Current Protocols in Cytometry
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Current Protocols in Cytometry
Table of Contents Foreword Preface Chapter 1 Flow Cytometry Instrumentation Introduction Unit 1.1 Overview of Flow Cytometry Instrumentation Unit 1.2 Fluidics Unit 1.3 Standardization, Calibration, and Control in Flow Cytometry Unit 1.4 Establishing and Maintaining System Linearity Unit 1.5 Optical Filter Sets for Multiparameter Flow Cytometry Unit 1.6 Laser Beam Shaping and Spot Size Unit 1.7 High-Speed Cell Sorting Unit 1.8 Principles of Gating Unit 1.9 Lasers for Flow Cytometry Unit 1.10 Techniques for Flow Cytometer Alignment Unit 1.11 Flow Cytometers for Characterization of Microorganisms Unit 1.12 Principles of Resonance Energy Transfer Unit 1.13 Light Scatter: Detection and Usage Unit 1.14 Compensation in Flow Cytometry Unit 1.15 Time-Resolved Fluorescence Measurements Unit 1.16 Simultaneous Analysis of the Cyan, Green, and Yellow Fluorescent Proteins Unit 1.17 Plug Flow Cytometry Unit 1.18 Dynamic Thermoregulation of the Sample in Flow Cytometry Unit 1.19 Excitation and Emission Spectra of Common Dyes
Unit 1.20 Characterization of Flow Cytometer Instrument Sensitivity Unit 1.21 Separation Index: An Easy-to-Use Metric for Evaluation of Different Configurations on the Same Flow Cytometer
Chapter 2 Image Cytometry Instrumentation Introduction Unit 2.1 Contrast Enhancement in Light Microscopy Unit 2.2 Microscope Objectives Unit 2.3 Cameras Unit 2.4 Optical Filtering Systems for Wavelength Selection in Light Microscopy Unit 2.5 Digital Fluorescence Microscopy Unit 2.6 Calibration: Sampling Density and Spatial Resolution Unit 2.7 Microscope Alignment Unit 2.8 Confocal Microscopy: Principles and Practices Unit 2.9 Multiphoton Imaging Unit 2.10 Scanning Laser Cytometry Unit 2.11 Shading Correction: Compensation for Illumination and Sensor Inhomogeneities Unit 2.12 Photobleaching Measurements of Diffusion in Cell Membranes and Aqueous Cell Compartments Unit 2.13 Optimizing Laser Source Operation for Confocal and Multiphoton Laser Scanning Microscopy
Chapter 3 Safety Procedures and Quality Control Introduction Unit 3.1 Principles of Quality Control
Unit 3.2 Components of Quality Control Unit 3.3 Testing the Efficiency of Aerosol Containment During Cell Sorting Unit 3.4 Safe Use of Hazardous Chemicals Unit 3.5 Method for Visualizing Aerosol Contamination in Flow Sorters Unit 3.6 Standard Safety Practices for Sorting of Unfixed Cells
Chapter 4 Molecular and Cellular Probes Introduction Unit 4.1 Titering Antibodies Unit 4.2 Conjugation of Fluorochromes to Monoclonal Antibodies Unit 4.3 Nucleic Acid Probes Unit 4.4 Cellular Function Probes Unit 4.5 Spectroscopic Analysis Using DNA and RNA Fluorescent Probes
Chapter 5 Specimen Handling, Storage, and Preparation Introduction Unit 5.1 Handling, Storage, and Preparation of Human Blood Cells Unit 5.2 Handling, Storage, and Preparation of Human Tissues Unit 5.3 Flow Analysis and Sorting of Plant Chromosomes
Chapter 6 Phenotypic Analysis Introduction Unit 6.1 Quality Control in Phenotypic Analysis by Flow Cytometry Unit 6.2 Immunophenotyping
Unit 6.3 High-Sensitivity Immunofluorescence/Flow Cytometry: Detection of Cytokine Receptors and Other Low-Abundance Membrane Molecules Unit 6.4 Enumeration of CD34+ Hematopoietic Stem and Progenitor Cells Unit 6.5 Immunophenotypic Analysis of Peripheral Blood Lymphocytes Unit 6.6 Immunophenotypic Analysis of Human Mast Cells by Flow Cytometry Unit 6.7 Measurement of CD40 Ligand (CD154) Expression on Resting and In Vitro–Activated T Cells Unit 6.8 Enumeration of Absolute Cell Counts Using Immunophenotypic Techniques Unit 6.9 Immunophenotypic Identification, Enumeration, and Characterization of Human Peripheral Blood Dendritic Cells and Dendritic-Cell Precursors Unit 6.10 Immunophenotypic Analysis of Platelets Unit 6.11 Immunophenotypic Analysis of PNH Cells Unit 6.12 Quantitative Flow Cytometric Analysis of Membrane Antigen Expression Unit 6.13 Immunophenotyping Using a Laser Scanning Cytometer Unit 6.14 Enzymatic Amplification Staining for Cell Surface Antigens Unit 6.15 Whole Blood Analysis of Leukocyte-Platelet Aggregates Unit 6.16 Flow Cytometric Assessment of HLA Alloantibodies Unit 6.17 Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood Unit 6.18 Identification of Human Antigen-Specific T Cells Using MHC Class I and Class II Tetramers Unit 6.19 ZAP-70 Staining in Chronic Lymphocytic Leukemia Unit 6.20 Multiparameter Analysis of Intracellular Phosphoepitopes in Immunophenotyped Cell Populations by Flow Cytometry Unit 6.21 Ten-Color Immunophenotyping of Hematopoietic Cells Unit 6.22 Flow Cytometric Screening for the HLA-B27 Antigen on Peripheral Blood Lymphocytes Unit 6.23 Immunophenotyping of Plasma Cells
Unit 6.24 Flow Rate Calibration for Absolute Cell Counting Rationale and Design
Chapter 7 Nucleic Acid Analysis Introduction Unit 7.1 Overview of Nucleic Acid Analysis Unit 7.2 Quality Control in Nucleic Acid Analysis Unit 7.3 Differential Staining of DNA and RNA Unit 7.4 Analysis of DNA Content and DNA Strand Breaks for Detection of Apoptotic Cells Unit 7.5 DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis Unit 7.6 Analysis of Nuclear DNA Content and Ploidy in Higher Plants Unit 7.7 Analysis of DNA Content and BrdU Incorporation Unit 7.8 Analysis of DNA Denaturation Unit 7.9 Bivariate Analysis of DNA Content and Expression of Cyclin Proteins Unit 7.10 Flow Cytometric Analysis of Reticulated Platelets Unit 7.11 Assessment of Viability, Immunofluorescence, and DNA Content Unit 7.12 Flow Cytometric Analysis of RNA Synthesis by Detection of Bromouridine Incorporation Unit 7.13 Sperm Chromatin Structure Assay for Fertility Assessment Unit 7.14 Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling and Multivariate Flow Cytometry Unit 7.15 Ultraviolet-Induced Detection of Halogenated Pyrimidines (UVID) Unit 7.16 Analysis of DNA Content and Green Fluorescent Protein Expression Unit 7.17 Analysis of Viral Infection and Viral and Cellular DNA and Proteins by Flow Cytometry Unit 7.18 Apoptosis Signaling Pathways
Unit 7.19 Flow Cytometry of Apoptosis Unit 7.20 Analysis of Fine-Needle Aspirate Biopsies from Solid Tumors by Laser Scanning Cytometry (LSC) Unit 7.21 Measurement of Cytogenetic Damage in Rodent Blood with a Single-Laser Flow Cytometer Unit 7.22 Analysis of Tissue Imprints by Scanning Laser Cytometry Unit 7.23 Cell Cycle Analysis of Budding Yeast Using SYTOX Green Unit 7.24 Detection of Mitotic Cells Unit 7.25 DRAQ5 Labeling of Nuclear DNA in Live and Fixed Cells Unit 7.26 Assessment of Telomere Length, Phenotype, and DNA Content Unit 7.27 Detection of Histone H2AX Phosphorylation on Ser-139 as an Indicator of DNA Damage (DNA Double-Strand Breaks) Unit 7.28 RNA and DNA Aptamers in Cytomics Analysis Unit 7.29 Nuclear DNA Content Analysis of Plant Seeds by Flow Cytometry Unit 7.30 Estimation of Relative Nuclear DNA Content in Dehydrated Plant Tissues by Flow Cytometry Unit 7.31 Assessment of Cell Proliferation by 5-Bromodeoxyuridine (BrdU) Labeling for Multicolor Flow Cytometry
Chapter 8 Molecular Cytogenetics Introduction Unit 8.1 Overview of Fluorescence In Situ Hybridization Techniques for Molecular Cytogenetics Unit 8.2 Basic Preparative Techniques for Fluorescence In Situ Hybridization Unit 8.3 Probe Labeling and Fluorescence In Situ Hybridization Unit 8.4 Immunocytochemical Detection Unit 8.5 Processing and Staining of Cell and Tissue Material for Interphase Cytogenetics
Unit 8.6 Advanced Preparative Techniques to Establish Probes for Molecular Cytogenetics Unit 8.7 Combination DNA/RNA Fish and Immunophenotyping Unit 8.8 Single-Nucleotide Sequence Discrimination In Situ Using Padlock Probes Unit 8.9 Tyramide Signal Amplification (TSA) Systems for the Enhancement of ISH Signals in Cytogenetics Unit 8.10 Molecular Combing Unit 8.11 Principles and Applications of PRINS in Cytogenetics Unit 8.12 Comparative Genomic Hybridization (CGH)—Detection of Unbalanced Genetic Aberrations Using Conventional and Micro-Array Techniques Unit 8.13 Combined Immunofluorescence and FISH: New Prospects for Tumor Cell Detection/Identification
Chapter 9 Studies of Cell Function Introduction Unit 9.1 Overview of Functional Cell Assays Unit 9.2 Assessment of Cell Viability Unit 9.3 Flow Cytometric Measurement of Intracellular pH Unit 9.4 Analysis of Intracellular Organelles by Flow Cytometry or Microscopy Unit 9.5 Reporters of Gene Expression: Enzymatic Assays Unit 9.6 Estimation of Membrane Potential by Flow Cytometry Unit 9.7 Oxidative Metabolism Unit 9.8 Measurement of Intracellular Calcium Ions by Flow Cytometry Unit 9.9 Intracellular Cytokines Unit 9.10 Assays of Natural Killer (NK) Cell Ligation to Target Cells Unit 9.11 Flow Cytometric Analysis of Cell Division by Dye Dilution
Unit 9.12 Reporters of Gene Expression: Autofluorescent Proteins Unit 9.13 In Vitro Invasion Assays: Phagocytosis of the Extracellular Matrix Unit 9.14 Flow Cytometric Analysis of Mitochondrial Membrane Potential Using JC1 Unit 9.15 Multiparameter Analysis of Physiological Changes in Apoptosis Unit 9.16 Signal Transduction During Natural Killer Cell Activation Unit 9.17 Assessment of Surface Markers and Functionality of Dendritic Cells (DCs) Unit 9.18 Stem Cell Identification and Sorting Using the Hoechst 33342 Side Population (SP) Unit 9.19 Assessment of Phagocyte Functions by Flow Cytometry Unit 9.20 Flow Cytometric Analysis of Calcium Mobilization in Whole-Blood Platelets Unit 9.21 Flow Cytometric Analysis of Cytokine Responses in Stimulated Whole Blood: Simultaneous Quantitation of TNF-α-Secreting Cells and Soluble Cytokines Unit 9.22 Optimized Whole-Blood Assay for Measurement of ZAP-70 Protein Expression Unit 9.23 Flow Cytometry of the Side Population (SP)
Chapter 10 Data Processing and Analysis Introduction Unit 10.1 Data Management Unit 10.2 Data File Standard for Flow Cytometry, FCS 3.0 Unit 10.3 Listmode Data Processing Unit 10.4 Multidimensional Data Analysis in Immunophenotyping Unit 10.5 Two-Dimensional Image Processing and Analysis Unit 10.6 Data Presentation Unit 10.7 Data Analysis Through Modeling
Unit 10.8 Multivariate Analysis Unit 10.9 Detection and Location of Hybridization Domains on Chromosomes by Image Cytometry Unit 10.10 Three-Dimensional Image Visualization and Analysis Unit 10.11 Image Processing and 2-D Morphometry Unit 10.12 Dial-In Flow Cytometry Data Analysis Unit 10.13 The Application of Data Mining to Flow Cytometry Unit 10.14 Intensity Calibration and Shading Correction for Fluorescence Microscopes Unit 10.15 A Software Method for Color Compensation
Chapter 11 Microbiological Applications Introduction Unit 11.1 Overview of Flow Cytometry and Microbiology Unit 11.2 Flow Cytometry and Environmental Microbiology Unit 11.3 Estimation of Microbial Viability Using Flow Cytometry Unit 11.4 Sorting of Bacteria Unit 11.5 Detection of Borreliacidal Antibodies by Flow Cytometry Unit 11.6 Flow Cytometric Detection of Pathogenic E. coli in Food Unit 11.7 Mycobacterium tuberculosis Susceptibility Testing by Flow Cytometry Unit 11.8 Antibiotic Susceptibility Testing by Flow Cytometry Unit 11.9 Determination of Bacterial Biomass from Flow Cytometric Measurements of Forward Light Scatter Intensity Unit 11.10 Flow Cytometry of Yeasts Unit 11.11 Enumeration of Phytoplankton, Bacteria, and Viruses in Marine Samples Unit 11.12 DNA/RNA Analysis of Phytoplankton by Flow Cytometry
Unit 11.13 Cell Cycle Analysis of Yeasts Unit 11.14 Flow Cytometric Assessment of Drug Susceptibility in Leishmania infantum Promastigotes Unit 11.15 Resolution of Viable and Membrane-Compromised Free Bacteria in Aquatic Environments by Flow Cytometry Unit 11.16 Functional Assays of Oxidative Stress Using Genetically Engineered Escherichia coli Strains Unit 11.17 Labeling of Bacterial Pathogens for Flow Cytometric Detection and Enumeration
