- and U373vIII-derived EVs from EV-depleted media. (a) Cells were incubated with 10% fetal bovine serum media or EV-depleted media for 3 days. MTS cell proliferation assay showed that EV-depleted media do not affect cell proliferation in either cell population (n ¼ 3). FBS, fetal bovine serum. (b) The particle concentration of each fraction was determined by NTA. (c) Fractions of iodixanol density gradients in U373 and U373vIII EVs were analyzed by Western blotting. CD81, TSG101, and syntenin-1, marker proteins of EVs, were mainly detected in Fraction 3. (d) Morphological features of EVs isolated from cultures of U373 and U373vIII cells are reminiscent of vesicle. (e) The size distribution of EVs as measured by NTA indicating an average diameter of 215.9  1.1 nm and 237.6  2.0 nm for U373 EVs and U373vIII EVs, respectively. (f) Table for the EV release characteristics of U373 and U373vIII cells in this condition (n ¼ 3). Reprinted and modified with permission from Mol. Cell. Proteomics, 17 (10), 1948–1964. Choi D et al., the impact of oncogenic EGFRvIII on the proteome of extracellular vesicles released from glioblastoma cells, with permission from The American Society for Biochemistry and Molecular Biology [5]

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12. Because EVs have lipid bilayers, denaturing reagents, such as urea, thiourea, and guanidine hydrochloride, cannot access intraluminal proteins and trypsin cannot digest them. Methanol/chloroform precipitation can effectively remove lipids as well as precipitate proteins without salts. With this method, it is possible to precipitate ~2 μg protein, but it is recommended that >10 μg of vesicular protein be used in methanol/chloroform precipitation for high yield. 13. About 20 μg vesicular protein is enough to identify several hundreds of proteins without peptide fractionation (e.g., ion-exchange chromatography, isoelectric focusing) using reverse-phase liquid chromatography coupled with mass spectrometry. To identify more proteins using peptide fractionation, appropriate amounts of vesicular proteins should be prepared. 14. Drying of the precipitated proteins for a long period results in difficulty in complete re-solubilization in protein denaturation buffer. 15. C18 spin columns can bind up to 30 μg of total peptide.

Acknowledgments We thank to Gyeongyun Go for helping in isolation and characterization of EVs and Jaewook Lee for analysis of the EVs in transmission electron microscope. This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MEST) (No. 2014023004, 2018R1A2A1A05079510, and 2012R1A1A2042534) and by the Foundation Grant (FDN 143322) from Canadian Institutes for Health Research to J.R. References 1. Choi DS, Kim DK, Kim YK, Gho YS (2013) Proteomics, transcriptomics and lipidomics of exosomes and ectosomes. Proteomics 13:1554–1571 2. Choi DS, Kim DK, Kim YK, Gho YS (2015) Proteomics of extracellular vesicles: exosomes and ectosomes. Mass Spectrom Rev 34:474–490 3. Kim DK, Kang B, Kim OY, Choi DS, Lee J, Kim SR, Go G, Yoon YJ, Kim JH, Jang SC, Park KS, Choi EJ, Kim KP, Desiderio DM, Kim YK, Lotvall JO, Hwang D, Gho YS (2013) EVpedia: an integrated database of high-throughput data for systemic analyses of

extracellular vesicles. J Extracell Vesicles 2:20384 4. Choi D, Spinelli C, Montermini L, Rak J (2019) Oncogenic regulation of extracellular vesicle proteome and heterogeneity. Proteomics 19:e1800169 5. Choi D, Montermini L, Kim DK, Meehan B, Roth FP, Rak J (2018) The impact of oncogenic EGFRvIII on the proteome of extracellular vesicles released from glioblastoma cells. Mol Cell Proteomics 17:1948–1964 6. Thery C, Amigorena S, Raposo G, Clayton A (2006) Isolation and characterization of exosomes from cell culture supernatants and

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biological fluids. Curr Protoc Cell Biol Chapter 3:Unit 3.22 7. Muralidharan-Chari V, Clancy J, Plou C, Romao M, Chavrier P, Raposo G, D’SouzaSchorey C (2009) ARF6-regulated shedding of tumor cell-derived plasma membrane microvesicles. Curr Biol 19:1875–1885 8. Kato K, Kobayashi M, Hanamura N, Akagi T, Kosaka N, Ochiya T, Ichik i T (2013) Electrokinetic evaluation of individual exosomes by on-chip microcapillary electrophoresis with laser dark-field microscopy. Jpn J Appl Phys 52:06GK10 9. Choi DS, Park JO, Jang SC, Yoon YJ, Jung JW, Choi DY, Kim JW, Kang JS, Park J, Hwang D, Lee KH, Park SH, Kim YK, Desiderio DM, Kim KP, Gho YS (2011) Proteomic analysis of microvesicles derived from human colorectal cancer ascites. Proteomics 11:2745–2751 10. Mathivanan S, Lim JW, Tauro BJ, Ji H, Moritz RL, Simpson RJ (2010) Proteomics analysis of A33 immunoaffinity-purified exosomes released from the human colon tumor cell line LIM1215 reveals a tissue-specific protein signature. Mol Cell Proteomics 9:197–208

11. Kang D, Oh S, Ahn SM, Lee BH, Moon MH (2008) Proteomic analysis of exosomes from human neural stem cells by flow field-flow fractionation and nanoflow liquid chromatography-tandem mass spectrometry. J Proteome Res 7:3475–3480 12. Dean WL, Lee MJ, Cummins TD, Schultz DJ, Powell DW (2009) Proteomic and functional characterisation of platelet microparticle size classes. Thromb Haemost 102:711–718 13. Looze C, Yui D, Leung L, Ingham M, Kaler M, Yao X, Wu WW, Shen RF, Daniels MP, Levine SJ (2009) Proteomic profiling of human plasma exosomes identifies PPARgamma as an exosome-associated protein. Biochem Biophys Res Commun 378:433–438 14. Hong BS, Cho JH, Kim H, Choi EJ, Rho S, Kim J, Kim JH, Choi DS, Kim YK, Hwang D, Gho YS (2009) Colorectal cancer cell-derived microvesicles are enriched in cell cycle-related mRNAs that promote proliferation of endothelial cells. BMC Genomics 10:556 15. Wessel D, Flugge UI (1984) A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal Biochem 138:141–143

Chapter 12 Isolation of Proteins from Extracellular Vesicles (EVs) for Mass Spectrometry-Based Proteomic Analyses Prabal Subedi, Michael Schneider, Michael J. Atkinson, and Soile Tapio Abstract Extracellular vesicles (EVs) are freely circulating nano/micrometer-sized membrane-bound vesicles released by various cell types. Their cargo consists of proteins, lipids, metabolites, and different types of RNA molecules reflecting the origin of the secreting cell type or tissue. Since the EV cargo is constantly changing in response to pathological status or different environmental stressors, it has been extensively studied in the quest for biomarkers, especially in the cancer research. Mass spectrometry (MS)-based proteome analysis is a powerful tool to elucidate the protein cargo in EVs. This chapter describes and characterizes three MS-compatible lysis methods, namely by using urea, guanidium hydrochloride, and radioimmunoprecipitation buffer for isolating proteins from EVs. Key words Extracellular vesicles (EVs), Exosomes, Mass spectrometry, Proteomics, Data-dependent acquisition, Sample preparation

1

Introduction Extracellular vesicles (EVs) are cell-secreted membrane-bound vesicles. They can be further subdivided into exosomes (30–150 nm) [1], microvesicles (50–1000 nm) [2], and apoptotic bodies (1–4 μm) [3] and contain a cargo of lipids [4], metabolites [5], nucleic acids [6], and proteins [7]. EVs are secreted by all forms of organisms [8, 9] and are present in most body fluids, including plasma [10], saliva [11], and cerebrospinal fluid [12]. They play a role in cellular signaling [13], homeostasis [2], and tumor metastasis [13]. Moreover, they contain a cargo of aggregated proteins, which could be involved in the propagation of neurodegenerative diseases [14]. Protein cargo of EVs was observed to be changed when the cells producing them were subjected to cellular stress [6]. Similarly, it has been observed that there is a difference in EV-proteome between cancer patients and healthy controls [15]. For studies related to EV-proteomics, it is imperative that these vesicles are lysed in a manner to enable the identification of as

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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many proteins as possible. Vesicles can be lysed using various methods such as mechanical force or introduction of hypo/hypertonic solutions or detergent-containing buffers. A good lysis buffer should be able to disrupt the phospholipid bilayer and keep the released proteins soluble [16]. We discuss here three different lysis methods that are compatible for mass spectrometry-based proteomic analyses.

2

Materials Use ultrapure water (resistance >18.2 MΩ/cm) for buffer preparation (see Note 1). 1. High-speed centrifuge and ultracentrifuge. 2. Cell counter. 3. Nanoparticle size and concentration counter. 4. T175 cell culture flasks. 5. 0.22 μm filter. 6. EV-depleted fetal bovine serum (FBS) (see Note 2). 7. EV-depleted cell culture medium (see Note 3). 8. Trypsin–EDTA solution (TE) solution: 0.025% trypsin and 0.01% EDTA in PBS, pH 7.4. 9. Sonicator (ultrasonic frequency of 37 kHz). 10. Thermomixer. 11. Vortex. 12. Dounce homogenizer. 13. Protease and phosphatase inhibitors: Add to the lysis buffers (see steps 15–17) just before use (see Note 4). 14. PBS (Phosphate-buffered saline): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4. Weigh 8 g of NaCl, 200 mg of KCl, 1.44 g of Na2HPO4, 240 mg KH2PO4, and add ~900 mL water. Use HCl to adjust the solution to pH 7.4. Fill up the volume to 1 L. 15. RIPA buffer: 25 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodiumdeoxycholate, 0.1% SDS. Weigh 30 mg of Tris-base and fill up to 7 mL of water in a falcon tube. Adjust the pH to 7.6 with HCl. Then add 88 mg of NaCl, 100 mg of NP-40, 100 mg of sodium deoxycholate, 10 mg of SDS, and dissolve well. Fill the volume up to 10 mL. 16. Urea-Thiourea buffer: 7 M Thiourea, 2 M Urea, 30 mM TrisHCl, pH 7.5. Weigh 5.33 g of thiourea, 1.20 g of urea, and 47 mg of Tris-base and dissolve well. Use HCl to adjust to

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pH 7.6 and then fill up to 10 mL. Slightly warm the solution to better dissolve the solutes. 17. 6 M Guanidium-hydrochloride (Gu-HCl) buffer. Weigh 5.73 g of Gu-HCl in 10 mL water and gently heat the solution to 35
C to completely dissolve Gu-HCl.

3 3.1

Methods EV-Cell Culture

1. Wash adherent cells (see Note 5) twice in T175 flask with PBS. 2. Add 2 mL TE solution to flask and incubate for 10–15 min (see Note 6). 3. Suspend the cells in 10 mL EV-depleted medium and determine the cell number. 4. Seed 6  106 cells, in a total of 25 mL EV-depleted medium, in each T175 flask and incubate for 72 h.

3.2

EV Isolation [17]

1. Combine conditioned medium from T175 cell culture flasks and keep on ice (4
C). 2. Centrifuge at 300  g (4
C) for 10 min to remove cells and transfer the supernatant into new tubes. 3. Centrifuge the collected supernatants at 10,000  g (4
C) for 10 min. 4. Pass the supernatant through a 0.22 μm filter for the removal of cellular fragments, big vesicles and apoptotic bodies (see Note 7). 5. Transfer the filtrate into ultracentrifuge-suitable tubes. 6. Centrifuge the tubes at 100,000  g (4
C) for 2 h (see Note 8). 7. Place the tubes on ice. 8. Carefully discard the supernatant in order not to destroy the vesicle pellet. 9. Add approx. 500 μL of PBS to the tubes and vortex vigorously for 15 s. 10. Combine the EV suspensions gathered from different tubes in one tube and fill with PBS. 11. Centrifuge again at 100,000  g (4
C) for 2 h and carefully discard supernatant. 12. Resuspend the pellet in the desired amount of PBS. 13. Sonicate the EVs on ice water for 2 min and store at 4
C (see Note 9).

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3.3 EV Lysis with Different Buffers 3.3.1 Lysis with RIPA Buffer

1. Add 50 μL of lysis buffer to 2 μL (see Note 10) of EV-pellet (~109 particles) in 1.5 mL Eppendorf tubes. 2. Place on thermomixer at 4
C and shake at 650 rpm for 30 min. 3. Sonicate the tubes on ice-cold water for 30 s. 4. Place them on the thermomixer again for another 15 min. 5. Submit the samples for FASP digestion [18] and subsequent LC-MS/MS analyses.

3.3.2 Lysis with UreaThiourea Solution

1. Add 50 μL of lysis buffer to 2 μL of EV-pellet. 2. Homogenize the pellet with a dounce homogenizer with 20 up and down movements while on ice (see Note 11). 3. Place the samples on ice-cold sonication bath and sonicate for 10 s. 4. Take them out and place on ice for 10 s. 5. Repeat the sonication procedure five times. 6. Submit for FASP digestion and subsequent LC-MS/MS analyses (see Notes 12 and 13).

3.3.3 Lysis with Guanidium-Hydrochloride (Gu-HCl) Solution

1. Add 50 μL of lysis buffer to 2 μL of EV-pellet. 2. Vortex the samples for 2 min. 3. Place the samples on a thermomixer at 25
C and 1000 rpm for 20 min. 4. Vortex the samples again for 2 min. 5. Sonicate the samples on ice-cold sonication bath for 30 s. 6. Cool the samples on ice-cold sonication bath for 2 min. 7. Repeat the sonication procedure for three times. 8. Submit for FASP digestion and subsequent LC-MS/MS analyses.

4

Notes 1. Only chemicals with analytical grade should be used. Prepare lysis buffers fresh and keep the buffers as well as EV-pellets on ice at all times unless otherwise specified. 2. For the removal of EVs present in FBS, centrifuge at 100,000  g (4
C, 14 h). Pass the supernatant through a 0.22 μm filter under sterile conditions. Use this filtrate for cell culture work. 3. The cell culture medium depends on the used cell line. For our work, a Dulbecco’s Modified Eagle Serum (Gibco™ high

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glucose, GlutaMAX supplement) was mixed with EV-depleted FBS in a 90:10 ratio, i.e., 90 mL DMEM and 10 mL EV-depleted DMEM. 4. Dissolve one tablet of PhosSTOP™ phosphatase inhibitor and one tablet of complete EDTA-free protease inhibitor in 10 mL of the lysis buffer. 5. We used cancer cell line BHY (ACC 404). 6. All incubations were performed in a humidified atmosphere with 5% CO2 and 37
C (depends on cell line). 7. The filtrate can be kept at 4
C for up to 7 days. 8. Mark the side of the tubes where the pellet is supposed to be since the pellet is not visible most of the times. 9. Long-term storage at 20
C or  80
C (avoid repeated freeze–thaw cycles). 10. Here, 230 mL conditioned media yielded 109 particles per μL. Use 50 μL of buffer for a maximum of 15 μL pellet. Scale accordingly. 11. Place the dounce homogenizer on ice when not in use. 12. FASP or filter-aided sample preparation is a standard technique at our research institute. Because FASP is explained in detail elsewhere [18], we do not elaborate it here. 13. We focus on small EVs (10% cross-reactivity to H3K4me2)

meaningful scale of histone modification density over a defined genomic locus [22, 23, 26]. This approach enables truly universal quantitative comparisons across experimental samples, and more broadly across research groups. Recombinant nucleosome spike-in controls can be used with either X-ChIP or N-ChIP for specific histone PTMs (Fig. 1). However, because these controls are native nucleosomes, they are best suited for N-ChIP workflows in order to most accurately quantitate endogenous chromatin. Additionally, since N-ChIP methods show

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higher IP efficiency, reduced artifacts, and improved resolution relative to X-ChIP, the protocol below describes detailed N-ChIP methods (with notes how this workflow may be adapted to capture transient chromatin interactions using X-ChIP).

2

Materials This protocol is adapted from previously described native ChIP methodology [26]. Starting with at least 30 million cells will allow sufficient material for five to twenty ChIP samples, depending on efficiency of the antibody and abundance of the target PTM. While some material is lost during processing steps such as sucrose gradient and hydroxyapatite (HAP) chromatography, the resulting chromatin is free of debris and associated proteins that can interfere with antibody binding [26]. In this protocol, native cells are used to prepare sucrose-purified nuclei (Fig. 3), which can be aliquoted and frozen for future IP. Chromatin is then digested into mononucleosomes using MNase (Fig. 4), and chromatin-associated proteins removed by HAP chromatography. The mononucleosomes are loaded onto bead-immobilized antibodies, immunoprecipitated, and the resulting target-associated DNA is isolated and prepared for qPCR or NGS. Using this workflow, relatively small cellequivalents (as little as 1
105) are sufficient to generate highquality sequencing tracks (see Note 1).

Fig. 3 Monitoring cell lysis and assessing isolated nuclei integrity is critical to ensure quality results. Images depict morphology of intact cells (top) and nuclei (middle). Representative examples of commonly observed problems are also shown (bottom), which would indicate that the nuclei preparation must be reattempted paying close attention to key protocol notes

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Fig. 4 Example experiment optimizing chromatin fragmentation by micrococcal nuclease (MNase) digestion. (a) Native nuclei isolated from K-562 cells were treated with the indicated concentrations of MNase for 10 min at 37  C. DNA was purified and subjected to chip-based capillary electrophoresis (Agilent Bioanalyzer) to assess fragment size distribution. The virtual gel shows enrichment of progressively smaller fragment sizes (bp DNA, left) as MNase concentration is increased. (b) Representative Bioanalyzer traces are shown for a case of over digestion caused by excessive MNase (top, fragmented peak with a left shoulder of smaller fragment sizes), optimal digestion (middle, clear peak at 150 bp showing >95% enrichment of mononucleosome-sized fragments), and under digested samples caused by insufficient MNase (bottom, little enrichment of 150 bp fragments and a strong peak of large bp fragments) 2.1 Nuclei Sample Preparation

1. Cells for study: grown to 80–85% confluency under recommended culture conditions. Ensure the population is in good health (viability > 95% when assessed by Trypan blue staining). 2. 0.4% Trypan Blue Stain. 3. Automated Cell Counter or hemocytometer and associated slides. 4. Microscope with brightfield or phase contrast capability. 5. Refrigerated Centrifuge, max. capacity 4
1000 mL. 6. 1
PBS: Dilute 10
PBS 1:10 in Molecular Biology Grade H2O (e.g., 50 mL 10
PBS in 450 mL water). Store at 4  C. 7. 10
Buffer N: 150 mM Tris Base, 150 mM NaCl, 600 mM KCl, 50 mM MgCl2, 10 mM CaCl2, pH 7.5. Filter-sterilize and store at 4  C. 8. 1
Buffer N: Dilute 10
Buffer N 1:10, add 8.5% (w/v) sucrose, 1 mM DTT, 200 mM PMSF, 1
Protease Inhibitor, 50 mg/mL BSA. Filter-sterilize, place on ice, and use fresh the day of preparation. 9. 22-gauge needle and 10 mL syringe (for cell trituration). 10. 2
Modified Lysis Buffer: 10 mL of 1
Buffer N, 1
Protease Inhibitor, 50 mg/mL BSA, 0.2% (v/v) NP-40 Alternative.

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Place on ice and use fresh the day of preparation. Important: NP-40 Alternative concentration should be optimized for the cell type of interest (see Note 2). 11. Sucrose Cushion: Dilute 10
Buffer N to 1
, add 30% (w/v) sucrose, 1 mM DTT, 200 mM PMSF, 1
Protease Inhibitor, 50 mg/mL BSA. Filter-sterilize, place on ice, and use fresh the day of preparation. 12. 2 M NaCl. 13. Vortex. 14. Sonicator. 15. NanoDrop™ Microvolume UV-Vis Spectrophotometer. 2.2 Prep Chromatin for IP

1. DNA barcoded recombinant nucleosome spike-in controls (use as a positive/negative spike-in control and normalization standard for histone PTMs) (see Note 3). 2. Spike-in Drosophila chromatin and primers. 3. Thermomixer, capacity 24
1.5 mL. 4. Micrococcal nuclease (MNase). 5. 10
MNase Stop Buffer: 110 mM EGTA, 110 mM EDTA. Autoclave and store at RT. 6. 5 M NaCl. 7. Refrigerated microcentrifuge. 8. Pan balance. 9. CHT Ceramic Hydroxyapatite (HAP) Type I, 20 μM. 10. Low bind microcentrifuge tubes. 11. HAP Wash Buffer 1: 5 mM NaPO4, 600 mM NaCl, 1 mM EDTA, pH 7.2. Filter-sterilize and store at 4  C. Add 200 μM PMSF fresh the day of use. If profiling histone acetylation targets, add 1 μL Trichostatin A (TSA) fresh the day of use (see Note 4). 12. Compact variable speed/angle vertical rotator. 13. Ultrafree MC-HV Centrifugal filter (0.45 μM). 14. HAP Wash Buffer 2: 5 mM NaPO4, 100 mM NaCl, 1 mM EDTA, pH 7.2. Filter-sterilize and store at 4  C. Add 200 μM PMSF fresh the day of use. 15. HAP Elution Buffer: 500 mM NaPO4, 100 mM NaCl, 1 mM EDTA, pH 7.2. Filter-sterilize and store at room temperature (RT). Add 200 μM PMSF fresh the day of use. 16. ChIP Buffer 1: 25 mM Tris–HCl, 5 mM MgCl2, 100 mM KCl, 10% (v/v) glycerol, 0.1% (v/v) NP-40 Alternative, pH 7.5.

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Filter-sterilize and store at 4  C. Add 100 μg/mL BSA and 200 μM PMSF fresh the day of use. 2.3 Prep Antibody for IP

1. Antibody to target of interest (see Note 5). 2. Dynabeads Protein A and G. 3. 8-Strip, 0.2 mL PCR-Tubes and Caps. 4. PCR magnetic stand. 5. 10 μL 16-channel pipette. 6. 200 μL 8-channel pipette.

2.4 Immunoprecipitation

1. ChIP Elution Buffer: 50 mM Tris–HCl, 1 mM EDTA, 1% (v/v) SDS, pH 7.5. Filter-sterilize and store at 4  C. 2. ChIP Buffer 2: 25 mM Tris–HCl, 5 mM MgCl2, 300 mM KCl, 10% (v/v) glycerol, 0.1% (v/v) NP-40 Alternative, pH 7.5. Filter-sterilize and store at 4  C. Add 100 μg/mL BSA and 200 μM PMSF fresh the day of use. 3. ChIP Buffer 3: 10 mM Tris–HCl, 250 mM LiCl, 0.5% (v/v) NaDeoxycholate, 0.5% (v/v) NP-40 Alternative, pH 7.5. Filter-sterilize and store at RT. Add 100 μg/mL BSA fresh the day of use. 4. 1
Tris-EDTA (TE) Buffer: 10 mM Tris–HCl, 1 mM EDTA, pH 8.0. Filter-sterilize and store at RT. 5. 0.5 M EDTA, pH 8.0. 6. Proteinase K, Solution DNase and RNase Free. 7. DNA purification reagents (e.g. phenol:chloroform, spin columns, or SPRI paramagnetic beads). 8. Real-Time PCR (qPCR) forward and reverse primers to gauge experimental success (e.g., known positive + negative enriched genomic regions and/or spike-in barcoded nucleosome control primers, Fig. 1) (see Note 6). 9. qPCR Master Mix. 10. qPCR Instrument. 11. Library Preparation Kit for Next-Generation Sequencing if proceeding to NGS.

3

Methods

3.1 Nuclei Sample Preparation

The following protocol describes bulk native nuclei preparation that can be aliquoted and used in multiple IPs. Accurate final quantification and preparation of intact nuclei (Fig. 3) is essential to ensure that the expected number of cell-equivalents are used in each IP and the resulting data is high quality.

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1. Decontaminate the tissue culture hood and all materials that will go into the hood, including gloved hands. 2. Count cells using Trypan blue staining (1:1 ratio of Trypan Blue and cells) and a hemocytometer or automated cell counter by following manufacturer’s recommendations (ideal viability >95%). 3. Collect 30–100 million cells into a 50 mL conical tube. For protocol adaptations to X-ChIP workflows see Notes 7–9. 4. Pellet cells for 5 min at 500
g at 4  C. 5. Pour off supernatant taking care not to disturb the pellet. Flick pellet to loosen it from the tube wall and reduce clump formation. 6. Wash cells with 10 mL of 1
PBS per 30–100 million cells and spin down for 5 min at 500
g at 4  C. 7. Repeat PBS wash for a total of three times. 8. Wash cells with 5 mL of 1
Buffer N, mixing using a 22-gauge needle and 10 mL syringe. 9. Spin down cells for 5 min at 500
g at 4  C, for a total of two times. Note pellet volume (packed cell volume, PCV). 10. Resuspend the cells in at least 2
PCV of 1
Buffer N (e.g., if pellet volume is 100 μL, resuspend in 200 μL). 11. Add an identical volume (e.g., 200 μL) of 2
Modified Lysis Buffer containing the optimized concentration of detergent for the experimental cell type (see Note 2) and gently pipet to mix. Cells will lyse to release nuclei (Fig. 3). 12. Submerge cells in ice for the optimized lysis time, to be determined for each experimental cell type (2–10 min) (see Note 2). 13. Spin down nuclei for 5 min at 500
g at 4  C, pour or pipet off supernatant, and note packed nuclei volume (PNV). 14. Resuspend nuclei in at least 6
PNV of 1
Buffer N. 15. Add 7.5 mL of the Sucrose Cushion to a 15 mL tube and slowly layer the nuclei atop the cushion. Ensure that nuclei are added slowly and the layers do not mix; a cloudy film should be visible at the interface. If no cloudy top layer and clear interface is visible, spin down the nuclei as described in the subsequent steps (5 min at 500
g at 4  C) and start over at resuspension in 6
PNV of 1
Buffer N (step 14), taking care to slowly pipet nuclei atop the sucrose without disturbing the layer. 16. Spin down nuclei for 12 min at 500
g at 4  C. 17. Take off supernatant by removing the top cloudy layer first. Pellet should be bright white and not viscous (see Note 10). Take note of the PNV and then gently flick the pellet to loosen/homogenize it.

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18. Resuspend nuclei in at least 2
PNV of 1
Buffer N. 19. Measure the nuclei concentration by diluting 2 μL of nuclei into 18 μL of 2 M NaCl. Vortex vigorously for 10 min or sonicate for 10–20 min (see Note 11). 20. Measure the nuclei concentration in triplicate with a NanoDrop using the nucleic acid program (A260) and 2 M NaCl as a blank. 21. Calculate the average of the triplicate readings and multiply by the dilution factor (10), then convert to μg/μL. 22. Calculate the amount of 1
Buffer N to bring the nuclei to a final concentration of 1 μg/μL. To avoid over-diluting, add a small portion of the calculated amount of 1
Buffer N and check concentration on NanoDrop again. Repeat this process in small increments to slowly approach the desired final concentration (see Note 12). 23. To confirm nuclei integrity, take a small aliquot and dilute 1:50 in 1
Buffer N. Mix with Trypan Blue at a 1:1 ratio and visualize under brightfield or phase contrast microscope. Confirm that morphology is consistent with intact, unclumped nuclei (Fig. 3) and that cell counts are within the expected ranges (see Note 13). 24. Dispense 100 μg aliquots into 1.5 mL tubes and slow freeze (e.g., isopropanol-buffered container placed at 80  C). 25. Store nuclei at 80  C until ready for IP. 3.2 Prep Chromatin for IP

The procedure outlined below uses MNase to digest chromatin to mononucleosomes (150 bp, Fig. 4). Chromatin under native conditions cannot withstand sonication (as commonly used in X-ChIP after cell fixation). After MNase digestion, HAP chromatography is performed to improve resolution of genomic binding occupancy. Because DNA has a very high affinity for HAP, mononucleosomes loaded onto HAP and washed under stringent high salt conditions will be retained, while chromatin-bound proteins are stripped away [26]. While this process decreases overall chromatin yield (30% of crude chromatin is recovered), the resulting mononucleosomes are largely depleted of chromatin binding proteins which can interfere with antibody binding to the target of interest. This process therefore results in a more thorough understanding of relative target enrichment across the genome. 1. Remove 100 μg nuclei aliquot (prepped in Subheading 3.1) from 80  C and thaw on ice. 2. Add spike-in control (Fig. 1): 2 μL SNAP-ChIP Spike-in Controls per 10 μg sample chromatin (use only for histone PTMs: defined controls confirm antibody specificity and enable

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quantitative normalization) or 750 ng of Spike-in Drosophila Chromatin added to 30 μg of human chromatin (use for any target of interest: enables experimental normalization by downsampling sequencing reads, but no insight into antibody behavior). 3. Incubate thawed nuclei at 37  C on Thermomixer for 1 min. 4. Incubate MNase and nuclei at 37  C on the Thermomixer, shaking at 900 RPM to facilitate digestion. Ensure that MNase conditions are optimized to produce the desired fragment sizes (Fig. 4) (see Note 14). For X-ChIP modifications see Note 15. 5. Stop MNase digestion by adding an appropriate volume of 10
Stop Buffer (to 1
final). Dropwise pipet the 10
Stop Buffer into the reaction while vortexing at the slowest speed to ensure Stop Buffer is evenly dispersed. 6. To lyse the nuclei and release the chromatin, add 5 M NaCl (to 0.6 M final) dropwise into the reaction while vortexing at the slowest speed. 7. Pellet insoluble material by centrifugation for 1 min at 17,720
g at 4  C. 8. Use a pan balance to weigh 66 mg of the CHT Ceramic HAP resin and transfer to a low-bind microcentrifuge tube. 9. Rehydrate the resin with 200 μL of HAP Wash Buffer 1. Vortex quickly and keep on ice. 10. Carefully pipet the soluble supernatant to the HAP CHT resin. 11. Incubate chromatin and HAP CHT resin for 15 min at 4  C on the Compact Variable Speed/Angle Vertical Rotator set to 15 RPM (see Note 16). 12. Pipet the chromatin–resin mixture to an Ultrafree MC-HV Centrifugal Filter (0.45 μM). 13. Centrifuge the sample for 30 s at 600
g at 4  C and discard flow-through. 14. Slowly pipet 200 μL of HAP Wash Buffer 1 to the resin and spin down for 30 s at 600
g and 4  C. 15. Repeat this process three times for a total of four HAP Wash Buffer 1 washes, discarding flow through after each wash. 16. Slowly pipet 200 μL of HAP Wash Buffer 2 and spin down for 30 s at 600
g, 4  C. 17. Repeat this process three times for a total of four HAP Wash Buffer 2 washes, discarding flow through after each wash. 18. Discard collection tube and place filter into a new low-bind microcentrifuge tube.

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19. Slowly pipet 100 μL of HAP Elution Buffer to the resin and spin down for 30 s at 600
g at 4  C. Save the flow-through (contains purified nucleosomes to be used in IP). 20. Repeat this process two times for a total of three elutions, changing the collection tube for each spin. 21. Combine all three elution collections into a single low-bind microcentrifuge tube and briefly vortex to mix into a homogeneous solution. 22. Measure chromatin concentration in triplicate using the NanoDrop Microvolume UV-Vis Spectrophotometer (260 nm). Use the HAP Elution Buffer as a blank. 23. Calculate average of NanoDrop readings and multiply by 300 μL, this equals total ng of DNA. A620 AVG
300 μL ¼ Total ng DNA 24. Divide total ng of DNA by 20 μg/mL. Subtract 300 μL from this calculated volume to derive the final volume of ChIP Buffer 1 needed to adjust the chromatin concentration to 20 μg/mL.  μg   300 μL ¼ μL ChIP Buffer 1 needed Total ng DNA  20 mL 25. Add this calculated volume of ChIP Buffer 1 to the 300 μL chromatin to prepare a 20 μg/mL solution. Keep on ice and use fresh for IP; do not freeze. 3.3 Prep Antibody for IP

Antibodies need to be immobilized onto Protein A or Protein G (depending on host species/heavy chain isotype; see Note 17). Magnetic beads are recommended, as they enable multichannel pipetting, thus increasing experimental throughput and reproducibility (toward this end, note that the subsequent protocol steps are conducted using 8-strip tubes and multichannel pipettes). Agarose beads can also be used, although they require centrifugation during target capture and wash steps, thus reducing experimental throughput. It is essential to saturate the beads with antibody (calculate with reference to the lot-specific bead capacity), as nonspecific chromatin adsorption to the bead surface can increase background. Alternatively, BSA or exogenous DNA can be used as a blocking agent. 1. If antibody is stored at 20  C, thaw on ice (see Note 18). 2. Determine the optimized/recommended amount of antibody and maximum volume of Dynabeads (Protein A or G coupled) that will be saturated by this amount (see Notes 17 and 19). In general 100 μL of Dynabeads will isolate approximately

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25–30 μg human IgG (therefore, a ratio of 3–5 μg antibody to 12.5–20 μL Dynabeads is common), but note the specific capacity of the lot in hand. 3. Thoroughly resuspend Dynabeads and pipet to an 8-strip PCR tube. 4. Place tube strip in the magnetic stand, allow beads to collect on magnet, and discard supernatant. 5. Pipet 150 μL of ChIP Buffer 1 to beads and remove from magnet. Pipet-mix ten times, place tube back in magnet, and discard supernatant. 6. Repeat steps 4 and 5 for a total of two washes. 7. Resuspend beads in 100 μL of ChIP Buffer 1, pipet-mix ten times. 8. Pipet required amount of antibody (typically 3–5 μg) to the washed beads (see Note 19). 9. Incubate antibody and beads for 1 h at 4  C on the rotator set to 15 RPM. 10. Place tube(s) in the magnetic stand. Once beads have collected on the magnet, discard the supernatant. 11. Pipet 200 μL of ChIP Buffer 1 to antibody-bead complex. Remove from magnet and pipet-mix ten times. 12. Place tube(s) in the magnetic stand. Once beads have collected on the magnet, discard the supernatant. 13. Resuspend beads in 25 μL of ChIP Buffer 1. 3.4 Immunoprecipitation

Now that the chromatin and antibody are prepped, immediately proceed to the immunoprecipitation. Incubate the chromatin with the bead-bound antibody followed by stringent washes to remove off-target binding. Finally, isolate the DNA, perform quality control check of the final yield and fragment size distribution (Fig. 4), and proceed to qPCR and/or NGS (Fig. 1). 1. Calculate the desired amount of diluted chromatin from Subheading 3.2 (see Note 1) to load onto antibody–bead conjugates. 2. Reserve 1:10 of this calculated volume. Bring to 50 μL with ChIP Elution Buffer. This is the Input sample, which is used in qPCR and NGS to control for differences in loading (see Note 20). Place on ice for later processing. 3. Add the calculated amount of diluted chromatin to the prepared antibody–bead conjugates prepared in Subheading 3.3. 4. Incubate the chromatin with antibody–bead conjugates for 20 min at 4  C on the rotator set to 15 RPM.