Chapter 12 Cellular and Molecular Imaging Introduction Unit 12.1 Comparative Overview of Flow and Image Cytometry Unit 12.2 Basics of Digital Microscopy Unit 12.3 Modern Confocal Microscopy Unit 12.4 Time-Lapse Microscopy Approaches to Track Cell Cycle Progression at the Single-Cell Level Unit 12.5 Three-Dimensional Visualization of Blood and Lymphatic Vasculature in Tissue Whole Mounts Using Confocal Microscopy Unit 12.6 Quantitative Fluorescence In Situ Hybridization (QFISH) of Telomere Lengths in Tissue and Cells Unit 12.7 Detecting Protein–Protein Interactions with CFP-YFP FRET by Acceptor Photobleaching Unit 12.8 Measuring FRET in Flow Cytometry and Microscopy
Chapter 13 Multiplexed and Microparticle-Based Analyses Introduction Unit 13.1 Multiplexed Microsphere-Based Flow Cytometric Immunoassays Unit 13.2 Microsphere Surface Protein Determination Using Flow Cytometry
Unit 13.3 Use of Microsphere-Supported Phospholipid Membranes for Analysis of Protein-Lipid Interactions Unit 13.4 Multiplexed SNP Genotyping Using Primer Single-Base Extension (SBE) and Microsphere Arrays Unit 13.5 BeadCons: Detection of Nucleic Acid Sequences by Flow Cytometry Unit 13.6 Characterization of Nuclear Receptor Ligands by Multiplexed Peptide Interactions Unit 13.7 Detection of Gene Fusions in Acute Leukemia Using Bead Microarrays Unit 13.8 Reagents and Instruments for Multiplexed Analysis Using Microparticles
Appendix 1 Abbreviations and Useful Data 1A Abbreviations Used in this Manual 1B Common Conversion Factors
Appendix 2 Stock Solutions, Equipment, and Laboratory Guidelines 2A Common Stock Solutions, Buffers, and Media
Appendix 3 Commonly Used Techniques 3A Cell Counting 3B Techniques for Mammalian Cell Tissue Culture 3C Diagnosis and Treatment of Mycoplasma-Contaminated Cell Cultures 3D Wright-Giemsa and Nonspecific Esterase Staining of Cells 3E Techniques for Bacterial Cell Culture: Media Preparation and Bacteriological Tools 3F Growing Bacteria in Liquid Media 3G Growing Bacteria on Solid Media
3H Importing Biological Materials 3I Production of Polyclonal Antisera 3J Production of Monoclonal Antibodies 3K Enzymatic Amplification of DNA by PCR: Standard Procedures and Optimization
Appendix Suppliers Selected Suppliers of Reagents and Equipment
CHAPTER 1 Flow Cytometry Instrumentation INTRODUCTION
T
he purpose of this chapter is to acquaint readers with the instrumentation utilized in flow cytometry by describing various elements of the technology, and to provide detailed information on specific methods of using the instruments. Introducing the chapter is UNIT 1.1, an overview of the instrumentation involved in flow cytometry. This unit describes the various parts of a flow cytometer, explains the basics of how each component works, and presents a brief history of the development of flow cytometry instrumentation. Later units in the chapter describe specific aspects of the instrumentation. The fluidics system of a flow cytometer transports objects through the instrument, positioning them for accurate measurement. The various aspects of fluidics—from the sample tube to the interaction with the illumination beam, including flow analyzers and cell sorters— are described in UNIT 1.2. Calibration of an instrument for stability in measurement results (such that the same objects yield the same results day after day) requires frequent use of calibration particles. Calibration methods and sources of calibration particles are described in UNIT 1.3; this unit also includes a discussion of the difference between “standardization” and “calibration” and the need for both. Methods for establishing and maintaining the linearity of the system in order to make reproducible and accurate measurements are presented in UNIT 1.4. Another significant aspect of a flow cytometry system is the optical filters. Flow cytometers and flow sorters use light beams to excite fluorescent dyes; optical filters are used to separate the fluorescent light from the incident light. They are especially important with the current emphasis on multivariate analysis, which requires the simultaneous use of several dyes and light sources. UNIT 1.5 describes the basic principles of optical filters and discusses how to select a filter for a specific measurement in a cytometer. Most flow cytometers and sorters utilize laser beams as the excitation source; UNIT 1.6 discusses the various lasers that are in use and the various methods for delivering a laser beam to the objects being measured. A particularly valuable aspect of flow cytometry is its capacity for analyzing thousands of cells per second. For the detection and sorting of rare cells, special versions of the machines (high-speed sorters) have been developed. UNIT 1.7 discusses the principles and practices of operating a cell sorter to maximize its sorting rate, including the tradeoffs between speed and resolution. All commercial flow cytometers have a built-in capability for “gating,” a feature required for cell sorting. UNIT 1.8 describes the principles under which data are processed in real time and off-line to select subpopulations of cells (or cell organelles) with specific characteristics. Most flow cytometers utilize lasers as their illumination source. UNIT 1.9 describes the wide variety of laser types available and explains how they work. Included are pure and mixed-gas ion lasers as well as diode and other solid-state lasers (e.g., YAG lasers). The unit also discusses future directions, particularly in the development of new solid-state devices that might replace gas lasers. To ensure that high-quality, accurate data are obtained, flow instruments are checked for system performance each time they are used. UNIT 1.10 describes two levels of system Contributed by Bill Hyun Current Protocols in Cytometry (2003) 1.0.1-1.0.3 Copyright © 2003 by John Wiley & Sons, Inc.
Flow Cytometry Instrumentation
1.0.1 Supplement 26
alignment—routine alignment checks and complete alignment—that can be applied to a variety of instrumental configurations. The measurement of microbes is becoming an increasingly important application in flow cytometry. The small size of these organisms can create special difficulties that must be carefully addressed. UNIT 1.11 describes the problems encountered in making such measurements, the instrument requirements, and some approaches that can be taken to optimize the instruments for such measurements. This material is essential background for the protocols found in Chapter 11, Microbiological Applications. Fluorescence resonance energy transfer (FRET) is a technique that helps make possible the simultaneous use of three or more fluorescent dyes for the structural analysis of proteins in biological membranes. UNIT 1.12 describes the theory behind FRET, characterizes available parameters and instruments, discusses the method’s limitations, and presents a few examples of its applications. Light scatter was the first parameter measured in a flow cytometer. This highly useful parameter is probably the most widely used and least well understood by laboratories today. UNIT 1.13 describes in general terms the interaction of light with small particles, how the light is measured, and some current applications of this parameter in flow cytometry. In general, fluorescent dyes have relatively broad excitation and emission spectra, and when two or more are used simultaneously, the emission spectra will usually overlap to some extent. The measured fluorescence for a given dye can actually contain a significant contribution from any or all of the other dyes. To obtain accurate flow results it becomes necessary to compensate (correct) for this spectral overlap. UNIT 1.14 explains why compensation is necessary, how it is accomplished, and how it affects the visualization of data. The author addresses popular misconceptions that result in incorrect compensation and provides extensive discussion of suitable controls—cells, beads, and gates. The measurement of time-resolved fluorescence is a relatively new technique still under development in both flow and image cytometry, and promises to add a new dimension to multiparameter cytometry. Properly used, this technique can provide information about fluorophore/cell-interactions at the molecular level. UNIT 1.15 describes the theory behind fluorescence lifetime measurements and presents some recent applications. The simultaneous measurement of several fluorescent proteins allows one to monitor such things as gene expression and the intracellular localization of proteins. UNIT 1.16 presents a protocol for using a single laser operating at 458 nm to detect three fluorescent proteins simultaneously in a single cell. This is accomplished using a simple optical configuration and real-time compensation. Rapid screening of large combinatorial libraries, for instance of potentially valuable compounds in drug discovery tests, requires reliable automated sample-handling techniques. UNIT 1.17 describes one such technique, “plug flow cytometry,” in which precisely defined volumes of sample suspensions are injected at regular intervals into the flow stream. Many functions of living cells as well as molecular interactions are temperature dependent. UNIT 1.18 presents a protocol for using a Peltier module to control the temperature of a sample being processed in a flow cytometer/cell sorter. UNIT 1.19 covers in fine detail the photophysical and chemical properties of the common probes routinely employed in current flow cytometry. The spectral characteristics and use of example fluorescent dyes, probes, and labels for various functional assays and applications are described.
Introduction
1.0.2 Supplement 26
Current Protocols in Cytometry
Units covering other aspects of flow cytometry, including additional information about multispectral compensation in both flow and image cytometry and the use of micromirrors in new generation systems, will be presented in future updates to this chapter. Bill Hyun
Flow Cytometry Instrumentation
1.0.3 Current Protocols in Cytometry
Supplement 26
Overview of Flow Cytometry Instrumentation Flow cytometry is a technology in which a variety of measurements are made on cells, cell organelles, and other objects suspended in a liquid and flowing at rates of several thousand per second through a flow chamber. Flow sorting is an extension of this technology in which any single cell or object measured can be selectively removed from the suspension based on the measurements made. Flow cytometry is a very broadly applicable methodology. A brief list of applications that use flow cytometers includes: Disease diagnosis Chromosome karyotyping Cell function analysis Cancer therapy monitoring Detecting fetal cells Cell kinetics Identifying tumor cells Cytogenetics Fundamental cell biology. In a flow cytometer, cells in suspension are made to flow one at a time through a sensing region of a flow chamber (flow cell) where measurements are made. An example of an early flow cytometer is the Coulter counter (APPENDIX 3A). In this device, cells pass through a small orifice across which an electric current is flowing. As a cell enters the orifice, the flow of current is reduced because the cells are largely nonconducting. Electronic circuits detect the decrease in current and thus the presence of the cell. In this way the device can count the number of cells per second passing through the orifice, and because the volume flow rate can be measured one can determine the number of cells per milliliter of sample. The Coulter counter has been in use since 1949 and is still a mainstay of the clinical laboratory. Under the right conditions (e.g., size and length of orifice, current magnitude), the reduction in current through the orifice is proportional to the size (volume) of the cell, as demonstrated at the Los Alamos Scientific Laboratory in 1962. In modern flow cytometers, cells flow through a light beam rather than through a Coulter orifice; a Coulter orifice can, however, be included in these devices. Many different types of measurements can be made on the cells, based on the size and shape of the light beam and on the dyes used to stain components of interest. The light beam can come from arc lamps (e.g., mercury), as in early flow cytomeContributed by Phillip N. Dean Current Protocols in Cytometry (1997) 1.1.1-1.1.8 Copyright © 1997 by John Wiley & Sons, Inc.
UNIT 1.1
ters, or from lasers. Methods of measurement include absorption and scattering of the light beam by the cell, fluorescence of attached fluorescent dyes, and shape of the detected signal. Some of the properties and components that can be measured by a flow cytometer using these various methods are listed in Table 1.1.1. In principle, any component of a cell to which a fluorescent dye can be attached can be measured in a flow cytometer. If the binding of the dye is stoichiometric (i.e., amount of dye is proportional to amount of component) then the measurement can be quantitative and highly accurate (to within a few percent or better). Table 1.1.1 Properties and Components of Cells Measured in Flow Cytometry
Properties
Components
Cell diameter
DNA
Dye distribution Internal structure
Nuclear antigens Enzymes
Membrane potential Nuclear diameter
Protein RNA
Surface area Volume
Hormones Surface antigens
A flow cytometer is made up of several parts, as shown diagrammatically in Figure 1.1.1. All components of the system are necessary; the weakest part of the system defines its limitations. Other chapter units discuss the different parts of the system in detail. This overview describes the technology in general to give the reader a feeling for the interplay between the various parts of a flow cytometer. It also contains a brief history of the development of flow cytometry instrumentation.
CELL PREPARATION Objects to be measured must be suspended in a liquid. This is simple for blood cells, for example, but cells from tissue must be disaggregated and removed from any noncellular material. For most tissues this can be accomplished by procedures as simple as mincing the tissue with a knife and pulling cells through a 19-gauge needle into a syringe, followed by passing the cell suspension through a 200-mesh nylon screen. Details for such procedures are
Flow Cytometry Instrumentation
1.1.1
found elsewhere in this publication in units that deal with specific measurement and analysis protocols (e.g., see UNIT 5.2 for general procedures for handling, storage, and preparing human tissues and APPENDIX 3B for procedures for disaggregating cultured cell monolayers). After a single-cell suspension is obtained, the cells
are stained with dyes that bind to the specific features that are to be measured.
FLOW CHAMBER After staining, cells are made to flow one at a time through the interrogating light beam; a laser beam is illustrated in Figure 1.1.2. To
light source (UNIT 1.5 )
cell preparation
fluidics control
flow chamber (UNIT 1.2 )
(UNIT 1.2 )
sorter module (this unit, UNIT 1.2 )
detectors and signal processing (UNITS 1.3 &1.4 & Chapter 10 introduction)
analysis (UNIT 10.1)
display (UNIT 10.4)
Internet (UNIT 10.2 )
Figure 1.1.1 Schematic diagram of a complete flow cytometer system.
sample sheath
flow chamber
focusing lens
laser beam
Overview of Flow Cytometry Instrumentation
Figure 1.1.2 Longitudinal cross-sectional view of the flow chamber of a flow cytometer. The sample stream is surrounded by the sheath fluid which confines the cells (black dots) to the center of the chamber. The laser beam is focused onto the cell stream.
1.1.2 Current Protocols in Cytometry
obtain the best resolution, every cell must flow through the middle of the beam and be exposed to the same intensity of illuminating light. However, the laser beam has a Gaussian intensity distribution (i.e., the intensity is at a maximum in the center of the beam and decreases exponentially in the radial direction), and this puts a severe constraint on the stability of the flow stream. The system includes two features to alleviate this problem. (1) The beam leaving the laser has a circular cross section, and a long-focal-length cylinder lens is used to spread the beam in the horizontal direction and to produce a large depth of focus, resulting in a relatively large region of constant intensity in the center of the flow stream. (2) A “sheath” stream is introduced to the flow chamber. This sheath has a higher flow rate (∼5 ml/min) than the sample (∼100 µl/min), which serves to compress the sample stream and confine it to the center of the overall flow stream. This technique, called “hydrodynamic focusing,” is explained in more detail in UNIT 1.2. The end result is that cells are constrained to flow through an expanded laser beam in the center of the flow chamber. An additional constraint on the flow chamber is that it must be constructed of a material that will pass the excitation beam without appreciable scattering or absorption; this is usually accomplished through the use of quartz glass, which must be kept scrupulously clean. This is especially true when using ultraviolet light for excitation. The flow chamber can take many configurations. If a small orifice (e.g., sapphire jewel with a 70- to 100-µm hole) is placed at the
chamber exit, the flow stream will be compressed and will leave the chamber at high velocity. If the chamber is then vibrated at high frequency (e.g., 20,000 Hz), the stream will break up into uniform droplets and the flow cytometer will become a flow sorter. In this configuration measurements on cells can still be made in the chamber, although the time interval between cell detection and sorting can be relatively long. However, it is more common to pass the laser beam through the fluid stream just below the jewel before the stream breaks up (see Fig. 1.1.6). Then the interval between cell detection and sorting is shorter. In the latter configuration, the material requirements on the chamber are considerably reduced; the chamber becomes what is often called a sorter nozzle and can be constructed of ceramic materials. Because the hydrodynamic focusing does take place in the nozzle, in some sense the nozzle is a chamber. The sorting configuration is described in more detail later (see Sorting).
DETECTORS As a cell flows through the beam, light scattered by the cell and fluorescence light from dyes added to the cell are collected by light detectors, usually photomultipliers and photodiodes (see UNIT 1.4 for further discussion of photodetectors). These devices convert the light signal to an electrical signal that can be processed by the data processing and analysis unit. Photomultipliers, being very sensitive to light, are used where the light signal is weak (fluorescence), and photodiodes are used where the signal is strong (small-angle light scatter). The simplest flow cytometer would have per-
photomultiplier
filter pinhole photodiode collection lens laser beam flow chamber
Figure 1.1.3 Arrangement for a simple flow cytometer, containing a single fluorescence detector (photomultiplier) and a photodiode for detecting laser light scattered by a cell.
Flow Cytometry Instrumentation
1.1.3 Current Protocols in Cytometry
haps one photomultiplier and one photodiode, as shown in Figure 1.1.3. With the appropriate electronics system, this permits one to make two simultaneous measurements on a cell. As a cell flows through the beam it scatters some of the incident light, and the light scattering is typically detected by the photodiode, which is less sensitive than the photomultiplier. This continues as long as the cell is within the beam. Thus, the length of time a cell is in the beam (and the width of the electrical pulse produced) is proportional to the width of the cell. If the cell is also stained with a DNA-specific dye, the photomultiplier is used to measure the amount of fluorescent light emitted by the cell while it is in the light beam, producing a signal proportional to the DNA content of the cell. In Figure 1.1.3, an optical filter is shown that passes the fluorescent light and blocks the scattered excitation (laser) light. Thus, two measurements are made simultaneously. UNIT 1.5 contains a comprehensive discussion on how optical filters for flow cytometry are made and selected. By using a dichroic mirror (beam splitter) in front of the photomultiplier and incorporating a second photomultiplier with a different filter, as illustrated in Figure 1.1.4, three measurements can be made: e.g., DNA, total protein,
and narrow-angle light scatter. A dichroic mirror is one that reflects light below a specific wavelength and passes longer-wavelength light. The requirement for using more than one dye with this configuration is that both dyes excite at the same wavelength but emit at different wavelengths. The mirror is selected to separate the two emissions. Each detector also has a filter to block scattered excitation light. Fluorescent light is always emitted at a wavelength longer than that of the excitation light. Many flow cytometers today use two laser beams operating at different wavelengths to excite four or more dyes simultaneously. Figure 1.1.5 illustrates how this is done. The laser beams are separated vertically by ∼200 µm so that a cell flows through the two beams with a separation time of a few microseconds. Thus the two pairs of signals are separated in time, making it easier to resolve them. Each laser beam interaction point has its own pair of photomultipliers, dichroic mirror, and filter arrangement. In addition to measuring fluorescence, these detectors can be used to measure scattered light at 90°. The latter signal can help to distinguish cells with different internal structures. In principle, more detectors can be added to make even more measurements on each cell, with the limitation being the number of dye combina-
photomultiplier P2 filter P1
dichroic beam splitter
pinhole
photodiode collection lens laser beam flow chamber
Overview of Flow Cytometry Instrumentation
Figure 1.1.4 Arrangement for a flow cytometer with dual fluorescence detectors and a scatter detector. Light from two fluorescent probes is separated by the dichroic mirror and optical filters. With the appropriate filters, photomultiplier P1 can also be used to measure light scattered at 90° to the laser beam.
1.1.4 Current Protocols in Cytometry
tions that can be used. The combinations of excitation and emission spectra must be significantly different (see UNIT 1.5).
ANALYSIS All modern flow cytometers incorporate computers to monitor and in some cases to control the instrument, and to provide a capability for on-line analysis of instrument data. The computers are mostly Macintosh and IBMcompatible personal computers, which now have the power to perform virtually any kind of analysis desired. As the method of analysis required is not always known during an experiment, the computers are also used for off-line analysis. Software packages are available from the instrument manufacturers and from independent software companies. For more details on data processing and analysis, see Chapter 10. Flow cytometers are capable of producing enormous quantities of data very rapidly. This presents a challenge to the user, who must provide a means for storing the data in such a fashion that they can be recalled on demand. Because most data are stored in “listmode,” data files can be very large. “Listmode” means that every measurement on every cell is stored in a list. Thus, if five measurements are made on each of 50,000 cells, with a maximum value of 1024 per measurement (2 bytes), space has to be found for 500,000 bytes of information per sample. With the current development of
ever larger and less expensive storage devices such as read/write optical disk cartridges, this is not a major problem. A data file standard has been developed for the storage of flow cytometry data to make it possible for different laboratories to share data. This topic is discussed in UNIT 10.2. Sharing of data and the results of data analysis has become an important part of research; access to the Internet has become a desirable attribute of flow cytometer systems. To accomplish this one needs an Internet service provider and a “browser,” a computer program that provides access to other sites on the network. There are several browsers available, notably Mosaic, Netscape, and Internet Explorer. Many flow cytometry laboratories throughout the world have established sites on the Internet and made them available to other researchers in the field. A convenient location to begin a journey through the Internet is the home page of the International Society for Analytical Cytology (http://nucleus.immunol.washington.edu/ISAC. html), which contains links to most of these sites as well as to other sources dealing with both flow and image cytometry.
SORTING Principles A flow sorter is a cytometer with the additional capability of selectively removing from
beam 1 photomultiplier P1 beam 2 filter P2
beam splitter
flow chamber
P3
P4
half mirror
Figure 1.1.5 Flow cytometer with two excitation beams (lasers) that are separated vertically by 200 µm. A half mirror is used to direct fluorescent light from each beam interaction to a different pair of photomultipliers, each of which has a beam splitter and filter arrangement as in Figure 1.1.4. A photodiode could be added for each beam to permit a total of six measurements per cell.
Flow Cytometry Instrumentation
1.1.5 Current Protocols in Cytometry
the suspension of cells any selected cell flowing through it. The physical arrangement of a sorter is illustrated in Figure 1.1.6; the detailed fluidics are discussed in UNIT 1.2. Basically, the fluid containing the cells passes through a narrow orifice (∼100 µm in diameter) into the air. At the same time, the flow chamber is vibrated by the attached piezoelectric crystal, at frequencies on the order of 20,000 Hz. The vibration produces a disturbance in the ejected stream. The disturbance grows very rapidly, and the stream eventually breaks up into drops (i.e., 20,000 drops per second in the example given). The steady vibration causes the drops to be very uniform in size and spacing. Each cell that flows through the system will end up in a drop.