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5. After incubation, quick spin tubes in benchtop microfuge to collect any liquid in the cap. Place tube(s) in the magnetic stand. Once beads have collected on the magnet, discard the supernatant. 6. Pipet 200 μL of ChIP Buffer 2 to the sample and remove from magnet. Pipet-mix ten times and transfer solution to a new prelabeled microcentrifuge tube. 7. Incubate for 10 min at 4  C on the rotator set to 15 RPM. Quick spin, place tube(s) in the magnetic stand, collect beads, and discard the supernatant as above. 8. Repeat steps 6 and 7 for a total of two washes. 9. Pipet 200 μL of ChIP Buffer 3 to the sample and remove from magnet. Pipet-mix ten times and transfer solution to a new prelabeled microcentrifuge tube. 10. Incubate for 10 min at 4  C on the rotator set to 15 RPM. Quick-spin, place tube(s) in the magnetic stand, collect beads, and discard the supernatant. 11. Pipet 200 μL of ChIP Buffer 1 to the sample, pipet-mix ten times, and transfer to a new microcentrifuge tube. Place tube back on magnet and discard supernatant. 12. Pipet 200 μL of 1
TE to the sample and pipet-mix ten times. Place tube back on magnet and discard supernatant. 13. Pipet 51.5 μL of room temperature ChIP Elution Buffer to the beads, pipet-mix ten times, and incubate at 55  C for 5 min in thermomixer, heat block, or thermocycler. 14. After incubation, place tubes on magnet and carefully transfer 50 μL of supernatant to a new prelabeled PCR tube. Discard the beads. 15. To all samples including Input from step 2, pipet 5 μL of 5 M NaCl (final 0.2 M) and 1 μL 0.5 M EDTA (final 10 mM) to each sample tube, and pipet-mix ten times. 16. Pipet 0.5 μL of Proteinase K (20 μg/mL, final 10 mM) to each tube and pipet-mix ten times. 17. Incubate samples for a minimum of 2 h (to overnight) at 55  C in a thermocycler, heat block, or thermomixer. 18. After incubation purify DNA (see Note 21). 19. Store samples at 20  C until further processing. 20. qPCR can be performed as a cost-control STOP/GO measure before investing in NGS (Fig. 1) for genome-wide analysis and normalization (see Notes 22 and 23).

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Notes 1. Standard amount of chromatin is 2–20 μg (1
105 to 1
106 cell equivalents) depending on target and antibody efficiency. For a high-abundance target (e.g., H3K27me3, 20% of total nucleosomes [27]) immunoprecipitated with a specific and efficient antibody (i.e., >10% of target recovered), 2 μg chromatin (1
105 cell equivalents) is often sufficient to generate high resolution data. Even for H3K4me3 (1.3 times including 35 genes encoding for secreted proteins. Among the top ten regulated genes with highest fold increases 7 were supposed to be secreted as proteins. In an immunobead based assay 12 of 23 analytes were significantly enriched in the EPS supernatant. Using a similar approach Raschke et al. analyzed conditioned medium of EPS and control cells and identified 48 contraction induced myokines [15]. These results indicate that the EPS model can be used to study the human muscle secretome by a global proteomics-based approach.

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Some critical considerations should be undertaken before applying EPS as an in vitro exercise model for the investigation of the exercised-induced human muscle secretome. In comparison to the murine skeletal muscle cell line C2C12, which responds with an easily observable and measurable contraction during EPS (after an initial phase of a few hours needed for the de novo sarcomere assembly [16]), the inducible contraction of primary human myotubes is less pronounced and visualization may be challenging [17]. Commercially available pulse generators (IonOptix C-Pace pulse generator, IonOptix, Dublin, Ireland) and self-designed systems may achieve different range of contractility. When establishing an EPS system for human myotubes, it is recommended to assess intracellular Ca2+ concentrations, sarcomeric structures, changes in glucose uptake and metabolism, and AMPK activation to validate the activation of exercise-related effects. We further recommend using electrodes without applying power as negative control for voltage-independent effects. One potential of using primary skeletal muscle cells for stimulation in an in vitro exercise model is the knowledge on the cell type specificity. To enable this, the purity of the cell culture (the percentage of myoblasts which can form myotubes during myodifferentiation) should be enriched and analyzed. Here, we provide a detailed protocol (see also flowchart Fig. 1) for the cultivation of human myoblasts, differentiation into myotubes before subsequent electric pulse stimulation, and sample preparation of the supernatant for targeted and untargeted proteomics-based analysis. For electric pulse stimulation, we refer to recent protocols [17, 18].

2

Materials

2.1 General Consumables

1. Pipetboy, pipette, and suitable tips. 2. 1.5 mL microcentrifuge tubes. 3. 150 cm2 cell culture dish. 4. Six-well cell culture dish (see Notes 1 and 2).

2.2

Cell Culture

1. Primary skeletal muscle cells are obtained from percutaneous needle biopsies performed on the lateral portion of quadriceps femoris (vastus lateralis) of human subjects. 2. Cloning medium for the propagation of myoblasts: 1:1 mixture of α-MEM and Ham’s F-12, 20% FBS, 2 mM glutamine, 1% chicken embryo extract, 100 U/mL penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 3. Fusion medium with 2%FBS: α-MEM, 2% FBS, 2 mM glutamine, 100 U/L penicillin, 100 μg/mL streptomycin, 0.5 μg/ mL amphotericin B.

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Satellite cell isolaon Power supply

Myoblasts

Incubator

Enrichment of CD56 posive myoblasts: e.g. MACS Mvo-differenaon for 5 days Electric pulse smulaon CD56+-Myoblasts

Myotubes

Cell culture

Fusion media

Fusion media - phenolred - FCS

Harvesng the secretome: Centrifugaon, concentraon

EPS / secretome collecon

Untargeted secretome profiling: Protein mass spectrometry approaches

Protein Profiling

Fig. 1 Workflow showing secretome analysis of human myotubes. Cell culture: Skeletal muscle cells are obtained from percutaneous needle biopsies performed on the lateral portion of quadriceps femoris (vastus lateralis) of human subjects. Satellite cells are isolated, CD56-positive myoblasts are enriched using, for example, magnetic cell sorting (MACS), cultivated and differentiated to multinucleated myotubes (immunostaining shows the fast-type skeletal muscle myosin in green; nuclei are shown in blue (DAPI staining)). Electric pulse stimulation (EPS) and secretome collection: After 5 days of differentiation myotubes are electrostimulated, for example, using the C-Pace EP system of IonOptix (detailed brochure: Culture Pacing System) and supernatants representing the secretome of working skeletal muscle cells are collected. Protein profiling: For quantitative comparison several targeted (e.g., Multiplex immunoassay) and untargeted (Mass spectrometry profiling) approaches are applicable. Especially, for MS-based analysis it is highly recommended to use media free of FBS and phenol red

4. Fusion medium without FBS: α-MEM, 2 mM glutamine, 100 U/L penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 5. Fusion medium without FBS and phenol red: α-MEM without phenol red, 2 mM glutamine, 100 U/L penicillin, 100 μg/mL streptomycin, 0.5 μg/mL amphotericin B. 6. Trypsin solution suitable for cell culture. 7. CO2 incubator (5% CO2), 37  C. 8. Phosphate buffered saline (PBS) with and without Mg2+ and Ca2+. 9. CD56 MACS microbeads and LS-columns (Miltenyi Biotech) (see Note 3). 2.3 Targeted Proteomics

1. Multiplex bead-based immunoassay (e.g., Bio-Plex Pro™ Assay; Milliplex® Multiplex Assay). 2. Suitable suspension array system and software for the measurement and data analysis.

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2.4 Untargeted Proteomics

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1. Falcon tubes (50 mL). 2. Protein concentrators (3 kDa cutoff). 3. Equipment for gel-based or gel-free protein separation technique and MS-based protein analysis [19].

3

Methods

3.1 Enrichment of CD56-Positive Myoblasts

1. Thaw, seed and proliferate human skeletal muscle cells in Cloning medium on 150 cm2 plates. Change medium every 2–3 days until cells are 50% confluent. Usually, cells from eight plates were used (approximately 10–20 million cells). 2. Trypsinate cells and purify CD56-positive cells by magnetic cell sorting (MACS) using CD56 MACS microbeads and LS-columns (see Note 3). Add 3 μL CD56 MACS microbeads per million cells. Take samples before isolation, from flowthrough and from eluate to assess content of CD56-positive cells by flow cytometry. Follow the manufacturer’s protocol, but increase bead incubation time to 30 min. 3. CD56-positive myoblasts are stored in the gaseous phase of liquid nitrogen or immediately used (see Note 4).

3.2 Fusion to Myotubes and Stimulation

1. Seed CD56-positive myoblasts at 1000 cells/cm2 on 6well dishes and proliferate them in Cloning medium. Change medium every 2–3 days until cells are 80–90% confluent. 2. When cells are 80–90% confluent change medium and differentiate cells to multinucleated myotubes with Fusion medium for 5 days (see Note 5). Change medium every 2–3 days. 3. Stimulate cells with electric pulses, for example, according to [17] or [18] (see Notes 6 and 7).

3.3 Collection of Medium for Targeted Secretome Studies

When using cell culture medium containing FBS, the high abundance of serum proteins in the FBS requires additional “enrichment” of low abundant proteins, for example, using ProteoMiner technology or metabolic pulse labeling [20, 21]. An efficient and sensitive way to analyze protein signatures in FBS-containing supernatant is via commercially available multiplex bead-based immunoassays (see Note 8). The following steps are necessary to collect conditioned medium and prepare it for measuring with multiplex bead-based immunoassays. 1. Directly after end of stimulation, take off medium and put on ice until proceeding to the next step (see Note 9). 2. To remove detached cells, spin supernatant with 2700  g for 4 min at 4  C.

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3. Transfer supernatant in a new tube and store at use for further analysis.

80  C until

4. When using a validated multiplex kit, proceed according to manufacturer’s instructions (see Notes 9–11). 5. Protein concentrations of the analytes can be calculated with the appropriate optimized standard curves using the appropriate software. 3.4 Collection of Medium for Untargeted Proteomics

An untargeted proteomic profiling approach enables the discovery of novel contraction-induced myokines and helps to identify specific and sensitive biomarkers secreted from the muscle [22]. For analysis diverse mass spectrometry–based approaches can be accessed allowing highly sensitive and quantitative investigation [23]. Nevertheless, due to the well-known challenges coming along with global secretome profiling, it is recommended to implement an appropriate sample quality control. Therefore, two-dimensional gel electrophoresis is still a powerful tool, allowing to visualize sample complexity, composition, and quality (see Note 12). This makes it easy to detect for example FBS contamination, which masks the real secretome prohibiting a meaningful analysis. For quantitative comparison analysis, also applied protein quantities have to be specifically standardized for each project. The following part describes a basic sample preparation procedure appropriate for most post analysis procedures. 1. For untargeted profiling Fusion medium without FBS has to be used during stimulation to prevent the contamination of proteins included in the serum. To decrease any serum carryovers wash cells 2 with PBS containing Ca2+/Mg2+ and 1 with standard PBS (see Note 13). Use also Fusion medium without phenol red (see Note 14). 2. Directly after stimulation end, take off supernatant and spin down detached cells with 2700  g for 5 min at 4  C to remove any cell debris. 3. Remaining insoluble material is separated by a further centrifugation step (45,000  g, 5 min, 4  C). Optionally, supernatants can be centrifuged at 100,000  g for 30 min in order to discriminate between the soluble part of the secretome (second supernatant) and microvesicles (pellet), which represent a significant part of the total secretome [24]. 4. Samples can be frozen at point.

80  C or further processed at this

5. Either continue directly with the unfrozen supernatant or thaw the supernatant on ice. Since the native protein concentration is in the lower μg/mL range, the supernatant can be concentrated by spin concentration devices (cutoff 3 kDa) (see Note 15).

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6. Concentrated supernatants are stored at 80  C until further processing for gel-based or gel-free secretome profiling. Use aliquots to assess protein concentrations, which is crucial for appropriate protein normalization.

4

Notes 1. Be sure to use six-well dishes that fit with the electrodes of the electric pulse stimulation system. The IonOptix handbook proposes six-well dishes of several companies. 2. Cell culture plates and dishes can be coated with Geltrex™ (Thermo, A1413302) according to the manufacturer’s thingel protocol for proliferation of stem cells: dilute Geltrex 1:300 in a 1:1 mixture of α-MEM and Ham’s F-12. Coating with Geltrex seems to improve proliferation of CD56-positive cells without improving proliferation of CD56-negative cells and thus leads to higher amounts and concentrations of CD56positive cells. The potential interference of Geltrex with EPS and proteomics analysis needs to be tested. 3. Human skeletal muscle-derived cells with the capacity to form multinucleated myotubes are characterized by the expression of the cell adhesion molecule CD56 (NCAM) and can be separated from CD56-negative fibroblasts [25]. 4. The content of approximately 95%.

CD56-positive

cells

should

be

5. We recommend Fusion medium without FBS. Removal of FBS during differentiation reduces the number of cells not fused to myotubes [26]. The addition of free fatty acids palmitate (50 μM) and oleate (50 μM) complexed to BSA and free carnitine (100 μM) to the Fusion medium better mimic physiological nutrient supply, but also adds 1.67% BSA and 0.05% ethanol. 6. For the proteomics-based investigation of secreted proteins it is necessary to minimize the contamination with proteins released from damaged cells. The absence of EPS-induced cell damage should be verified by the quantification of cytosolic or mitochondrial localized proteins in the supernatant, for example, by creatine kinase (CK) activity, also used as plasma marker for muscle fiber damage. Lactate dehydrogenase (LDH) is another marker for muscle fiber damage. However, after 24 h of EP stimulation, LDH is significantly increased in both, the total cell lysates of EPS cells and also in the supernatant [13]. These data suggest that elevated levels of cytosolic proteins in the secretome do not necessarily indicate higher percentage of damaged cells but can be due to increased production of this protein.

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7. For the selection of candidate myokines it might be helpful to do a whole genome transcriptome analysis or quantitative Realtime PCR prior to secretome study to find EPS-regulated proteins. 8. Diverse antibody-based approaches can be used for specific and sensitive quantification. In this context multiplex immunoassays (MIA) have several advantages. Compared to commercially available enzyme-linked immunosorbent assay (ELISA) MIA enable the measurement of several proteins simultaneously from very small sample amounts in one assay. It combines easy handling with high sensitivity, allowing for exact quantification in a concerted system. 9. During immunoassay antibodies might cross-react with proteins of FBS, hence a sample of unconditioned medium (reference medium) is urgently needed for all immunoassay-based measurements. Spin down the reference medium according to Subheading 3.3, step 2 and store supernatant at 80  C until used for further analysis. 10. When mixing individual singleplexes check x-Plex Bead Regions and avoid using 2 singleplexes with similar x-Plex Bead Regions in one assay. 11. For the conditioned medium of human myotubes, neat samples were used (50 μL sample/well). Make sure that intensities of measured analytes are in the assay working range. If values are above the upper limit of quantification dilute the samples properly. 12. Although recent mass spectrometry profiling approaches require only ng amounts of protein, for project planning it has to be considered, that comprehensive analysis including quality assessment, for example utilizing two-dimensional gel electrophoresis, requires much more material (at least several micrograms). 13. To get rid of the complex protein mixture included in FBS, cells are washed two times with PBS containing Ca2+/Mg2+. This is used to wash adherent cell cultures when one merely wishes to wash and have the cells remain adherent because adhesion proteins require divalent cations to function. The third washing step is done with PBS without Ca2+/Mg2+ to decrease salt concentration in the medium. 14. For untargeted proteomics the medium should be free of phenol red because it might interfere with protein assays performed to determine protein concentration for subsequent analysis. 15. Since the native protein concentration is very low it is essential to concentrate the supernatant to achieve a working concentration for subsequent analysis. It might be necessary to

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transfer the supernatant during concentration process in an even smaller protein concentrator tube to gain smaller volumes and the required protein concentration. Be aware that with every change of the device material gets lost. To minimize the loss, rinse the membranes of the larger device with flow through and add it to the eluent of the smaller device. Additionally, take care of protein precipitations that might form.

Acknowledgments This work was supported, in part, by a grant from the German Federal Ministry of Education and Research (BMBF) to the German Center for Diabetes research (DZD e.V.; No. 01GI0925). References 1. Pedersen BK (2009) The diseasome of physical inactivity--and the role of myokines in muscle-fat cross talk. J Physiol 587(Pt 23):5559–5568. https://doi.org/10.1113/jphysiol.2009. 179515 2. Gleeson M (2007) Immune function in sport and exercise. J Appl Physiol 103(2):693–699. https://doi.org/10.1152/japplphysiol. 00008.2007 3. Goldstein MS (1961) Humoral nature of the hypoglycemic factor of muscular work. Diabetes 10:232–234 4. Ostrowski K, Rohde T, Zacho M, Asp S, Pedersen BK (1998) Evidence that interleukin-6 is produced in human skeletal muscle during prolonged running. J Physiol 508(Pt 3):949–953 5. Febbraio MA, Hiscock N, Sacchetti M, Fischer CP, Pedersen BK (2004) Interleukin-6 is a novel factor mediating glucose homeostasis during skeletal muscle contraction. Diabetes 53(7):1643–1648 6. Pedersen BK, Febbraio MA (2012) Muscles, exercise and obesity: skeletal muscle as a secretory organ. Nat Rev Endocrinol 8 (8):457–465. https://doi.org/10.1038/ nrendo.2012.49 7. Hoffmann C, Weigert C (2017) Skeletal muscle as an endocrine organ: the role of Myokines in exercise adaptations. Cold Spring Harb Perspect Med 7(11):a029793. https://doi.org/ 10.1101/cshperspect.a029793 8. Weigert C, Lehmann R, Hartwig S, Lehr S (2014) The secretome of the working human skeletal muscle-a promising opportunity to combat the metabolic disaster? Proteomics Clin Appl 8(1-2):5–18. https://doi.org/10. 1002/prca.201300094

9. Nikolic N, Gorgens SW, Thoresen GH, Aas V, Eckel J, Eckardt K (2017) Electrical pulse stimulation of cultured skeletal muscle cells as a model for in vitro exercise—possibilities and limitations. Acta Physiol (Oxf) 220 (3):310–331. https://doi.org/10.1111/apha. 12830 10. Fujita H, Nedachi T, Kanzaki M (2007) Accelerated de novo sarcomere assembly by electric pulse stimulation in C2C12 myotubes. Exp Cell Res 313(9):1853–1865. https://doi. org/10.1016/j.yexcr.2007.03.002 11. Lambernd S, Taube A, Schober A, Platzbecker B, Gorgens SW, Schlich R, Jeruschke K, Weiss J, Eckardt K, Eckel J (2012) Contractile activity of human skeletal muscle cells prevents insulin resistance by inhibiting pro-inflammatory signalling pathways. Diabetologia 55(4):1128–1139. https://doi. org/10.1007/s00125-012-2454-z 12. Manabe Y, Miyatake S, Takagi M, Nakamura M, Okeda A, Nakano T, Hirshman MF, Goodyear LJ, Fujii NL (2012) Characterization of an acute muscle contraction model using cultured C2C12 myotubes. PLoS One 7 (12):e52592. https://doi.org/10.1371/jour nal.pone.0052592 13. Scheler M, Irmler M, Lehr S, Hartwig S, Staiger H, Al-Hasani H, Beckers J, de Angelis MH, Haring HU, Weigert C (2013) Cytokine response of primary human myotubes in an in vitro exercise model. Am J Physiol Cell Physiol 305(8):C877–C886. https://doi.org/ 10.1152/ajpcell.00043.2013 14. Nikolic N, Skaret Bakke S, Tranheim Kase E, Rudberg I, Flo Halle I, Rustan AC, Thoresen GH, Aas V (2012) Electrical pulse stimulation

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of cultured human skeletal muscle cells as an in vitro model of exercise. PLoS One 7(3): e33203. https://doi.org/10.1371/journal. pone.0033203 15. Raschke S, Eckardt K, Bjorklund Holven K, Jensen J, Eckel J (2013) Identification and validation of novel contraction-regulated myokines released from primary human skeletal muscle cells. PLoS One 8(4):e62008. https:// doi.org/10.1371/journal.pone.0062008 16. Fujita H, Nedachi T, Kanzaki M (2007) Accelerated de novo sarcomere assembly by electric pulse stimulation in C2C12 myotubes. Exp Cell Res 313(9):1853–1865 17. Nikolic N, Aas V (2019) Electrical pulse stimulation of primary human skeletal muscle cells. Methods Mol Biol 1889:17–24. https://doi. org/10.1007/978-1-4939-8897-6_2 18. Scheler M, de Angelis MH, Al Hasani H, Haring HU, Weigert C, Lehr S (2015) Methods for proteomics-based analysis of the human muscle secretome using an in vitro exercise model. Methods Mol Biol 1295:55–64 19. Wohlbrand L, Trautwein K, Rabus R (2013) Proteomic tools for environmental microbiology--a roadmap from sample preparation to protein identification and quantification. Proteomics 13(18-19):2700–2730. https://doi. org/10.1002/pmic.201300175 20. Eichelbaum K, Winter M, Berriel Diaz M, Herzig S, Krijgsveld J (2012) Selective enrichment of newly synthesized proteins for quantitative secretome analysis. Nat Biotechnol 30 (10):984–990. https://doi.org/10.1038/ nbt.2356 21. Hartwig S, Czibere A, Kotzka J, Passlack W, Haas R, Eckel J, Lehr S (2009) Combinatorial hexapeptide ligand libraries (ProteoMiner): an innovative fractionation tool for differential

quantitative clinical proteomics. Arch Physiol Biochem 115(3):155–160. https://doi.org/ 10.1080/13813450903154224 22. Hartwig S, Raschke S, Knebel B, Scheler M, Irmler M, Passlack W, Muller S, Hanisch FG, Franz T, Li X, Dicken HD, Eckardt K, Beckers J, de Angelis MH, Weigert C, Haring HU, Al Hasani H, Ouwens DM, Eckel J, Kotzka J, Lehr S (2014) Secretome profiling of primary human skeletal muscle cells. Biochim Biophys Acta 1844(5):1011–1017 23. Li X, Wang W, Chen J (2017) Recent progress in mass spectrometry proteomics for biomedical research. Sci China Life Sci 60 (10):1093–1113. https://doi.org/10.1007/ s11427-017-9175-2 24. Hartwig S, De Filippo E, Goddeke S, Knebel B, Kotzka J, Al-Hasani H, Roden M, Lehr S, Sell H (2019) Exosomal proteins constitute an essential part of the human adipose tissue secretome. Biochim Biophys Acta Proteins Proteom 1867(12):140172. https://doi. org/10.1016/j.bbapap.2018.11.009 25. Stewart JD, Masi TL, Cumming AE, Molnar GM, Wentworth BM, Sampath K, McPherson JM, Yaeger PC (2003) Characterization of proliferating human skeletal muscle-derived cells in vitro: differential modulation of myoblast markers by TGF-beta2. J Cell Physiol 196 (1):70–78 26. Hoffmann C, Hockele S, Kappler L, Hrabe de Angelis M, Haring HU, Weigert C (2018) The effect of differentiation and TGFbeta on mitochondrial respiration and mitochondrial enzyme abundance in cultured primary human skeletal muscle cells. Sci Rep 8(1):737. https://doi.org/10.1038/s41598-01718658-3

Chapter 28 Western Blotting Using In-Gel Protein Labeling as a Normalization Control: Advantages of Stain-Free Technology Roˆmulo Lea˜o Silva Neris, Andrea Marie Chua Dobles, and Aldrin V. Gomes Abstract Western blotting is one of the most used techniques in research laboratories. It is popular because it is an easy way of semiquantifying protein amounts in different samples. In Western blotting, the most commonly used method for controlling the differences in the amount of protein loaded is to independently quantify housekeeping proteins (typically actin, GAPDH or tubulin). Another less commonly used method is total protein normalization using stains, such as Ponceau S or Coomassie Brilliant Blue, which stains all the proteins on the blots. A less commonly used but powerful total protein staining technique is stain-free normalization. The stain-free technology is able to detect total protein in a large linear dynamic range and has the advantage of allowing protein detection on the gel before transblotting. This chapter discusses the theory, advantages, and method used to do total protein quantification using stain-free gels for normalization of Western blots. Key words Immunoblotting, Loading control, Stain-free technology, Total protein normalization, Western blotting

1

Introduction Western blotting is widely used in life sciences to access specific protein levels in a sample. It relies on protein separation by size in an acrylamide matrix in the presence of an electric current, followed by the transfer of proteins from the gel to a membrane, the blocking of sites on the membrane that were not exposed to proteins transferred from the gel, the incubation of the membranes with antibodies with high affinity for a target protein, and the detection of these primary antibodies by different methods [1]. Given Western blotting relevance to describe mechanisms for metabolism, disease states, and several other cellular processes, it is important to have both robust and reproducible protocols. Since Western blotting techniques are very sensitive to variations in protein amounts loaded on gels, which can occur from pipetting errors

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_28, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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and transfer steps, it is important to ensure the proper normalization of the amount of the target protein through a loading control. The most common loading controls are the housekeeping proteins [2, 3]. These housekeeping proteins are highly abundant proteins that are constitutively expressed by cells in many different states and conditions. Actin, glyceraldehyde 3-phosphate dehydrogenase (GAPDH), and tubulin are some of the most frequent housekeeping proteins used in the literature. However, the use of housekeeping proteins as loading controls has some limitations, and in several cases is not a reliable method to normalize Western blotting data. It has been shown that in many cell states, like proliferation, differentiation, and disease, significant changes in the amount of these housekeeping proteins occur [4–8]. In some reports, the saturation of housekeeping proteins at relatively low total protein amounts leads to misleading results for the target protein amounts, especially when the target proteins are present in low amounts [6, 7]. It has been suggested by several reports that the use of total protein amounts is more appropriate to normalize Western blotting data, leading to less variation, more reproducibility, and decreased potential errors associated with different cellular states. The most used techniques to determine total protein amounts are based on using dyes to stain all proteins on the blot membrane. This can be accessed before the membrane blocking, with Ponceau S [9], of after imaging it, like Coomassie blue and Amido-black staining [10]. These total protein staining methods have their own limitations as shown in the table below (Table 1). Stain-free imaging can be a powerful tool to overcome some of the staining protocol limitations. The concept of stain-free imaging of gels and membranes is based on the ability of aromatic amino acids like tryptophan to undergo changes under UV light stimulation in the presence of trihalo compounds. Tryptophan present on proteins covalently bind to the trihalo compounds in the gel, resulting in these labeled proteins emitting detectible fluorescence, and enabling the detection of amounts as low as 20 ng of protein [11]. This method is very reliable but requires a gel that was polymerized together with trihalo compound. In this chapter, we demonstrate the effectiveness of the stain-free method to accurately quantify the total amount of protein in mouse heart homogenates and to normalize the amount of the proteasome subunit β1 by Western blotting. Hence, the stain-free technology appears to be a robust technique to ensure proper loading controls for protein analysis.

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Table 1 Advantages and disadvantages of different total protein staining methods Detection Method

Pros

Ponceau S

l l l

Coomassie blue

l

l

l

Amido black

l

l

l

Stain-free

l

l l

l l

l

2

Cons

Easily reversible Water soluble Staining solutions can be reused many times

l

Good sensitivity (~50 ng minimum) Suitable both for gels and membranes Staining solutions can be reused many times

l

Good sensitivity (~50 ng minimum) Suitable both for gels and membranes Staining solutions can be reused many times

l

Greater sensitivity (~20 ng minimum) Does not require solutions No additional steps of washing/shaking No incubation Suitable both for gels and membranes No reagent waste generated

l

l l l

l l

l

l l

l

l

References

Additional steps of washing and shaking Low sensitivity (~200 ng minimum) Recommended only for membranes Increased background

[9, 10, 12, 13]

Toxic/harmful waste Additional steps of washing/shaking Requires the incubation of weak acid and organic alcohols Irreversible staining

[10, 12–14]

Toxic/harmful waste Additional steps of washing/shaking Requires the incubation of weak acid and organic alcohols on the membrane Irreversible staining

[10, 12, 13]

Requires a gel prepared with trihalo compound Antibodies that bind to tryptophan residues in the target protein antibody epitope may have its efficiency affected

[11, 15, 16]

Materials Prepare all solutions with ultrapure water (18 MΩ) and reagents that are analytical grade. Prepare and store all reagents at room temperature unless otherwise indicated. Do not add sodium azide to any of the solutions and follow all local waste disposal protocols for disposal of Western blotting waste.

2.1 Tissue Sample Preparation

1. 100 mg mouse heart. 2. Homogenization buffer: 150 mM NaCl, 5 mM MgCl2, 1 mM EDTA, and 50 mM Tris–HCl, pH 7.5. To prepare 250 mL homogenization buffer, add together 7.5 mL 5 M NaCl, 1.25 mL 1 M MgCl2, 0.5 mL 0.5 M EDTA, and 6.25 mL 2 M Tris base. Dissolve all reagents in about 200 mL of water. After adjusting the pH to 7.5 with HCl, top up the solution to 250 mL with water (see Note 1). 3. Spectrophotometer for small samples: NanoDrop 2000c.

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2.2 SDS Polyacrylamide Gel Electrophoresis (SDSPAGE)

1. Laemmli sample buffer (4
): 131.6 mM Tris–HCl, pH 6.8, 4.2% SDS, 52.6% (w/v) glycerol, and 0.02% (w/v) bromophenol blue. Add β-mercaptoethanol to the sample buffer when ready to use (100 μL for each 900 μL of 4
Laemmli sample buffer) (see Note 2). 2. SDS-PAGE gels with stain-free capabilities (e.g., 4–20% 26-well Criterion TGX Stain-Free Precast Gel). 3. Protein standards (e.g., Precision Plus Protein Dual Color Standards) (see Note 3). 4. 10
SDS-PAGE running buffer: 192 mM glycine, 25 mM Tris base, and 0.1% SDS. To prepare 1 L of 10
SDS-PAGE running buffer, add 144 g glycine, 30.2 g Tris base, and 50 mL 20% SDS solution. Mix the reagents in around 900 mL of water until dissolved, and then fill to 1 L with water (see Note 4).