Measurements are made on the cells while they are either in the flow chamber or in the stream just below the orifice, before the disturbance of the stream has grown significantly. If the measurement result indicates that the cell is to be sorted, a voltage is applied to the stream just as the cell of choice reaches the end of the stream and a droplet is forming. When the drop separates from the stream it will carry electrical charge. The voltage on the stream is immediately reduced to zero so other drops will not be charged. Charges can be negative or positive, leading to the possibility of sorting two categories of cells simultaneously. As the drops continue to move downward they pass between two metal plates charged to a high voltage. Because
nozzle sapphire jewel laser beam
droplets
– –
deflection plates
–
charged droplets
– – collection beakers
Overview of Flow Cytometry Instrumentation
Figure 1.1.6 Diagram illustrating the principle of cell sorting. Cells flowing through the system are represented by small black dots. As cells to be sorted approach the end of the solid stream, a charge is applied to the stream. As the drop carrying the cell separates from the stream, the drop carries the charge. Passing between the high-voltage plates, charged drops containing desired cells are deflected into separate collection beakers. Deflection can be left or right, allowing for the simultaneous sorting of two classes of cells. In this illustration, two drops are sorted for each cell.
1.1.6 Current Protocols in Cytometry
the drops containing the selected cells are charged, they are deflected from the main stream of drops and collected in tubes or onto microscope slides for visual examination. Drops containing undesired cells are not charged and go directly into a waste tube.
Sort Purity Efficient and accurate sorting requires that the charge be applied just as the cell reaches the end of the stream. To compensate for variation in the flow velocity of the stream and to be certain the desired cell is sorted, typically two or three drops are sorted for each cell; Figure 1.1.6 shows two drops per cell. The sorting electronics are capable of detecting other cells
in the vicinity of the desired cell; if an unwanted cell might be sorted along with the desired cell, the sort is aborted. In some cases, particularly in the detection of very rare cells, one might want to accept some impurity in the sorted cell population to guarantee collection of the wanted cells. In that case, the sort purity requirement can be eased and the abort circuit disabled.
CHRONOLOGY OF FLOW CYTOMETRY DEVELOPMENT Table 1.1.2 is a list of significant events in the development of flow cytometry instrumentation. It is by no means comprehensive but illustrates the long history of the field. For a
Table 1.1.2
Development of Flow Cytometry
Year
Event
1934
Moldovan measures red blood cells in microscope with capillary flow and photodetector
1941
Kielland patents device like that developed by Moldovan
1947
Gucker uses Reynolds work to design laminar flow system for aerosols
1949
Coulter files for patent, “Means for Counting Particles Suspended in a Fluid”: the Coulter counter
1953
Crosland-Taylor designs system for aqueous suspension of cells flowing with a sheath: hydrodynamic focusing Parker and Horst apply for patent on device to do blood cell differentials using absorption of two colors of light
1956
“Model A” Coulter counter introduced
1962
Van Dilla uses Coulter counter to measure cell volume distribution
1965
Kamentsky measures UV absorbance and visible light scatter (500 cells/sec), generates bivariate plots Fulwyler develops electrostatic cell sorter, based on volume measurement Kamentsky develops fluidic switch cell sorter
1966
Van Dilla et al. introduce orthogonal measurements and laser excitation; prove DNA measurements were accurate; first to show discrete G1, S, and G2/M phase populations; show the possibility of quantitative cell kinetics studies
1968
Dittrich and Gohde patent microscope-based flow system with flow parallel to optic axis
1969
Hulett et al. introduce sorting based on cell fluorescence
1970
Wheeless et al. patent cell classification combining size and fluorescence
1971
Dittrich and Gohde introduce dual staining for DNA/protein, using ethidium bromide/FITC stains
1972
Wheeless et al. patent “slit-scan cytofluorometry” for automated cell recognition
1975
Gray et al. introduce flow karyotyping
1975 and beyond
Various investigators develop indexed sorting, high-speed sorting Flow Cytometry Instrumentation
1.1.7 Current Protocols in Cytometry
more complete discussion of the history, the interested reader is directed to Melamed et al. (1990).
SUMMARY Flow cytometry as a technology is still developing. New instruments with new or improved capabilities are constantly being introduced. This overview will be updated frequently to keep the reader apprised of developments in flow cytometry. A number of books provide excellent summaries of flow cytometry instrumentation (see Key References). For more details on particular techniques, the reader is referred to the articles in these books and to their extensive lists of references.
LITERATURE CITED Melamed, M.R., Lindmo, T., and Mendelsohn, M.L. (eds.) 1990. Flow Cytometry and Cell Sorting, 2nd ed. Wiley-Liss, New York.
Melamed, Lindmo, and Mendelsohn (eds.) 1990. See above. Second edition with all-new papers, producing true update of the first edition. Also contains extensive history of the field. Shapiro, H.M. 1995. Practical Flow Cytometry, 3rd ed. Wiley-Liss, New York. Comprises user’s reference manual for the laboratory. Also includes extensive list of current literature and list of key suppliers of instruments, parts, and reagents. Van Dilla, M.A., Dean, P.N., Laerum, O.D., and Melamed, M.R. 1985. Flow Cytometry: Instrumentation and Data Analysis. Academic Press, London. Contains papers by leading experts in the field on both subjects.
INTERNET RESOURCES http://nucleus.immunol.washington.edu/ISAC.html Homepage of the International Society for Analytical Cytology (ISAC).
KEY REFERENCES Melamed, M.R., Mullaney, P.F., and Mendelsohn, M.L. (eds.) 1979. Flow Cytometry and Cell Sorting, 1st ed. John Wiley & Sons, New York.
Contributed by Phillip N. Dean Livermore, California
First overall summary of the field, with many authors describing the state of the art as of 1979. Covers applications of the technology as well as instrumentation.
Overview of Flow Cytometry Instrumentation
1.1.8 Current Protocols in Cytometry
Fluidics
UNIT 1.2
Flow cytometry is so named because in this technique cells or subcellular particles suspended in a fluid are made to flow past sensors that take measurements from the cells. The primary function of the fluidics of a flow cytometric instrument is to deliver cells to the sensing area in single file and well aligned with the sensors. In a sorting flow cytometer, the fluidics must additionally be able to physically isolate cells chosen on the basis of measurements made by the instrument.
PRIMARY FLUIDIC FUNCTIONS Sample and Sheath Flow Figure 1.2.1 illustrates the main fluidic elements of a flow cytometer. Two primary lines feed fluid to the sensing region: the sample flow line and the sheath flow line. The sample flow line delivers the cell sample to the sensing region, and the sheath flow line provides a carrier fluid that helps position the sample flow and ultimately cells in the sensing region.
Hydrodynamic Focusing Most flow cytometry measurements are made optically, and it is important to keep particles well positioned in the flow stream in order to make accurate optical measurements. Flow cytometers use a fluidic method called hydrody-
namic focusing to control the position of particles in the flow (Fig. 1.2.2). In this technique, a flow of carrier fluid, called the sheath fluid, is established in the cytometer. The sheath fluid, which is usually normal physiological saline with perhaps a few additives, originates from a supply tank under pressure and flows through tubing to a sensing region where the detectors are located. Just before its arrival at the sensing region, the sheath fluid flows into a chamber of relatively large diameter, then out through a tapered conical section that reduces the diameter of the flow to the dimensions of the sensing region. The sample-containing fluid is introduced into the middle of this chamber through a tube positioned on the central axis of the sheath flow. Under laminar flow conditions, the sample and sheath fluids do not mix but join together to form a coaxial flow. This combined flow then passes through the tapered section, which reduces the diameter (and increases the velocity) of both the sheath and sample flows simultaneously before they reach the sensing region. This technique confines the cells to a very narrow central core so that the path cells follow through the sensing region is very consistent. The central area of combined flow that originated from the sample flow is called the sample core. It should be noted that the size of the sample
sample sheath pressure
sheath fluid
sample pressure
hydrodynamic focusing section sensing region
sheath tank
sample
sample tube
Figure 1.2.1 The basic flow system of a flow cytometer consists of sample and sheath forced under pressure through tubing to a hydrodynamic focusing section where the flows are combined in a coaxial flow prior to arrival at the sensing region. Flow Cytometry Instrumentation Contributed by Richard Stovel Current Protocols in Cytometry (1997) 1.2.1-1.2.7 Copyright © 1997 by John Wiley & Sons, Inc.
1.2.1 Supplement 1
sample sheath fluid sheath fluid
Figure 1.2.2 Flow cytometers use the principle of hydrodynamic focusing to align cells (represented here as black dots) in the center of the flow prior to their passage through the sensing region. The sample is injected into the middle of a sheath flow and the combined flow is reduced in diameter in a tapered section, forcing the cell into the center of the stream as shown.
to sensing region
core in relation to the diameter of the combined flow at the sensing area is a function of the relative sample and sheath flow rates, not the relative diameters of the sample and sheath introduction tubes. The effects of increasing or decreasing sample flow rate relative to sheath flow rate are shown in Figure 1.2.3. The reason that this point must be stressed is that the size of the sample core can be important for achieving data consistency. It is difficult optically to achieve spatially uniform illumination across the sensing region, so variation in particle position will result in variation of illumination intensity and therefore more variation in measurements due to position. Sample flow rates must be kept low relative to the sheath flow rate to maintain a narrow sample core and high data consistency.
Flow Cell
For most flow cytometers, the fluidic configuration of the sensing region can be classified as belonging to one of two types: jet-in-air or flow cell. Each type has its advantages and disadvantages. Some cytometers allow either type to be installed, and hybrids of the two types exist.
A second sensing configuration is the flow cell, in which the hydrodynamically focused coaxial flow passes through a transparent closed chamber wherein excitation light is brought into the chamber, and scattered and emitted light from the cells passes back out of the chamber to the detectors (Fig. 1.2.5). Generally, the inner diameters of flow cells are larger than the nozzles of jet-in-air systems, and the particles flow more slowly and spend more time in the sensing region. Higher-quality optical resolution can be achieved with flow-cell designs; moreover, the closed-flow system is advantageous when biohazardous samples are being handled.
Jet-in-Air
Hybrid
In a jet-in-air cytometer, the sheath and sample flow are combined in a nozzle, which tapers down to an orifice. The fluid in the nozzle is under enough pressure to form a continuous cylinder of fluid, or jet, as it emerges from the orifice (Fig. 1.2.4). Although some hydrodynamic focusing occurs in the tapered section, further focusing takes place as the fluid flow
Some cytometers combine advantages of both configuration types by sensing within a flow cell, then passing the flow through a jetforming orifice for droplet sorting. In this combination the exit section shown in Figure 1.2.5 is replaced by the orifice shown in Figure 1.2.4. This configuration provides a combination of high-quality optical measurement and droplet-
TYPES OF SENSING AREA
Fluidics
diameter narrows at the entrance to the orifice. The jet emerges from the orifice with the cells confined to the center of the jet. A light source, usually a laser, is focused onto the jet near the orifice, and optical elements collect the light scattered or emitted by cells as they pass through the light source and transmit this light to detectors, which convert it to an electronic signal. The jet-in-air configuration is well suited to droplet sorting (see discussion of Sorting).
1.2.2 Supplement 1
Current Protocols in Cytometry
A
sample sheath fluid sheath fluid
broad sample core
B
sample sheath fluid sheath fluid
narrow sample core
Figure 1.2.3 Diameter of the sample core is not dependent on the relative diameters of the sheath and sample tubes, but rather on the relative flow rates of the two fluids. (A) A sample at relatively high flow bulges after leaving the sample injection tube and is focused to a relatively broad sample core in the sensing region. (B) Low flow produces a narrow sample core that results in more accurate positioning of the cells in the sensing region.
sorting capabilities, but the sorting is less precise due to the longer transit time of the cells between sensing and sorting, and the biohazard containment advantage of a closed-flow system is lost.
Other Configurations In some cases flow cytometry techniques have been adapted to existing microscope technology by devising various flow configurations that use hydrodynamic focusing and pass cells through the focal plane of a microscope objective. One cytometer designed on these principles uses a jet-forming orifice to squirt the flow onto a microscope slide. The jet forms a continuous sheet of fluid on the slide, and flow conditions can be kept stable enough for cytometric measurements to be made through the microscope. Another type uses flow along the optical axis of the microscope; cells are measured as they pass axially through the focal plane of the microscope objective.
TYPES OF SAMPLE DELIVERY Standard Setup Samples of suspended cells are typically loaded into the flow cytometer in test tubes which are pushed on over an uptake tube that reaches to the bottom of the sample tube (see Fig. 1.2.1). The test tube pushes against a seal, and air pressure is applied through a port that projects through the seal. The pressure forces
the sample through the sample tubing to be combined with the sheath flow in the flow cell or nozzle. The pressure in the sample line at the point where the sample is injected into the sheath flow need be only marginally higher than the pressure in the sheath flow. Actual pressures applied at the sheath supply tank and the sample tube may differ by a larger amount due to pressure drops in the tubing and filters and differences in the height of the sample tube and sheath tank.
Alternative Techniques Other methods and devices, such as syringe pumps, are sometimes used to introduce samples into the cytometer. Automated delivery devices that can pick up and deliver samples sequentially from a rack of many samples are available. These are usually based on motordriven syringe pumps. There are also semiautomated pickup devices that are designed to pick up and deliver small-volume samples from microtiter plates. Special sample chambers are also available for certain specific purposes, such as mixing reagents with samples just prior to delivery for timed-reaction experiments.
POTENTIAL PROBLEMS AFFECTING SAMPLE FLOW Transient Flow Effects Liquid flowing through small tubing tends to travel slowly near the edges of the tubing,
Flow Cytometry Instrumentation
1.2.3 Current Protocols in Cytometry
Figure 1.2.4 A jet-in-air flow cytometer forces the coaxial flow of sheath and sample fluid through an orifice under sufficient pressure to form a jet. Illumination and detection take place in the jet after it has left the orifice.
orifice illumination
jet
due to drag by the walls, and faster in the center. The fluid develops a parabolic velocity profile in which the center of the flow travels twice as fast as the average flow. When a flow cytometric sample enters the sample tubing, the tubing is already filled with the sheath fluid that has been used to backflush the sample line. Because the outer fluid travels more slowly than the inner, the interface between sample and sheath fluid does not remain flat, but rather forms a curved surface; therefore, the first part of the sample to reach the sensing area will be what traveled in the center of the sample tubing, not the outside. This can lead to unusual effects. The first cells arriving at the sensing area may be more accurately positioned than following cells. Also, the first cells through have an opportunity to mix with and become diluted by the sheath fluid that was already in the tube. In situations where fluorescent dye is loosely associated with cells and the amount attached to cells depends on an equilibrium with the amount of free dye in the sample, this first-cell-through effect can cause measurements to vary over time until a stable sample flow that completely fills the sample tubing is achieved.
Settling
Fluidics
Cells will tend to settle out of suspension at a rate that varies with cell size and density. A typical settling rate for 10-mm-diameter lymphocytes is ~1 cm/hr. Thus, cells in a tube mounted on a cytometer for long periods require occasional resuspension. If the flow rate is very low, settling in the sample delivery tube can also be a concern.
The adherence properties, or stickiness, of cells influence their tendency to adhere to tubing walls or to each other. Differential settling of cells with different properties, in either the sample tube or the delivery tubing, can lead to changes in the distribution of cells reaching the sensing area.
Clogging Clogging of the small-diameter sensing region is frequently a problem in flow cytometry, especially in the case of jet-in-air systems, which have small jet-forming orifices; to avoid this problem, filtration of the cell sample is often advisable. Instruments with closed-flow sensing and no sorting orifice are more forgiving, because flow cells typically have a larger inner diameter than does the orifice of a sorting machine and therefore do not clog as easily. Filters are typically installed in the sheath fluid line both to guard against particulates that might clog the orifice or flow cell and to sterilize the fluid. Cell samples may or may not require filtration as they are introduced into the flow system of the cytometer, depending on the reliability of the sample preparation and the tendency of the cells to aggregate. If a filter is used, it can be a source of cross-contamination between samples; where high purity is needed, such as during reanalysis of sorted samples, the filter should be changed between samples.
Random Cell Arrival In a well-mixed cell suspension, the cells are, in principle, randomly distributed throughout the sample. Although it would be advanta-
1.2.4 Current Protocols in Cytometry
Figure 1.2.5 In a flow cell, the sensing region is contained within a chamber with transparent sides. Although not shown here, the cell usually has a square or rectangular cross section. The flow cell optical configuration may be combined with jet-in-air sorting by replacing the exit section with a jet-forming orifice.
illumination
transparent flow cell
geous to control the interval at which cells arrive at the sensing region, there is no way to do so, and in fact cell arrival time tends to follow a random distribution called the Poisson distribution. A number of effects may cause the actual arrival rate to deviate from the theoretical Poisson distribution: for instance, cells in the sample tube may not be mixed adequately, or may adhere to each other and tend to clump. The random arrival of cells at the sensing area presents problems for the signal processing electronics of the cytometer. A cell may arrive too soon after the previous cell so that the cytometer is not ready to measure it, or even worse, two cells may be so close together that the cytometer sees them as one. Inevitably, a certain percentage of measurements cannot be made satisfactorily. Such problems are alleviated if the average cell arrival rate is kept low, but this means that samples take longer to process.
SORTING The utility of flow cytometry in scientific experimentation is greatly enhanced by the ability of the instruments to isolate, or sort, cells on the basis of measurements made by the device. Some flow cytometers have this added capability and others do not. Most sorting flow cytometers use the electrostatic drop deflection method, which employs a jet-in-air configuration; some use other fluidic methods.
Electrostatic Drop Deflection A liquid jet in air emerges from its orifice as a column of fluid, but surface tension eventually causes it to break up into drops. If the jet is allowed to break up on its own, the sizes of the resulting drops will vary randomly over some range. In a sorting instrument, this breakup is brought under precise control by applying a periodic vibration at the orifice: this causes the breakup to occur in a very regular way so that the stream of drops is very uniform. The vibration is produced by means of a piezoelectric transducer attached to the nozzle: a periodic electrical signal applied to the transducer causes a small periodic variation in the diameter of the jet. Surface tension amplifies this variation as the jet progresses, and eventually the wave grows big enough to sever the jet, creating drops. The vibration may be applied either to the nozzle as a whole or to the fluid inside the nozzle. The main features of a sorting jet are illustrated in Figure 1.2.6. If the cell-sensing region of the cytometer is in the jet, a band of light is reflected and scattered perpendicular to the jet. The presence of a small surface wave on the jet causes this light band to be deflected up or down periodically by an amount that depends on the strength of the vibration that is applied to the jet. This effect causes additional optical noise that must be blocked in the measurement optical system usually accomplished with an obscuration bar. The vibration amplitude required to initi-
Flow Cytometry Instrumentation
1.2.5 Current Protocols in Cytometry
illumination
Figure 1.2.6 A jet-in-air sorter generates drops by vibrating the jet at a suitable frequency. Drops form with a uniform separation distance, called the drop wavelength.
satellite drop
drop wavelength
Fluidics
ate breakup of the jet by surface tension is quite small, however, and the optical noise problem is manageable. After measurements are made on a cell passing through the sensing region, the cytometer makes a decision on the basis of these measurements whether or not to sort the cell. The cytometer keeps track of the timing of desired cells, and when a desired cell arrives in the breakoff region of the jet, an electrical charging pulse is applied to the jet (which is an electrically conducting fluid) in such a way that the drop that contains the cell becomes charged and surrounding drops do not. The drop stream then passes through a strong, steady electric field that deflects the charged drops containing desired cells out of the stream of drops and into a collecting vessel. A drop may be either positively or negatively charged, allowing two populations to be sorted simultaneously, one to each side of the uncharged drop stream. In order to isolate chosen cells successfully, the rate at which cells arrive at the sensing region must be kept well below the drop formation rate. Otherwise, the probability of an unwanted cell being in the same drop as a wanted cell would be unacceptably high. Be-
cause sorting speed often limits the scope of experiments that can be performed with sorted cells, it is advantageous to generate drops at as high a frequency as possible. Drops are produced at the rate of the imposed vibration, but physical principles limit the size of the drops that can be produced by a jet of a particular diameter. For a cylinder of fluid, such as a sorting jet, the shortest section of fluid that can be made to form drops is a wavelength of about three times the diameter of the jet. A wavelength of 4.5 times the diameter is most favorable for drop formation. The frequency of the applied vibration must remain within these bounds. Given this limitation, there are only two ways to increase the rate of drop formation: to use a smaller-diameter jet (smaller orifice size) or to increase the velocity of the jet (higher nozzle pressure). Orifice size is limited by the tendency of small orifices to become clogged and by the size of the objects being sorted. Objects with diameters approaching that of the orifice will interfere with sorting in two ways: they will interfere with the regular formation of the surface wave at the orifice, and with the regular breakup of the jet at the breakoff point. Jet velocity is limited by the ability of cells to
1.2.6 Current Protocols in Cytometry
withstand the mechanical rigors of the sorting process and by the ability of the fluidic plumbing to withstand the higher pressures required. For historical reasons, commercial sorters have been limited to operating pressures of ~1 atm (15 psi) and nozzle sizes of ~70 to 100 µm, which meant that drop frequencies have been limited to ~25,000–35,000/sec. More recently, sorters have become available that operate at significantly higher pressure and correspondingly higher jet velocity and drop frequency. Sorting jets have the interesting property of forming a satellite drop between the main drops. Normally this smaller drop soon merges with an adjacent larger drop and is of little consequence. Under some circumstances, however, it may not merge, and a separate stream of smaller satellite drops may be formed; this can have the undesirable effects of interfering with sorting and causing an additional biohazard.
Closed-System Mechanical Sorting An alternative sorting method available in some commercial machines employs a mechanical actuator situated downstream from the sensing region of a closed-system flow cell. The actuator moves a collection tube into the flow of particles and picks off desired cells as they flow by. This method has the significant advantage of maintaining a closed flow path, which makes it more suitable for biohazardous experiments than jet-in-air methods. Its disadvantages include a slower sorting rate and a more dilute sorted fraction.
OTHER FLUIDIC FUNCTIONS In addition to its primary functions of delivering samples to the sensing region and combining sample and sheath flow for hydrodynamic focusing, the fluidic system of a flow cytometer generally has secondary functions built in. A boost function advances the sample from the sample tube to the sensing region at high speed at the start of a sample run to reduce the waiting time before the leading edge of the sample reaches the sensing region. This is done by switching the sample pressure momentarily to a higher level. Depending on the cytometer, this may be done either automatically or by the operator by means of a boost air-switch on the flow-control panel.
Another function that is often provided is a fill function, which allows the fluidic system to be filled more quickly after it has been drained. To do this, an extra port near the sensing region is opened to allow fluid from the sheath tank to enter the system at a much faster rate than if the sheath were flowing normally through the small-diameter sensing region. It is important to backflush the sample line between samples to remove all trace of the previous sample from the fluid system before introducing the next one. To do this, the uptake end of the sample line is left open to the atmosphere while the sheath flow and pressure is maintained at the sensing end. The resulting pressure differential causes sheath fluid to flow backward through the sample line, clearing out the remaining sample. Waste removal capability is provided to receive the sample after it has passed the sensing region and to remove fluid during backflushing. Waste fluids may drain into a waste tank under gravity or may be sucked into the waste lines by means of a vacuum applied to the waste tank. Jet-in-air systems usually have an aspirator to catch the unsorted droplet stream.
KEY REFERENCES Kachel,V., Fellner-Feldegg, H., and Menke, E. 1990. Hydrodynamic properties of flow cytometry instruments. In Flow Cytometry and Sorting (M. R. Melamed, T. Lindmo, and M.L. Mendelsohn, eds.) pp. 27-44. Wiley-Liss, New York. Excellent review of published work in fluidics relevant to flow cytometry. Pinkel, D. and Stovel, R. 1985. Flow chambers and sample handling. In Flow Cytometry: Instrumentation and Data Analysis (M. A. Van Dilla, P.N. Dean, O.D. Laerum, and M.R. Melamed, eds.) pp. 77-128. Academic Press, London. Detailed treatment of both theoretical and practical flow cytometric fluidics. Shapiro, H.M. 1995. Practical Flow Cytometry, 3rd ed. Wiley-Liss, New York. Includes thorough description of fluidics as well as many other aspects of flow cytometry.