2.3

Protein Transfer

1. Precut filter paper (e.g., Trans-Blot Turbo Midi-size Transfer Stacks). 2. Blotting membrane (e.g., Trans-Blot Turbo Nitrocellulose membrane) (see Note 5). 3. Transfer buffer (e.g., Trans-Blot Turbo 5
Transfer Buffer [Bio-Rad]). Prepare 1 L of 1
transfer buffer by mixing 200 mL 5
transfer buffer with 600 mL water and 200 mL 200 proof pure ethanol. 4. Blotting apparatus: Trans-Blot Turbo Transfer System. 5. Tris-buffered saline (TBS): 10 mM Tris–HCl, 150 mM NaCl, pH 7.4 (see Note 6). To prepare 1 L of 1
TBS, add 2.4 g Tris base and 8.8 g NaCl to around 800 mL water. Mix thoroughly to dissolve and adjust the pH to 7.4 with HCl. Fill to 1 L with water. 6. TBST: TBS with 0.05% (v/v) Tween 20. Add 0.5 mL Tween 20 to 1 L TBS to make TBST (see Note 7). 7. Blocking solution: 3% nonfat milk in TBST. To make 50 mL of blocking solution, weigh out 1.5 g of powdered milk in a 50 mL conical centrifuge tube, and fill to 50 mL with TBST. Vortex the solution until completely mixed (see Note 8). 8. Antibody diluent solution: 1% nonfat milk in TBST. To make 15 mL of antibody diluent solution, dissolve 0.15 g of powdered milk in 15 mL TBST in a conical centrifuge tube (see Note 8). 9. Substrate for enhanced chemiluminescence (ECL): Clarity Western ECL Substrate, containing Clarity Western peroxidase reagent and Clarity Western luminol/enhancer that should be mixed at a ratio of 1:1 when ready to use (see Note 9). 10. Blotting box (see Note 10).

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11. Forceps for gentle handling of membranes. 12. Roller. 13. Delicate wiper paper (see Note 11). 14. Imaging equipment with stain-free technology capabilities: ChemiDoc MP. 15. Computer with Image Lab software, version 6.01 (see Note 12). 2.4 Antibodies for Immunostaining

1. Primary antibody solution: monoclonal mouse 20S Proteasome β1 (D-9) antibody (# sc-374,405, Santa Cruz Biotechnology), used at a concentration of 1:2000 in 1% (v/v) milk/ TBST. 2. Secondary antibody solution: peroxidase-conjugated rabbit anti-mouse IgG antibody (# A9044, Sigma-Aldrich), used at a concentration of 1:10000 (v/v) in 1% milk/TBST.

3

Methods Perform the following procedures at room temperature, unless otherwise noted.

3.1 Sample Preparation

1. Thaw the mouse heart on ice and chop approximately 50 mg with a razor blade. Use 25 strokes to homogenize the chopped heart in 0.5 mL cold homogenization buffer on ice using a Dounce homogenizer (see Note 13). 2. Transfer the homogenate into 1.5 mL centrifuge tubes and centrifuge at 12,000
g for 15 min to separate the debris, mitochondria and nucleus. Transfer the supernatant to a new tube and measure the concentration using a NanoDrop Spectrophotometer. Dilute the homogenate to 16 mg/mL with homogenization buffer. Store lysate at 80  C if not using immediately. 3. Prepare the samples for SDS-PAGE. Combine equal parts 16 mg/mL homogenate with 2
Laemmli sample buffer (containing β-mercaptoethanol) to create 8 mg/mL heart homogenate. 2
Laemmli sample buffer is made by diluting 4
Laemmli sample buffer with water in a 1:1 ratio. Vortex the mixture, briefly spin down, and heat at 96  C for 5 min. Cool to room temperature.

3.2

SDS-Page

1. Dilute the protein standard to 1/10 volume with 1
Laemmli sample buffer (see Note 3). 2. After removal of the comb and tape from the precut stain-free gel, prepare the electrophoresis tank by placing the gel inside the tank filled with SDS-PAGE running buffer. Load 1 μL

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Table 2 Gel loading pattern. Various volumes of the mouse heart homogenate sample (8 μg/μL) were loaded in order to result in 10–80 μg of the sample into the wells. Each amount was loaded into three different wells. The first set is from lanes 3–10, the second set is from lanes 11–18, and the third set is from lanes 19–26

Lane

1

2

3 11 19

4 12 20

5 13 21

6 14 22

7 15 23

8 16 24

9 17 25

10 18 26

Sample amount (μg)

N/Aa

N/Aa

10

20

30

40

50

60

70

80

Volume (μL)

1

2

1.25

2.5

3.75

5

6.25

7.5

8.75

10

N/A: Not applicable. Lanes 1 and 2 were not loaded with the mouse heart homogenate. Lane 1 was loaded with 1 μL of the 1/10 diluted protein standard. Lane 2 was loaded with 2 μL 1
sample buffer

a

(tenfold diluted) of the diluted protein standard in lane 1. Add 2 μL of 1
Laemmli sample buffer in lane 3. In the subsequent lanes, add increasing amounts of the heart homogenate (10–80 μg), and repeat this three times (Table 2) (see Note 14). 3. Run the electrophoresis at 120 V constant voltage until the dye front reaches the agarose layer of the gel (see Note 15). Remove the gel from the cassette and cut away the wells and the agarose with a razor blade. Make a small cut at the top left side of the gel for orientation. 4. Lay the gel in the Bio-Rad ChemiDoc MP Imaging system. Image the gel using the stain-free gel setting. For the first image, be sure to check the setting to activate the gel for 1 min. This activation setting exposes the gel to UV light. Under UV light, the tryptophan resides in the proteins will react with the trihalo compounds in the gel and become fluorescent, enabling viewing of the total protein content in the gel (see Note 16). Manually adjust the exposure time if needed to avoid overexposed bands (Fig. 1a). 3.3

Protein Transfer

1. Soak one stack of filter paper in transfer buffer and place it in the Trans-Blot Turbo Transfer System cassette. Wet the nitrocellulose membrane with transfer buffer and lay on top of the filter paper stack. Allow at least 2 min for the filter paper and membrane to soak in the transfer buffer. Place the gel on top of the membrane, followed by another stack of filter paper soaked in transfer buffer. Firmly roll out the air bubbles with a roller. Use your fingers to apply mild pressure on the whole transfer stack while closing the lid of the cassette to ensure no additional air bubbles form. Wipe the outside of the cassette with tissue paper and place it inside the Trans-Blot Turbo Transfer System machine. Run with the Turbo midi setting of 7 min to transfer the proteins from the gel to the membrane.

Stain-Free Technology

A kDa

Heart homogenate (μg) 10 20 30 40 50 60 70 80

449

B

5

Relative Intensity

250 150

75

37 25

4 3 2 1 0 0

20

40

60

80

Total Protein Amount (µg)

10

Stain-Free Gel

Fig. 1 Detection of total mouse heart protein on stain-free gel. (a) Detection of 10–80 μg of mouse heart homogenate using stain-free technology. The Criterion stain-free gel was exposed to 1 min of UV transillumination using the ChemiDoc MP to activate it before imaging. (b) Quantification of the relative intensity of total protein detected on the gel versus the total protein amount loaded onto the gel (μg) (n ¼ 3 for each data point). Data are represented as mean  standard deviation

2. Take the membrane from the cassette and image it while still wet in the ChemiDoc MP Imaging system using the stain-free membrane setting and optimized exposure time. There is no need to activate the membrane again. Adjust the exposure time as needed to avoid overexposed images (Fig. 2a). If necessary, the membrane can be cut into smaller sizes at this stage (see Notes 17 and 18). 3. Dry the membrane on top of a piece of tissue paper. Allowing the membrane to dry before proceeding to immunostaining has been suggested to help the proteins adhere better to the membrane (see Note 19). 3.4

Immunostaining

1. Place the membrane in a blotting container and add enough blocking solution (3% milk/TBST) to cover the entire membrane surface. Pour solutions into the corner of the blotting container and not directly onto the membrane so that the proteins on the membranes are not disturbed. Incubate the membrane in a shaker for 1 h. Pour out the blocking solution and wash three times with TBST for 3 min each in a shaker. 2. Add the primary mouse anti-20S proteasome β1 antibody solution and incubate at 4  C overnight in a shaker, followed by incubation with shaking for 1 h at room temperature (see

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A kDa

Heart homogenate (μg)

B

10 20 30 40 50 60 70 80

7

Relative Intensity

250 150

75

37

6 5 4 3 2 1 0

25

0

20

40

60

80

Total Protein Amount (µg) 10

Stain-Free Membrane

Fig. 2 Quantification of total mouse heart protein detected on nitrocellulose membrane using stain-free technology. (a) Detection of 10–80 μg of mouse heart homogenate on nitrocellulose membrane using stainfree technology. (b) Quantification of the relative intensity of total protein detected on the membrane versus the total protein amount loaded onto the gel (μg) (n ¼ 3 for each data point). Data are represented as mean  standard deviation

Note 20). Pour out the primary antibody solution and wash three times with TBST for 3 min each on a shaker. 3. Add the anti-mouse secondary antibody solution and incubate with shaking for 1 h. (see Note 21) Pour out the secondary antibody solution and wash three times with TBST for 3 min each on a shaker. 4. In a 1.5 mL centrifuge tube, mix together equal parts of Clarity western peroxide agent and Clarity western luminol/enhancer reagent. This creates the substrate for enhanced chemiluminescence (ECL). Pipette enough ECL substrate to evenly cover the surface of the membrane where the proteins are attached (see Note 22). Incubate the ECL covered membrane in a dark condition for 2 min before removing the excess ECL reagent (see Note 23). 5. Pick up the membrane from one side with a pair of forceps and drain away the excess ECL by gently touching the edge of the membrane to a delicate wiper paper. Arrange the membrane on the imaging surface, taking care not to create bubbles on the surface. 6. Image the blot in the Bio-Rad ChemiDoc MP Imaging system with the Chemi Hi Sensitivity setting and optimized exposure time. For a higher-resolution image, the blot can be imaged

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451

with the Chemi Hi Resolution setting with the optimized exposure time setting or with the exposure time set at approximately two times the time used for the Chemi Hi Sensitivity setting. The exposure time can be adjusted as needed to obtain the best image that contain no overexposed bands (Fig. 3a). Take a multichannel image with the stain-free and either the Chemi Hi Sensitivity or Hi Resolution setting to obtain an image of the chemiluminescent protein bands of interest overlaid with the stain-free prestained protein markers. Although the other proteins on the membrane will also show up with the stain-free image, the chemiluminescent signal shows up on a different channel. Verify the molecular weight of the band of interest based on the multichannel image. 7. Quantify the total proteins from the gel and membrane imaged with the image analysis software (Figs. 1b and 2b). Quantify the immunostained protein band and normalize the intensity of the band against the stain-free total protein (Fig. 3b, c). The data should be normalized against the stain-free total protein from the membrane and not the gel (see Notes 24 and 25).

4

Notes 1. Unused homogenization buffer can be stored at 4  C for up to 3 months. 2. Sample buffer lacking β-mercaptoethanol can be made and stored at RT after aliquoting. After β-mercaptoethanol is added the sample buffer is ready to use and unused sample buffer with β-mercaptoethanol can be stored at 20  C for at least 3 months or at 80  C for at least 6 months. 3. Many prestained protein standards are available and can be used. However, some protein standards are better for stainfree gels as some standard proteins with high tryptophan content could result in very strong signals which affect the quantification of the lanes nearby. Standards can be diluted if the signals are too strong. 4. The SDS-PAGE running buffer should not be adjusted with acid or base. The pH of the final 1
buffer should be between 8.3 and 8.9. We prefer to use 20% SDS stock solution instead of powdered SDS to reduce the potential for SDS particles getting in suspension into the lab environment. 5. Nitrocellulose membranes have a slightly lower background as compared to polyvinylidene difluoride (PVDF) membranes, though PVDF membranes can also be used together with stain-free gels. In particular, low fluorescence PVDF membranes are recommended when the secondary antibodies are fluorescently labeled.

B

0

1

2

3

4

0

40

60

80

Total Protein Amount (µg)

20

Not normalized Normalized to total protein

25

C

0

1

2

3

0

20

30

40

Total Protein Amount (µg)

10

Not normalized Normalized to total protein

10 20 30 40 50 60 70 80 μg protein

Fig. 3 Quantification and normalization of 20S proteasome β1 levels in mouse heart homogenates. (a) Western blot of mouse heart homogenate (10–80 μg) using an antibody against the β1 subunit of the 20S proteasome. (b) Quantification of 20S proteasome β1 levels (10–80 μg), not normalized and normalized to total protein detected using stain-free technology on the nitrocellulose membrane (n ¼ 3 for each data point). Data are represented as mean  standard deviation. (c) Quantification of 20S proteasome β1 levels (10–40 μg), not normalized and normalized to total protein detected using stain-free technology on the nitrocellulose membrane (n ¼ 3 for each data point). Data are represented as mean  standard deviation

Relative Intensity

A

Relative Intensity

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453

6. 10
TBS can be prepared and diluted to 1
TBS when needed. Unused 10
TBS can be kept up to 6 months at room temperature, as storing it at 4  C will cause it to crystallize. 7. To add extremely viscous solutions like Tween 20 to other solutions, cut off the tip of the pipette with clean scissors to create a larger opening and pipette slowly. 8. Blocking and diluting antibodies can be done with either nonfat milk or BSA in TBST. For better results, dilute antibodies in 1% BSA/TBST or 1% milk/TBST, as these will usually give a better signal than 3–5% BSA or milk. Depending on the antibody, milk or BSA may give a stronger signal with lower background noise. In general, BSA works better with biotinand alkaline phosphatase-labeled antibodies, as well as antiphosphoprotein antibodies. Unused BSA can be stored for up to 1 week at 4  C. 9. A small aliquot of ECL substrate can be stored at room temperature, as cold ECL reagent results in lower signal intensities of the target bands. 10. In order to save on antibodies, different blotting box sizes can be used depending on the size of the membrane. For Criterion gels, the most common blotting box size is 15.5 cm (l)
10.5 cm (w)
3.5 cm (h). 11. Wiper papers do not leave traces of fiber or lint after being used to clean surfaces such as glass. 12. The Image Lab software comes with the ChemiDoc MP imager, as well as with several other Bio-Rad imagers, and can be installed on an unlimited number of lab computers. 13. Different ratios of tissue sample and homogenization buffer can be used depending on the concentration of protein sample required. 14. You should use the same type of tip and the same micropipette for all samples to minimize the difference in loading errors between the samples. Prewet the tip by sucking and dispensing the maximum volume of sample that will be loaded. In addition, a gel loading tip is recommended to prevent spillage of large sample volumes into other wells, as these tips are long with thinner openings than regular tips. Dispense the sample into the bottom of the well while slowly raising the tip toward the top of the well, ensuring that air bubbles do not get pipetted into the well. Keep the pipette plunger depressed until the tip is removed from the wells to prevent reuptake of running buffer and sample into the pipette tip. 15. When running one Criterion SDS-PAGE at 120 V, milliamps (mA’s) of less than 35 typically indicates that the SDS-PAGE running buffer is too dilute, while mA’s of more than

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80 sometimes suggests that the running buffer is too concentrated. Use new SDS-PAGE buffer. If the samples are not for quantification (such as for testing antibody specificity) you can adjust as needed with small amounts of 10
running buffer (if too dilute) or water (if too concentrated). 16. Longer activation times of 2 or 5 min do not increase the linearity of the total protein detected, although they may yield higher intensity bands as a result of more tryptophan residues being modified. However, activation times of longer than 1 min are not recommended if the primary antibody being used detects tryptophan residues, as the modification of the residues may interfere with antibody binding and subsequent detection of the protein of interest. There is no need to activate the gel again. 17. Image the membrane while still wet, as some image quality is decreased if the membrane is allowed to dry before imaging. 18. Using scissors or a razor blade, cut the membrane based on the molecular weight markers to obtain the region with the desired size of the protein of interest. If using diluted protein standards, it may be difficult to accurately cut the membrane to the desired size. In this case, stain the membrane with 0.1% Ponceau S stain for 30 s or until the proteins become clear enough. One corner of each piece of membrane can also be cut to indicate the orientation of the membrane. After cutting, remove the Ponceau S stain by washing multiple times with water or TBST. 19. After drying, the membrane can be stored at 4  C for up to 1 week. Wet the membrane with TBST before reimaging, if needed. 20. The primary antibody solution may be reused and stored at 4  C for up to 1 week. Do not use if the BSA or milk has gone bad. 21. Try using a different secondary antibody if poor results are obtained during Western blotting after multiple trials. Good quality secondary antibodies are essential for high-quality Western blots. 22. The volume of ECL substrate needed varies depending on the size of the membrane. Uncut midi membranes will need around 5–6 mL of ECL substrate, though less ECL substrate is needed if a clear plastic support (e.g., Saran wrap) is placed on top of the membrane, spreading the ECL evenly. 23. If the signal is weak, the ECL reagent can be left for longer than 2 min, but when the ECL reagent is left on the membrane for longer than 5 min, the length of time that the maximum

Stain-Free Technology

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signal lasts is shorter than when the ECL reagent is left for 2 min. Therefore, we recommend removing the excess ECL reagent and leaving the ECL reagent on for no more than 5 min (however, this will depend on your specific ECL reagent used). 24. Various image analysis software have different tools and methods to quantify gels and blots. Here, all quantification and normalization was done using Image Lab software, version 6.0.1, and the following tips are for that software. The Lane Finder tool can be used to automatically find all the lanes, though these should be further manually manipulated to better fit each lane. Do not change the width of just one lane, though individual lanes may be moved or bent. The Band Finder tool can be used to detect all the bands in each lane with the custom sensitivity set to 100%. In addition, the bands should be added, deleted, and/or adjusted to maintain consistency across lanes and samples by checking the Lane Profile tool, which shows the signal intensities of the bands above the variable background level. 25. All graphs were plotted with the SigmaPlot software, version 12. The results show that the total protein for the mouse heart homogenate was linear up to 80 μg of total protein (Fig. 2b), while the antibody against the 20S proteasome β1 was only linear up to 40 μg of total protein (Fig. 3b, c). Automatically fitting a linear line to the data can give the false impression that the antibody is linear up to 80 μg. However, we can determine when the line is no longer linear by looking for a significant reduction in the increase in signal intensity between each additional 10 μg of sample. The values for the normalized protein start dropping off after 40 μg, and the values that were not normalized begin to curve the line after this point. It is best to take a look at the raw data. In order for the normalized data to be truly linear, the relative intensity of the values should stay close to 1. If the differences between each data point varies by more than 10%, we cannot consider it to be linear. We recommend stain-free western blotting for the quantification and normalization of proteins, with the caveat that the stain-free total protein may have a higher linear detection range than the antibody used, depending on the protein samples and antibodies utilized.

Acknowledgments This work was supported by grants from the National Institutes of Health Superfund Research Program (P42 ES004699) and the American Heart Association (16GRNT31350040).

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References 1. Bass JJ, Wilkinson DJ, Rankin D, Phillips BE, Szewczyk NJ, Smith K, Atherton PJ (2017) An overview of technical considerations for Western blotting applications to physiological research. Scand J Med Sci Sports 27(1):4–25. https://doi.org/10.1111/sms.12702 2. Ghosh R, Gilda JE, Gomes AV (2014) The necessity of and strategies for improving confidence in the accuracy of western blots. Expert Rev Proteomics 11(5):549–560. https://doi. org/10.1586/14789450.2014.939635 3. Thellin O, Zorzi W, Lakaye B, De Borman B, Coumans B, Hennen G, Grisar T, Igout A, Heinen E (1999) Housekeeping genes as internal standards: use and limits. J Biotechnol 75 (2-3):291–295. https://doi.org/10.1016/ s0168-1656(99)00163-7 4. Hu X, Du S, Yu J, Yang X, Yang C, Zhou D, Wang Q, Qin S, Yan X, He L, Han D, Wan C (2016) Common housekeeping proteins are upregulated in colorectal adenocarcinoma and hepatocellular carcinoma, making the total protein a better "housekeeper". Oncotarget 7 (41):66679–66688. https://doi.org/10. 18632/oncotarget.11439 5. Pandey N, Ganapathi M, Kumar K, Dasgupta D, Das Sutar SK, Dash D (2004) Comparative analysis of protein unfoldedness in human housekeeping and non-housekeeping proteins. Bioinformatics 20(17):2904–2910. https://doi.org/10. 1093/bioinformatics/bth344 6. Dittmer A, Dittmer J (2006) Beta-actin is not a reliable loading control in Western blot analysis. Electrophoresis 27(14):2844–2845. https://doi.org/10.1002/elps.200500785 7. Liu NK, Xu XM (2006) Beta-tubulin is a more suitable internal control than beta-actin in western blot analysis of spinal cord tissues after traumatic injury. J Neurotrauma 23 (12):1794–1801. https://doi.org/10.1089/ neu.2006.23.1794 8. Lowe DA, Degens H, Chen KD, Alway SE (2000) Glyceraldehyde-3-phosphate dehydrogenase varies with age in glycolytic muscles of rats. J Gerontol A Biol Sci Med Sci 55(3):

B160–B164. https://doi.org/10.1093/ gerona/55.3.b160 9. Sander H, Wallace S, Plouse R, Tiwari S, Gomes AV (2019) Ponceau S waste: Ponceau S staining for total protein normalization. Anal Biochem 575:44–53. https://doi.org/10. 1016/j.ab.2019.03.010 10. Goldman A, Harper S, Speicher DW (2016) Detection of proteins on blot membranes. Curr Protoc Protein Sci 86:10 18 11. https:// doi.org/10.1002/cpps.15 11. Gilda JE, Gomes AV (2013) Stain-free total protein staining is a superior loading control to beta-actin for Western blots. Anal Biochem 440(2):186–188. https://doi.org/10.1016/j. ab.2013.05.027 12. Dsouza A, Scofield RH (2015) Protein stains to detect antigen on membranes. Methods Mol Biol 1314:33–40. https://doi.org/10.1007/ 978-1-4939-2718-0_5 13. Yonan CR, Duong PT, Chang FN (2005) High-efficiency staining of proteins on different blot membranes. Anal Biochem 338 (1):159–161. https://doi.org/10.1016/j.ab. 2004.11.010 14. Westermeier R (2006) Sensitive, quantitative, and fast modifications for Coomassie blue staining of polyacrylamide gels. Proteomics 6 (Suppl 2):61–64. https://doi.org/10.1002/ pmic.200690121 15. Rivero-Gutierrez B, Anzola A, MartinezAugustin O, de Medina FS (2014) Stain-free detection as loading control alternative to Ponceau and housekeeping protein immunodetection in Western blotting. Anal Biochem 467:1–3. https://doi.org/10.1016/j.ab. 2014.08.027 16. Vigelso A, Dybboe R, Hansen CN, Dela F, Helge JW, Guadalupe Grau A (2015) GAPDH and beta-actin protein decreases with aging, making stain-free technology a superior loading control in Western blotting of human skeletal muscle. J Appl Physiol (1985) 118 (3):386–394. https://doi.org/10.1152/ japplphysiol.00840.2014

Chapter 29 Technical Considerations for Contemporary Western Blot Techniques Kenneth Oh Abstract Western blotting continues to be a workhorse assay in laboratories throughout the world. The utility, low cost and accessibility of western blotting have allowed the technique to remain in practice, despite being developed over 40 years ago. Advances in antibody specificity, chemiluminescent formulations, properties of fluorescent molecules and imaging techniques provide gains in sensitivity, dynamic range, and ease of use. Here we discuss such aspects for the users’ consideration when planning and executing western blots, to take full advantage of contemporary practices. Key words Western blot, Immunoblotting, Immunodetection, Chemiluminescence, Enhanced chemiluminescence, Fluorescence, Multiplex, Digital imaging, CCD, Dynamic range, Binning, Bit depth, Resolution, Quantitation, Total protein normalization, Housekeeping normalization

1

Introduction The most commonly used protein blotting technique, western blotting (immunoblotting), was developed for specific detection of proteins that are inaccessible to antibodies in polyacrylamide gels. Western blotting involves the transfer of proteins that have been separated by gel electrophoresis onto a membrane, followed by immunological detection of these proteins. Western blotting combines the resolution of gel electrophoresis with the specificity of immunoassays, allowing individual proteins in mixtures to be identified and analyzed. As there are many excellent technical references for the electrophoresis and blotting steps during the workflow [1], presented here is an overview of methods, techniques, and considerations starting with the immunological detection of the target proteins (immunodetection). Because the membrane surface has a high affinity for proteins, the blot must be incubated with a solution of inert protein(s) or passivating agents before any antibody is added. This blocking step coats the unoccupied area of the blot to prevent nonspecific

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_29, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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binding of primary antibodies. Blocking and primary antibody incubation require selection of appropriate methods and reagents. The next step is the detection step, referred to here as the signal acquisition step. Signal acquisition converts biological abundance on the blot to a measurable output that can be recorded electronically. This may be performed using several different methods, including chemiluminescent, fluorescent, and colorimetric detection. The last step is image analysis. Signal interpretations are performed in order to accurately calculate protein quantities from signal intensities from multiple samples, multiple targets within the same sample, or both. Normalization is typically performed in order to accurately report intensity differences arising from biological events versus error from manual manipulation or other technical variations.

2

Immunodetection The purpose of immunodetection (immunological detection) is to identify specific proteins blotted to membranes. After the proteins have been transferred to the membrane, the steps used for immunological detection vary little. In short, the membrane is blocked, incubated with a primary antibody, and then incubated with a secondary antibody, with washes after each of the antibody incubation steps. When selecting an immunodetection protocol, many factors must be considered. For the blocking step, the blocking buffer composition and incubation time must be selected. For each primary or secondary antibody, consider the species that the target protein was produced in, the species the antibody was produced in, the quality of the antibody, the quantity of antibody required, and the incubation time and buffer composition. The conjugate must also be considered for secondary antibodies, taking into account the desired detection method. Finally, the number and length of the wash steps must be determined, as well as the composition of the wash buffer(s). In general, the primary antibody is specific for the protein of interest and the secondary antibody enables its detection. The secondary antibody can be labeled with a fluorescent compound or conjugated to an enzyme, such as alkaline phosphatase (AP) or horseradish peroxidase (HRP), for subsequent detection. For many years, radiometric secondary antibodies were the method of choice for detection, but newer methods have evolved that are equally sensitive, easier to use, and less hazardous. Available detection methods now also include chemiluminescent, fluorescent, colorimetric, bioluminescence, chemifluorescent, and immunogold detection.

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2.1

Blocking

459

A blocking step is necessary to coat the remaining membrane surface not occupied by proteins transferred from the sample. This prevents nonspecific binding of the antibodies to the membrane, which has a high affinity for proteins. Blocking reagents must be chosen carefully to avoid interfering with target-antibody binding. When choosing a blocking procedure, consider the following: 1. The composition of the blocking agent (protein- or nonprotein-based). 2. The percentage detergent (if applicable). 3. Blocking time. The ideal time is 1 h at room temperature. 4. Volume of blocking buffer. This is not a place to skimp. Failing to completely block unoccupied sites can lead to high background. A variety of blocking reagents are available, including gelatin, nonfat milk, and BSA, which are compared in Table 1. Optimize the detection system for minimal background with no loss of signal by testing several blocking agents. The type of membrane also affects the selection of blocking reagent. After blocking, a typical immunodetection protocol utilizes two rounds of antibody incubation. The first round contains the primary antibody, which is directed against the target antigen(s). Possible antigens include proteins, protein ligands, specific epitopes on a protein, or carbohydrate groups. The second round of incubation contains the secondary antibody, which recognizes and binds to the primary antibody. The secondary antibody is usually conjugated to an enzyme, such as AP or HRP, which is used for detection. Antibody incubations are generally carried out in antibody buffer containing Tris-buffered saline with Tween and a blocking reagent.

Table 1 Comparison of blocking reagents

Blocking Reagent

Membrane Recommended Compatibility Concentration Notes

Gelatin

Nitrocellulose 1–3%

Requires heat to solubilized

Nonfat dry milk, BLOTTO, blottinggrade blocker

All

0.5–5%

PVDF and LF-PVDF require higher concentrations of nonfat milk than nitrocellulose

Bovine serum albumin (BSA)

All

1–5%

PVDF and LF-PVDF require higher concentrations of nonfat milk than nitrocellulose

Tween 20

All

0.05–0.3%

May strip some proteins from the blot

EveryBlot blocking buffer

All

N/A

Universal blocking buffer, supplied at 1


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2.2 Wash Volume and Agitation

Washing the blot between the two antibody incubations and prior to detection removes excess antibodies and prevents nonspecific binding. Though washing solutions and times may vary depending on the antibodies and detection systems used, washes generally utilize Tris-buffered saline (TBS) or phosphate-buffered saline (PBS). A detergent, such as Tween 20, may be added to decrease background; however, detergents may inhibit certain detection reactions.

2.3 Antibody Selection and Dilution

An antibody is an immunoglobulin protein, such as IgG, which is synthesized by an animal in response to exposure to a foreign substance, or antigen. Antibodies have specific affinity for the antigens that elicited their synthesis. Structurally, most IgG antibodies contain four polypeptide chains (two identical heavy chains of ~55 kD and two identical light chains of ~25 kD) oriented in a “Y” shape (Fig. 1). These four chains are held together by disulfide bridges and noncovalent interactions. These proteins contain a region with specific affinity for the antigen that elicited their synthesis, known as the Fab region. In addition, a constant region (Fc) on the antibody provides binding sites for proteins needed during an immune response.

en e tig sit An ing d bin

bin Anti din gen g sit e

Variable Constant

Fab Light chain

Fc

Heavy chain

Fig. 1 Antibody structure. The components of a typical IgG molecule are highlighted. The Fab fragment contains the variable region responsible for antigen binding, while the Fc (constant) region is necessary for binding other proteins involved in the immune response

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The primary antibody recognizes and binds to the target antigen on the membrane. For incubations with primary antibody, the entire blot must be covered with antibody-containing solution. The optimum antibody concentration is the greatest dilution of antibody that still yields a strong positive signal without background or nonspecific reactions. Instructions for antibodies obtained from a manufacturer typically suggest a starting dilution range. For custom antibodies or for those where a dilution range is not suggested, good starting points are as follows: 1. 1:100–1:1000 dilution when serum or tissue culture supernatants are the source of the antibody. 2. 1:500–1:10,000 dilution of chromatographically purified, specific antibodies. 3. 1:1000–1:100,000 dilution may be used when ascites fluid is the source of the antibody. Note: Determine the appropriate concentration or dilution (titer) of the primary antibody empirically for each new lot of primary antibody. The Bio-Rad Mini-PROTEAN II multiscreen apparatus and mini incubation trays are useful tools for determining antibody titer.

2.3.2 Secondary Immunodetection

Secondary antibodies are specific for the isotype (class) and species of the primary antibody; for example, a goat anti-rabbit secondary antibody is an antibody generated in goat for detection of a primary antibody that was generated in rabbit. Secondary antibodies bind to multiple sites on primary antibodies to increase detection sensitivity. For immunodetection, use only blotting-grade, species-specific secondary antibodies. Secondary antibodies can be labeled and detected in a variety of ways. The antibody may be labeled with a fluorescent protein or metallic particles, or, most commonly, conjugated to an enzyme, such as AP or HRP. If the secondary antibody is biotinylated, biotin-avidin-AP or -HRP complexes can be formed. Because the purity of the reagents is critical to the success of the experiment, the following steps are critical if the antibodies used are not of blotting grade: 1. Purify all sera by affinity chromatography to obtain only those antibodies directed against the particular IgG of interest; otherwise, background staining and false positive reactions due to nonspecific antibody binding may occur. 2. Cross-adsorb the purified antibody solution against a solution of IgG from an unrelated species; for example, human IgG for anti-rabbit or anti-mouse antibodies and bovine IgG for antihuman antibodies, to remove antibodies that are not specific for the species of interest.

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Blotting-grade antibodies are directed to both heavy and light chains of the IgG molecules, so the reagents can also be used to identify other classes and subtypes of immunoglobulins. Some protein molecular weight standards include specific tags which can be recognized by an HRP-conjugate so that the bands can be detected with a chemiluminescence system. If applicable, add the appropriate HRP conjugate(s) during incubation with the secondary antibody. 2.3.3 Antibody-Specific Ligands

3

Protein A and protein G are bacterial cell surface proteins that bind to the Fc regions of immunoglobulin molecules [2–4]. The advantage of using protein A or protein G is their ability to bind to antibodies of many different species (Table 2). This is often desirable for laboratories using antibody probes from many different species or for those using one of the less common primary antibody systems in their experiments, such as rat, goat, or guinea pig. In addition, these reagents bind only to antibody molecules. This can reduce the background from nonspecific binding of antibodies to membrane-bound proteins when a low titer, poorly purified secondary antibody is used. The major limitation of protein A and protein G conjugates is their lower sensitivity. Because only one ligand molecule binds to each antibody, the enhancement of a multiple-binding detection system, such as a species-specific polyclonal antibody, is lost. Generally, the species-specific antibody is 10–50 times more sensitive than the ligand reagent when the same detection system is used.

Image Acquisition The image acquisition step converts biomolecular identity and abundance into identifiable and measurable signals. This is accomplished by, quite simply, converting photons into electrons. The single most important aspect of image acquisition is optimizing the signal-to-background ratio (S/B), also referred to as the signal-tonoise ratio (S/N). Thoughtful considerations and careful execution at all manual steps in the western blotting workflow contribute to both the maximization of signal and minimization of background. It is the balance of these two factors that will generate the highest quality image with the most accurate results.