Contributed by Richard Stovel Stanford University Stanford, California
Flow Cytometry Instrumentation
1.2.7 Current Protocols in Cytometry
Standardization, Calibration, and Control in Flow Cytometry Standardization, control, and calibration provide different degrees of certainty about the data acquired with an instrument. Each process is aimed at assuring that results from the instrument have the quality required for the intended purpose (Horan et al., 1990; NCCLS, 1998a; Muirhead, 1993a,b; Schwartz and Fernandez-Repollet, 1993; Owens and Loken, 1995; Schwartz et al., 1996; Owens et al., 2000). The purpose may be an individual research experiment or a clinical result that determines the course of patient treatment. In the terminology used in this commentary, an instrument is standardized at certain time points and subsequently operated under quality control conditions (see UNIT 3.1–3.2). These processes maintain the instrument within predetermined bounds and ensure that results will vary only within certain limits. If a result is also calibrated when the instrument is standardized, then future results can be objectively and quantitatively compared with those from other laboratories. Quantitation of results should be considered. Most results from flow cytometers are expressed either in terms of “percent positive” or in qualitative terms such as “dim” or “bright.” These terms are relative: what is considered “negative,” “dim,” and “bright” in one laboratory may be quite different in another laboratory. When visualizing fluorescence using a fluorescence microscope, such relative terms are necessary. Flow cytometers can measure the amount of fluorescence and provide more objective criteria for expressing results. As flow cytometers are designed to measure particle characteristics (see UNIT 1.1 for an overview of flow cytometry), particles are the most common materials used to calibrate, control, and standardize the instruments. This commentary describes how various types of particles are used for these purposes. It also briefly reviews the status of standardization and quality control for flow cytometry (see Chapter 3 for further discussion of quality control). UNIT 1.4 covers calibration of detection system components (e.g., linear and logarithmic amplifiers) to ensure linearity of the flow cytometer response. The first section of this unit focuses on how the term “standard” has been used in flow
UNIT 1.3
cytometry (see Standards, Standardization, and Jargon). The intent is to alert readers of flow cytometry literature that they must always interpret critically how “standard” is being used in a particular context. The next section defines terms and also includes comments to put the term in context or to highlight issues (see Definitions). After providing extensive background on particle types and cautions (see Overview of Standardization in Flow Cytometry), this unit describes practical aspects of methods to standardize and calibrate flow cytometers (e.g., in terms of optical alignment, fluorescence and light scatter resolution, and sensitivity; see Standardization and Calibration section). Finally, suggestions are given for analyzing particles used as calibrators, including how to assign to fluorescent beads a value for molecules of equivalent soluble fluorochrome (MESF) and how to determine the inherent fluorescence coefficient of variation (CV) of a dim bead sample (see Characterizing Particles for Calibration and Control of a Flow Cytometer).
STANDARDS, STANDARDIZATION, AND JARGON It is common in flow cytometry to combine words that describe use of a particle with the word “standard.” Examples are “calibration standard” and “alignment standard” (Horan et al., 1990; Schwartz and Fernandez-Repollet, 1993; Shapiro, 2003; Schwartz et al., 1996). Rarely is there any indication of who has set the “standard” and by what authority or consensus. There can be many levels of “standards,” depending on the size and authority of the group that establishes them. For example, an individual laboratory or investigator may have standard practices or materials. A large clinical or research study may have standard practices and materials that are agreed to by all investigators involved in the study. A professional organization may establish standard methods or identify standard materials for specific purposes. If the word “standard” is not modified by a term such as “laboratory,” “clinical trial,” or “study XYZ,” it may imply something that is generally and widely accepted by acknowledged authorities. In that authoritative Flow Cytometry Instrumentation
Contributed by Robert A. Hoffman Current Protocols in Cytometry (2005) 1.3.1-1.3.21 C 2005 by John Wiley & Sons, Inc. Copyright
1.3.1 Supplement 32
sense, however, there are few “standards” in flow cytometry. Clear and common understanding of what is meant by a term is important, especially as flow cytometry is used by increasing numbers of investigators. The verb “standardize” means to cause to be without variation. Early use of the noun “standard” in flow cytometry seems to have been in the sense of a particle used to standardize (make consistent) one instrument in one laboratory (Fulwyler, 1979). This is much different from the authoritative sense of “standard.” In this commentary, other terms are used to describe more specifically what type of particle or material is being used for a particular purpose. For example, “calibration particle” or “calibrator” is used instead of “calibration standard,” and “alignment particle” rather than “alignment standard.”
DEFINITIONS
Standardization, Calibration, and Control in Flow Cytometry
Concern with terminology and its evolution is not just semantics, but reflects what has been important in flow cytometer technology and how the technology has grown and changed. More precise and generally accepted terminology should clarify communication and understanding among flow cytometrists as well as scientists in other fields. A very useful set of definitions that generally cover the field of clinical diagnostics is NCCLS publication NRSCL8-A, Terminology and Definitions for use in NCCLS Documents (NCCLS, 1998a). The definitions for limits below are from the NCCLS definitions. Additional definitions related to quantitative fluorescence cytometry are provided by Henderson et al. (1998). The definitions below should be considered a reasonable point along the way toward authoritative and broadly accepted and understood terminology. Some definitions include comments and references that may help put them in context. Accuracy: degree to which a measurement agrees with the true or expected value. Alignment particle: particle with uniform size, fluorescence, and light scatter characteristics that is used to check the alignment (or, in some instruments, adjust the alignment) of the excitation and emission optics in the flow cytometer. It is desirable that the alignment particle emits fluorescence in all detector channels, as this allows all channels to be checked simultaneously. Alignment of the optics is optimal when signals from the particles have maximum intensity and minimum variation or CV. The more uniform the particles, the better the degree to which small deviations from optimal
alignment can be detected. Optimal alignment is most critical for measuring DNA, because of the very low inherent variation in DNA content from cell to cell. Antibody binding capacity (ABC): number of antibodies of a particular type that can bind to a cell under saturating staining conditions. Autofluorescence: inherent fluorescence from a cell or particle to which no stain or fluorochrome has been added. Manufactured particles (such as plastic beads) can be prepared to have nearly the same autofluorescence as lymphocytes. Background (noise, fluorescence, scatter): signal present when no particles are flowing in the sample stream. Background noise is one factor that limits the sensitivity of fluorescence detection (see definitions of fluorescence sensitivity and light-scatter sensitivity below). Depending on how low are the signals that one is trying to detect in the sample, different factors are dominant contributors to the background. When no light is coming from the flow cell (e.g., lasers turned off), detector noise is the background limit. For photomultiplier tubes (PMTs), the detector background noise is called dark current and is due to random emission of electrons from the photocathode. For photodiodes and other solid-state detectors, which have no or low signal amplification, the limiting factor under best conditions is noise from the amplifier required to raise the signal to a useful level. Sources of fluorescence noise include Raman scatter from water and optical components; fluorescence from unbound fluorochrome, reagent, or contaminants in the sample or sheath stream; and fluorescence from optical components. Calibration: process of adjusting an instrument so that the analytical result is accurately expressed in some physical measure. Calibrator: material that has been manufactured or assayed to have known, measured values of one or more characteristics. The assayed values are provided with the material. Fluorescent manufactured particles can be assayed for diameter or for the amount of fluorescence they produce. A practical measure of particle fluorescence is the number of fluorochrome molecules in solution that produce the same amount of fluorescence as one bead (see definition of MESF). Coefficient of variation (CV): statistical measurement of the broadness of a distribution of values, usually defined as CV = σ /µ, where the standard deviation σ = [(xi − µ)2 / (N − 1)]1/2 , with the sum over N measurements of xi (where xi is the ith measurement of
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variable x), and the mean µ = ( xi )/N. Shapiro (2003) gives an excellent discussion of CV and other, more robust statistics for flow cytometry. Another excellent reference for statistical methods is Bevington (1969). Control particle or material: stable material (e.g., sample of manufactured particles) that gives reproducible results when analyzed. Particles used to set up a flow cytometer are used as a control even if they do not have an assayed value assigned to a physical characteristic. Controls can be used to monitor the stability of an instrument and determine whether it is acceptably within calibration. A calibrator can be used as a control material, but a control material does not have to have an assigned value for a characteristic. Control sample: sample prepared in the same or nearly same way as a test or unknown sample and which should give an expected, predetermined result. In immunofluorescence analysis a positive control sample may use known cells (characterized for reactivity to a panel of antibodies) and the same antibody reagents as the test sample. A negative control sample may use the test cells but without antibody reagent or with an irrelevant antibody reagent. Fluorescence sensitivity: In flow cytometry there are two different aspects to the notion of sensitivity: threshold and resolution. The first has to do with the smallest amount of light that can be detected (Wood, 1993; Owens and Loken, 1995; Schwartz et al., 1996; Shapiro, 2003). This notion has also been given the name “detection threshold” (Schwartz et al., 1996). The second has to do with the ability to resolve dimly stained cells from unstained cells in a mixture (Brown et al., 1986; Horan et al., 1990; Shapiro, 2003). These concepts do not measure the same thing. The second notion incorporates a measure of the broadness of the fluorescence distributions for dim and unstained particles, not just the average fluorescence. Two instruments can have the same detection threshold but differ significantly in ability to resolve a dimly stained population. This is illustrated by example later (see Standardization and Calibration Section). 1. Degree to which a flow cytometer can measure dimly stained particles and distinguish them from a particle-free background (threshold). Threshold is important when the mean fluorescence of a dimly fluorescent population is measured. The greater the number of particles analyzed, the more accurately and precisely will the mean fluorescence be measured.
2. Degree to which a flow cytometer can distinguish unstained and dimly stained populations in a mixture of particles (resolution). Resolution is important for immunofluorescence analysis of subpopulations and is strongly affected by the measurement CVs for dim and unstained particles. Inherent sample CV: actual variability in the characteristics of a sample; for example, the actual variation in the amount of fluorochrome per bead in a sample of beads. Because the measurement process is not perfect and itself adds variation, the CV of the measured fluorescence will be greater than the inherent sample CV. The inherent CV of a sample can be estimated within a small uncertainty if the measurement variability added by the flow cytometer is well characterized (see Determining Inherent Fluorescence CV of a Dim Particle Sample). Light-scatter sensitivity: degree to which small particles can be detected above “particlefree” fluid. In practice, forward-scatter sensitivity is usually limited by optical noise caused by the excitation source, and side-scatter sensitivity is usually limited by submicron particles in the sheath fluid. Limit of detection: the lowest amount of analyte in a sample that can be detected but not quantified as an exact value. Limit of quantitation: the lowest amount of analyte in a sample that can be quantitatively determined with acceptable precision and accuracy under stated experimental conditions. Manufactured particles (beads, plastic beads, latex particles, microspheres, microbeads): particles made of synthetic polymers (plastics). Sizes range from submicron to over 100 µm, which generally covers the range of cells analyzed in flow cytometry. Most manufactured particles are made by bulk polymerization, but very uniform beads can be made employing the same droplet generation principle used for flow cytometric cell sorting (Fulwyler et al., 1973). Colored or fluorescent particles can be made by staining the beads with dyes or fluorochromes. Nonfluorescent beads, as well as many fluorescently stained beads, seem to be stable for many years. Two methods, namely, solvent (or “hard”) dying and surface staining, are used to stain particles. In solvent staining, non-water-soluble dyes are mixed with the particles in an organic solvent. The particles take up the dye and are then suspended in aqueous solution. The dye is trapped in the beads, which essentially become a “hard-dyed” plastic material. In some cases, hard-dyed particles can be synthesized
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directly using fluorescent monomers (Rembaum, 1979). As most dyes or fluorochromes used to stain cells are water soluble, solvent staining cannot generally be used for them. When solvent staining is possible for watersoluble fluorochromes, the spectral characteristics can differ significantly from those of fluorochrome in aqueous solution. Surface staining allows many common fluorochromes— especially those used as tags on fluorescent antibodies—to be used for particle staining. In this case a chemical group on the particle surface (e.g., amino group) is covalently bound to a reactive group on the fluorochrome. MESF (molecules of equivalent soluble fluorochrome): measure of particle fluorescence in which the signal from a fluorescent particle is equal to that from a known number of molecules in solution. This is a practical measure because a known concentration of particles can be compared directly with a solution of fluorochrome in a spectrofluorometer (see Calibrating Particle Fluorescence in MESF). Nonfluorescent particle: particle whose fluorescence distribution is the same as that of a particle-free sample. In practice, the concept of nonfluorescence is dependent on the sensitivity of the instrument making the measurement. A particle that is not measurably fluorescent in one instrument may be so in a more sensitive instrument. Fluorescence (or other luminescence or Raman scatter) from otherwise unstained manufactured particles depends on the material and treatment with which the beads are made. With all other factors equal, the “fluorescent” signal from microbeads will be proportional to the volume of a single bead. Precision or reproducibility: degree to which repeated measurements of the same thing agree with each other. In flow cytometry, precision of a measurement is estimated by the CV obtained when measuring a sample of particles (biological or nonbiological) with very uniform characteristics. Resolution: degree to which a flow cytometry measurement parameter can distinguish two populations in a mixture of particles that differ in mean signal intensity. Fluorescence sensitivity (see above) can be considered a special case of fluorescence resolution for which the signals are very dim. Note that the resolution will appear different when data are acquired and/or displayed on a logarithmic rather than linear intensity scale. Depending on the maximum number of channels into which the signal intensity is acquired (e.g., 256 or 1024),
a logarithmic display of the data may not have sufficient resolution to display populations that can actually be resolved by the instrument. Standard: 1. noun. a. acknowledged measure of comparison for quantitative or qualitative value. b. something recognized as correct by common consent or by those most competent to decide. 2. adj. a. serving as a standard of measurement or value. b. commonly used and accepted as an authority. Standardize: a. cause to conform to a given standard. b. cause to be without variation. Test pulse–triggered background fluorescence: measurement of background fluorescence in a flow cytometer by using an electronic pulse to trigger the pulse detection electronics and acquire data from the fluorescence detector(s) (see UNIT 1.4). As no particle is present to emit light, the fluorescence signals acquired are due only to instrument background light and noise and thus establish the lowest signal that can be measured. The duration of a test pulse usually simulates a signal from a particle of typical size. Larger particles would have signals of longer duration and produce more background signal and noise. If equipped with a test pulse function, the flow cytometer can provide a measurement equivalent to running a sample of truly nonfluorescent particles. The background fluorescence distribution produced by a test pulse should provide a measure of the “detection threshold” described by Schwartz et al. (1996). In many instruments the test pulse signal produces a pulse of light from a light-emitting diode that is detected and processed by one detector. When the test pulse signal is applied only to the forward scatter detector, the response of all other detectors to background light and noise can be measured. When a sample is run under normal conditions, any signal from particles above this background and noise level actually comes from the particles. There is no guarantee, however, that the particle signal is from particle fluorescence; for example, light scattered by the particles may not be totally blocked by the optical filters, or in some cases the light scatter may actually induce the filter to fluoresce. The possibility of scatter-induced “fluorescence” signal can be checked by running unstained cells and looking for a signal in the fluorescence channel. Because such a signal can also come from autofluorescence, one should also look at the side scatter versus fluorescence histogram.
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OVERVIEW OF STANDARDIZATION IN FLOW CYTOMETRY Standardization (see Definitions) is the foundation of flow cytometry and allows investigators to have confidence in instrument performance. This section surveys characteristics of particles used in flow cytometry, for example, to standardize immunofluorescence and to check alignment and measurement precision (see Types of Particles). Specific types of particles are compared. Standardization can be complicated, however, by factors other than particle type (see General Cautions for Using Particles in Standardization and Calibration; see What the Instrument Cannot Control: Sample, Reagent, and Data Analysis), but prospects for formalizing flow cytometry standards are encouraging (see Standard-Setting Organizations). The next section (see Standardization and Calibration) reviews various parameters of flow cytometers that can be standardized, such as resolution and sensitivity, and the final section (see Characterizing Particles for Calibration and Control of a Flow Cytometer) describes procedures and cautions for characterizing particles.
Types of Particles Manufactured particles and biological particles may be used to standardize flow cytometers. Beads may be spectrally matched to the fluorochromes used to stain cells, or they may simply fluoresce to a useful extent in the spectral range of interest. Spectrally matched beads allow standardization or even calibration across instruments that do not have exactly the same emission filters and/or excitation wavelengths. Biological particles may be stained with the same fluorochromes used in experiments to stain cells. Examples of data for some of the more common types of particles follow. Classification schemes for various types of particles used for standardization in flow cytometry have been proposed (Schwartz et al., 1996, 1998).
Comparison of spectrally matched and unmatched fluorescent particles Figures 1.3.1 and 1.3.2 show emission spectra from three types of particles: fluorochrometagged beads (CaliBRITE beads, BD Biosciences), stained with either fluorescein isothiocyanate (FITC) or phycoerythrin (PE); broad-spectrum hard-dyed beads (Rainbow beads, Spherotech); and glutaraldehyde-fixed chicken red blood cells (gCRBC, BioSure).
Figure 1.3.1A compares FITC-stained CaliBRITE beads with Rainbow beads and gCRBC, which are not spectrally matched to FITC. Figure 1.3.2A makes the same comparison with PE-stained CaliBRITE beads. Figure 1.3.1B compares quantitatively the fluorescence signal of each particle through filters of differing spectral bandwidth placed in front of PMT1, with data being normalized to the signal from FITC CaliBRITE beads for each filter. The gCRBC varied by about 35% over the range of filters used. Rainbow beads, however, varied by nearly 200% with the same filters. The differences in relative fluorescence with different filters should be considered when comparing different instruments. Even for a particular flow cytometer type or model, filters vary slightly due to manufacturing tolerances. Figure 1.3.2B shows a similar comparison for the relative fluorescence with different filters placed in front of PMT2. In this case, both gCRBC and Rainbow beads vary only slightly from PE-stained CaliBRITE beads. The fluorescence could be standardized with a maximum difference of 40% with any of these particles.
“Nonfluorescent” and autofluorescent particles Figure 1.3.3 shows emission spectra for particles with very low fluorescence. Unstained CaliBRITE beads have fluorescence comparable to autofluorescence from lymphocytes. Osmium-fixed chicken red blood cells (CRBC) had no fluorescence detectable above background in the fluorometer. Such “negative” particles are useful for estimating how well low level signals can be detected, as discussed later (see Sensitivity or Signal/Noise for Dim Fluorescence).
Comparison of particles for standardizing immunofluorescence analysis Figure 1.3.4 shows light-scatter dot plots (panel A) and green (515 to 545 nm) fluorescence histograms (panels B-F) for several types of particles used to standardize flow cytometers for immunofluorescence analysis. Fluorescence from the stained particles is in the range observed for immunofluorescence from cell-surface markers. All data were acquired using the same instrument settings, and panels A-D were obtained from the same sample acquisition of a mixture containing (1) unstained (autofluorescence) and FITC-stained CaliBRITE beads, shown in
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Figure 1.3.1 (A) Fluorescence emission spectra of FITC CaliBRITE beads (Becton Dickinson Immunocytometry), Rainbow beads RFP-30-5K (Spherotech), and glutaraldehyde-fixed chicken red blood cells (gCRBC, BioSure). Excitation at 488 nm was used. (B) Percentage of fluorescence signal through different optical filters for Rainbow beads and gCRBC normalized to signal from FITC CaliBRITE beads. Data in B are also scaled relative to the signal through the 505 to 525–nm filter.
Standardization, Calibration, and Control in Flow Cytometry
region R1 in panel A; (2) a combination of unstained and multiple levels of stained Rainbow beads, shown in region R2 in panel A; (3) gCRBC, shown in region R3; and (4) forwardscatter (FS) test pulses (no particle, R4 in panel A). Fluorescence histograms in Figure 1.3.4 are from unstained and FITC CaliBRITE beads (panel B), Rainbow beads (panel C), FS test pulses and gCRBC (panel D), Quantum 24 beads (Flow Cytometry Standards; panel E), and osmium- and glutaraldehyde-fixed CRBC (panel F). Panels B, D, and F illustrate different pairs of particles or signals at the low and high ranges of a scale for immunofluorescence. The Quantum 24 beads shown in Figure 1.3.4E had calibration values for the stained beads (upper four peaks in the histogram) of 4,201, 16,936, 37,466, and 65,797 fluorescein MESF (molecules of equivalent soluble fluorochrome).
Particles for aligning and checking measurement precision Figure 1.3.5 shows scatter and fluorescence data for a uniform 2.49-µm-diameter fluorescent bead that is useful for checking or adjusting optical alignment. All fluorescence CVs were
10× is used with a covered specimen, the image quality will be poor. At a magnification of ≤10×, objectives designed for use with and without cover slips may be used interchangeably for routine applications. Microscope cover glasses come in several thicknesses, as indicated by a number (e.g., 1, 1.5, or 2). Each number represents a defined
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eyepiece integrated observation tube
intermediate image
tube lens
infinity space
parallel light beam
intermediate attachments
objective
Figure 2.2.14 Infinity space: the distance between the back of an infinity-corrected objective and the tube lens (schematic). Reproduced from Abramowitz (1994) by courtesy of Olympus America.
thickness range. For example, #1.5 cover glasses are typically 0.16 to 0.19 mm in thickness. Typical dry biological objectives are designed and optically corrected for a cover-glass thickness of precisely 0.17 mm. Dry objectives with NA greater than ~0.75 will suffer noticeable image degradation if the cover glass differs even by a few hundredths of a millimeter from the specified thickness. Because cover glass thickness may vary by several hundredths of a millimeter even within a package, “high dry” (40× high-NA) objectives are available with an adjustable correction collar and scale that permits them to be adjusted for different cover glass thicknesses (e.g., from 0.11 to 0.23 mm). Turning the correction collar to match the actual thickness of the individual cover glass in use prevents the introduction of spherical aberration and its consequence, image degradation. Using too much mounting medium on the tissue will create an additional “cover glass–like” optical layer that must be added to the thickness of the cover glass to determine the total “effective” cover glass thickness. If the effective cover glass thickness is different from that specified for the objective, spherical aberration will be introduced into the image. For this reason, many experienced microscopists do not rely on the correction collar’s numbered scale
to set the proper correction. Rather, they choose a suitable area of the specimen and repeatedly refocus the microscope while moving the correction collar to different positions, finally reaching the setting that provides the best image. On objectives used for inverted tissue culture studies with flasks or other relatively thick culture vessels, the correction collar may have a range of correction from 0 to 2 mm; on standard upright microscope objectives, the range is usually from 0.11 to 0.22 mm.
OBJECTIVES FOR OTHER MICROSCOPY APPLICATIONS Phase Contrast For phase-contrast microscopy, an annular “phase plate” is installed by the manufacturer inside the back of the objective. This plate serves to “speed up” the undiffracted light passing through it and also to reduce its intensity. Phase specimens, such as unstained cells and tissues, are almost invisible in standard brightfield microscopy. The phase plate in the objective, when aligned with the annular opening of a phase condenser, optically renders small phase objects visible without the use of stains. Because phase-contrast observation is often done through glass or plastic culture vessels,
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eyepiece diaphragm plane
objective front focal plane of objective specimen
Figure 2.2.15 Objective system of finite tube length, showing the projection of the image by a finite objective to the intermediate image plane within the eyepiece tube. Reproduced from Abramowitz (1994) by courtesy of Olympus America.
some manufacturers offer interchangeable accessory lenses or “caps” that attach to the front lens of the objective (one set for use with plastic vessels, one set for glass vessels) to avoid distortion of images.
Polarization Techniques
Microscope Objectives
In polarization microscopy, it is important that the objective itself not contribute to the alteration of polarization effects induced by the specimen. Because glass that is physically strained affects polarized light, microscope manufacturers carefully select objectives in which the glass elements and their mountings are strain-free. The barrel of strain-free objectives supplied with polarizing microscopes is usually marked with a “P”, “SF,” or “POL” and is sometimes inscribed in a color different from the usual inscription color. Differential interference contrast (DIC) microscopy is also invaluable for making small phase objects readily visible. It has further advantages in that it (1) yields a pseudo-three-
dimensional image, in which the object appears shadowed—brighter on one side and darker on the other—displaying “elevations” and “depressions” within the specimen; (2) permits the use of high-NA optics; and (3) makes possible “optical staining” and “optical sectioning” of the specimen. In DIC microscopy, the distance from the back focal plane of the objective to the upper Wollaston prism (a special prism positioned above the objective) is usually critical, and microscope companies may therefore designate particular objectives for use in DIC microscopy. These objectives are relatively strainfree, because interference microscopy also involves the use of polarized light, and may be labeled DIC or NIC (for Nomarski interference contrast, a particular type of DIC).