3.1 Visualization Methods

Blotted proteins are generally detected using secondary antibodies that are labeled with enzymes, such as horseradish peroxidase (HRP) or alkaline phosphatase (AP), or fluorescent molecules (fluorophores). Historically, colloidal gold or radioisotope 125 I-labeled reagents similar to those used in radioimmunoassays have also been utilized. These systems provide sensitive results but

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Table 2 Immunoglobulin-binding specificities of protein A and protein G Immunoglobulin

Protein A

Protein G

Human IgG1

ll

ll

Human IgG2

ll

ll

Human IgG3

–

ll

Human IgG4

ll

ll

Mouse IgG1

l

l

Mouse IgG2

ll

ll

Mouse IgG3

ll

ll

Mouse IgG4

ll

ll

Rat IgG1

l

l

Rat IgG2a

–

ll

Rat IgG2b

ll

l

Rat IgG2c

ll

ll

Pig IgG

l

ll

Rabbit IgG

ll

ll

Bovine IgG1

–

ll

Bovine IgG2

ll

ll

Sheep IgG1

–

ll

Sheep IgG2

l

ll

Goat IgG1

l

ll

Goat IgG2

ll

ll

Horse IgG(ab)

l

ll

Horse IgG(c)

l

ll

Horse IgG(t)

–

l

Dog IgG

ll

l

ll

¼ Strong binding, l ¼ Weak binding, — ¼ No binding

the special handling and disposal problems of 125I reagents have discouraged continued use of this technique. Instead, a number of enzyme systems and detection reagents have evolved. For practical purposes, there are three common types of signal visualization: chemiluminescent, fluorescent, and colorimetric. The most commonly used detection methods use secondary antibodies conjugated to horseradish peroxidase (HRP) or alkaline phosphatase (AP). These enzyme-based methods emit light

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a Colorimetric

b Chemiluminescence Substrate

Substrate Light

c Fluorescence

Product Product

Light

Fig. 2 Detection chemistry mechanisms. In each method of western blot detection, a detectable signal is generated following binding of an antibody specific for the protein of interest; (a) in colorimetric detection, the signal is a colored precipitate; (b) in chemiluminescence the reaction itself emits light; (c) in fluorescence detection, the antibody is labeled with a fluorophore

(chemiluminescence detection) or generate a colored precipitate (colorimetric detection) when the appropriate substrate is added. The light signal from a chemiluminescent reaction can be captured with a digital imaging system or film (Fig. 2). Secondary antibodies conjugated to fluorophores are gaining popularity and can be directly visualized on the blot with a compatible imager, without the need for additional liquid substrate. Although colorimetric blots have been used in the past, they suffer from a limited dynamic range and have been outstripped by advancements in chemiluminescence and fluorophore synthesis. Today, colorimetric detection is used less and less frequently, and therefore will not be discussed here. The signal of each of these methods is acquired via several methods, including camera-based digital imagers and flatbed scanners. Due to the preponderance of use in laboratories today, we will only be discussing camera-based digital imagers. 3.1.1 Chemiluminescence Detection

Chemiluminescence detection uses an enzyme-catalyzed chemical reaction which produces light as a byproduct. AP and HRP are the enzymes most commonly used for chemiluminescence detection (Fig. 3). The light signal can be captured on X-ray film, a chargecoupled device (CCD) imager, or a complementary metal-oxide semiconductor (CMOS) imager. The principal advantages of chemiluminescent western blotting over other methods are its speed and sensitivity (Table 3). Chemiluminescence detection works well with CCD imaging, which avoids the slow film step. Exposure times for average blots are usually 5 s to 5 min, depending on the sensitivity of the substrate. Detection of protein down to femtogram amounts is possible with these systems. The detection sensitivity depends on

Technical Considerations for Contemporary Western Blot Techniques Luminol

CDP-Star O

465

NH2

O

O

CH3

O

NH

Cl

OPO3Na2 NH

Cl

AP + H2O

O HRP + H2O2

– OPO32– O

O

O-

NH2

CH3

O Cl

O-

N

O O

N

Peroxy intermediate

Cl OO O

O

CH3 O-

+ Cl

NH2 COO-

Product in excited state

Cl Product in excited state

COO-

Light emission Light emission

Fig. 3 Chemiluminescence detection. The secondary antibody is linked to an enzyme which catalyzes a reaction leading to light emission. (Left) CDP-Star or another 1,2-dioxetane AP substrate is dephosphorylated by AP, resulting in formation of an anion in an excited state that emits light. (Right) luminol oxidized by HRP in the presence of H2O2 leads to the formation of 3-aminophthalate dianion and the release of light

the affinity of the protein, the primary and secondary antibodies, and the HRP substrate, and can vary from one sample to another. A more extensive review of chemiluminescence reaction principles and applications is available elsewhere [5]. 3.1.2 Fluorescence Detection

In fluorescence detection, a primary or secondary antibody labeled with a fluorophore is used for immunodetection. A light source excites the fluorophore and the emitted fluorescent signal is captured by a CCD or CMOS imager to produce the final image. In

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Table 3 Chemiluminescence detection systems Trade Detection method name

Substrate Sensitivity

Advantages

Disadvantages

Luminol Chemiluminescent Clarity HRP Western ECL substrate

Azide inhibits MidShort exposure (30 s); enzyme femtogram compatible with PVDF activity and nitrocellulose; signal duration up to 24 h; stable at room temperature

Luminol Chemiluminescent Clarity HRP max Western ECL substrate

LowCompatible with PVDF femtogram and nitrocellulose; optimized for CCD imagers; high sensitivity; stable at room temperature

Azide inhibits enzyme activity

Chemiluminescent ImmunAP star AP

10 picogram 30 s to 5 min exposure; signal continues for 24 h after activation; blot can be reactivated

Endogenous phosphatase activity may lead to false positives

CDPStar

S' 1

Energy

S1 hνex

hνem

S0 Fig. 4 Jablonski diagram of fluorophore excitation and emission

fluorescence, a high-energy photon (hνex) excites a fluorophore, causing it to leave the ground state (S0) and enter a higher energy state (S0 1). Some of this energy dissipates, allowing the fluorophore to enter a relaxed excited state (S1). A photon of light is emitted (hνem), returning the fluorophore to the ground state. The emitted photon is of a lower energy (longer wavelength) due to the dissipation of energy while in the excited state (Fig. 4). When using fluorescence detection, consider the following optical characteristics of the fluorophores to optimize the signal:

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1. Quantum yield is the efficiency of photon emission after absorption of a photon. Processes that return the fluorophore to the ground state but do not result in the emission of a photon lower the quantum yield. Fluorophores with higher quantum yields are generally brighter. 2. Extinction coefficient is a measure of how well a fluorophore absorbs light at a specific wavelength. Since absorbance depends on path length and concentration (Beer’s law), the extinction coefficient is usually expressed in cm1 M1. As with quantum yield, fluorophores with higher extinction coefficients are usually brighter. 3. Stokes shift is the difference between the maximum excitation and emission wavelengths of a fluorophore. Since some energy is dissipated while the fluorophore is in the excited state, emitted photons are of lower energy (longer wavelength) than the light used for excitation. Larger Stokes shifts minimize overlap between the excitation and emission wavelengths, increasing the detected signal. 4. Excitation spectra are plots of the fluorescence intensity of a fluorophore over the range of excitation wavelengths; emission spectra show the emission wavelengths of the fluorescing molecule. Choose fluorophores that can be excited by the light source in the imager and that have emission spectra that can be captured by the instrument. When performing multiplex western blots, choose fluorophores with minimally overlapping spectra to avoid channel crosstalk (Fig. 5).

Relative intensity

Stokes shift Absorption (excitation) Emission Spectral overlap

400

500

600 Wavelength, nm

700

800

Fig. 5 Spectral considerations for fluorophore excitation and emission

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Fig. 6 Multiplex fluorescence detection of a two-fold dilution series of two proteins, GST (red) and soybean trypsin inhibitor (green). Starting concentration was 500 ng of each protein

Several fluorophores spanning a wide range of excitation and emission wavelengths are now available, including some based on organic dyes (e.g., cyanine and fluorescein), nanocrystals of semiconductor material (e.g., Qdot nanocrystals), and naturally fluorescent proteins (e.g., phycobiliproteins such as phycoerythrin and allophycocyanin). Fluorescence detection (Fig. 6) offers several advantages over other methods: 1. Multiplexing—use of multiple distinct fluorophores for simultaneous detection of several target proteins on the same blot. When detecting multiple proteins in a fluorescent multiplex western blot, ensure the fluorescent signals generated for each protein can be differentiated. Use primary antibodies from different host species (e.g., mouse and rabbit) and secondary antibodies that are cross-adsorbed against other species to avoid cross-reactivity. Use fluorophores conjugated to secondary antibodies with distinct spectra to avoid cross-channel fluorescence. 2. Dynamic range—fluorescence detection offers a ten-fold greater dynamic range over chemiluminescence detection, and therefore better linearity within detection limits. 3. Stability—many fluorescent molecules are stable for a long period of time, allowing blots to be stored for imaging at a later date—often weeks or months later—without significant signal loss. Most fluorescence techniques are also compatible with stripping and reprobing protocols (provided the blots are not allowed to dry out between successive western detection rounds).

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A historical drawback of fluorescence detection is its reduced sensitivity compared to chemiluminescence methods, such that detection using low-affinity antibodies or of low-abundance proteins may yield lower signals. However, the recent development of photostable fluorophores, improved instrumentation, and membranes with low autofluorescence characteristics allows for fluorescence detection to approach the sensitivities seen with chemiluminescence techniques. 3.2 Stripping and Reprobing

Membranes that have been detected with noncolorimetric methods, such as chemiluminescent or fluorescent techniques, can be stripped of antibodies for use in subsequent rounds of Western detection (Fig. 7). This allows for reuse of the same blot for investigation of different proteins and saves both time and sample material. Blots can be stripped and reprobed several times; however, each round of stripping removes sample protein from the blot [6]. This decreases the sensitivity of later rounds of detection and may necessitate longer exposure times or more sensitive detection methods. When probing a blot multiple times: 1. If detecting proteins of different abundance or when using antibodies with very different binding affinities, first detect the protein with the lower expected signal intensity. 2. Comparisons of target protein abundance across different rounds of detection will be unreliable, as some sample is removed during the stripping process. 3. If possible, use PVDF membranes. PVDF is more durable and resists loss of sample better than nitrocellulose membranes. After stripping a blot, test it for complete removal of the antibody. If chemiluminescent detection methods were used, confirm removal of the secondary antibody by incubation with fresh chemiluminescent substrate. Detect any remaining primary antibodies by incubation with an HRP-labeled secondary antibody followed by incubation with fresh chemiluminescent substrate. If any antibody is detected using these tests, restrip the blot before subsequent experiments.

3.3

Imaging Systems

Several methods are employed to document western blotting results (Table 4). CCD cameras are versatile systems that image both gels and blots. CCD systems operate with either transillumination provided by light boxes (visible or UV) positioned underneath the gel for imaging a variety of stains (Coomassie, silver, fluorescence) or epi-illumination of blots detected using colorimetric or fluorescence techniques. Different illumination wavelengths are available for multiplex fluorescence immunodetection. CCD cameras can also be used without illumination to detect

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Fig. 7 Stripping and reprobing PVDF membranes. E. coli lysate containing human transferrin and a GST-tagged protein was loaded on a gel and blotted onto PVDF membranes; (a, Upper left) the blot probed with an anti-GST antibody and developed with chemiluminescent substrate; (b, Upper right) the same blot subsequently stripped of antibody and reprobed with the secondary antibody and developed with chemiluminescent substrate to demonstrate removal of primary antibody. This blot was also reprobed with StrepTactin-HRP to visualize the ladder; (c, Lower left) the stripped blot reprobed with an antihuman transferrin antibody; (d, Lower right) a control blot that did not undergo the stripping procedure probed with the anti-human transferrin antibody

luminescent signals. Supercooled CCD cameras reduce image noise, allowing detection of faint luminescent signals. Historically, X-ray film was the most prevalent method of recording luminescent signals; however, modern digital imaging instruments have essentially replaced the use of film, particularly given the maintenance and environmental costs associated with maintaining X-ray film developing facilities.

Technical Considerations for Contemporary Western Blot Techniques

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Table 4 Comparison of western blot documentation and analysis methods Imaging System ChemiDoc MP™

ChemiDoc™

ChemiDoc™XRS+

Chemiluminescence

l

l

l

Fluorescence

l

—

—

Colorimetric

l

l

l

Stain-Free

l

l

l

Colorimetric

l

l

l

Fluorescent

l

UV only

UV only

l

l

l

Trans UV/Vis

l

l

l

Epi white

l

l

l

LED RGB

l

—

—

LED NIR

l

—

—

Immunodetection

Total Protein Stain

Imager Type Supercooled CCD Excitation Type

The ChemiDoc MP Imaging System is capable of up to three-channel multiplex fluorescence detection in a single exposure

3.3.1 Digital Imaging for Chemiluminescence Detection

For chemiluminescence detection, CCD imaging is the easiest, fastest, and most accurate method. Traditionally, the chemiluminescent signal from blots was detected by X-ray film. Film is a sensitive medium for capturing the chemiluminescent signal but suffers from a sigmoidal response to light with a narrow region or linear response, which limits its dynamic range. To gather information from a blot that has both intense and weak signals, multiple exposures are required to produce data for all samples in the linear range of the film. Additionally, quantitation of data collected by exposure to film requires digitization (that is, scanning of X-ray film with a densitometer). CCD cameras have a linear response over a broad dynamic range (2–5 orders of magnitude) depending on the bit depth of the system. Unlike film, CCD cameras do not require continuous purchase of consumables, and provide a convenient digital record of experiments for data analysis, sharing, and archiving. CCD cameras also approach the limit of signal detection in a relatively short

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time. For example, the Bio-Rad ChemiDoc MP Imaging System can reach the limit of detection of a given experiment in Rhod interior mean intensity geometric mean and 0.05 mm < radius < 0.25 mm to automatically pick and deposit HCP-binding beads into the deposit plates. Perform the pick run in suspension mode with a 20 μL aspiration volume and 60 μL expel volume. 12. Elute the proteins from the selected beads by incubation with 20 μL of bead elution buffer on an orbital shaker at room temperature for 1 h. 13. Wash the beads three times with 100 μL of bead equilibration buffer. Remove the supernatant (e.g., by centrifugation or vacuum). 14. Seal the plates and store at 2–8  C until ready for sequencing. 15. Wash the beads five times with 200 μL of deionized water and sequence the peptides (e.g., Edman degradation or LC/MS/ MS). 3.3 Confirmation of HCP Binding by Selected Peptides

This section describes the capture of HCPs from a cell culture harvest. 1. Determine the values of Load Factor (i.e., mass of protein to be loaded per volume of resin) to be evaluated in static binding mode (see Note 24). 2. Calculate the values of volume of harvest load required to attain the desired load factor assigned at step 1 when using >50 μL volume of settled resin. V Load ¼

Load Factor  V Resin , C

where VLoad is the required volume of cell culture harvest, VResin is the volume of resin to be aliquoted, and C is the concentration of HCPs referenced in the load factor in the production harvest material to be loaded. 3. Aliquot a volume of VResin of test resin into SPE tubes. For resins provided dry, initially suspend the resin in resin equilibration buffer and allow the beads to swell at room temperature for 2–24 h. For resins provided as a slurry, aliquot the desired volume using centrifugation to pellet the resin and ensure that the volume utilized is accurate. Care should be taken, when transferring the resin slurry, to include washing

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steps to recover the resin beads that adhere to the walls of the containers (e.g., centrifuge tubes or SPE tubes). 4. Using a vacuum manifold or syringe adaptor, wash the resin: add equilibration buffer at 3–5 the resin volume, cap the SPE tube and gently mix by inversion, and remove the supernatant. Repeat three times, leaving the final wash in the tube until ready to apply the cell culture load. 5. Remove the wash solution and add the volume of cell culture harvest calculated in step 2 above. Cap the tube and incubate on a rotator at room temperature for 1 h. Retain 1–2 mL for the production harvest solution for further analysis. 6. Collect the spent harvest. 7. Wash the resin with resin equilibration buffer (2–5 the resin volume). Gently invert to mix and collect the wash. Pool the spent harvest and all the wash fractions. If the samples are not analyzed immediately, aliquot them to prevent the requirement for multiple freeze-thaws, and store at 20  C until ready for use. 8. Measure the concentration of total protein, total HCP, and protein product in the load and pooled samples using preferred quantification assays. The selectivity of HCP removal is determined by comparing the partition coefficient (Kp, the ratio of mass of protein bound vs. unbound) for both HCP and protein product, where Kp,HCP  Kp,Product denotes selective binding of HCPs. 3.4 Proteomic Analysis of the Chromatographic Fractions [14] 3.4.1 Filter-Aided Sample Preparation (FASP) [16]

1. Using the total protein concentration (see step 8 in Subheading 3.3), calculate the volume to be FASP-prepared (VFASP ) for a total protein mass of ~150 μg (see Note 13). 2. Transfer the sample volume calculated at step 1 to a 10 kDa MWCO 0.5 mL spin filter. 3. Add DTT solution (0.05 the sample volume), mix by pipetting, and incubate the samples at 56  C for 30 min. 4. Add 200 μL UA solution, mix by pipetting, and centrifuge the filter unit at 14,000  g for 15 min. Discard the filtrate. 5. Add a second 200 μL UA solution, mix by pipetting, and centrifuge the filter unit at 14,000  g for 15 min. Discard the filtrate. 6. Add 100 μL IAA buffer, mix by pipetting, and incubate protected from light at room temperature for 20 min. 7. Centrifuge at 14,000  g for 10 min at room temperature. Discard the filtrate.

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8. Add 100 μL UA solution, mix by pipetting, and centrifuge at 14,000  g for 10 min at room temperature. Repeat twice, and discard the filtrate. 9. Add 100 μL ABC solution, mix by pipetting, and centrifuge at 14,000  g for 10 min at room temperature. Repeat twice, and discard the filtrate. 10. Replace the filtrate collection tube with a fresh tube. Add 50 μL trypsin solution, mix by pipetting, and incubate at 37  C for 4–18 h. 11. Add 40 μL ABC solution, mix by pipetting, and centrifuge at 14,000  g for 10 min at room temperature. Repeat once. 12. Discard the retentate, and concentrate the filtrate to dryness using a vacuum concentrator. Store at 20  C until ready for LC/MS/MS analysis. 3.4.2 nanoLC-MS/MS Analysis

1. Reconstitute the sample in 200 μL mobile phase A (MPA) prior to injection. 2. Load 2 μL microliters of sample/control on the Q-Exactive HF-X nanoLC-MS/MS system, subjected to online desalting and reverse-phase nano-LC separation (“trap and elute” configuration) and perform reverse-phase (RP) separation using an EASY-Spray™ C18nanoLC column at a flow rate of 300 nL/ min. 3. Collect the data using a data-dependent acquisition (DDA) method for discovery proteomics (see Note 25). 4. Perform post-acquisition data analysis (see Note 26). 5. As quality control analyze standard HeLa digest at the beginning and end of experiments to verify proteome coverage (# protein groups identified >3000 is considered good coverage). 6. Sample queue for quality control: blank (MPA), QC (standard BSA digest), blank, Sample-01, Sample-02, Sample-03, blank, QC, blank, Sample-04, Sample-05, Sample-06, blank, QC, blank, and so on.

3.4.3 Statistical Analysis of the DDA Proteomics Data

1. Calculate the spectral abundance factor (SAF) for each protein identified in each sample. Exclude any proteins whose sum total spectral count across three replicates is 4 across triplicate samples) in the feed, but not in the pooled sample, OR species with statistically significantly higher abundance ( p < 0.05 by ANOVA) in the feed compared to the pooled samples.

4

Notes 1. ChemMatrix resins were selected based upon extensive work in previous publications [17–20]. Marani et al. noted that ChemMatrix enables the detection of false positives in screening, although other resins may be used [17]. Solid-phase peptide libraries are commercially available, or can be synthesized on-site using well-documented methods for library synthesis, such as the “split-couple-recombine” method [21–24]. 2. The sequencing of peptides carried on library beads can be performed either by liquid chromatography tandem mass spec (LC/MS/MS) or N-terminal sequencing, also known as Edman degradation. Sequencing by LC/MS/MS requires cleavage of the peptides from the solid support prior to sequencing. Accordingly, the synthesis of the peptide library must be preceded by the resin functionalization with a linker that enables peptide cleavage under conditions that are orthogonal to the deprotection step conducted by acidolysis after chain elongation. In prior work we have utilized alkaline-labile hydroxymethylbenzoic acid (HMBA)-ChemMatrix (PCAS Biomatrix, Saint-Jean-sur-Richelieu, Quebec, Canada) as resin of choice for synthesis and screening of OBOP libraries. Peptide sequencing by Edman degradation, on the other hand, may be performed irrespectively of the peptide linkage to the solid phase. Accordingly, aminomethyl-ChemMatrix resin is suitable for this application. 3. The bead-sorting technologies discussed herein are currently best suited for resins with particle diameter ranging between 100 and 300 μm, such as ChemMatrix resins (PCAS Biomatrix, Saint-Jean-sur-Richelieu, Quebec, Canada). However, other resins may be used with modification of bead sorting technologies for size ranges. 4. The fluorescent labels were selected based on the fluorescence detectors available on-site that minimized overlapping excitation/emission wavelengths. However, other fluorescent dyes may be utilized for protein labeling.

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5. Many peptide synthesis resins show some autofluorescence in green range (e.g., FITC or Alexa Fluor 488). We therefore recommend using this dye for nontarget species to minimize the false positive identification of target-binding peptides, particularly when the target species is presence at lower abundance. 6. Density can be reasonably assumed to be ~1 g/mL for these solutions. 7. Minimal to no titration is required for these solutions. 8. Custom peptides can be commercially sourced for conjugation to chromatographic resins, or may be synthesized in-place on Toyopearl AF-Amino-650 M resins [24, 25]. 9. The equilibration buffer for static binding tests should reproduce the pH, buffer capacity, and conductivity conditions of the null clarified harvest to be loaded. If cationic amino acids (arginine, lysine, or histidine) feature prominently in the HCP-binding peptide candidates, it is recommended to select a positively charged buffer ion, such as Bis-Tris buffer. Conversely, negatively charged buffer ions such as phosphate are recommended for peptides rich in anionic residues (aspartate and glutamate). 10. ELISAs for HCP quantification are commercially available and are specific to cell type. 11. Quantification methods that are specific to a given protein product may vary. ELISAs are common for this application, however other means of quantification may be more appropriate (e.g., analytical Protein A or Protein G for mAb quantification). 12. The requirements of the total protein assay may vary from system to system, and components of the cell culture harvest should be taken into account based on reagent assay compatibility. For example, some cell culture media include phenol red in the formulation, a component that causes elevated signal for bicinchoninic acid (BCA)-based assays, making them unfit to that quantification. 13. Trypsin concentration should be based on the amount of total protein loaded onto the filters, where the target ratio is 1:100 trypsin–protein. The limits of this procedure can range between 2–250 μg of total protein loaded on the filters in a volume between 20 and 200 μL. 14. Buffer exchange may be performed using spin filters with an appropriately sized molecular weight cutoff (MWCO) for the competitor species (typically, 3–5 times smaller than the molecular weight of the target protein). Following centrifugation,

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the desired buffer is added to the retentate and centrifuged repeatedly to replace the initial buffer. Typically, 3 or more diavolumes of exchange are suitable for this purpose. 15. Concentration factors at this centrifugation speed and time may vary drastically depending on the feed source and should be monitored closely. A 90-min centrifugation cycle for our harvest material resulted in 3–4-fold concentration factor. 16. Concentration of the harvest may result in precipitation of some host cell proteins, and monitoring the retentate to detect precipitation is recommended. If precipitation is observed, repeat the concentration step with unconcentrated harvest and reduce the target concentration factor. 17. The volume of library beads for incubation is dependent on the throughput of the screening device used. With HMBAChemMatrix bead libraries, we have found that a settled volume of library beads of ~1 μL corresponds to ~100 beads for screening. 18. Load factor range is approximate and may need optimization for other systems. 19. Exposure and intensity settings may vary based on the target protein, fluorophore system, and microscope and camera. It is recommended that a pool comprising >100 test beads is initially tested on the camera to refine the choice of settings that prevent saturation. 20. If resins other than ChemMatrix are used for the library, the settled volume must be verified. 6-well plates should be seeded with Export>Export for Analysis). This creates 16 bit tiff images. 2. Open 2-DE image analysis software, import 16 bit tiff images and perform data analysis as briefly describes in Fig. 1. Calculate match rate that correspond to the ratio of spots detected on the immune-stained blot to the total number of unique spots detectable on both the Oriole stained gel and the immunostained blot (Table 1). The Stain-free blot image (identical physical size as the immunostained blot image) has a “hinge” function to enable the matching of the Oriole stained gel image and the immunostained blot image. Usually gel images and blot images do not have identical physical dimensions making spot alignment without SF blot image more difficult.

Table 1 Generation of match rate for anti-HEK293 HCP antibody from 2D image analysis data Number of features detected on Oriole stained gel

360

Number of features detected with anti-HCP (western blot)

130

Total number of unique features detected between the two methods

415

Match rate (ratio of features detected with anti-HCP to total detected features)

31%

Host Cell Protein Characterization

4

521

Notes 1. Cells should be grown in protein-free medium so that proteins that do not derive from cultured cells will be found in the cell culture medium. 2. EDTA is a broad-spectrum metalloproteinase inhibitor. 3. AEBFS (trade name: Pefabloc SC) is an irreversible serine protease inhibitor. Inhibition constants are similar to those of PMSF. As an alternative to PMSF, AEBSF offers lower toxicity, improved solubility in water and improved stability in aqueous solutions. 4. The 0.2 M Tris stock solution has a pH of 10.5–10.6. Do not titrate with HCl. 5. The sample solubilization buffer I has a pH of 9.0–9.1. If necessary, adjust pH with diluted HCl or NaOH. This solution should be prepared shortly before use. 6. The IEF instrument provides single lane control and this new concept enables concurrent monitoring of the current and voltage profiles of each individual IPG-strip during separation. This design enables to process samples of different conductivity or IPG-strips with different pH-ranges and/or protocols in parallel. Since the first-dimension separation is often the most critical with respect to its influence on the quality and resolution of the 2-DE separation this new IEF-concept will improve data quality and furthermore help to identify strips/samples, which did not perform properly in order to exclude them from the current 2-DE experiment. 7. Solutions containing urea are prepared immediately before use. 8. It is convenient to use a concentrated stock solution of Bromophenol Blue, but this dye will not dissolve in unbuffered water. It is therefore prepared in Tris base solution. 9. Other pH ranges may be used. pH 3–10 NL (nonlinear) was chosen for this experiment because it covers most of the range of protein pIs. This nonlinear gradient is flatter in the region of the gradient where most proteins are found, thereby spreading them out and allowing for resolution of more individual proteins. 10. Mineral oil prevents evaporation of the sample solution. 11. Equilibration buffer is highly viscous and it may be difficult to stir efficiently. Allow several hours for the urea to dissolve into the liquid components. 12. The amount of Equilibration solution with DTT or iodoacetamide may be scaled up or down depending on the number of IPG strips run. These solutions may be prepared from

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aliquoted Equilibration buffer (see step 5). Frozen tubes of Equilibration buffer should be allowed to thaw slowly in a beaker of water (do not heat). 13. Stain-free gel chemistry use a unique trihalo compound, which, when activated with UV light, reacts with tryptophan residues allowing for quick visualization of proteins without any staining. 14. Gel casting with consistently good quality is a challenging process. Due to uneven polymerization local or global distortions can occur within the gel matrix, which subsequently leads frequently to distortions within the protein spot pattern potentially hindering reasonable data analysis by image analysis software. The utilization of standardized precast gels allows circumventing those problems. 15. Use low melting, low electroendosmosis (EOO) agarose. 16. This solution contains SDS and is highly prone to boiling over. It should be microwaved at low power with constant monitoring. When the solution starts to boil over, turn off the power, remove from the microwave oven and swirl before returning to the microwave oven. Repeat until the agarose is fully dissolved. Keep solution at about 60  C for usage in second-dimension SDS-PAGE or store at 4  C. 17. EveryBlot Blocking Buffer provides 5 min blocking and maximum sensitivity for all western blots regardless of detection method. Alternative blocking reagents may be substituted. 18. This primary detection antibody is from goat, necessitating the use of an anti-goat secondary detection reagent. 19. This imaging instrument allows for both fluorescent and chemiluminescent imaging at the same resolution with the same camera. This simplifies alignment and matching of the total protein (fluorescent) gel image, total protein (fluorescent) blot image, and immunodetected (chemiluminescent) blot image. 20. Software developed for host cell protein coverage analysis. Tracks the percentage of the total protein that is being detected by the anti-HCP antibodies in 2-DE Western blot experiments. 21. Protease-inhibitor cocktails containing protein- and peptidebased inhibitors are not recommended. 22. Protein is most accurately quantitated in solution lacking detergent or reductant, so concentrated protein samples are first prepared without these additives. The detergent (CHAPS) and reductant (DTT) are supplied by sample solubilization buffer II in which the sample is diluted prior to first-dimension IEF.

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23. IEF is conducted in 8 M urea, the addition of thiourea (2 M) is usually not necessary. Secreted proteins are generally very soluble. Each of the sample solution components has a specific role in promoting high resolution IEF separation. Urea is a protein denaturant that promotes the complete unfolding of polypeptide chains so that all ionizable groups are exposed to the solution. CHAPS is a detergent that prevents protein aggregation and promotes solubility. DTT is a thiol reductant that breaks disulfide bonds within and between polypeptide chains, and maintains proteins in a fully reduced state. Carrier ampholytes enhance protein solubility by minimizing protein aggregation due to charge-charge interactions. Bromophenol blue is a tracking dye. Its inclusion is not necessary, but clearance of the dye from the IPG strip during electrophoresis provides a visual confirmation that electrical current is being delivered to the IPG strip appropriately. 24. Each sample is run in replicate and separated on two seconddimension gels, one of which is stained and the other blotted. 25. Recommended IPG-strip rehydration volumes depend on the length of the IPG-strips used. 26. Handle IPG strips from the ends using forceps. The sample is applied to the entire length of the immobilized pH gradient by rehydration into a dry IPG strip. As voltage is applied, each peptide in the sample focuses to a narrow zone in the pH gradient corresponding to its isoelectric point. 27. Recommended maximum storage time is 6 months at 80  C. Additional details of the sample application and IEF procedure may be found in the user manual for the PROTEAN i12 IEF System. 28. Although size standards are not strictly required for this application, they may be applied to the gel at this stage if desired. Apply 5–7 μL of standards solution into the standards well (already filled with agarose overlay solution, not yet solidified) of the SDS-PAGE gel using a syringe or gel loading pipet tip. 29. The 15 min low voltage step is to allow gradual transfer of the proteins from the IPG strip to the second-dimension gel. This reduces vertical streaking and gives a higher-resolution seconddimension separation. 30. The Oriole fluorescent stain used is resistant to photobleaching and shielding the staining reaction from normal room light should not be necessary. If the gel is under bright light, or in sunlight, the staining tray may be covered with aluminum foil. 31. LF (low fluorescence) PVDF membranes are protein blotting membranes optimized for fluorescent (e.g., Stain-free imaging after UV activation) and multiplex fluorescent applications.

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They offer high signal-to-background ratio, low autofluorescence, and superior protein retention to maximize blot detection sensitivity and enable downstream quantitation. 32. Wear gloves and avoid touching the gel anywhere besides the edges and corners. The gel is best held by the bottom corners. Place the gel on the anode stack and membrane by draping the gel over the membrane starting from the top of the gel. Once the gel is in place on the membrane, it should not be lifted again or moved. 33. The gel is stained following transfer to verify completeness of transfer. This step is optional. 34. At this stage, the membrane may be allowed to dry and can be stored for several weeks at room temperature between sheets of blotting paper. 35. Alternatively, this step may be carried out for 1–3 h at room temperature. 36. The substrate solution should form a puddle that covers the blot. Ensure that the surface of the blot is completely covered with substrate solution. 37. The default settings for capture of chemiluminescence images take advantage of binning in order to shorten exposure times at the expense of image resolution. In order to capture images at sufficient resolution for analysis of 2-DE gels, and to generate images at the same resolution as the fluorescent images generated for total protein, binning is turned off, or set to 1  1. References 1. Wurm FM (2004) Production of recombinant protein therapeutics in cultivated mammalian cells. Nat Biotechnol 22(11):1393–1398 2. Follman DK, Fahrner RL (2004) Factorial screening of antibody purification processes using three chromatography steps without protein A. J Chromatogr A 1024(1–2):79–85 3. Kornecki M, Mestm€acker F, Zobel-Roos S, Heikaus de Figueiredo L, Schlu¨ter H, Strube J (2017) Host cell proteins in biologics manufacturing: the good, the bad, and the ugly. Antibodies 6:45–62 4. Tscheliessnig AL, Konrath J, Bates R, Jungbauer A (2013) Host cell protein analysis in therapeutic protein bioprocessing - methods and applications. Biotechnol J 8(6):655–670 5. Posch A, Franz T, Hartwig S, Knebel B, Al-Hasani H, Passlack W, Kunz N, Hinze Y,

Li X, Kotzka J, Lehr S (2013) 2D-ToGo workflow: increasing feasibility and reproducibility of 2-dimensional gel electrophoresis. Arch Physiol Biochem 119(3):108–113 6. Gurtler A, Kunz N, Gomolka M, Hornhardt S, Friedl AA, McDonald K, Kohn JE, Posch A (2013) Stain-free technology as a normalization tool in Western blot analysis. Anal Biochem 433 (2):105–111 7. Berkelman T, Harbers A, Bandhakavi S (2015) 2-D Western blotting for evaluation of antibodies developed for detection of host cell protein. Methods Mol Biol 1295:393–414 8. Ambort D, Lottaz D, Sterchi E (2008) Sample preparation of culture medium from MadinDarby canine kidney cells. Methods Mol Biol 425:113–130

Chapter 33 Quantitative Proteomic Analysis Using Formalin-Fixed, Paraffin-Embedded (FFPE) Human Cardiac Tissue Omid Azimzadeh, Michael J. Atkinson, and Soile Tapio Abstract Clinical tissue archives represent an invaluable source of biological information. Formalin-fixed, paraffinembedded (FFPE) tissue can be used for retrospective investigation of biomarkers of diseases and prognosis. Recently, the number of studies using proteome profiling of samples from clinical archives has markedly increased. However, the application of conventional quantitative proteomics technologies remains a challenge mainly due to the harsh fixation process resulting in protein cross-linking and protein degradation. In the present chapter, we demonstrate a protocol for label-free proteomic analysis of FFPE tissue prepared from human cardiac autopsies. The data presented here highlight the applicability and suitability of FFPE heart tissue for understanding the molecular mechanism of cardiac injury using a proteomics approach. Key words Formalin-fixed, paraffin-embedded (FFPE), Heart, Label-free, Proteomics, Protein extraction, Cross-linking

1

Introduction Formalin-fixed, paraffin-embedded (FFPE) tissue has been considered for more than a decade as a potential source of material in clinical investigations due to its excellent capacity and suitability for long-term storage [1–7]. This is particularly valid in cancer research. Recently, the use of FFPE tissue for cardiovascular disease research has become interesting since the sampling of fresh-frozen heart material is complicated for ethical reasons and therefore inefficient. Proteomic analysis using FFPE material as an alternative to fresh-frozen tissue has achieved much attention since it was first presented [8–15]. However, the quantitative proteomic analysis of archival samples remains challenging. The long storage and harsh fixation procedure frequently prohibit successful protein identification and reproducible protein quantification [12, 13, 16, 17]. Several studies have described a variety of methods to extract and

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_33, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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separate proteins from FFPE samples by testing different extraction factors such as buffer components, detergents, pH, temperature, pressure, and others [3, 18–20]. Indeed, several protein-extraction protocols have been quite successful in analyzing tumor FFPE tissues [3, 21]. In contrast, only a few proteomic studies have been able to successfully examine human FFPE tissue in cardiac pathologies [22, 23]. In this chapter, we provide a methodological protocol for the label-free proteomic analysis of the FFPE tissue prepared form human cardiac autopsies exposed to external low-dose gamma-ray irradiation. The analysis highlights the applicability of FFPE cardiac tissue for investigation of heart injury using a proteomic approach. The data presented here are in good agreement with the results from comparable fresh-frozen tissues [24]. This methodology is applicable to any kind of tissue and therefore provides a tool for the validation in archival samples in the search for biomarkers of prognosis and disease.

2

Materials All solutions are prepared using HPLC grade water.

2.1 Sample Preparation and Protein Digestion

1. Protein LoBind tubes. 2. Tissue homogenizer. 3. Ultrafiltration spin columns (500 μL, 30 kDa cutoff filter). 4. Xylene. 5. Rehydration buffer: graded series of ethanol (100%, 95% and 70%). 6. Tris-buffered saline (TBS): 50 mM Tris–HCl, 150 mM NaCl, pH 7.6. 7. Extraction buffer: 8 M urea in 0.1 M Tris–HCl, pH 8. 8. 10% SDS in water. 9. Urea buffer (UA): 8 M urea in 0.1 M Tris–HCl, pH 8. 10. Heating block with agitation. 11. 50 mM ammonium bicarbonate (ABC) solution in water. 12. Reducing solution: 1 M DTT in 50 mM ABC. 13. Alkylating solution: 300 mM iodoacetamide in 50 mM ABC. 14. Proteases for protein digestion: 0.5 μg/μL Lys-C in 50 mM ABC; 0.5 μg/μL trypsin in 50 mM ABC. 15. Incubator set to 37  C. 16. 5% acetonitrile (ACN) in water. 17. 0.5% trifluoroacetic acid (TFA) in water. 18. Bradford protein assay.

Quantitative Proteomics of FFPE Human Heart

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1. Q-Exactive HF-X mass spectrometer online coupled to an UltiMate 3000 RSLCnano UHPLC System. 2. C18 trap column: 300 μm  5 mm; Acclaim PepMap100 C18, 5 μm, 100 A˚. 3. C18 reversed-phase analytical column: 75 μm  250 mm; nanoEase MZ HSS T3, 100 A˚, 1.8 μm. 4. Mobile Phase A: 3% (v/v) acetonitrile, 0.1% formic acid. 5. Mobile Phase B: 0.1% formic acid in acetonitrile. 6. Proteome Discoverer 2.3 software. 7. INGENUITY Pathway Analysis (IPA) software.

3

Methods

3.1 Protein Extraction from FFPE Tissue

1. Cut a section (20 μm) from FFPE tissue blocks after initial trimming to remove air-exposed surfaces (see Notes 1 and 2). 2. Place FFPE tissue sections on microscope slides and deparaffinize by incubating twice with xylene for 10 min at room temperature before rehydration in a graded series of ethanol (100%, 95%, and 70%) for 10 min each (see Note 3). 3. Scrape the tissue sections from the slides and transfer to a Protein LoBind tube (see Note 1). 4. Incubate the FFPE tissue samples in 100 μL TBS for 60 min at 99  C on the heating block for decrosslinking (see Notes 4 and 5). 5. Lyse the tissue using 150 μL of extraction buffer by homogenization in a tissue homogenizer. 6. Add to 250 μL lysate, 5 μL of 10% SDS to the final concentration of 0.2% before further incubation for 30 min at room temperature. 7. Clear lysate by centrifugation (14,000  g) for 30 min at 4  C. 8. Transfer supernatant to a new Protein LoBind tube. 9. Estimate the protein concentration in the lysate by the Bradford protein assay (see Notes 6 and 7).

3.2

Protein Digestion

The alterations in the protein expression are analyzed by mass spectrometry. Digest the protein lysate using Filter Aided Sample Preparation (FASP) protocol [15] (see Notes 4 and 5). 1. Dilute the protein lysate with 100 μL ABC. 2. Add 1 μL of reducing solution. 3. Incubate the reaction shaking for 30 min at 60  C.

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4. Let the solution cool down to room temperature for 1 min before adding alkylating buffer. 5. Incubate the solution with 5 μL of alkylating buffer for 30 min at room temperature in dark. 6. Equilibrate filters with 100 μL UA buffer and centrifuge (14,000  g) for 5 min at room temperature. 7. Transfer the sample to the cutoff filter. 8. Centrifuge (14,000  g) and discard flow-through. 9. Wash 3 times with 200 μL UA buffer and discard flow-through. 10. Wash 2 times with 200 μL 50 mM ABC and discard flowthrough. 11. Put the cutoff filter in a new Protein LoBind tube. 12. Add 40 μL ABC to the cutoff filter. 13. Add 0.5 μg Lys-C (enzyme–protein 1:20) (see Note 8). 14. Incubate for 2 h at room temperature. 15. Add 0.5 μg trypsin (enzyme–protein 1:20) (see Note 8). 16. Incubate overnight at 37  C (sealed with Parafilm). 17. Centrifuge peptides through the cutoff filter. 18. Transfer the pipette flow-through to a new Protein LoBind tube. 19. Add 20 μL ABC solution containing 5% ACN on the cutoff filter. 20. Centrifuge (16,000  g) for 15 min at room temperature. 21. Combine the with ACN eluted peptides with the first eluate. 22. Acidify combined eluate with 0.5% TFA (pH of ~2 needed) before injection to high-performance liquid chromatography (HPLC) coupled with the mass spectrometry (LC-MS/MS). 3.3 LC-MS/MS Analysis, Quantification, and Functional Analysis

Perform the LC-MS/MS analysis on a Q-Exactive HF-X mass spectrometer coupled to an UltiMate 3000 RSLCnano UHPLC System (see Notes 9 and 10). 1. The tryptic peptides are automatically loaded on a C18 trap column prior to C18 reversed-phase chromatography separation on the analytical column at a flow rate of 250 nl/min in a 95 min nonlinear ACN gradient from 3% to 40% in 0.1% formic acid. 2. Record profile precursor spectra from 300 to 1500 m/z at 60,000 resolution with automatic gain control (AGC) target value of of 3e6 and maximum injection time of 30 ms. Subsequently, record top 15 fragment spectra of charges 2 to 7 at 15,000 resolution with an AGC target of 1e5, the maximum injection time being 50 ms, the isolation window being 1.6 m/

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z, and the normalized collision energy 28 and the dynamic exclusion being 30 s. 3. For label-free proteomics, identify and quantify the acquired spectra using the Proteome Discoverer 2.3 software (see Note 9). 4. Load the individual raw-files to the Proteome discoverer 2.3 software allowing for peptide identification and label-free quantification using the Minora node. 5. Perform the searches using Sequest HT as search engine in the Swissprot human database with the following search settings: 10 ppm precursor tolerance, 0.02 Da fragment tolerance. 6. Allow two missed cleavages and carbamidomethyl on cysteine as a fixed modification, deamidation of glutamine and asparagine as variable modifications, as well as oxidation of methionine and Met-loss combined with acetylation at the N-terminus of the protein. 7. After filtering for a 5% peptide and protein false discovery rate (FDR < 0.05), quantify the proteins by summing up abundances of allocated unique and razor peptides. 8. Use the resulting normalized protein abundances for calculation of fold-change and ANOVA values of the individual proteins. 9. To interpret the observed alterations in the proteome, analyze the significantly differentially expressed proteins by the software tool INGENUITY Pathway Analysis. 3.4 Quantitative Proteomics Using FFPE Cardiac Tissue: An Example

The described methodology was used to analyze the FFPE tissue prepared form human cardiac autopsies of nuclear workers who were chronically exposed to external gamma rays using label-free quantitative proteomics. These samples were compared to the nonirradiated controls [25]. The proteomics data were further analyzed by bioinformatics. A total number of 2663 proteins were identified and quantified, of which 1855 proteins were quantified with at least two unique peptides across multiple samples. Both quantity and quality of protein identification and quantification were comparable to the proteomics data observed previously from fresh-frozen human cardiac tissue [24]. The functional interactions and biological pathways among significantly deregulated proteins in the irradiated group were further analyzed using IPA. Fatty acid metabolism, sirtuin signaling, protein ubiquitination, cardiac fibrosis, mitochondrial dysfunction and oxidative stress were the most affected pathways in the irradiated heart compared to the control group (Fig. 1a). The majority of significantly altered proteins were associated with heart pathologies including cardiac dilation, cardiac enlargement, cardiac dysfunction, heart failure, and cardiac fibrosis (Fig. 1b).

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Fig. 1 Pathway and disease analysis of significantly differentially expressed proteins in irradiated FFPE cardiac autopsy. Bars indicate top canonical pathways (a) and cardiac diseases (b) and the y-axis displays the – (log) significance. Tall bars are more significant than short bars. The analyses of signaling pathways and diseases were performed by the software tools INGENUITY Pathway Analysis (IPA)

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Notes 1. During all steps of the experiment always wear gloves to avoid keratin contamination. 2. The number and thickness of the slices depend on the sample type and can be optimized based on the tissue and amount of yield protein. 3. The xylene may harm the eyes and skin, so work under a chemical hood. 4. Use ultrapure proteomics grade reagents to prepare buffers for lysis and digestion. 5. The buffers and solutions must be freshly prepared within a day of the experiment. 6. It is important to measure the protein concentration before you start with the protein digestion. 7. If you use another protein quantification method, consider that some chemicals in the lysis buffer may be causing interference in the protein measurement. Before protein measurement, look at the compatibility of protein assay. 8. A final protease–protein ratio of 1:20 to 1:100 (w/w) is recommended. 9. LC-MS/MS setting and proteomics quantification need to be guided by trained researchers. 10. Analyze at least three replicates of one control sample by repetitive LC-MS/MS runs to evaluate the technical variability of mass spectrometry runs of label-free peptide quantifications of FFPE samples.

Acknowledgments This work was supported by grants from Federal Office for Radiation Protection (BfS) (# 3611S30022); from the European Community’s Seventh Framework Program (EURATOM), # 295823 PROCARDIO; and from Federal Medical Biological Agency (FMBA Russia). References 1. Bayer M, Angenendt L, Schliemann C, Hartmann W, Konig S (2020) Are formalinfixed and paraffin-embedded tissues fit for proteomic analysis? J Mass Spectrom 55:e4347. https://doi.org/10.1002/jms.4347 2. Zhu Y, Weiss T, Zhang Q, Sun R, Wang B, Yi X, Wu Z, Gao H, Cai X, Ruan G, Zhu T, Xu C, Lou S, Yu X, Gillet L, Blattmann P,

Saba K, Fankhauser CD, Schmid MB, Rutishauser D, Ljubicic J, Christiansen A, Fritz C, Rupp NJ, Poyet C, Rushing E, Weller M, Roth P, Haralambieva E, Hofer S, Chen C, Jochum W, Gao X, Teng X, Chen L, Zhong Q, Wild PJ, Aebersold R, Guo T (2019) High-throughput proteomic analysis of FFPE tissue samples facilitates tumor stratification.

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Mol Oncol 13(11):2305–2328. https://doi. org/10.1002/1878-0261.12570 3. Giusti L, Angeloni C, Lucacchini A (2019) Update on proteomic studies of formalinfixed paraffin-embedded tissues. Expert Rev Proteomics 16(6):513–520. https://doi.org/ 10.1080/14789450.2019.1615452 4. Tapio S, Hornhardt S, Gomolka M, Leszczynski D, Posch A, Thalhammer S, Atkinson MJ (2010) Use of proteomics in radiobiological research: current state of the art. Radiat Environ Biophys 49(1):1–4. https://doi.org/ 10.1007/s00411-009-0263-7 5. Tapio S, Atkinson MJ (2008) Molecular information obtained from radiobiological tissue archives: achievements of the past and visions of the future. Radiat Environ Biophys 47 (2):183–187 6. Tapio S, Schofield PN, Adelmann C, Atkinson MJ, Bard JL, Bijwaard H, Birschwilks M, Dubus P, Fiette L, Gerber G, Gruenberger M, Quintanilla-Martinez L, Rozell B, Saigusa S, Warren M, Watson CR, Grosche B (2008) Progress in updating the European radiobiology archives. Int J Radiat Biol 84 (11):930–936. https://doi.org/10.1080/ 09553000802460214 7. Gustafsson OJ, Arentz G, Hoffmann P (2015) Proteomic developments in the analysis of formalin-fixed tissue. Biochim Biophys Acta 1854(6):559–580. https://doi.org/10.1016/ j.bbapap.2014.10.003 8. Nirmalan NJ, Harnden P, Selby PJ, Banks RE (2009) Development and validation of a novel protein extraction methodology for quantitation of protein expression in formalin-fixed paraffin-embedded tissues using western blotting. J Pathol 217(4):497–506 9. Tanca A, Addis MF, Pagnozzi D, CossuRocca P, Tonelli R, Falchi G, Eccher A, Roggio T, Fanciulli G, Uzzau S (2011) Proteomic analysis of formalin-fixed, paraffinembedded lung neuroendocrine tumor samples from hospital archives. J Proteome 74 (3):359–370. https://doi.org/10.1016/j. jprot.2010.12.001 10. Azimzadeh O, Scherthan H, Yentrapalli R, Barjaktarovic Z, Ueffing M, Conrad M, Neff F, Calzada-Wack J, Aubele M, Buske C, Atkinson MJ, Hauck SM, Tapio S (2012) Label-free protein profiling of formalin-fixed paraffin-embedded (FFPE) heart tissue reveals immediate mitochondrial impairment after ionising radiation. J Proteome 75 (8):2384–2395. https://doi.org/10.1016/j. jprot.2012.02.019 11. Tanca A, Pagnozzi D, Burrai GP, Polinas M, Uzzau S, Antuofermo E, Addis MF (2012)

Comparability of differential proteomics data generated from paired archival fresh-frozen and formalin-fixed samples by GeLC-MS/MS and spectral counting. J Proteome 77:561–576. https://doi.org/10.1016/j. jprot.2012.09.033 12. Giusti L, Lucacchini A (2013) Proteomic studies of formalin-fixed paraffin-embedded tissues. Expert Rev Proteomics 10(2):165–177. https://doi.org/10.1586/epr.13.3 13. Azimzadeh O, Barjaktarovic Z, Aubele M, Calzada-Wack J, Sarioglu H, Atkinson MJ, Tapio S (2010) Formalin-fixed paraffinembedded (FFPE) proteome analysis using gel-free and gel-based proteomics. J Proteome Res 9(9):4710–4720. https://doi.org/10. 1021/pr1004168 14. Wisniewski JR, Dus K, Mann M (2013) Proteomic workflow for analysis of archival formalin-fixed and paraffin-embedded clinical samples to a depth of 10 000 proteins. Proteomics Clin Appl 7(3–4):225–233. https://doi. org/10.1002/prca.201200046 15. Wisniewski JR, Zougman A, Nagaraj N, Mann M (2009) Universal sample preparation method for proteome analysis. Nat Methods 6 (5):359–362. https://doi.org/10.1038/ nmeth.1322 16. Metz B, Kersten GF, Hoogerhout P, Brugghe HF, Timmermans HA, de Jong A, Meiring H, ten Hove J, Hennink WE, Crommelin DJ, Jiskoot W (2004) Identification of formaldehydeinduced modifications in proteins: reactions with model peptides. J Biol Chem 279 (8):6235–6243 17. Steiner C, Ducret A, Tille JC, Thomas M, McKee TA, Rubbia-Brandt L, Scherl A, Lescuyer P, Cutler P (2014) Applications of mass spectrometry for quantitative protein analysis in formalin-fixed paraffin-embedded tissues. Proteomics 14(4–5):441–451. https://doi.org/10.1002/pmic.201300311 18. Azimzadeh O, Atkinson MJ, Tapio S (2015) Qualitative and quantitative proteomic analysis of formalin-fixed paraffin-embedded (FFPE) tissue. Methods Mol Biol (Clifton, NJ) 1295:109–115. https://doi.org/10.1007/ 978-1-4939-2550-6_10 19. Foll MC, Fahrner M, Oria VO, Kuhs M, Biniossek ML, Werner M, Bronsert P, Schilling O (2018) Reproducible proteomics sample preparation for single FFPE tissue slices using acid-labile surfactant and direct trypsinization. Clin Proteomics 15:11. https://doi.org/10. 1186/s12014-018-9188-y 20. Quesada-Calvo F, Bertrand V, Longuespee R, Delga A, Mazzucchelli G, Smargiasso N, Baiwir D, Delvenne P, Malaise M, De Pauw-

Quantitative Proteomics of FFPE Human Heart Gillet MC, De Pauw E, Louis E, Meuwis MA (2015) Comparison of two FFPE preparation methods using label-free shotgun proteomics: application to tissues of diverticulitis patients. J Proteome 112:250–261. https://doi.org/10. 1016/j.jprot.2014.08.013 21. Gaffney EF, Riegman PH, Grizzle WE, Watson PH (2018) Factors that drive the increasing use of FFPE tissue in basic and translational cancer research. Biotech Histochem 93(5):373–386. https://doi.org/10.1080/10520295.2018. 1446101 22. Kakimoto Y, Ito S, Abiru H, Kotani H, Ozeki M, Tamaki K, Tsuruyama T (2013) Sorbin and SH3 domain-containing protein 2 is released from infarcted heart in the very early phase: proteomic analysis of cardiac tissues from patients. J Am Heart Assoc 2(6): e000565. https://doi.org/10.1161/jaha. 113.000565 23. Holub D, Flodrova P, Pika T, Flodr P, Hajduch M, Dzubak P (2019) Mass spectrometry amyloid typing is reproducible across

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multiple organ sites. Biomed Res Int 2019:3689091. https://doi.org/10.1155/ 2019/3689091 24. Azimzadeh O, Azizova T, Merl-Pham J, Subramanian V, Bakshi MV, Moseeva M, Zubkova O, Hauck SM, Anastasov N, Atkinson MJ, Tapio S (2017) A dose-dependent perturbation in cardiac energy metabolism is linked to radiation-induced ischemic heart disease in Mayak nuclear workers. Oncotarget 8 (6):9067–9078. https://doi.org/10.18632/ oncotarget.10424 25. Azimzadeh O, Azizova T, Merl-Pham J, Blutke A, Moseeva M, Zubkova O, Anastasov N, Feuchtinger A, Hauck SM, Atkinson MJ, Tapio S (2020) Chronic Occupational Exposure to Ionizing Radiation Induces Alterations in the Structure and Metabolism of the Heart: A Proteomic Analysis of Human FormalinFixed Paraffin-Embedded (FFPE) Cardiac Tissue. Int J Mol Sci 2020 21(18):E6832. https://doi.org/10.3390/ijms21186832. PMID: 32957660

Chapter 34 Chloroplast Isolation and Enrichment of Low-Abundance Proteins by Affinity Chromatography for Identification in Complex Proteomes Roman G. Bayer, Simon Stael, and Markus Teige Abstract Comprehensive knowledge of the proteome is a crucial prerequisite to understand dynamic changes in biological systems. Particularly low-abundance proteins are of high relevance in these processes as these are often proteins involved in signal transduction and acclimation responses. Although technological advances resulted in a tremendous increase in protein identification sensitivity by mass spectrometry (MS), the dynamic range in protein abundance is still the most limiting problem for the detection of low-abundance proteins in complex proteomes. These proteins will typically escape detection in shotgun MS experiments due to the presence of high-abundance proteins. Therefore, specific enrichment strategies are still required to overcome this technical limitation of MS-based protein discovery. We have searched for novel signal transduction proteins, more specifically kinases and calcium-binding proteins, and here we describe different approaches for enrichment of these low-abundance proteins from isolated chloroplasts from pea and Arabidopsis for subsequent proteomic analysis by MS. These approaches could be extended to include other signal transduction proteins and target different organelles. Key words Chloroplast isolation, Affinity chromatography, Mass spectrometry, Proteomics, Organelle proteome, ATP-binding protein, Calcium-binding protein

1

Introduction Organelle proteomics has become a strong focus in biology over the last years due to several aspects: Organelles perform key activities for cellular metabolism and provide the basis for growth and production of valuable compounds [1]. Moreover, the use of isolated organelles as starting material does already significantly reduce the complexity of the sample as compared to total cell extracts. However, in the case of plant leaves, the difference between isolated chloroplasts and total leaves becomes rather small in terms of their overall protein composition, thus calling for additional enrichment strategies for the identification of low-abundant proteins such as signaling components. With the

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_34, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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notion that organelles are increasingly being seen as important signal transduction hubs in the cell, for example in retrograde signaling, oxidative stress signaling, hormone signaling and defense response [2–4], this latter aspect becomes more and more important. The study of chloroplasts, as a central organelle to these pathways and specific to plants, has strong implications for agricultural research, and consequently, chloroplast proteomics from a number of different plant species has gained a lot of attention during the last decade [5–8]. However, despite all efforts, the knowledge of regulatory components in chloroplasts such as protein kinases is still very limited [9], underpinning the need for application of specifically targeted enrichment strategies [10]. This is even more important as it turned out that particularly for those components the application of prediction algorithms, which are based on the genome sequence, is strongly biased [10]. Traditionally, highly pure and intact chloroplasts used for functional studies and protein import experiments, were obtained from Spinacia oleracea or Pisum sativum. Since their genomes have not been sequenced, chloroplast isolation protocols were later on also adapted for Arabidopsis thaliana. Our chloroplast isolation protocol from Arabidopsis is based on the method from Kunst [11], which yields reasonable amounts of intact chloroplasts with stromal content sufficient for downstream purification steps and/or analyses. To circumvent the labor-intensive isolation of Arabidopsis chloroplasts for mass spectrometry–based proteomic approaches, the isolation of chloroplasts from pea provides a useful alternative, since the release of an expressed sequence tag (EST) database, which could be used for protein identification [12] and recently the pea genome was published [13]. Therefore we describe both methods here. The chloroplast proteome can further be divided into several subcompartments: (a) the soluble part, the chloroplast stroma; (b) the thylakoid membrane, and (c) the chloroplast envelope membrane. As for the latter two recently proteomics protocols have been published [14, 15] and were recently refined [8], we focus here exclusively on the stromal proteins, which become easily accessible after osmotic disruption of isolated chloroplasts. A general flow-scheme of our experimental strategy is displayed in Fig. 1. After extraction of stromal proteins we perform size exclusion chromatography to remove the highly abundant RuBisCO protein complex (~540 kDa) which generally presents the biggest problem for MS-based identification of low-abundant proteins. In addition, also ribosomes are removed during this step. After size exclusion chromatography, all fractions of interest can be pooled and either subjected to mass spectrometry or to additional downstream purification approaches (e.g., affinity chromatography) in order to further reduce the sample complexity.

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a

b

c [kDa] 95 72 55

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9 10 11 12 13 14 15 16

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Fig. 1 Scheme of the workflow and typical profile of a size exclusion chromatography (SEC) of stroma extracts from isolated chloroplasts on a Superdex S200 gel filtration column. (a) Scheme of the work-flow to obtain stroma extract. (b) Scheme of a typical elution profile of stromal proteins on a Superdex S-200 size exclusion column. X-axis shows mL of eluting sample. Y-axis shows OD280 indicating protein content. Fractions eluting after the large peak of Ribulose 1,5 Bisphosphate Carboxylase/Oxygenase (RuBisCO) are pooled and subjected to: MS, mass spectrometry; IEX, ion exchange chromatography; AC, affinity chromatography; HIC, hydrophobic interaction chromatography. (c) SDS PAGE of chromatography fractions (1–16) collected from 73 mL to 96 mL of a Superdex S200 gel filtration column. T ¼ total stromal protein extract

2

Materials

2.1 Chloroplast Isolation from Pea (Pisum sativum)

1. Leaves from Pea (see Note 1): Pea seedlings are grown for 8–9 days under long day conditions (photoperiod of 8 h dark:16 h light at ~70 μmol m
2 s
1; 21  C  5; humidity 70–90%) on vermiculite.

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2. P-ISO buffer: 330 mM sorbitol, 20 mM MOPS, 13 mM Trisbase (see Note 2), 0.1% BSA, 3 mM MgCl2 (store at 4  C < 1 week). 3. P-WASH buffer: 330 mM sorbitol, 50 mM HEPES-KOH, pH 7.6, 3 mM MgCl2 (store at
20  C). 4. 80% Percoll solution: 330 mM sorbitol, 50 mM HEPES-KOH, pH 7.6, 80% v/v Percoll (store at
20  C). 5. 40% Percoll solution: 330 mM sorbitol, 50 mM HEPES-KOH, pH 7.6, 40% v/v Percoll (store at
20  C). 6. Percoll step gradient: 7 mL of 80% Percoll solution are placed carefully below 12 mL 40% Percoll with a 10 mL pipette in a round-bottom tube. 7. Miracloth. 8. Waring blender (see Note 3). 9. Ultracentrifuge with swing-out rotor. 2.2 Chloroplast Isolation from Arabidopsis

1. Leaves from Arabidopsis (see Note 1): Arabidopsis plants are grown for ~6–8 weeks under short day conditions (photoperiod of 16 h dark:8 h light at 100–150 μmol m
2 s
1; 22  C  5; humidity 60%  20%) on soil. 2. Homogenization buffer (HB buffer): 450 mM sorbitol, 20 mM Tricine–KOH, pH 8.4, 10 mM EDTA, 5 mM NaHCO3, 0.1% BSA, 10 mM isoascorbate, 1 mM reduced glutathione (see Note 4). 3. Resuspension buffer (RB buffer): 300 mM sorbitol, 20 mM Tricine–KOH, pH 7.6, 5 mM MgCl2, 2.5 mM EDTA (store at 4  C). 4. Percoll continuous gradient: Per gradient mix 15 mL Percoll with 15 mL 2 RB buffer. Store at 4  C for less than 1 week in centrifugation tubes. Centrifuge for 30 min at ~53,000  g in a swing-out rotor (brake “slow”) to form a continuous gradient. 5. Miracloth. 6. Waring blender (see Note 3). 7. Ultracentrifuge with swing-out rotor.

2.3 Chloroplast Stroma Extraction

1. Osmotic lysis buffer: 10 mM Tricine–KOH, pH 8.0, 10 mM MgCl2, 1 mM DTT and protease inhibitor cocktail (Complete Mini EDTA–free). 2. THY buffer: 25 mM Tricine–KOH, pH 8.0, 10 mM MgCl2, 10 mM isoascorbate, 2 mM β-mercaptoethanol (store at
20  C).

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1. Gel filtration buffer: 50 mM Tris–HCl, pH 7.8, 50 mM NaCl, 10 mM MgCl2. Filter buffer through a 0.22 μm membrane and degas. 2. PD-10 desalting columns. 3. Centriprep Centrifugal Filter Units (molecular weight cutoff: 10 kDa). 4. Superdex 200 (S200) gel filtration column. 5. Superdex 75 (S75) gel filtration column.

2.5 Ion Exchange Chromatography

1. MonoQ (MQ) column for anion exchange chromatography. 2. MonoS (MS) column for cation exchange chromatography. 3. MQ-A buffer: 20 mM Tris–HCl, pH 8.0. Filter the buffer through a 0.22 μm membrane and degas. 4. MQ-B buffer: 20 mM Tris–HCl, pH 8.0, 1 M NaCl. Filter the buffer through a 0.22 μm membrane and degas. 5. MS-A buffer: 20 mM MES–NaOH, pH 6.0. Filter the buffer through a 0.22 μm membrane and degas. 6. MS-B buffer: 20 mM MES–NaOH, pH 6.0, 1 M NaCl. Filter the buffer through a 0.22 μm membrane and degas. 7. Centrifugal Filter Units (molecular weight cutoff: 10 kDa).

2.6 Hydrophobic Interaction Chromatography (HIC)

1. Phenyl-Superose (PS) column. 2. PS-A buffer: 50 mM sodium phosphate buffer, pH 7.0. Filter the buffer through a 0.22 μm membrane and degas. 3. PS-B buffer: 50 mM sodium phosphate buffer, pH 7.0, 1.5 M (NH4)2SO4. Filter the buffer through a 0.22 μm membrane and degas. 4. PD-10 desalting columns. 5. Centrifugal Filter Units (molecular weight cutoff: 10 kDa).

2.7 ATP/Purvalanol B Affinity Chromatography

1. C10-linked Aminophenyl-ATP-Sepharose. 2. Purvalanol B (PurB) affinity Sepharose [16]. 3. Column preparation: pour 500 μL of affinity Sepharose slurry into disposable polystyrene columns and always run by gravity flow at room temperature. 4. ATP buffer: gel filtration buffer (see item 1 in Subheading 2.4) + 100 mM NaCl, 0.05% NP-40. 5. PurB buffer: gel filtration buffer (see item 1 in Subheading 2.4) + 350 mM NaCl, 0.5% Triton X-100. 6. 0.5% w/v SDS in ultrapure water.

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2.8 Heat Treatment of Isolated Chloroplasts and Protein Extraction

1. Chloroplast lysis buffer: 20 mM DTT, 0.1% Triton X-100, protease inhibitor cocktail (Complete Mini EDTA-free).

2.9 Eu3+-IDA Column Affinity Chromatography

1. Equilibration buffer: 100 mM Tris–HCl, pH 7.5, 2 M NaCl, 200 mM CaCl2.

2. Thermomixer. 3. Ultracentrifuge with swing-out rotor.

2. IDA column-loading buffer: 100 mM Tris–HCl, pH 7.5, 3 M NaCl, 200 mM CaCl2. 3. Sulfate buffer: 600 mM Na2SO4, 100 mM Tris–HCl, pH 7.5, 2 M NaCl. 4. Malonate buffer: 40 mM malonate, 600 mM Na2SO4, 100 mM Tris–HCl, pH 7.5, 2 M NaCl. 5. Citrate buffer: 0.2 M phosphate buffer, pH 7.5, 3 M NaCl, 200 mM citrate. 6. IDA- Sepharose. 7. Column packing: fill 1 mL of IDA Sepharose in a disposable polystyrene column and wash with 5 mL of 100 mM EDTA solution, pH 7.0, followed by 10 mL of double distilled water. Load the column with 5 CV of 50 mM EuCl3 solution, wash with 25 mL double-distilled water, and equilibrate with 10 mL of equilibration buffer. 8. PD-10 desalting columns. 9. 100% acetone, stored at 4  C.

3

Methods (See Note 5)

3.1 Chloroplast Isolation from Pea (See Note 6)

The chloroplast isolation protocol for pea is adapted from the method published by Schleiff [17]. 1. Cut about 100–120 g leaves from the shoots and homogenize in about 150 mL P-ISO buffer in a Waring blender using three pulses: low–high–low for 3 s each (see Notes 3 and 7). 2. Filter the homogenate through four layers of Miracloth into four 50 mL round-bottom tubes and centrifuge for 2 min at 2800  g (brake “on”). 3. Gently resuspend the pellets by pipetting in 1 mL P-WASH buffer (see Note 8), load this on top of two Percoll step gradients and centrifuge for 5 min at 8000  g (brake “off”) using a swing-out rotor (see Note 9). 4. Remove broken chloroplasts on top of the 40% Percoll layer with a vacuum pump together with excess 40% Percoll layer but

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leave some of the 40% Percoll on the 40–80% interphase, not to expose intact chloroplasts to air, while recovering them. 5. Recover intact chloroplasts after centrifugation from the 40–80% interphase and transfer them into two 50 mL tubes. 6. Wash the isolated chloroplasts with ~30 mL P-WASH buffer and precipitate by centrifugation for 2 min at 2800  g (brake “on”). For use of the chloroplasts in protein import experiments, the washing step should be repeated. 7. Finally, resuspend the pellets in ~500 μL P-WASH buffer, pool and either use directly for further analysis (e.g., chlorophyll content estimation, see Subheading 3.3) or freeze in liquid nitrogen for storage at
80  C (see Note 10). 3.2 Chloroplast Isolation from Arabidopsis (See Note 6)

The experimental procedure for the isolation of chloroplasts from Arabidopsis was adapted from the protocol of Kunst [11]. Prepare Percoll gradients prior to the harvesting of leaves! 1. Cut ~90 g of leaves and homogenize them in 500 mL of homogenization buffer (HB buffer) using a Waring blender and three pulses: low–low–high, 2–3 s each (see Notes 3 and 7). 2. Filter the homogenate through four layers of Miracloth into a suited centrifuge tube and centrifuge for 5 min at 1519  g (brake “on”). 3. Gently resuspend the pellet in 20 mL RB buffer using a fine paintbrush and distribute the chloroplast suspension to the four preformed Percoll gradients (see Notes 8 and 9), 1 mL on top of each, using a pipette. 4. Centrifuge for 6 min at ~10,700  g (brake “slow”), recover intact chloroplasts (lower green band of each gradient) and transfer them into two 50 mL tubes (see Note 8). Take care not to touch the pellet of the tube containing starch granules. 5. Wash the chloroplasts with 1 RB buffer by centrifugation for 3 min at 1700  g (brake “on”) and discard the supernatant. Resuspend each pellet in ~300 μL 1 RB buffer, pool the isolated chloroplasts, and either use directly for further analysis (e.g., chlorophyll content estimation, see Subheading 3.3) or immediately freeze them in liquid nitrogen and store at
80  C (see Note 10).

3.3 Estimation of the Chlorophyll Content [18]

1. Mix 5 μL isolated chloroplast suspension with 5 mL 80% acetone and centrifuge for 2 min at 3000  g. 2. Measure OD645 and OD663 of the supernatant and determine chlorophyll concentration using the following formula (see Note 11): (OD645  20.2 + OD663  8.02)  1000 ¼ μg/mL chlorophyll in the sample. If the yield is to low see Notes 12 and 13.

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3.4 Chloroplast Stroma Extraction

1. Incubate chloroplasts (amount equivalent to ~20 mg of chlorophyll) in ~1.5 volumes of hypotonic osmotic lysis buffer for 5 min on ice to achieve osmolytic bursting of the envelopes. 2. Centrifuge for 6 min at 12,000  g at 4  C and transfer the supernatant containing the stromal content to a 50 mL tube. 3. Resuspend the chloroplast pellet in ~5 mL osmotic lysis buffer and repeat the extraction step once. 4. Pool the stromal protein extracts and keep on ice until further treatment. 5. Resuspend the thylakoid pellet in ~5 mL THY buffer and store at
80  C for further analysis.

3.5

Gel Filtration

1. Prior to size exclusion chromatography (SEC), exchange the buffer of the stromal protein extracts to gel filtration buffer with PD-10 desalting columns. 2. Concentrate the protein solution to a volume of ~500 μL using Centrifugal Filter Units. 3. Clear the extract by centrifugation for 10 min at 16,100  g and 4  C, and apply the supernatant to an S200 gel filtration column. 4. Perform S200 size exclusion chromatography at a flow rate of 0.8 mL/min using the gel filtration buffer. 5. Store the eluate fractions at 4  C for further analysis (see Note 14). 6. Pool the fractions of interest from S200 chromatography and concentrate them to a volume of ~500 μL using a Centrifugal Filter Unit. 7. Centrifuge the sample for 3 min at 16,100  g and 4  C. 8. Apply the supernatant to a S75 gel filtration column and perform size exclusion chromatography at a flow rate of 0.4 mL/ min using gel filtration buffer. 9. Perform protein content measurements and/or functional assays of the individual chromatography fractions as desired. 10. Store protein samples at 4  C until further analysis.

3.6 Ion Exchange Chromatography

1. Concentrate protein samples obtained from size exclusion chromatography to a volume of ~500 μL using a Centrifugal Filter Unit. 2. Centrifuge for 10 min at 16,100  g at 4  C and apply the supernatant to a MonoS or MonoQ column. 3. Perform ion exchange chromatography at a flow rate of 2 mL/ min.

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4. Elute proteins by applying a linear gradient over five column volumes (CV) of buffer B from 0% to 100% (MQ-B for MonoQ, and MS-B for MonoS column, respectively). 5. Collect the individual eluate fractions and conduct protein content measurements and/or functional assays as desired. 6. Store protein samples at 4  C until further analysis. 3.7 Hydrophobic Interaction Chromatography (HIC)

1. Buffer exchange the protein extract of interest to PS-B buffer using PD-10 Desalting columns. 2. Concentrate the sample to a volume of ~500 μL using a Centrifugal Filter Unit. 3. Centrifuge sample for 10 min at 16,100  g at 4  C, and apply the supernatant to a Phenyl-Superose column. 4. Perform hydrophobic interaction chromatography at a flow rate of 2 mL/min. 5. Elute proteins by applying a gradient of PS-B and PS-A buffer. 6. Collect the individual eluate fractions and conduct protein content measurements and/or functional assays as desired. 7. Store protein samples at 4  C until further analysis.

3.8 ATP/Purvalanol B Affinity Chromatography

1. Equilibrate ATP or PurB chromatography column with 10 column volumes (CV) of ATP buffer or PurB buffer, respectively. 2. Pool the protein extracts of interest (e.g., eluates from size exclusion or ion exchange chromatography; amount equivalent up to ~1.5 mg protein). 3. Adjust the ionic content of the sample solution to ATP buffer or PurB buffer by dilution with an appropriate stock solution and apply the sample to the column. 4. Wash the column with 20 CV of ATP buffer or PurB buffer and elute bound proteins with 6 CV of 0.5% SDS. 5. Collect the individual eluate fractions and store at 4  C for further analysis (see Note 15).

3.9 Heat Treatment of Chloroplasts and Protein Extraction

Some small proteins like calmodulins are particularly resistant to heat treatment, which allows a rapid and easy enrichment from complex protein mixtures, because larger proteins will typically denature upon heating and precipitate after centrifugation. 1. Lyse 3 mL isolated chloroplasts (containing ~4 mg/mL chlorophyll) by adding 7 mL of chloroplast lysis buffer and incubate for 10 min on ice. 2. Divide the chloroplast suspension into 1 mL aliquots and heat rapidly to 75  C for 5 min on a heat block, and immediately cool on ice.

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3. Pellet the heat-denatured proteins and thylakoid membranes by centrifugation at 20,000  g for 10 min. 4. Clear the supernatant further by centrifugation for 30 min at 100,000  g at 4  C. 3.10 Eu3+-IDA Column Affinity Chromatography

Ca2+ binding proteins can be enriched by affinity chromatography on immobilized Eu3+ ions using an iminodiacetic acid (IDA) column. This method has previously been described by Chaga et al. [19], and was adapted to chloroplast proteins. Protein obtained from chloroplast stroma SEC or from heat-treated isolated chloroplasts was loaded on the column. 1. For heat-treated samples, rebuffer the sample to IDA columnloading buffer, using PD-10 desalting columns. For SEC fractionated stroma extracts, add salts to the extracts to match the composition of the IDA column-loading buffer. 2. After sample loading, wash the column with 10 mL equilibration buffer, 5 mL of sulfate buffer, and 2.5 mL of malonate buffer. 3. Elute proteins with citrate buffer. Finally, strip the column with 100 mM EDTA. 4. Buffer exchange the citrate and EDTA eluates to 50 mM Tris– HCl, pH 7.5 with a PD-10 column. 5. Perform protein precipitation by addition of four volumes of ice-cold acetone (see Note 15).

4

Notes 1. Starch granules in the chloroplast will contribute to breaking of chloroplasts during handling. Therefore, prepare chloroplasts in the morning, when starch levels in the chloroplast are lowest. 2. No pH adjustment necessary. The desired pH is reached by mixing the two buffer components (MOPS and Tris base). 3. To gain better yields of intact chloroplasts and to allow for a shorter and gentler homogenization the blades of the Waring blender should be as sharp as possible (can usually be detached for sharpening). 4. Add isoascorbate and glutathione always freshly prior to use and readjust pH to 8.4 with KOH. 5. Isolation of chloroplasts requires a certain amount of practice in order to achieve reproducible results and optimal yield in terms of intact organelles. However, based on empirical experience there is a natural variation and sometimes the yield of intact chloroplasts is much lower than expected, although the

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plants have been grown under the same conditions and the same experimental procedures have been followed. 6. Carry out all steps at 4  C (in a cooling chamber or on ice). 7. If the homogenization appears to be a problem, younger plant material should be used. 8. When pipetting chloroplast suspensions always use CUT TIPS in order to minimize mechanical damage to the organelles. 9. One “typical” error that occurs frequently is that no sorbitol is added in the Percoll solutions. This will result to osmotic burst of the chloroplasts. 10. For easy storage and handling, chloroplast suspensions are pipetted drop by drop directly in liquid nitrogen and afterward collected in Eppendorf tubes. In this way only part of a tube can be used, while the rest is kept frozen at
80  C. 11. Expected chloroplast yields at the volumes mentioned in these protocols are in the range of 1–2 mg chlorophyll for Arabidopsis and 7–10 mg for pea. 12. The stromal content of Arabidopsis chloroplasts is frequently lost during the isolation procedure [20]. This becomes visible in a comparison of the total protein content of isolated chloroplasts from different species by SDS-PAGE as for example shown in Fig. 2 for pea and Arabidopsis and could present a serious problem for proteomic approaches. Therefore, it is crucial to check the amount of stromal proteins by SDS-PAGE before further usage when using Arabidopsis leaves as source for chloroplast isolation. 13. To spot at which step problems might occur in cases where the yield of intact chloroplasts is to low, just check a drop of the chloroplast suspension from the different steps by light microscopy (intact chloroplast can easily be recognized by application of phase contrast). 14. The protein content of the eluting fractions can be determined for example by photometry (e.g., measuring OD280), protein assay or by SDS-PAGE after protein precipitation (using a standard protocol). Functional assays such as in vitro protein kinase assays can also be easily performed on the protein fractions. 15. Perform protein precipitation and analyze protein content by SDS-PAGE. Regions of interest on the protein gel (e.g., after Coomassie or silver staining) can subsequently be excised and subjected to MS analysis.

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Fig. 2 Comparison of the protein content of isolated chloroplasts from different species. Ps, Pisum sativum. At, Arabidopsis thaliana. Stroma and thylakoid proteins extracted from isolated chloroplasts were analyzed by SDS-PAGE. The relative amount of the large subunit of RuBisCO (52 kDa) can be used as a measure of chloroplast intactness

Acknowledgments Work in the authors’ lab is supported by grants from the Austrian Science Fund (FWF) to MT (P 28491-B29) and the FP7 Marie Curie Initial Training Network (ITN) “CALIPSO” (GA ITN 2013-607 607) from the European Union. References 1. Rolland N, Curien G, Finazzi G, Kuntz M, Marechal E, Matringe M et al (2012) The biosynthetic capacities of the plastids and integration between cytoplasmic and chloroplast processes. Ann Rev Genet 46:233–264 2. Jarvis P, Lopez-Juez E (2013) Biogenesis and homeostasis of chloroplasts and other plastids. Nat Rev Mol Cell Biol 14:787–802 3. Kmiecik P, Leonardelli M, Teige M (2016) Novel connections in plant organellar signalling link different stress responses and signalling pathways. J Exp Bot 67:3793–3807 4. Stael S, Kmiecik P, Willems P, Van Der Kelen K, Coll NS, Teige M et al (2015) Plant innate immunity--sunny side up? Trends Plant Sci 20:3–11 5. Zybailov B, Rutschow H, Friso G, Rudella A, Emanuelsson O, Sun Q et al (2008) Sorting signals, N-terminal modifications and

abundance of the chloroplast proteome. PLoS One 3:e1994 6. Ferro M, Brugiere S, Salvi D, SeigneurinBerny D, Court M, Moyet L et al (2010) AT_CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins. Mol Cell Proteomics 9:1063–1084 7. Huang M, Friso G, Nishimura K, Qu X, Olinares PD, Majeran W et al (2013) Construction of plastid reference proteomes for maize and Arabidopsis and evaluation of their orthologous relationships; the concept of orthoproteomics. J Proteome Res 12:491–504 8. Bouchnak I, Brugiere S, Moyet L, Le Gall S, Salvi D, Kuntz M et al (2019) Unraveling hidden components of the chloroplast envelope proteome: opportunities and limits of

Chloroplast Isolation and Affinity Chromatography better MS sensitivity. Mol Cell Proteomics 18:1285–1306 9. Bayer RG, Stael S, Rocha AG, Mair A, Vothknecht UC, Teige M (2012) Chloroplastlocalized protein kinases: a step forward towards a complete inventory. J Exp Bot 63:1713–1723 10. Bayer RG, Stael S, Csaszar E, Teige M (2011) Mining the soluble chloroplast proteome by affinity chromatography. Proteomics 11:1287–1299 11. Kunst L (1998) Preparation of physiologically active chloroplasts from Arabidopsis. Methods Mol Biol 82:43–48 12. Brautigam A, Shrestha RP, Whitten D, Wilkerson CG, Carr KM, Froehlich JE et al (2008) Low-coverage massively parallel pyrosequencing of cDNAs enables proteomics in non-model species: comparison of a speciesspecific database generated by pyrosequencing with databases from related species for proteome analysis of pea chloroplast envelopes. J Biotechnol 136:44–53 13. Kreplak J, Madoui MA, Capal P, Novak P, Labadie K, Aubert G et al (2019) A reference genome for pea provides insight into legume genome evolution. Nat Genet 51:1411–1422 14. Simm S, Papasotiriou DG, Ibrahim M, Leisegang MS, Muller B, Schorge T et al (2013)

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Defining the core proteome of the chloroplast envelope membranes. Front Plant Sci 4:11 15. Tomizioli M, Lazar C, Brugiere S, Burger T, Salvi D, Gatto L et al (2014) Deciphering thylakoid sub-compartments using a mass spectrometry-based approach. Mol Cell Proteomics 13:2147–2167 16. Wissing J, Jansch L, Nimtz M, Dieterich G, Hornberger R, Keri G et al (2007) Proteomics analysis of protein kinases by target classselective prefractionation and tandem mass spectrometry. Mol Cell Proteomics 6:537–547 17. Schleiff E, Soll J, Kuchler M, Kuhlbrandt W, Harrer R (2003) Characterization of the translocon of the outer envelope of chloroplasts. J Cell Biol 160:541–551 18. Arnon DI (1949) Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta Vulgaris. Plant Physiol 24:1–15 19. Chaga GS, Ersson B, Porath JO (1996) Isolation of calcium-binding proteins on selective adsorbents. Application to purification of bovine calmodulin. J Chromatogr A 732:261–269 20. Halliwell B (1978) The chloroplast at work. A review of modern developments in our understanding of chloroplast metabolism. Prog Biophys Mol Biol 33:1–54

Chapter 35 Principles of Protein Labeling Techniques Christian Obermaier, Anja Griebel, and Reiner Westermeier Abstract Protein labeling methods prior to separation and analysis have become indispensable approaches for proteomic profiling. Basically, three different types of tags are employed: stable isotopes, mass tags, and fluorophores. While proteins labeled with stable isotopes and mass tags are measured and differentiated by mass spectrometry, fluorescent labels are detected with fluorescence imagers. The major purposes for protein labeling are monitoring of biological processes, reliable quantification of compounds and specific detection of protein modifications and isoforms in multiplexed samples, enhancement of detection sensitivity, and simplification of detection workflows. Proteins can be labeled during cell growth by incorporation of amino acids containing different isotopes, or in biological fluids, cells or tissue samples by attaching specific groups to the ε–amino group of lysine, the N-terminus, or the cysteine residues. The principles and the modifications of the different labeling approaches on the protein level are described; benefits and shortcomings of the methods are discussed. Key words Protein labeling, Quantification, Protein detection, Stable isotopes, Mass tags, Fluorophores, Difference gel electrophoresis, Mass spectrometry, Fluorescence imager

1

Introduction Proteomics profiling employs various methods for identification, monitoring of structural changes, and quantification of expression levels of proteins in complex mixtures. The most commonly applied techniques are the separation by gel electrophoresis and chromatography, and further investigation by image analysis, Western blotting, and mass spectrometry. Lately, a comprehensive overview on these tools has been published for the example of environmental microbiology [1]. The different proteomic profiling workflows possess various methodological shortcomings, which could partly be prevented or at least reduced by prelabeling proteins prior to the analysis. Furthermore, labeling of specific protein moieties offers the possibility of studying some dedicated protein modifications. In the following we try to give a small overview on methodical issues,

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_35, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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where labeling of proteins can improve the quality of the results and enable methodical developments: The conventional approach for protein detection in gel electrophoresis is staining the proteins in the gels subsequently to the separation. A number of poststaining methods exist using organic and fluorescent dyes, and silver staining [2]. While the latter shows the highest sensitivity of detection, it has a very limited capacity for quantification. Labeling of proteins prior to the separation with an appropriate and bright fluorescent dye could increase sensitivity and quantification features at the same time. Furthermore this would enable direct detection after separation, and save time and work. Proteomics samples are highly complex: biological fluids, cell lysates, and tissue extracts contain thousands of different proteins and peptides with very high structural diversities. In the original profiling approach this challenge is met as follows: With two-dimensional gel electrophoresis the protein mixture is “individualized” into several hundred to a few thousand fraction spots, of which each contains one or a few different proteins. After punching out the spots of interest and digesting the proteins with trypsin inside the gel plug, the resulting peptides are eluted and subjected to mass spectrometry analysis. It is still an undisputed fact, that two-dimensional electrophoresis affords the highest resolution of all protein separation methods. The technology using immobilized pH gradients for isoelectric focusing in the first and high-resolution SDS–polyacrylamide gel electrophoresis in the second dimension has meanwhile been further developed to a robust method with a high level of reproducibility [3]. However, there exist still little experimental variations, which cause partial protein losses [4]. The incorporation of an internal control into the samples during separation would equalize such effects and improve the quantification data. This can, however, only be realized by labeling the proteins of different samples with different fluorescent tags and working according to a multiplexing concept. Redox proteomics has gained great importance: Oxido-redox protein modifications, especially of cysteines, link redox metabolism to biological structure and function [5]. One of the strategies would be labeling cysteine thiol residues of differently oxidized samples with different fluorescent dyes to compare and determine the oxido-redox status of biological material by using 2D electrophoresis. In the complementary gel-free workflow the complete initial protein mixture is digested with endopeptidases. This means that thousands of protein fractions are converted into an even more complex mixture: each protein is cleaved into about 30 peptides on an average. In most cases the workflow is performed in an automated mode; reversed-phase liquid chromatography of the peptides online connected to electrospray ionization mass

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spectrometry. Due to this enormous heterogeneity of the peptide mixture the contents of the chromatography peaks are very complex, resulting in undersampling in the mass spectrometer: During the chromatography separation the mass spectrometer has only a limited time window to produce the related peptide mass spectra, which are therefore incomplete. In practice, repeated runs of the same sample show varying peak spectra; reproducibility is compromised. It is therefore very complicated to find differences in protein expression between different biological samples: Several technical replicates would be needed to cope with adequate statistical requirements. Furthermore, the gel-free quantification procedures based on mass spectrometry peaks of peptides suffer from the limitation that the peak intensities of the peptides do not correlate with their abundances [1]. However, because the mass spectrometry peak intensities of identical peptides correlate with their relative abundances in different samples [6], labeling of the sample proteins with different stable isotopes or mass tags would facilitate the qualitative and quantitative assessment of relative protein abundances in a multiplexing approach. These quantification issues can also be solved by labeling the peptides after protein digestion using nonisobaric isotope coded amino-reactive groups or isobaric mass tags, which allow quantification during a tandem MS step. Thus protein or peptide labeling is indispensable for protein quantification. The following review describes the principles of different labeling techniques which are applied on intact proteins, not on the digestion products, the peptides.

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Materials Generally, the use of high-quality electrophoresis/proteomic-grade chemicals is paramount to achieving successful experiments. Fluorescent dyes for prelabeling are available from a number of specialist suppliers and some fine chemicals companies. Also the stable isotope labeling kits and the mass tag kits are marketed by several fine chemical and mass spectrometry companies.

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3.1 Labeling with Fluorescent Dyes Prior to Electrophoretic Separations

The easiest accessible targets for covalently attaching a fluorescent group to a protein are the ε-amino groups of the lysines and the N-terminus. According to the UniProt database lysine represents 6% of amino acids in the average proteins, which—statistically— means that lysines are present in almost every protein. Coupling of a reagent via the N-hydroxy-succinimide (NHS) ester to primary amines is a robust and well established procedure. Furthermore, it

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is not necessary to derivatize the proteins prior to the reaction. In order to ensure accessibility to the ε-amino groups of the lysines, ideally the protein samples are denatured with high molar chaotropes like urea or a combination of urea and thiourea. Labeling of proteins prior to electrophoresis with a fluorogenic amino reactive label has several advantages over poststaining: no washing and staining procedures are needed, the image can be recorded without fixing of the proteins, thus the gels can be further processed with Western blotting. In this way the results are obtained much faster; the method is more cost and environment friendly and easier, no buffers and organic solvents for staining and washing are required. € ¨ et al. [7] have applied the concept of prelabeling the Unlu proteins in their original method “difference gel electrophoresis” (DIGE), which has become one of the most successful methods in comparative proteomics. In the DIGE technique, cyanine-NHS CyDyes™ with different excitation and emission wavelengths are covalently bound to the protein lysine moieties for multiplex analysis with two-dimensional electrophoresis. After the different protein samples have been prelabeled with Cy2, Cy3, or Cy5 they are combined together, and the mixture is submitted to isoelectric focusing followed by SDS–polyacrylamide gel electrophoresis. The three dyes have been selected to have sufficiently different fluorescent properties to prevent “cross-talking” during imaging. For perfect comigration of the same proteins to the identical gel positions the dyes are charge-matched and have similar molecular weights. For labeling the lysine moieties with cyanine dyes “minimal labeling” is applied: Only a small portion, about 5%, of the proteins in the sample are labeled by limiting the amount of dye per protein, which is about 400 pM dye per 50 μg of protein. This prevents the proteins from becoming too hydrophobic, and insures that only singly labeled proteins will be detected. In this minimal labeling concept, the amount of multiple labeled proteins is statistically below the detection level. The resulting spot pattern is the same like the pattern obtained from nonlabeled proteins. The limit of detection of a single protein is about 25 pg. This means that the sensitivity is similar to silver staining. Downstream analyses with mass spectrometry or Western blotting are not affected, because 95% of the separated proteins are unlabeled, hence not modified. Imaging is performed by exciting the fluorophore at the optimum wavelength with laser, LED, or white light with excitation filter and collecting the light with higher wavelength emitted from the fluorophore using an emission filter with narrow band pass. The patterns created from the different samples can be displayed on the computer screen with false color representation. By superimposing these images with an image editing program a preview of the result is obtained, which shows upregulation and downregulation of protein expression levels and serves as a control for sample

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preparation and labeling, as well as appropriate comigration of the proteins. For accurate qualitative and quantitative evaluation dedicated 2D software has to be employed. Advanced software packages perform the image analyses almost automatically, for instance by image warping from different gels. The principle of the DIGE method is displayed in Fig. 1a, b (from ref. 8). Because up to three samples are run in the identical matrix, gel-to gel-variations between the sample patterns are excluded, and the individual spots are codetected across the samples for the calculation of protein abundance ratios. However, the most powerful feature of DIGE is the possibility of the incorporation of an internal control. Therefore, a pooled internal standard is created by mixing aliquots taken from each sample of the experiment. This sample pool is labeled with one of the fluorophores, mostly Cy2, and coelectrophoresed with each sample. Thus every protein in the population appears on each gel. Inherent gel-to-gel variations between gels are compensated. Gel-to-gel matching becomes much easier than in conventional gels, because the patterns of the internal standard need to be matched, which are resulting from the identical sample mixture. For quantitative comparison, the spot volume ratios of the samples are calculated related to the internal standard spot volume. This approach makes it possible to compare protein abundances across different gels with high accuracy. Appropriate sample cleanup and the adherence to exact labeling conditions are very important prerequisites to achieve optimum results (see Note 1). The principle of the DIGE method is displayed in Fig. 1a, b. Meanwhile a quadruplex kit with an additional dye molecule for the detection close to infrared light is available, which allows for comparison of up to four different samples within one 2D gel. Alternatively, the fluorescent dyes can be attached to the thiol group of the cysteine via a maleimide reactive group. Cysteines are less abundant than lysine, which allows saturation labeling without the risk of creating protein modifications with too high hydrophobicity. This offers a very high sensitivity of detection. Cysteine labeling dyes are charge-neutral, and they have also similar molecular weights. Because all available cysteines will receive a label, the proteins containing many cysteines will become multiply labeled: migrate markedly slower in the SDS gel than unlabeled proteins, and their light emission will be disproportionally high. Proteins without any cysteine cannot be detected. Thus the spot patterns are different from those achieved with unlabeled or lysine-labeled proteins. The efforts are higher than in lysine labeling: Prior to labeling the disulfide bridges of the proteins must be cleaved with a reductant like tris(carboxyethyl)phosphine (TCEP) or dithiothreitol (DTT) (see Note 2). For cysteine labeling only two dyes are available: Cy3 and Cy5. The internal standard is usually labeled

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Fig. 1 (a) Schematic drawing of the DIGE 2-D electrophoresis method principle employing protein labeling with fluorophores. The sample proteins are prelabeled with Cy3 and Cy5. An internal standard is created by applying a mixture of all samples labeled with the third dye Cy2. (b) False color representation of the images of samples and an internal standard, which have been coseparated in the same gel by DIGE. The images have

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with Cy3, the sample with Cy5. But the detection sensitivity of this method is very high, which makes it particularly useful for scarce samples or detection of low-abundance proteins [9]. Furthermore, cysteine labeling offers a new opportunity for the specific detection of redox-modified proteins. The procedure has lately been optimized [10]: Free protein thiols in sample A can be directly labeled with Cy3 maleimide. However, since some protein thiols may have become oxidized and will no more react with the Cy3 maleimide, the selective labeling of the oxidized subproteome (sample B), in particular its reversible modified thiols, which can become reduced with a reductant such as DTT, might be also of great interest. Thus, in order to maintain the overall spot pattern the specific labeling of sample A requires the exclusion of any nonreacted labeling reagent included in sample B and vice versa. Therefore, subsequently to the labeling of sample A, excessive label is removed via gel filtration, which excludes the labeling of components of sample B, the none labeled protein thiols, and their irreversible alkylation with the Cys-interacting compound (CinC). In contrast, for sample B, free protein thiols are first irreversibly alkylated with the Cys-interacting compound (CinC), the excessive alkylating reagent removed via gel filtration, the eluent (eluted/ recovered sample) treated with a reductant like DTT, and subsequently labeled with the Cy3 maleimide. The individual samples (A or B) are combined with the internal pooled standard (saturated labeled sample) and submitted to a 2D electrophoresis. When the false-color images, where Cy 3 (sample A) is represented in green and Cy3 (sample B) in red, are overlaid, green spots indicate redox-modified proteins, yellow spots show proteins containing occluded thiols. By using CinC instead of the formerly employed N-ethylmaleimide the pattern matching is improved, because the molecular sizes of the blocking reagent is more similar to the sizes of the dye maleimides. Irreversibly oxidized proteins can be visualized by poststaining of the gel. These differential multiplexing methods have initiated great acceptance of protein labeling with fluorophores prior to the separation [11]. Their features improve the statistical basis for relative quantitative measurements of protein expression levels considerably. These advantages have justified the relatively high investments for dyes and fluorescent imagers. ä Fig. 1 (continued) been acquired with a Typhoon™ multifluorescent laser scanner (GE Healthcare LifeSciences). (A) Cy2 channel: pooled standard composed of the two samples; (B) Cy3 channel: mouse liver proteins, control; (C) Cy5 channel: mouse liver proteins, treated; (D) overlay of all three images; the arrows point to a protein spot which is upregulated in the treated sample; (E) three-dimensional view of this spot in the Cy3 channel (left) and Cy5 channel (right). first dimension: Isoelectric focusing in IPG pH 3–10, 18 cm; second dimension: SDS–polyacrylamide gel electrophoresis in a 12.5% T gel. From ref. 8

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The vast majority of gels are still poststained, because of the high price level of the labeling reagents and the investments for fluorescent imagers. Lately new types of fluorescent dye labels for proteins have been introduced [12], which are less expensive and can also be applied in presence of carrier ampholytes and reductants (see Note 3). However, it should be noted that gel-based separation techniques have several limitations: very large and very small, very hydrophobic proteins—like membrane proteins—and those with very high isoelectric points are not included, LC/MS and CE/MS possess superior sensitivity of detection features. 3.2 Protein Labeling with Stable Isotopes

Nonradioactive isotope labeling is a very useful approach for comparing samples by mass spectrometry analysis, because the mass differences introduced into the sample components by different isotope compositions can be easily detected, while the chemical properties of the molecules are not altered. Thus, all the commonly used physicochemical sample preparation techniques can be applied for reduction of the complexity of the sample. Such an approach was first time utilized for proteins by Gysi et al. [13], introducing isotope-coded affinity tags (ICAT): the protein cysteine residues are chemically labeled with a reagent comprised of a thiol-reactive group (iodoacetamide), biotin, and a linker containing either eight hydrogen or eight deuterium atoms. Typically the control sample is labeled with the light and the test sample with the heavy tag and combined in a 1:1 ratio. The mixture is digested with an endopeptidase, the—biotin-containing— labeled peptides are purified by affinity chromatography on a streptavidin column, and submitted to reversed-phase HPLC coupled with mass spectrometry. Peptide pairs originating from differential samples are identified by a mass difference of 8 Da, their abundances are compared for relative quantification. Protein identification is performed by MS/MS analysis of the respective peptides. In order to overcome isotopic shift effects caused by the deuterium interaction with reversed phase chromatography media, and improve MS/MS spectra, the tags were later modified by using 13 C labels [14] and a cleavable biotin moiety [15]. Figure 2 shows a schematic representation of the ICAT workflow. A great advantage of ICAT is the very efficient reduction of sample complexity, because it isolates the few cysteine containing peptides. This makes it very useful for automated LC/MS systems. However, because not all proteins and peptides contain cysteines, the ICAT method does unfortunately not result in global labeling; this leads to reduced sequence coverage in mass spectrometry analysis and missing values. An isotopic label can also be introduced metabolically by salts and amino acids containing different compositions of deuterium, 13 C or 15N during cell growth. The mostly applied in vivo labeling

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Biotin „heavy“ linker (2H or 13C) IAA

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sample 2

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Fig. 2 Schematic drawing of the principle of ICAT. The different samples are reduced with a thiol reagent and labeled at the free SH groups of their cysteines with the ICAT reagents (light and heavy) via the iodoacetamide group (IAA). After tryptic digestion of the sample proteins the labeled (biotin-containing) peptides are purified with a Streptavidin affinity column and submitted to LC/MS for relative quantification and protein identification

method is SILAC (stable isotope labeling with amino acids in cell culture) which has been introduced by Ong et al. [16]. Whereas the growth medium for one cell population contains normal amino acids, the second cell population is fed with a growth medium containing amino acids with heavier isotopes. A number of SILAC variations exist, differing by the choice of amino acids and isotope combinations. After a couple of cell divisions, the labeled amino acids are completely incorporated into the cellular proteins. The samples are mixed and can be directly digested with a protease, or previously separated by electrophoresis or chromatography for

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sample 1 grown with 12

sample 2 grown with

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Fig. 3 Schematic drawing of the principle of SILAC. The cells are grown under different conditions in media containing different isotope compositions of hydrogen, carbon, and nitrogen. The samples are then mixed together. The duplex mixture can be fractionated by any protein separation method. After tryptic digestion of the protein fractions, the peptides are submitted to LC/MS for relative quantification using the defined mass shifts of the related peptide peaks, and protein identification

reduction of complexity. The peptide mixture is then analyzed by LC-MS, relative protein amounts are quantified by comparison of the abundance of respective peptide doublets. The proteins are identified by MS/MS analysis of the peptides. The SILAC principle is shown in Fig. 3. By adding the labeled amino acids to the growth medium for only a short period of time, also differences in de novo protein turnover can be studied. This method is called Pulsed SILAC [17]. SILAC generally allows global labeling on the protein level,

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which is a great advantage of the method. SILAC can only be used for labeling growing cells. Meanwhile SILAC has already been applied to labeling multicellular model organisms, such as mouse, Caenorhabditis elegans, Drosophila melanogaster, and Arabidopsis thaliana. In order to overcome limitations of the approaches described above, Schmidt et al. have developed Isotope-Coded Protein Labeling (ICPL), which is based on the well-established reaction of N-nicotinoyloxy-succinimide (Nic-NHS) with the ε-amino group of lysine residues and the available N-termini of intact proteins [18]. The ICPL technology is available with up to four labels, which contain different combinations of hydrogen /deuterium and 12 C and 13C atoms, and create mass shifts of 0, 4, 6, and 10 Da. In order to facilitate access to all free ε-amino groups of the lysines, the samples are denatured, reduced, and alkylated prior to the labeling reaction. The tagged samples are mixed in equal amounts and separated by electrophoretic or chromatographic techniques. Because all lysines are blocked by the ICPL labels, protein cleavage is ideally performed by double digest with trypsin and endoproteinase GluC. This measure ensures improved sequence coverage. The use of Nic-NHS as a label enhances the ionization of the labeled peptides, and hence the sensitivity of mass spectrometry, which allows also the detection of low-abundance proteins. Figure 4 shows a schematic drawing of the ICPL principle. The method is compatible with both MALDI and electrospray ionization mass spectrometry, for quantification with ICPL MS/MS is not required. The evaluation software selects only the peptide peak sets with the defined mass shifts of 0, 4, 6, and 10 showing differing expression levels. The ratios of peak intensities of the peptide pairs, triplets or quadruplets are used to determine the related protein abundances originating from the different samples. Analyzing quadruplets instead of triplets or doublets considerably reduces the number of false-positive multiplets, because randomly occurring sets with the four defined mass differences are considerably less frequent than sets with only two or three defined mass shifts (see also Note 4). ICPL has the advantage, that it yields high sequence coverage, also when the complexity of the labeled samples has been reduced with high-resolution separation techniques. This feature is very important for adequate detection of posttranslational modifications and protein isoforms. It offers multiplex analysis of up to four samples, and it can be applied on any species, cell lysates, or tissue types.

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+10

+6

NICNHS

NICNHS

NICNHS

NICNHS

sample 1

sample 2

sample 3

sample 4

multiplex sample

protein separation

digestion with trypsin/Glu-C liquid chromatography peptide separation

mass spectrometry quantification on the MS level m/z

Fig. 4 Schematic drawing of the principle of ICPL. The protein samples are labeled with the ICPL reagents containing different isotope compositions of hydrogen and carbon at the ε-amino groups of the lysines. The samples are then mixed together. The multiplex mixture can be fractionated by any protein separation method. After tryptic digestion of the protein fractions, the peptides are submitted to LC/MS for relative quantification using the defined mass shifts of the related peptide peaks, and protein identification

4

Notes 1. For optimum results, a sample cleanup based on protein precipitation prior to labeling is recommended. It is essential for labeling that the sample solution have a pH value between 8 and 9. In order to avoid any interference with the labeling procedure, the sample solution must not contain any primary amines, like carrier ampholytes, and reductant, like dithiothreitol. These reagents are added after labeling has been completed.

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2. Saturation labeling is performed for 1 h at a pH value of 8.0 and a temperature of 37  C. Like with minimal labeling, the sample must not contain any carrier ampholytes and reductant. Also here these compounds are added prior to application on the first dimension. 3. SERVA HPE™ Lightning Red is compatible with all additives typically used for sample solubilization and protein extraction, including carrier ampholytes and reductants like dithiothreitol (DTT) and dithioerythritol (DTE). For efficient labeling the pH value of the sample must be 8 or higher. 4. The ICPL software cannot identify incomplete quadruplets, which sometimes occur due to the complete absence of a protein in a sample as a result of a complete downregulation. It is therefore advised to prepare an internal standard, which is composed of equal amounts of all samples to be analyzed. This mixed sample is divided into four aliquots, which receive the four labels before they are combined again. The internal standard is treated in the same way like the samples. This will create equally abundant quadruplets, which are used in the software database for the detection and quantification of incomplete quadruplets. References 1. Wo¨hlbrand L, Trautwein K, Rabus R (2013) Proteomic tools for environmental microbiology—a roadmap from sample preparation to protein identification and quantification. Proteomics 13:2700–2730 2. Miller I, Crawford J, Gianazza E (2006) Protein stains for proteomic applications: which, when, why? Proteomics 6:5385–5408 3. Moche M, Albrecht D, Maaß S et al (2013) The new horizon in 2D electrophoresis–new technology to increase resolution and sensitivity. Electrophoresis 34:1510–1518 4. Zhou S, Bailey MJ, Dunn MJ et al (2005) A quantitative investigation into the losses of proteins at different stages of a two-dimensional gel electrophoresis procedure. Proteomics 5:2739–2747 5. Go Y-M, Jones DP (2013) The redox proteome. J Biol Chem 288:26512–26520 6. Silva JC, Denny R, Dorschel CA et al (2005) Quantitative proteomic analysis by accurate mass retention time pairs. Anal Chem 77:2187–2200 € ¨ M, Morgan ME, Minden JS (1997) Dif7. Unlu ference gel electrophoresis: a single gel method for detecting changes in protein extracts. Electrophoresis 18:2071–2077

8. Westermeier R, Scheibe B (2008) Difference gel electrophoresis based on Lys/Cys tagging. In: Posch A (ed) Sample preparation and fractionation for 2-D PAGE/proteomics. Methods in molecular biology, vol 424. Humana, New York, pp 73–85 9. Berendt FJ, Fro¨hlich T, Bolbrinker P et al (2009) Highly sensitive saturation labeling reveals changes in abundance of cell cycleassociated proteins and redox enzyme variants during oocyte maturation in vitro. Proteomics 9:550–564 10. Lennicke C, Rahn J, Heimer N et al (2016) Redox proteomics: methods for the identification and enrichment of redox-modified proteins and their applications. Proteomics 16:197–213 11. Cramer R, Westermeier R (eds) (2012) Difference gel electrophoresis (DIGE): methods and protocols, vol 854. Springer GmbH, Heidelberg 12. Griebel A, Obermaier C, Westermeier R et al (2013) Simplification and improvement of protein detection in two-dimensional electrophoresis gels with SERVA HPE™ LightningRed. Arch Physiol Biochem 119:94–99

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13. Gygi SP, Rist B, Gerber SA et al (1999) Quantitative analysis of complex protein mixtures using isotope-coded affinity tags. Nat Biotechnol 17:994–999 14. Hansen KC, Schmitt-Ulms G, Chalkley RJ et al (2003) Mass spectrometric analysis of protein mixtures at low levels using cleavable 13C-isotope-coded affinity tag and multidimensional chromatography. Mol Cell Proteomics 2:299–314 15. Li J, Steen H, Gygi SP (2003) Protein profiling with cleavable isotope-coded affinity tag (cICAT) reagents: the yeast salinity stress response. Mol Cell Proteomics 2:1198–1204

16. Ong SE, Blagoev B, Kratchmarova I et al (2002) Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol Cell Proteomics 1:376–386 17. Schwanh€ausser B, Gossen M, Dittmar G et al (2009) Global analysis of cellular protein translation by pulsed SILAC. Proteomics 9:205–209 18. Schmidt A, Kellermann J, Lottspeich F (2005) A novel strategy for quantitative proteomics using isotope-coded protein labels. Proteomics 5:4–15

Chapter 36 Mechanical/Physical Methods of Cell Disruption and Tissue Homogenization Stanley Goldberg Abstract This chapter covers the various methods of mechanical cell disruption and tissue homogenization that are currently commercially available for processing small samples s < 1 mL) to larger multikilogram production quantities. These mechanical methods of lysing do not introduce chemicals or enzymes to the system. However, the energies required when using these “harsh,” high mechanical energy methods can be enough to damage the very components being sought. The destruction of cell membranes and walls is effected by subjecting the cells (a) to shearing by liquid flow, (b) to exploding by pressure differences between inside and outside of cell, (c) to collision forces by impact of beads or paddles, or (d) a combination of these forces. Practical suggestions to optimize each method, where to acquire such equipment, and links to reference sources are included. Several novel technologies are presented. Key words Cell disruption, Bead mills, Bead beater, BioNeb cell disruption, Cell disruption vessel, Dounce tissue grinder, DYNO-MILL, Electro Water Separation, French Press G-M, Gaulin high pressure homogenizer, Heartbreaker, High pressure homogenizer, MEGATRON , Microfluidics, Microwave assisted centrifugation, Mixer-Mill, Mortar, Pestle, Parr nitrogen vessel, Opposed jet homogenization, Parr nitrogen vessel, POLYTRON, Potter-Elvehjem tissue grinders, Pressure vessel, Pulsed electrical field, Rotor–stator homogenizer, Sonicator, SONITUBE , Tissue grinders, Tissue homogenization, Ultrasonic processor

1

Introduction Mechanical methods of processing cells or tissues to release intracellular components avoids introduction of encumbering chemicals or enzymes. The destruction of cell membranes and walls by these “harsh” methods is effected by subjecting the cells (a) to shearing by liquid flow, (b) to exploding by pressure differences between inside and outside of cell, (c) to collision forces by impact of beads or paddles, or (d) a combination of these forces. Generally speaking, any of the techniques described here can, to some degree, disrupt any cell or tissue. For more difficult materials, just increasing mechanical force or increasing the time of

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9_36, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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exposure will improve breakage. However, in some case, use of excessive force is to be limited due to the generation of detrimental heat and/or shear that can ruin the desired proteins or intracellular organelles. In addition, excess force will accelerate wear and ultimately damage the equipment being used. By judicious use of the equipment one can select from a gentle nicking of the cell to release intact organelles up to a vigorous action to release membrane bound proteins. Some methods are suitable to handle tissue only, others for free cells only, and some are suitable for both. Some techniques are capable of processing only small quantities of material while others are limited to handling larger amounts. Tissues that are difficult to break down include the tough outside membrane surrounding heart muscle, lung, intestine, and skin tissues. On the other hand, some fragile mammalian cells can be broken by just a moderate shaking of the suspended cells. Free cells that are difficult to process include those that are extremely small size (below 0.25 μm) bacteria, and the tough yeasts and spores. Plant materials and seeds will need higher energy inputs for proper maceration. Table 1 provides an overview of the mechanical methods with at least one example of commercial equipment and related websites, while Table 2 describes suitable subjects and capacity for each method.

2 2.1

Bead Impact Methods: Shaking Vessel Theory

All bead milling devices disrupt the cells or homogenize tissues by the crushing action of thrown beads (also called “grinding media”) against the cells/tissue. Also the accelerated beads generate localized shear in the liquid buffer surrounding the cells/tissues, which helps to pull them apart. Two methods to accelerate the grinding media (beads) are (a) by shaking the entire container or (b) by a spinning agitator within a container (see next section). The shaking container method is commonly used for tissues as well as free cells. For extremely small samples of ~0.2 mL (these may be placed in microvials or microplates) to somewhat larger quantities of 50 mL (in sealed containers or tubes) the shaking of the vessel is the method of choice. The motion can be of differing geometries depending upon what equipment is selected. Shaking can only be done in batch operation, thus limiting the amount of materials that can be processed. The equipment is low cost, durable, and simple to operate, requiring minimal training. When viewed under a microscope, cells lysed by this method often appear to still be intact cells. However, when stained with trypan blue it is see that the cells are empty “ghosts,” and the contents have been liberated.

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Table 1 Methods overview, trade names, websites Technique

Trade name(s)

Websites

Bead impact—shaking vessel [1, 2]

MIXER MILL Mini-BeadBeater™ Bullet Blender® Heartbreaker

www.RETSCH.de www.BIOSPEC.com www.NEXTADVANCE.com www.GLENMILLS.com

Bead impact—agitator shaft [3]

DYNO®-MILL

www.WAB-GROUP.com www.GLENMILLS.com

Rotor–stator—shear by spinning shaft [2, 4]

POLYTRON® TissueTearor

www.KINEMATICA.ch www.BIOSPEC.com

Mortar–pestle—shear by mechanical pressure [5]

Potter-Elvehjem tissue grinders

www.WHEATON.com

High pressure batch—liquid expansion [6] French Press G-M®

www.GLENMILLS.com

High pressure batch—gas expansion [7]

PARR® Vessel

www.PARRINST.com

High pressure flow—high velocity liquid shear [8]

High Pressure Gaulin and Rannie

www.SPXFLOW.com

High pressure—opposed liquid streams [9]

MICROFLUIDIZER®

www.MICROFLUIDICSMPT.com

Droplet—low pressure flow droplet nebulizing [10]

BioNeb®

www.GLASCOL.com

Ultrasonic—shear by collapsing bubbles [11]

SONCIATOR® SONITUBE®

www.SONICATOR.com www.SYNETUDE.com

Electromotive force [12]

Electro Water Separation

www.ORIGINOIL.com

Pulsed electric field [13, 14]

n/a

n/a

Microwave-assisted centrifugation [15, 16]

n/a

www.IFTS-SLS.com www.GLENMILLS.com

Materials needed to operate any of these shaking bead mills include the cells/tissue, grinding media (beads), liquid phase such as buffer, the container, and the equipment. Variables include the bead selection (density, diameter, and quantity), speed of agitation, cell concentration, and duration of run (Figs. 1 and 2). 2.2

Practical Aspects

l

Denaturing of proteins due to high temperature or excessive shear is to be considered in all mechanical disruption/homogenization equipment. Also, debris from the wear of the grinding media and container into the samples need to be evaluated.

l

Hints for successful temperature control include (1) prechilling of samples and containers; (2) repeated runs of short duration with rest time to allow for rechilling samples on ice; (3) use of fewer beads and/or extra buffer to act a heat sink; (4) reduced degree of shaking vigor.

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Table 2 Suitable subjects and capacity for each method Type of biomaterial to be lysed !

Yeast, algae, fungi, Bacteria spores

Seeds Plants Tissues Capacity

Bead impact—shaking vessel

Y

Y

Y

Y

Y

S/M

Bead impact—agitator shaft

Y

Y

?

Y

Y

M/L

Rotor–stator—shear by spinning shaft

N

N

Y

Y

Y

S/M/L

Mortar–pestle—shear by mechanical pressure

Y

Y

Y

Y

Y

S/M

High pressure batch—liquid expansion

Y

Y

?

?

?

S/M

High pressure batch—gas expansion

Y

N

N

Y

Y

S/M

High pressure flow—high velocity Y shear

Y

N

N

N

S/M/L

High pressure flow—opposed liquid streams

Y

Y

N

N

N

S/M/L

Droplet—low pressure droplet nebulizing

Y

Y

N

N

N

S/M

Ultrasonic—shear collapsing bubbles

Y

Y

N

Y

Y

S/M/L

Electro water separation

N

?

N

N

N

M/L

Pulsed electrical field

Y

Y

N

N

Y

S/M

Microwave—centrifugal

N

N

?

Y

Y

S/M

Technique #

Suitability

Quantities

Y—general good practice

S ¼ small 0.1–25 mL

N—not recommended

M ¼ medium 10–500 mL

?—not known or marginal success

L ¼ 250 mL to many liters

l

Detrimental contamination of the batch due to wear of either the grinding media or container walls is usually rare due to the insignificant amounts involved. A hint to mitigate any contamination problem is to use inert materials of construction such as using beads and containers made of zirconium oxide stabilized with yttria (95%/5%; specific gravity 6.0), glass, and plastics.

l

Beads size: For small diameter cells (e.g., bacteria) use beads of 0.10–0.5 mm diameter. For larger cells (e.g., yeast, algae, hyphae) use beads of 0.5–1.25 mm. Glass (sp. gr. 2.5) is a

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Fig. 1 Bead impact—shaken vessel—Mini-BeadBeater-1 equipment

Fig. 2 Bead impact—shaken vessel—inside view—beads moving in jar

good starting material due to low cost. For homogenizing plant or animal tissues beads of 1.0–5.0 mm diameter are used, and the samples may also need to be prechopped with a razor. l

Bead density: If addition energy is needed to improve breakage/ homogenization of tough cells, then use higher density materials. These material types include ceramic (zirconium oxide family) with specific gravities from 3.8 to 6.0, stainless steel of sp. gr. 7.0+, and tungsten carbide of sp. gr. 14.2+. Also, use of larger diameter beads from 2 to 20 mm can improve breakage. Recently, success has been reported with garnet and SiC grit with their sharp edges, and with stainless-steel ballcones that have a wedged edge at the equator.

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Time savings can be achieved by ganging several samples into 96-well or larger microtiter plates rather than running one sample at a time. The larger equipment models can accommodate these plates.

l

A simple way to evaluate the suitability of bead shaking can be done as follows. In a test tube place some glass beads and buffered cells/tissues. Hold the tube against a vortex shaker for up to 3 min. If breakage/homogenization is realized, then bead shaking has promise. Switching to suitable equipment as described above will reduce strain to the user holding the test tubes.

Bead Impact Methods: Stirred Agitated Beads

3.1

Theory

For modest sample quantities of 50 mL and scaling up to industrial amounts of several thousand liters, the agitation of the beads by a turning agitator within the vessel is the method of choice. In this class of agitated bead mills, the beads and the cell suspension are loaded into a chamber. Into the mix is placed one or more spinning discs that accelerate the beads. The beads striking the cells combine with the shearing by the moving liquid phase will disrupt the cells. These units are normally used for disruption of free microorganism cells (bacteria, yeasts, hyphae, mycelia) and not for tissue samples. For lesser quantities, see previous section on shaking container method. Materials needed to operate the agitated bead mills include the cells/tissue, grinding media (beads), liquid phase such as buffer, cooling ice or jacket fluids, and the mill equipment. Variables include the bead selection (density, diameter, and quantity), speed of agitation, cell concentration, and duration of run (Figs. 3 and 4).

3.2

Practical Aspects

l

Denaturing of proteins due to high temperature or excessive shear is to be considered in all mechanical disruption/homogenization equipment. Also, the wear of the grinding media and container into the samples need to be evaluated.

l

Successful temperature control hints include (1) prechilling of samples to below 5 degrees Celsius (5  C), (2) reducing residence time by increased feed rates, and then run a second pass with interstage cooling, (3) slow tip speed of agitator discs to 6 m/s, and (4) lowering the concentration of cells if viscosity contributes to excessive heating.

l

Detrimental contamination of the batch due to wear of either the grinding media or container walls is usually rare due to the insignificant amounts involved. A hint to mitigate any contamination problem is to use inert materials of construction. For

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Fig. 3 Bead impact—agitated beads—DYNO-MILL with 600 mL chamber equipment

Fig. 4 Bead impact—agitated stirred beads—inside view—spinning agitators moving beads against suspended cells. (1) input, (2) output, (3) agitator discs, (4) beads, (5) cooling jacket, and (6) bead screen

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example when iron would be harmful, do not use steel beads, switch to glass or ceramics. The least wear is reportedly seen when both the ceramic beads and containers are fabricated from zirconium oxide stabilized with yttrium (95%/5%; specific gravity 6.0). l

4 4.1

For small diameter cells (e.g., E. coli of 0.25–1.0 μm) use beads of 0.10–0.5 mm diameter. For larger cells (e.g., yeast, algae, hyphae) use beads of 0.5–1.25 mm. Glass (sp. gr. 2.5) is a good starting material due to low cost. If addition energy is needed to improve breakage/homogenization, then switch to ceramic (zirconium oxide family) with specific gravities from 3.8 to 6.1.

l

A quick preliminary test can be run with standard lab equipment. Load a beaker with the buffered cells/tissue along with an equal volume of beads. Place on a magnetic stirrer (or use an overhead stirrer) and spin the beads. If some cell breakage is seen then the bead mill is a good candidate for processing the cells in question.

l

Degree of cell disruption can be controlled also by the time the sample is subjected to bead beating. For recovery of larger components such as intact nuclei, mitochondria, and other organelles accept only 60–70% breaker since a longer run may open more of the cells, but at the cost of damage to the components wanted. For strong stable molecules, such as the precursors for biodiesel fuels, higher breakage rates are warranted.

Rotor–Stator Homogenizer Theory

Rotor–stator homogenizers consist of a rapidly spinning paddle (rotor) contained within a nonmoving open-ended tube with slots (stator) near the working end. The space between the rotor and the stator has close tolerances. The turning paddle pushes liquid out the slots, creating a low-pressure region that draws fresh suspension up from the open end. As the tissue is pulled up, there is a stretching/shearing action. Then, as the material is forced between the narrow gaps, there is a cutting action. The working part of the equipment that contacts the samples is collectively called the generator. The rotor–stator design consists of two or more coaxial interlocking rows of teeth. The internal rotor(s) is driven with a motor running at speeds between 3000 rpm for the larger, industrial units to 27,000 rpm for the smaller lab units. The size of the motor is chosen based on the size of the equipment and application. Rotor tip speeds of up to 50 m/s can be achieved. Usually the gap between rotor and stator is around 300–500 μm (Figs. 5 and 6).

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Fig. 5 Rotor–stator—shear by spinning shaft—POLYTRON equipment 4.2

Practical Aspects

l

Choice of the rotor–stator geometry depending upon the application. Some examples of specialized generators include extended knives to cut into a larger tissue sample, low foaming generators, and easy-cleaning units that may be autoclaved.

l

Next choose the correct size generator for the sample volume to be homogenized. Factory tables provide the volume ranges suitable for each generator thus selecting the correct parts is quite easy. For example, a sample of size from 0.5 mL to 5 mL

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Fig. 6 Rotor–stator—shear by spinning shaft—inside view

would be best processed with a generator of 5 mm diameter. As the quantity of material to be processed at one time increases, larger sizes of generators and motors are needed.

5

l

Interchangeable generators are designed to fit onto only certain motors. Therefore, one must decide which motor is the most suitable for the particular range of applications and then be sure to select only generators designed to fit onto that motor.

l

The homogenizers currently available can process sample volumes anywhere from less than 0.5 mL up to 150,000 L. Batch equipment (e.g., POLYTRON®) is use for small sample quantities from less than 0.5 mL through larger units can handle several hundred liters. For larger industrial in-line systems for either continuous or recirculation flow there are single, double, or even triple stage rotor–stator configurations (e.g., MEGATRON®).

Mortar and Pestle Tissue Grinders: Shear by Mechanical Pressure This class of simple devices disrupts cells and homogenizes tissues by the pressure and friction generated when a moving pestle pinches the samples against the wall of the mortar. Though usually operated manually, there are electrically driven units available. This equipment class differs from rotor–stator in that the sample is pressed against the container’s outer walls, not pulled up into the stator tube and turned through the slots. Selection of proper mortar depends upon samples to be processed. Materials of

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construction include glass, stainless steel, Teflon, and plastics. An example of this equipment is the Wheaton Potter-Elvehjem tissue grinders for samples of 2 mL and larger. One practical hint when using these devices for bacteria is to freeze the sample with liquid nitrogen.

6 6.1

High Pressure Batch: Expanding Fluids Theory

There are two widely used cell disruption methods that employ rapidly expanding fluids from within the cell to explode the cell membranes. The French Press G-M® uses liquid under pressure, and the Parr Cell Disruption Vessel uses compressed gases. Since these are bath operations, they are only suitable for small quantities of less than about a liter.

6.1.1 The French Press G-M®

The French Press G-M design consists of a stainless-steel cylinder (called pressure cell) fitted with an exit valve at one end and a piston/plunger at the other end. Up to 35 mL of suspended free cells in buffer are loaded into the pressure cell. With the exit valve closed, the piston is pressed against the liquid by a hydraulic press, called the FRENCH PRESS G-M. Once a suitable pressure (up to 40 kpsi) is achieved throughout the liquid and within the cell body, then the outlet valve is opened to allow the cell suspension to drip out at a slow rate of about 1 mL per min (9 to 20 drops per min). This exposure to atmospheric pressure, being much lower than the pressure that was forced within the microorganism’s body causes the liquid to burst out, and thereby rupturing the cell membrane. Normally bacteria (at 40 kpsi) and yeast (at 20 kpsi) are handled in the French Press G-M, not tissues, plants, or seeds (Fig. 7).

6.1.2 The Parr Cell Disruption Vessel

The Parr Cell Disruption Vessel is a pressure vessel into which the sample to be disrupted is placed along with a dip tube fitted with an exit valve. Suitable gas such as nitrogen at 2 kpsi is forced into the Vessel and dissolves into the cells. When the exit valve is opened the gas pressure is suddenly released causing the nitrogen to come out of solution within the cells as expanding bubbles. This action stretches the membranes of each cell until they rupture and release the contents of the cell. Although sometimes referred to as “explosive decompression,” nitrogen decompression is actually a gentle method. This method is suited for treating mammalian and other membrane-bound cells, for treating plant cells, for releasing virus from fertilized eggs, and for treating fragile bacteria. It is not recommended for untreated bacterial cells, unless using various pretreatment procedures to weaken the cell wall. Yeast, fungus, spores, and other materials with tough walls do not respond well to this method (Figs. 8 and 9).

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Fig. 7 High pressure batch—liquid expansion French Press G-M and Pressure Cells (35 and 3.7 mL) Equipment 6.2

Practical Aspects

6.2.1 French Press G-M®

l

Keeping the samples cold is achieved by prechilling the pressure cell. After the cells exit the valve, immediately cool by keeping the collection beaker on ice. There are no provisions to chill during the disruption process.

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Fig. 8 High pressure batch—gas expansion—Parr Vessel Equipment

6.2.2 PARR Cell Disruption Vessel

l

Removal of air prior to closing the exit valve will minimize the amount of oxygen degradation of the released proteins.

l

During the paced release of materials, the pressure in the pressure cell will drop. This needs to be balanced by periodically repressurizing the system by running the hydraulic press.

l

Individual cells such as lymphocytes, leukocytes, tissue culture cells, or very fragile bacterial cells will not require pretreatment. Tissues must usually be preminced to ensure that they not plug the exit dip tube and discharge valve.

l

The intended use of a homogenate generally determines the composition of the suspending medium. Isotonic solutions are commonly used. Solutions with higher concentrations will tend to stabilize the nucleus and organelles. Conversely, very dilute solutions will prestretch the cells by osmotic pressure and will render them more susceptible to disruption by the Vessel method.

l

Low concentrations of calcium chloride, magnesium acetate, or magnesium chloride added to the suspending medium will stabilize the nuclei when differential rupture is desired. Ratios of

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Fig. 9 High pressure batch—gas expansion—inside view

approximately 10 mL of suspending medium to 1 g of wet cells are commonly used to prepare the cell suspension. l

Small sample quantities can be held inside of a smaller test tube or beaker placed in the vessel. The inner container should be approximately twice the volume of the suspension to be treated, and the dip tube adjusted to reach the bottom of the container.

l

To cool a small inner vessel, it can be floated on ice–water bath within the vessel. The dip tube will hold it in place. Alternatively, the entire external unit may be chilled.

l

Degree of disruption can be controlled by the amount of gas pressure introduced. The greater the pressure, the more homogenization. Use of moderate pressures will reduce the disruptive forces and thus leave nuclei, active mitochondria, and other organelles intact.

Mechanical/Physical Cell Disruption Methods

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577

HIGH Pressure Flow: Shear Through a Valve or Tube Theory

In high-pressure homogenizers the liquid stream of suspended cells is forced at high pressure down a narrow channel or across the small gap of a valve. This accelerates its speed, thereby stretching and shearing cells. In some designs the moving stream is subsequently and abruptly impacted against an obstacle to further damage the cells’ membrane. Two versions of impact are (a) where the stream is directed to slam against an impingement wall (trade name Gaulin®), and (b) where the stream is split into two legs of a “Y,” and these lines are then directed at one another in an interaction chamber where they collide, further disrupting the cells (trade name Microfluidics®). These devices are used for freely suspended cells, not for tissue samples.

7.1.1 High Pressure Valve with Impingement Wall: Gaulin and Rannie

The construction of the high-pressure homogenizer consists of a positive displacement pump, a homogenizing valve, and sometime an impingement wall or ring. Though pumps may consist of one, two, three, or five plungers, most smaller laboratory high-pressure homogenizers for cell breakage have only one plunger. Attached to the pump is a homogenizing valve assembly that may consist of one or two stages. For cell disruption a single-stage valve is needed, typically consisting of three parts: a seat (bottom part), a valve (top part), and an impact (wear) wall or ring. By adjusting the gap or clearance between the valve and seat, the flow area in the homogenizing valve is controlled. When the flow area is reduced, pressure within the pump discharge manifold increases. When the flow area is increased, the pressure is reduced. The high pressure generated by the pump is converted to fluid velocity and heat as the fluid is discharged from the restricted area in the homogenizing valve. For cell disruption the microorganisms are disrupted due to various mechanisms associated with the fluid velocity. At 100 MPa, the fluid velocity can be as high as 450 m/s (Figs. 10 and 11).

7.1.2 High Pressure Flow Narrow Tubes or Opposed Jets: Microfluidics

These processors disrupt cells by applying a combination of shear and impact forces onto the cells. The media that contains the cells is forced through the narrow channels of the proprietary interaction chamber of the processor. Inside these channels, with typical dimensions of 75–300 μm, the fluid achieves velocities up to 400 m/s. High pressures (up to 275 MPa) are required to generate the high velocities inside the interaction chamber. The resulting shear rates can be up to 10,000,000 s1 and are the highest commercially available. An alternative configuration splits the stream of pressurized cell suspension fluid into two legs. These are then directed at one another in an interaction chamber where they collide, disrupting the cells.

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Stanley Goldberg

Fig. 10 High pressure flow—liquid shear—SPX equipment

This equipment is suitable for a variety of free cells (bacteria, yeasts, monocellular mammalian cells), but not seeds, tissue samples, or plant materials. The design of identical fluid channels in both laboratory scale and large-scale units allows for direct scale-up from the smallest laboratory unit (14 mL batch) directly to large production units (tens of L/min) (Fig. 12). 7.2

Practical Aspects

7.2.1 Practical Aspects Using High Pressure Valve with Impingement Wall: Gaulin and Rannie

l

To process a small sample size, first a liquid compatible with the continuous phase of the cell slurry is added to the feed hopper of the homogenizer. The machine is started and the pressure is set. When the liquid level reaches the very bottom of the feed hopper, the cell slurry can be added quickly. The cell slurry will push the liquid ahead of it. When the slurry is observed in the discharge, a sample can be taken. In this way limited sample is not lost while waiting for pressure to rise to operational levels.

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Fig. 11 High pressure flow—liquid shear—inside view

Fig. 12 High pressure flow—opposed liquid streams—MICROFUIDIZER Inside View l

If the product is very sensitive to heat, then it may be necessary to cool the equipment prior to introducing cell suspension and to cool the suspension immediately after it discharges from the homogenizer. A cooling coil is connected to the discharge tube of the homogenizer and is immersed in an ice–water bath. Since the homogenizer is a positive displacement pump there must be no valves or restrictions in this discharge line that could potentially shut off flow. Rapid chilling will minimize losses due to a typical temperature rise of 2.5  C per 10 MPa of pressure generated during the nearly adiabatic heating during pressurization.

l

To improve breakage, higher pressures are used up to the pump’s limits. High pressure can shorten valve life.

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Stanley Goldberg

7.2.2 Practical Aspects: High Pressure Narrow Tubes/Opposed Jets

8

l

The sample material should be fully thawed before processing; ice may plug the chambers.

l

Increased process pressure and multiple passes may increase amount of cell breakage, but also decrease the size of cell wall debris. These small particles may compromise downstream column purification steps.

l

The operating pressure and number of passes are to be evaluated to obtain maximum breakage without damaging the released materials or too much cell wall debris.

l

Most models can be autoclaved and are air driven, though some are electric-hydraulic systems. In many circumstances, especially when samples are processed multiple times, the Microfluidizer® processors require sample cooling before and/or after processing.

Low Pressure: Shear by Droplet’s Impingement Invented at Indiana University the BioNeb® disruption system disrupts cells by a low-pressure droplet method. In the process of droplet formation, large molecules or cells suspended in the liquid being nebulized are forcefully distributed from the liquid into the forming secondary droplet. This creates a transient laminar flow in the microcapillary “nebulization” channel formed between the surface of the liquid and the forming secondary droplet. The laminar flow in the capillary channel exerts sufficient shearing forces to break cells. The shearing force created depends on the gas pressure applied (10–250 psi), the type of gas (nitrogen, argon, etc.) and the viscosity of the liquid. By varying these parameters it is possible to precisely regulate the magnitude of the force applied during nebulization. Throughput using this method is very limited (Fig. 13).

9 9.1

Ultrasonic Processors: Shear by Collapsing Bubbles Theory

The use of sound waves in fluids can disrupt cells. The operation starts with normal electrical current (50 Hz or 60 Hz) being transformed to 20,000 Hz. This electrical signal is fed to a piezoelectric crystal causing it to oscillation at this high frequency. The vibrations move a titanium metal HORN about 5–15 μm. The shape of the horn amplifies this motion to 100–150 μm per cycle. By placing the horn’s end—the tip—into fluid, the tip moves the liquid forward (away) and then retracts (back) quicker than the liquid can return. During the return stroke, the pressure in the system drops below the vapor pressure of the liquid so boiling

Mechanical/Physical Cell Disruption Methods

581

Fig. 13 Droplet low pressure nebulizer—BioNeb equipment

occurs (“cavitation”). As the liquid flows back, the bubbles collapse. This bubble collapsing imparts the energy needed to disrupt the cells (Fig. 14). 9.2

Practical Aspects

l

Denaturing of proteins due to high temperature or excessive shear is often noted. The equipment has ON–OFF–ON cycle adjustments available. During the OFF periods, the samples can cool. Lowered amplitude settings may reduce protein damage.

l

Hint to improve bubble formation is to keep the system as cold as possible with ice.

l

Power density at the generator is a way to compare different equipment designs, and to see what levels offer the best cell breakage with good recoveries of target components. Higher output power is required to sustain good performance in large size probes. For cell disruption, probe densities should be at least 100 W/cm2 or more to increase tip amplitude movement (30–250 μm).

l

Time savings can be achieved by ganging several samples into 96-well or larger microtiter plates in place of running one sample at a time. Multi models can accommodate these plates. Also, some models have several tips fitted on a single horn that can handle several samples simultaneously.

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Stanley Goldberg

Fig. 14 Ultrasonic——shear by collapsing bubbles—SONICATOR equipment

10 10.1

l

For microorganisms adding 0.1–0.5 mm diameter glass beads in a 1:2 ratio of beads to liquid volume will enhance cell breakage. Tough tissues such as skin or tendon should be prechopped with a blade or pulverized in liquid nitrogen.

l

Keep the probe deep into the sample to prevent foaming.

l

Free radical can be generated by sonication and these will oxidize and damage most biomolecules. This can be minimized by flooding with nitrogen and/or add scavengers such as cysteine, dithiothreitol, or other –SH compounds.

l

Larger quantities up to 100 L per h flow rates can be processed in the SONITUBE. Here the entire pipe for 15 in. (35 cm) oscillates and ultrasonically processes all fluids pumped through.

Extraction Across an Electromotive Field Theory

Extracting nonpolar lipids from microalgae is achieved using a lipid extraction device having an anode and a cathode that forms a channel and defines a fluid flow path through which aqueous slurry

Mechanical/Physical Cell Disruption Methods

583

Electro Water SeparationTM The OriginOil Two-Stage Process

Electro-Flotation Concentrate

Electrical Pulses

Contaminated Water

Clear Water

Electro-Coagulation Electrical Pulses

Fig. 15 Electro Water Separation

is passed. An electromotive force is applied across the channel at a gap distance in a range from 0.5 mm to 200 mm to cause the nonpolar lipids to be released from the algae cells (Fig. 15). 10.2 Practical Aspects

11 11.1

l

Still experimental for cell lysing or product extracting. The manufacturer has minimized commercialization of this technology.

l

The nonpolar lipids can be extracted at a high-throughput rate and with low concentrations of polar lipids such as phospholipids and chlorophyll.

l

Used to concentrate algae and other microorganisms from aqueous systems for harvesting and concentrating cell masses.

l

Clearing water streams.

Lysis with Pulsed Electric Fields Theory

11.2 Practical Aspects

Pulsed electric field (PEF) processing uses short, high-voltage pulses. These pulses induce electroporation of cells leading to cell disintegration that allows contained components to be released. This method has numerous applications in biology and biotechnology and has become an important technique in molecular medicine. l

Low energy cell breakage technology for the assisted extraction of different molecules (proteins, plasmid DNA, lipids) from bacterial and yeast cells.

584

12 12.1

Stanley Goldberg l

Medical applications in cell biology. Research underway to detect intracellular biomarkers in viable cells.

l

Food applications for product recovery. Also to kill cells to increase shelf life.

l

By creating pores in algal cell walls this allows for use of less solvent to extract lipids from algae.

Microwave-Assisted Centrifugation Theory

12.2 Practical Aspects

Microwave-assisted centrifugation (MW/C) combines microwave heating to disrupt biological cells of plant materials combined with centrifugation to intensify diffusion, collection and separation of biological material. MW/C is usually performed at atmospheric pressure without adding any solvent or water. l

Similar applications as with pulsed electric field, but by combining heat with the centrifugal forces. This can allow for the reduction in the amount of solvents needed.

l

Enrichment of phosphopeptides via NiO nanoparticles using a microwave-assisted centrifugation on-particle ionization/ enrichment approach in MALDI-MS.

l

Recovery of polyphenols from lettuce.

Acknowledgments Superb assistance was rendered by the following people, especially Tim Hopkins, who are most conversant in their noted equipment areas. Bead Milling by Tim Hopkins (BIOSPEC PRODUCTS) and Harald Frommherz (W. A. BACHOFEN AG); Rotor–Stator by Roger Munsinger (KINEMATICA USA/AG); High Pressure Flowing Liquid by Toni Parker (SPX) and Peter Sokol (MICRO FLUIDICS/IDEX); High Pressure Batch Liquid by Robert Borgon (University of Central Florida); High Pressure Batch Gas by Lisa Randolph (PARR INSTRUMENTS); Low Pressure Flowing Gas by James Jacso (GLAS-COL); Ultrasonic Processors by Andrea Coppola and Marc Lusting (QSONICA); Electro Water Separation by Riggs Eckelberry (ORIGIN OIL); Microwave Assisted Centrifugation by Gerard Lynch (Sigma Design Company). My book editor Dr. Anton Posch has been exceptionally supportive in both technical and motivational matters. My wife Robin, sons Evan and Adam, and granddaughter Sadie are the sparkle of my life.

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References 1. Reinkemeier M, Rocken W, Leitzmann C (1996) A rapid mechanical lysing procedure for routine analysis of plasmids from lactobacilli, isolated from sourdoughs. Int J Food Microbiol 29:93–104 2. Rezwan M, Laneelle MA, Sander P, Daffe M (2007) Breaking down the wall: fractionation of mycobacteria. J Microbiol Methods 68:32–39 3. Jung J, Xing X, Matsumoto K (2001) Kinetic analysis of disruption of excess activated sludge by Dyno Mill and characteristics of protein release for recovery of useful materials. Biochem Eng J 8:1–7 4. Lizotte E, Tremblay A, Allen BG, Fiset C (2005) Isolation and characterization of subcellular protein fractions from mouse heart. Anal Biochem 345:47–54 5. Lindeskog P, Haaparanta T, Norgard M, Glaumann H, Hansson T, Gustafsson JA (1986) Isolation of rat intestinal microsomes: partial characterization of mucosal cytochrome P-450. Arch Biochem Biophys 244:492–501 6. Borgon RA, Verity N (2012) Quantitative biological methods. Pearson Learning Solutions, Boston, MA 7. Autuori F, Brunk U, Peterson E, Dallner G (1982) Fractionation of isolated liver cells after disruption with a nitrogen vessel and sonication. J Cell Sci 57:1–13 8. Kelly WJ, Muske KR (2004) Optimal operation of high-pressure homogenization for intracellular product recovery. Bioprocess Biosyst Eng 27:25–37 9. Mehlhorn I, Groth D, Stockel J, Moffat B, Reilly D, Yansura D, Willett WS, Baldwin M, Fletterick R, Cohen FE, Vandlen R, Henner D,

Prusiner SB (1996) High-level expression and characterization of a purified 142-residue polypeptide of the prion protein. Biochemistry 35:5528–5537 10. Vinatier J, Herzog E, Plamont MA, Wojcik SM, Schmidt A, Brose N, Daviet L, El Mestikawy S, Giros B (2006) Interaction between the vesicular glutamate transporter type 1 and endophilin A1, a protein essential for endocytosis. J Neurochem 97:1111–1125 11. Jaki BU, Franzblau SG, Cho SH, Pauli GF (2006) Development of an extraction method for mycobacterial metabolome analysis. J Pharm Biomed Anal 41:196–200 12. Eckelberry N (2013) US patent application publication, US 2013/0211113 A1, 15 Aug 2013 13. Bleakley S, Hayes M (2017) Algal proteins: extraction, application, and challenges concerning production. Foods 6(5):33 14. Kasˇe˙ta V, Kausˇyle˙ A, Kavaliauskaite˙ J, Petreikyte˙ M, Stirke˙ A, Biziulevicˇiene˙ G (2020) Detection of intracellular biomarkers in viable cells using millisecond pulsed electric fields. Exp Cell Res 25:111877 15. Angoy A, Valat M, Ginisty P, Sommier A, Goupy P, Caris-Veyrat C, Chemat F (2018) Development of microwave-assisted dynamic extraction by combination with centrifugal force for polyphenols extraction from lettuce. LWT 98:283–290 16. Hasan N, Wu H (2011) Highly selective and sensitive enrichment of phosphopeptides via NiO nanoparticles using a microwave-assisted centrifugation on-particle ionization/enrichment approach in MALDI-MS. Anal Bioanal Chem 400:3451–3462

INDEX A Acetone precipitation.................................................... 544 Adipokines ............................................................ 421, 422 Adipose tissues ................................................6, 421–430, 433, 434 Affinity chromatography....................................... 56, 461, 535–546, 552 Alkylation...........................................................14, 18, 22, 27, 29, 112, 126, 139, 159, 168, 180, 196, 200, 232, 239, 281, 282, 555 Alzheimer disease ......................................................79–91 Antibody anti-biotherapeutic......................................... 294, 300 anti-drug................................................ 291, 296, 297 anti-idiotypic .................................................. 291–305 primary........................................................... 110, 111, 124, 125, 223, 237, 291, 361, 364, 447, 449, 454, 458, 459, 461, 462, 465, 466, 470, 485, 517, 518, 522 secondary ....................................................... 110, 111, 124, 125, 166, 221, 361, 365, 447, 450, 454, 458, 459, 461, 462, 465, 466, 470, 517, 522 Arabidopsis .......................................................... 346, 348, 536, 538, 541, 545, 559 Assay bead-based ............................................................... 278 bridging .........................................293, 298, 300, 303 cell activation .................................................. 114, 132 immunogenicity ...................................................... 294 invasion .................................................. 106, 114, 132 ligand binding ........................................291–304, 402 mass spectrometry.......................................... 278, 382 peptides............................................................ 62, 112, 128, 140, 160, 169, 180, 278, 279, 304 proteins ...............................................................83, 89, 135, 157, 198, 199, 202, 242, 263, 264, 277–286, 296, 297, 304, 382, 390, 395–409, 434, 440, 481, 482, 494, 502, 507, 514, 527, 545 proximity ................................................................. 382

B Balch homogenizer ....................................................... 412 Bead-based array ........................................................... 276 Bead beater ................................................................27, 29 Bead impact .......................................................... 564–570

Bead mills .................................................... 565, 566, 570 Benzonase.................................................... 384, 389, 392 Bicinchoninic acid (BCA) protein assay....................... 502 Biomarkers............................................................... 25, 26, 152, 153, 230, 264, 267, 276, 278, 307, 438, 526, 584 BioNeb cell disruption................................ 565, 580, 581 Bio-Plex ............................................................... 249, 256, 258, 260, 264–268, 270, 272–274, 276, 436 Biosimilars ........................................................94, 95, 101 Biotinylation identification (BioID) ................... 357–377 Bradford protein assay ........................................ 491, 493, 511, 514, 526, 527

C Carrier ampholytes ................................................ 94, 102, 487, 511, 523, 556, 560, 561 cDNA displays ..................................... 307–312, 315–318 Cell counter ......................................................... 208, 215, 216, 329, 332 Cell disruption bead impact methods..................................... 564, 568 electromotive field.......................................... 582–583 high pressure batch ....................... 565, 566, 573–576 high pressure flow ......................... 565, 566, 577–580 low pressure................................................... 565, 566, 570, 580, 581 mortar and pestle tissue grinders ........................... 572 rotor-stator homogenizer .............................. 570–572 ultrasonic processors ...................................... 580–582 Cell lysis ............................................................... 215, 216, 223, 328, 338, 366, 374 Cells CHO ............................................................... 489, 509 HEK293 ......................................................... 242, 509 HeLa ................................................................. 62, 242 primary skeletal muscle ........................................... 435 SW480 ..................................................................... 197 U373............................................................... 197, 202 Cell walls.................................................. 8, 573, 579, 584 Chaotropes ........................................................... 239, 552 ChIP, see Chromatin immunoprecipitation (ChIP) Chloroplasts arabidopsis ............................................. 536, 541, 545 pea ................................................................... 536, 545

Anton Posch (ed.), Proteomic Profiling: Methods and Protocols, Methods in Molecular Biology, vol. 2261, https://doi.org/10.1007/978-1-0716-1186-9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

587

PROTEOMIC PROFILING: METHODS

588 Index

AND

PROTOCOLS

Chromatin immunoprecipitation (ChIP) ..................................................6, 323–342, 346, 348–355 Chromatography affinities.................................................. 332, 377, 552 flow-through (FT) ..............................................57, 69 hydrophilic interaction (HILIC).................. 7, 37–39, 56, 57, 60, 61, 66–68, 70 hydrophobic interaction (HIC) .................... 539, 543 ion exchange (IEX) ........................................ 539, 542 reverse-phase (RP) ............................ 57, 70, 194, 205 size exclusion (SE) ................................ 377, 536, 542 ultra-high performance (UHP).............................. 170 CID, see Collision induced dissociation (CID) Co-immunoprecipitation (Co-IP) ............................... 230 Co-IP, see Co-immunoprecipitation (CO-IP) Collision induced dissociation (CID) ............................ 68 Coomassie Brilliant Blue G .......................................... 383 Coomassie Brilliant Blue R.................................. 397, 399 Coomassie staining ............................................... 17, 383, 397, 399, 444, 469, 482, 545 Crosslinking................................................................... 358 Culture medium.................................................. 107, 108, 114, 115, 119, 132, 136, 178, 194, 195, 197, 202, 208, 210, 257, 271, 423, 426, 427, 437, 502, 521 CyDyes........................................................................... 552 Cysteine labeling .................................................. 553, 555

D Data normalization stain-free technology...................................... 443–455 western blotting ............................................. 443–455 Detergents ........................................................... 136, 141, 194, 230, 231, 241–243, 281, 283, 332, 338, 376, 459, 460, 522, 523, 526 Dimethylformamide (DMF) .................................. 37, 38, 235, 241, 308, 491, 494, 495 Disulfide bridges ......................................... 282, 460, 553 Dithioerythritol (DTE) ................................................ 561 Dithiothreitol (DTT).............................................. 14, 15, 23, 26, 27, 48, 50, 58, 82, 83, 86, 87, 108, 112, 124, 126, 136, 139, 158, 159, 167, 168, 175, 180, 242, 311, 314, 315, 360, 361, 384, 391, 476, 484, 493, 499, 511, 512, 515, 521–523, 526, 538, 540, 553, 555, 560, 561, 582 DMEM, see Dulbecco’s modified Eagle medium (DMEM) DMF, see Dimethylformamide (DMF) Dounce homogenizer .........................208, 210, 211, 447 Down Syndrome .......................................................79–91 Droplet low pressure nebulizer .................................... 581 Droplet’s impingement................................................. 579

DTT, see Dithiothreitol (DTT) Dulbecco’s modified Eagle medium (DMEM).................................................. 107, 108, 115, 120, 134, 195, 211, 360–363, 384, 388, 423 Dyno-Mill ...................................................................... 569

E Ectosomes............................................................. 193, 194 Electric pulse stimulation ........................... 434, 435, 439 Electron microscopy ........................................... 134, 139, 154, 203, 204 Electrophoresis 2-D electrophoresis......................................... 85, 511, 514, 515, 518 differential in gel electrophoresis (DIGE) ..................................................... 552–554 imaged capillary isoelectric focusing (icIEF)..........................................................93–102 isoelectric focusing .................................................. 550 SDS-PAGE ...................................................... 74, 158, 165, 446, 447, 515, 518 Electrospray ionization (ESI) ..........................2, 550, 559 Endotoxemia ........................................................ 248, 249 Escherichia coli (E. coli) ......................................... 74, 247, 358, 369, 372, 383, 386, 387, 397, 470 ESI, see Electrospray ionization (ESI) Exosome density-gradient separation..................................... 116 electron microscopy ....................................... 111, 125 immunoaffinity capture methods .................. 109, 124 ultracentrifugation ........................................ 107, 109, 114–116, 119, 121–123, 134, 136

F Filter aided sample preparation (FASP)................. 14, 15, 19, 25, 27, 30, 210, 211, 493, 499, 527 Flow cytometry cluster of differentiation (CD) ............................... 214 forward scatter (FSC) ........................... 213, 214, 220 immunophenotyping ..................................... 214, 221 side scatter (SSC) .................................................... 226 Formalin fixed paraffin embedded (FFPE) human heart ................................................... 525–531 Fractionation ........................................................... 26, 56, 57, 60, 65, 66, 175, 194, 205, 230 French press.......................................................... 573, 574

G Gel imaging binning............................................................ 475, 524 bit depth ......................................................... 471, 473

PROTEOMIC PROFILING: METHODS CCD............................................................... 167, 264, 464, 465, 469–472, 474 dynamic range ............................................... 464, 468, 472, 473, 477 housekeeping normalization .................................. 477 quantitation ........................................................87, 93, 121, 473 resolution................................................................. 121 total protein normalization .................................... 477 Glycomics ..................................................................1, 3, 6 Glycosylation ..............................................................9, 35, 55–71, 93, 482, 485–487 Gradient centrifugation exosomes.................................................................. 116 mitochondria ........................................................... 413

H Heparin ................................................................ 173, 215, 216, 223, 257, 266 High mobility group box 1 protein (HMGB1) ................................................. 277–288 HILIC, see Hydrophilic interaction liquid chromatography (HILIC) Homogenizers......................................................... 81, 83, 208, 211, 415–418, 447, 526, 527, 569–572, 577–579 Horseradish peroxidase (HRP) western blotting ............................................. 513, 519 Host cell protein (HCP) antibody coverage ................................................... 520 CHO cells ................................................................ 489 image analysis ........................................ 513, 520, 522 peptide ligands ............................................... 489–504 Western blotting............................................. 507–523 2-D electrophoresis........................................ 514, 518 Human Combinatorial Antibody Library (HuCAL) .................................................. 292, 294 Human plasma .............................................................. 434 Human serum ....................................................... 96, 175, 215, 217, 291–293, 295, 297, 298, 300 Hydrophilic interaction liquid chromatography (HILIC) ................................................... 7, 37–39, 56, 57, 60, 61, 66–68, 70

I Iceman ......................................................................... 1–10 ICPL, see Isotopecoded protein label (ICPL) IEF, see Isoelectric focusing (IEF) Image acquisition ................................................. 462–475 Immobilized metal affinity chromatography (IMAC) ........................................................ 56, 57, 60, 63, 69, 71 Immobilized pH gradient (IPG)............................ 82, 86, 511, 512, 514–517, 521, 523

AND

PROTOCOLS Index 589

Immunoaffinity ................................................... 153, 176, 194, 231, 232, 278 Immunoassay bio-Plex..................................................249, 253–256, 258, 260, 264–276 multiplexing................................... 247–260, 263–276 Immunoblotting, see Western blotting Immunophenotyping, see Flow cytometry Immuno-polymerase chain reaction (IPCR) ...................................................... 307, 308 Immunoprecipitation (IP) biotin.............................................................. 231, 232, 237, 238, 242–244 crosslinking .............................................................. 358 GFP-trap ........................................ 382, 385, 388–393 magnetic beads .............................................. 231, 235, 238, 239, 243, 244, 281 mass spectrometry.......................................... 229–244 protein A/G .................................................. 232, 242, 331, 335, 349 protein-protein interaction ........................... 358, 382, 383, 385, 388, 392 streptavidin .................................................... 231, 232, 235, 238, 239, 243, 244 Inflammation .........................................80, 248, 278, 433 In-gel protein digest ............................................ 443–455 In-solution protein digest............................................. 196 Iodixanol.............................................................. 109, 121, 122, 137, 194, 196, 198, 199, 203 IPG, see Immobilized pH gradient (IPG) Isobaric Tags for Relative and Absolute Quantitation (iTRAQ) ..................................61, 70 Isoelectric focusing (IEF) ....................................... 82, 86, 89, 90, 93, 95, 482, 511, 514, 515, 521–523, 552 Isoelectric point (pl) ............................................. 87, 490, 508, 523 Isotope-coded protein label (ICPL) ................... 559–561

L Label-free proteomics ................................. 130, 171, 529 Lipidomics .................................................................1, 3, 5 Liquid biopsies ..................................................... 152–181 Liquid chromatography coupled to mass spectrometry (LC-MS) ............................. 2, 6, 10, 14, 15, 17, 18, 31, 34, 36–38, 49, 60, 67, 71, 111–113, 159, 160, 210, 211, 235, 236, 239, 241, 243, 281, 282, 284, 285, 411, 527, 528, 531, 558 Loading buffer .........................................................58–60, 62, 63, 65, 66, 70, 71, 112, 113, 127, 128, 159, 160, 169, 243, 244, 282, 285, 384, 389, 390 Lymphocytes ............................................... 217, 226, 575 Lysis solution .......................................................... 27, 29, 30, 32, 120, 165

PROTEOMIC PROFILING: METHODS

590 Index

AND

PROTOCOLS

M

O

MALDI, see Matrix assisted laser desorption ionization (MALDI) Mass spectrometry (MS) acetylation.................................................................. 56 carbamidomethylation ..................................... 68, 504 electrospray ionization (ESI).....................2, 551, 559 elemental bio imaging............................................. 2, 9 FASP ...................................................... 13, 14, 15, 19 immunoaffinity ............................................... 277–288 inductively coupled plasma mass spectrometry (ICP-MS)......................... 2, 7, 9, 10 internal standard triggered PRM (isPRM) .................................................... 230, 234 MALDI ................................................ 2, 15, 559, 584 N-glycan analysis ....................................................... 36 on-bead digestion ................................................... 230 parallel reaction monitoring (PRM) ...................................................... 230, 234, 235, 277, 284, 286 phosphorylation .....................................56, 60, 67–69 selective Reaction Monitoring (SRM) ...................................... 229–232, 234, 286 StageTip..................................................15, 16, 20–23 suspension trapping (STrap)..................28–30, 32, 33 tandem ....................................................................... 81 Matrix assisted laser desorption ionization (MALDI) .......................................................2, 559 Membrane proteins.............................................. 486, 556 Metabolomics ............................................................1–3, 6 Metallomics ................................................................... 3, 8 Microarrays .................................................................... 346 Microfluidizer................................................................ 579 Microvesicles isolations ......................................................... 136, 176 Microwave assisted centrifugation ............................... 584 Mini bead beater .................................................. 565, 567 Mitochondria isolations ........................................412, 415, 417, 418 Mortar and pestle tissue grinders ........................ 572–573 Multiple myeloma ................................................ 151–181 Multiplexing ............................................................ 2, 232, 264, 468, 550, 551, 555 Muscle human ...................................................... 433–441

Oetzi ............................................................................ 1–10 Organelle isolation chloroplast ............................................................... 536 mitochondria ........................................................... 570

N Nanoparticle tracking analyses ........................... 137, 154, 158, 166–167, 203 Nuclei isolation ......................................................... 105, 113, 114, 132, 133, 152, 153, 193, 207, 225, 276, 278, 329, 351

P Paraffin embedding .............................................. 525–531 Peptide libraries.................................................... 490, 501 Peptide ligands ..................................................... 489–504 Peptides desalting................................................. 15, 20, 59, 66 glycopeptides ................................................ 60, 63, 70 identifications .......................................................2, 57, 81, 371, 434, 529, 552 phosphopeptides .......................................... 56, 60, 69 reversed-phase chromatography.................... 528, 552 shotgun proteomics ............................................ 30–32 Peroxisomes................................................................... 138 Phenylmethylsulfonyl fluoride (PMSF) .................................................... 281, 283, 287, 329–331, 349, 396, 397, 511, 521 Phosphatase inhibitor ............................................. 33, 58, 208, 211, 242 Phosphopeptides enrichments ......................................... 56, 60, 69, 584 identifications ............................................................ 69 Phosphoproteins enrichment...................................................... 234, 235 fluorescent gel staining ....................................... 74–77 Phosphorylation .......................................................55–71, 73, 74, 93, 242, 338, 357, 434, 482 Phos-tag technology .................................................73–77 Plasma proteins ............................................................. 277 Polytron ......................................................................... 572 Polyubiquitination ............................................. 80, 81, 87 Posttranslational modifications (PTM) acetylation.................................................................. 56 glycosylation .............................................................. 56 phosphorylation ........................................................ 56 ubiquitination.......................................................... 382 Pre-adipocytes ..................................................... 422–424, 427, 428, 430 Protease inhibitors .................................................. 26, 29, 33, 58, 157, 211, 349, 361, 366, 384, 391, 424, 428, 521, 522, 538, 540 Proteinase K ........................................................ 331, 337, 350, 353, 355 Protein assays bicinchoninic acid (BCA) ............................. 108, 157, 242, 397, 399

PROTEOMIC PROFILING: METHODS bradford ......................................................... 491, 493, 511, 514, 526 detergent compatible (DC) .......................... 115, 116, 118, 119, 135 NanoDrop ..........................................................15, 16, 20, 310, 315, 330, 333, 335, 339, 384, 445, 447 Protein complexes ............................................... 194, 229, 358, 371, 377, 536 Protein depletion .......................................................... 325 Protein digestion alkylation.............................................................14, 18, 22, 58, 62, 111, 127, 139, 159, 167 filter aided sample preparation (FASP) ..........................................................14, 15, 19, 28, 211, 493, 499, 527 in-gel digest ..........................................................4, 14, 17, 22, 23 in-solution digest ............................................ 14, 200, 230, 242 Lys-C .............................................................. 139, 180 OASIS .................................................................14, 17, 18, 22, 23, 58, 70 reduction ............................................................14, 18, 58, 62, 111, 126, 139, 159, 167, 176, 180, 196, 200, 232, 239, 281, 282, 301, 314, 455, 556, 558, 584 STAGETip ..........................................................20, 21, 32, 32, 126, 140, 159, 180 trypsin .................................................................14, 15, 18, 23, 31, 32, 58, 62, 68, 83, 244, 371, 376, 504 Protein-DNA interaction....................323, 324, 345–355 Protein extraction ................................................. 70, 526, 527, 540, 543, 561 Protein fractionation ....................................57, 60, 65–67 Protein interactions............................................. 131, 267, 358, 359, 369, 372, 373, 377, 381, 382 Protein labeling dimethyl labeling....................................................... 70 fluorescent dyes ............................................. 501, 550, 551, 553, 556 stable isotopes ......................................................... 552 tandem mass tag (TMT)....................................14, 17, 21, 22, 61, 70, 140, 180, 233 Protein microarray ........................................................ 346 Protein precipitation acetone..................................................................... 544 clean-up kit ..................................................... 310, 315 methanol/chloroform .........................................7, 28, 33, 58, 62, 82, 108, 109, 158, 166, 194, 196, 200, 205, 244, 331, 355, 516 Protein profiling ..................................................... 4, 8, 10 Protein-protein interaction........................ 131, 357–377, 381–393

AND

PROTOCOLS Index 591

Proteomics............................................................. 1–5, 25, 26, 33, 81, 83, 89, 106, 111–112, 126–131, 133, 136, 139, 140, 152–181, 193–205, 207–226, 233, 244, 276, 277, 307, 371, 411–419, 421–431, 434, 436–440, 493–495, 499–501, 525–531, 535, 536, 545, 549, 550, 552 Proximity labeling ................................................ 357–377

Q Quantitative polymerase chain reaction (qPCR)..................................................... 312, 318, 328, 331, 336–338, 340, 341, 346, 350, 353, 355, 430

R Reducing agents dithiothreitol (DTT)............................. 136, 242, 391 tris(carboxyethyl) phosphine (TCEP) ...............................................50, 242, 391 Rotor-stator homogenizer................................... 570–572

S Sample clean-up ..................................112, 159, 553, 560 Sample preparations ........................................3, 8, 13–23, 25, 35, 36, 38–43, 81, 111, 139, 211, 216, 266, 268, 329, 331, 422, 435, 438, 445, 447, 477, 493, 499, 509, 513, 526, 552 Secretome analysis......................................................... 424 Shotgun proteomics..................................................25–34 SILAC, see Stable isotope labeling with amino acids in cell culture (SILAC) Silver staining .............................................. 545, 550, 552 Solid phase extraction (SPE) .....................................6, 16, 20, 23, 36–39, 50, 492, 495, 496, 498, 499 Sonication ................................................................ 19, 20, 27, 83, 89, 133, 210, 324, 332, 339, 340, 346, 349, 352, 355, 366, 386, 388, 392, 582 Sonicator bath sonicator................................................... 83, 210 tip sonicator............................................................... 88 SPE, see Solid phase extraction (SPE) SPR, see Surface plasmon resonance (SPR) Stable isotope labeling with amino acids in cell culture (SILAC) ..................................... 23, 233, 557–559 Stain-free technology ..........................443–455, 477, 478 Streptavidin.......................................................... 138, 231, 232, 235, 237, 238, 243, 244, 248, 252, 256, 264, 302, 310–312, 316, 318, 358, 361, 365–368, 370–373, 384, 552 Surface plasmon resonance (SPR)................................ 377 Sypro Ruby ......................................................... 82, 86, 89

PROTEOMIC PROFILING: METHODS

592 Index

AND

PROTOCOLS

T Thermal shift assay (TSA) fatty acid binding protein .............................. 395–408 real-time PCR................................................ 396, 398, 400, 401, 408 SYPRO Orange dye ....................................... 398, 407 Tissue grinders .............................................................. 573 Tissue homogenization........................................ 563–584 Tris(carboxyethyl) phosphine (TCEP) ..................................................22, 27, 29, 48, 82, 108, 196, 236, 242, 281, 329, 361, 384, 397, 415, 445, 446, 511, 512, 521, 544, 553 Trypsin ...............................................................14, 15, 18, 20, 23, 27, 30, 32, 58, 62, 68, 70, 83, 88, 112, 127, 128, 130, 139, 159, 168, 172, 180, 194–196, 201, 205, 208, 222, 232, 235, 236, 239–242, 278, 281, 283, 358, 371, 376, 436, 468, 493, 500, 502, 504, 526, 528, 550, 559 2D-PAGE ...................................................................... 177

U Ultracentrifugation ............................................. 107, 114, 121–123, 134, 136, 153, 176, 194, 195, 198, 199, 202, 203 Ultrafiltration ....................................................... 176, 526 Urine proteomics .......................................................... 307

V Vacuum centrifuge ...................................................15–17, 19–22, 58, 60, 63, 65, 66, 70, 112, 160, 309, 315

Vesicle extracellular ............................................152–180, 428

W Western blotting antibody selection ................................................... 457 blocking ................................................................... 443 chemiluminescence ................................................. 463 data normalization .................................................. 354 fluorescence ............................................................. 466 housekeeping proteins .................................. 444, 477, 478, 484 image acquisition .................................................... 462 image analysis ................................................ 451, 455, 458, 475, 513, 520, 522, 549 imaging systems ...................................................... 469 immunostaining .................................... 436, 447, 449 loading controls ...................................................... 444 molecular weight (size) estimation ........................ 475 multiplexing................................................... 468, 550, 551, 555 nitrocellulose membrane ............................... 450–452 PVDF membrane .......................................... 451, 469, 470, 512, 516–518, 523 reprobing ............................................... 468, 469, 470 stain-free technology............................................... 477 stripping.......................................................... 468–470 visualization methods ............................................. 462 Wheaton Potter-Elvehjem tissue grinder ............................................ 565, 573