Dark-Field Microscopy In transmitted-light dark-field microscopy, the illumination is directed obliquely so that the specimen appears bright on a dark background. For dark-field microscopy with high-NA objec-
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tives (≥1.00), the NA of the objective must be reduced below that of the oil darkfield condenser. Manufacturers therefore provide highNA objectives with built-in iris diaphragms (see Fig. 2.2.12). For dark-field use, the diaphragm is closed down to yield an NA below 1.1. For general use, the diaphragm must be fully open or optical performance will be degraded.
Ultraviolet (UV), Fluorescence, and Infrared (IR) Applications Standard glass objectives are relatively opaque to wavelengths in the lower UV range, below ∼380 nm. Special objectives are manufactured with special glasses to achieve greater transmission of these lower wavelengths, which are used to excite certain fluorescent dyes for measurement of intracellular ions. The cements used in complex lens elements for fluorescence microscopy are nonfluorescing, and the best fluorescence objectives are made using quartz optics. Other investigations may be carried out using longer, IR wavelengths (>750 nm), which offer poorer resolution (see Abbe’s equation in the discussion of Resolving Power) but greater depth of penetration into biological (and other) materials. Several companies offer objectives specially designed to more efficiently transmit wavelengths up to 1800 nm. The technical departments of the major microscope companies can provide transmission and spectral data for their objectives upon request to aid in selecting the proper objectives for special applications.
OTHER CONSIDERATIONS IN CHOOSING OBJECTIVES Other considerations may prove valuable in understanding the performance of objectives and in guiding the selection, purchase, and use of suitable objectives. Numerical aperture, the ability of the objective to capture a cone of light of wider angle, has a crucial effect on resolution. Although intuitively it may seem that resolving power should increase with increasing magnification, it can be shown that the ability to distinguish closely spaced details within a specimen is directly proportional to the twice the working NA. However, the use of objectives with higherthan-necessary magnification and NA for a given application can be detrimental not only because they are more expensive, but also because the specimen area observed within a field of view will be smaller and both the depth of
field (the vertical distance above and below the plane being observed that is still in acceptable focus) and working distance are shallower. When the finest specimen details need to be observed, high-NA objectives are required. High-NA objectives are also indicated when maximum throughput of light is needed. The light transmittance for an objective, using visible wavelengths, typically varies with the square of the NA of the objective. In reflectedlight and epifluorescence microscopy, light passes through the objective twice (first the illuminating light, and then the reflected or fluorescent signal), and so the intensity varies with the fourth power of the NA. In situations where the light level is low, NA is a critical factor in obtaining brighter images. A question often asked is why higher magnification cannot be achieved simply by using higher-magnification eyepieces with a given objective. Because of limitations due to the size of light waves themselves and the phenomenon of diffraction, higher and higher magnifications unaccompanied by increased NA will result in images that are less and less clear. The limiting factor in ensuring usable, as opposed to empty, magnification is the NA of the objective (more precisely, the average NA of the objective and the condenser). Eyepieces and accessory lenses are designed for use with certain objectives and condensers, and should not be switched to increase magnification except as recommended by the manufacturer. An oft-cited rule of thumb is that the user should limit the total optical magnification (the objective magnification multiplied by the eyepiece magnification and that of any other lenses) to between 500 and 1000 times the NA of the objective. At 1000 times the NA, the likely result is empty magnification. In the favored method of Koehler illumination, the condenser diaphragm is partially closed down, slightly lowering the overall NA in order to improve contrast. Hence, a total magnification of ∼750 times the NA will usually produce excellent images with satisfactory contrast. All of the foregoing discussion of objective design, features, and performance assumes that the optics (and the rest of the microscope) remain forever in the pristine state in which they presumably arrived. Proper care of the objectives, including handling, storage, and cleaning, are essential prerequisites to keeping them in proper working order. The authors have often noted that the best microscopy is not necessarily performed by those with the best equipment,
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and quite often the performance of superior optics is profoundly or subtly degraded by a lack of care in choosing and maintaining the optics.
Delly, J.G. 1988. Photography Through The Microscope. Eastman Kodak, Rochester, N.Y. Inoue, S. 1986. Video Microscopy. Plenum Press, New York.
LITERATURE CITED
Leitz, E. 1938. The Microscope And Its Application. Ernst Leitz, Wetzlar, Germany.
Abramowitz, M. 1994. Optics: A Primer. Olympus America Inc., New York.
Mollring, F.K. 1976. Microscopy From The Very Beginning. Carl Zeiss, Oberkochen, Germany.
KEY REFERENCES
Spencer, M. 1982. Fundamentals of Light Microscopy. Cambridge University Press, Cambridge, UK.
Abramowitz, M. 1985. Microscope Basics and Beyond. Olympus Corporation, New York. Abramowitz, M. 1987. Contrast Methods in Microscopy: Transmitted Light. Olympus Corporation, New York. Abramowitz, M. 1993. Fluorescence Microscopy: The Essentials. Olympus America Inc., New York. Abramowitz, 1994. See above. Bradbury, S. 1984. An Introduction to the Optical Microscope. Oxford University Press, Oxford, UK.
Contributed by Mortimer Abramowitz Olympus America Inc. Melville, New York Marc M. Friedman AccuMed International Chicago, Illinois
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Cameras One of the most critical components of an image cytometer is the camera, the unit that converts the optical image into electrical form so that it can be viewed on a TV monitor, recorded, or digitized for subsequent analysis. Any image degradation introduced at this stage will affect the quality and accuracy of the system’s output. Compared to those of the past, modern cameras offer an array of reasonably inexpensive alternatives. The proper choice of camera depends on the usage for which the instrument is designed. The camera should be matched both to the other system components and to the problems that will be addressed by the instrument. The camera is the eye of an image cytometer. It is called upon to convert a two-dimensional spatial distribution of light intensity into a corresponding electrical signal that is a faithful representation of the specimen. This, in turn, can be displayed, recorded, or processed. How accurately the camera can conduct this transformation affects the quality of results one can obtain from the instrument. Image sensing is a complex technology that harnesses a variety of phenomena. There are numerous sources of noise, distortion, and loss of resolution in the process. Any particular camera represents a series of design tradeoffs, and its performance will be higher in some areas and lower in others when compared with another unit. A poor choice of camera can severely limit the accuracy and usefulness of an image cytometry system. The camera with the best specifications or the highest price is not always the right one for a particular job. It is necessary to understand the fundamentals of camera phenomena in order to design an image cytometer or use it to best advantage. Fortunately, many of the image degradations that are introduced by camera shortcomings can be reduced or eliminated by subsequent image processing. Thus, the camera’s performance is interwoven with that of the software.
DEFINITIONS Image sensing involves three steps: sampling, transduction, and scanning. The image is divided up into an array (usually a rectangular grid) of small regions, called “picture elements” or “pixels,” that can be considered one at a time. Sampling is the process of measuring
Contributed by Kenneth R. Castleman Current Protocols in Cytometry (1998) 2.3.1-2.3.15 Copyright © 1998 by John Wiley & Sons, Inc.
UNIT 2.3 the light intensity at an individual pixel location. It requires a sampling aperture that defines the size and shape of the pixel (usually circular, square, or rectangular). Transduction is the process of converting the light intensity at a particular pixel location into a corresponding voltage. Scanning is the process of selectively addressing the picture elements in order. This creates the data stream that represents the image. If the image is to be processed digitally, it must be digitized. Quantization is the process of generating an integer that reflects the brightness of the image at a particular pixel location. Digitization is the process of sampling and quantizing an image. The degree to which a camera can reproduce small objects is its resolution. If it warps the objects in the image, this is distortion. Any undesirable additive components of the image are called noise.
IMAGE SENSING Light Sensing Light-sensing devices produce an electrical signal proportional to the intensity of light falling upon them. Different physical phenomena can be employed for this purpose, giving rise to different types of light sensors. Photoconductors, such as selenium, show a drop in their electrical resistance when exposed to light. Semiconductor devices made from pure silicon crystals generate free electrons in response to incident photons. Both these phenomena have been harnessed to sense images.
Photometry Photometry is the technology of quantifying light intensity, and there are many ways to do this. For example, photons of different wavelength have different energy. Thus, if incident light energy flux is measured, the spectrum of the light affects the intensity. Commonly used image sensors, however, merely count photons, so wavelength considerations do not directly affect the measured intensity. Although the sensors do have different sensitivities at different wavelengths, this is best accounted for separately. A quality image digitizer will produce an array wherein each gray level is proportional to the number of photons that landed on that pixel during the exposure time.
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The linearity of an image sensor specifies how accurately its output reflects the incident photon flux. Modern charge-coupled device (CCD) image sensor chips are quite linear over their entire range, and thus linearity is seldom a problem, as long as saturation (overload) of the sensor is avoided.
Scanning Conventions Video scanning conventions Figure 2.3.1 illustrates the Electronic Industries Association (EIA) RS-170 scanning convention, which is the standard for monochrome broadcast television in the United States (Fink, 1957; Fink and Christiansen, 1989; Hutson et al., 1990; Castleman, 1996). The beam scans the entire image in 525 horizontal scan lines, 30 times each second. The lines are not scanned in sequential order, however, because if the TV screen were to be refreshed at only a 30/sec rate, the eye would perceive an annoying flicker. Instead, an interlaced scanning convention is used to yield an apparent 60/sec refresh rate on the screen. Each frame is made up of two interlaced fields, each consisting of 262.5 lines. The first field of each frame scans all the odd-numbered lines, while the second field scans the intervening even-numbered lines. Interlacing yields a 60/sec field rate to minimize perceived flicker, while the 30/sec frame rate reduces the frequency bandwidth as required for broadcast television channels.
1 2 3 4
Each horizontal line scan requires 63.5 µsec, of which ~50 µsec (83%) is active, containing image information. Of the 525 lines per frame, 16 are lost in the vertical retrace of each field, leaving about 483 active lines per frame. The bandwidth of the standard video signal extends up to 4.5 megahertz (MHz), which allows 225 cycles, or about 550 pixels worth of information, across the active portion of each line. The NTSC (National Television Standards Committee) timing standard for color television in the USA differs only slightly from the RS-170 convention. It was designed to accommodate color transmission while maintaining compatibility with existing monochrome receivers. Different scanning conventions are used in other countries. For example, the CCIR (Comité Consultatif International des Radiocommunications) standard used in much of Europe employs a frame of 625 interlaced scan lines of about 768 pixels each and runs at 25 frames/sec. New broadcast scanning conventions, offering more lines per frame, more pixels per line, and higher image quality, are being developed in several countries under the name “high-definition television” (HDTV). Two proposals now under consideration in the USA are 1280 pixel × 720 line progressive (noninterlaced) scanning, and 1920 pixel × 1080 line interlaced scanning; both of these standards employ a 16:9 aspect ratio. One can use a video camera as an image digitizer simply by sampling the video signal with a fast analog-to-digital converter operat-
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30 frames/sec 525 lines/frame 15,750 lines/sec 63.5 µsec/line 2 fields/frame 262.5 lines/field 60 fields/sec
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524
483 active lines/frame
Figure 2.3.1 The RS-170 scanning convention, used for monochrome broadcast television transmission in the United States. The CCIR convention, used in much of Europe, employs 625 lines and operates at 25 frames/sec.
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ing at ∼14 MHz. A frame grabber is a digitizer that stores this high-speed data stream in a solid-state memory and then feeds it out at a slower rate to a more permanent storage device, such as a disk drive. Other scanning conventions Cameras designed specifically for scientific image sensing can be made to scan by any set of timing rules. Typically scientific cameras have larger image formats (more scan lines and pixels per line), noninterlaced scanning, and slower readout rates (to reduce the noise associated with readout). They also may incorporate variable-length frame-integration periods (to increase sensitivity) and sensor-chip cooling (to reduce thermal noise).
Camera Performance Although cameras differ in the approach they use to sense an image, they can be compared on the basis of their performance. Image size An important parameter is the size of the image a camera produces. Image size is specified by the maximum numbers of scan lines and of pixels per line. Pixel size and spacing Two important characteristics are the size of the sampling aperture and the spacing between adjacent pixels. These parameters, specified at the image plane, scale down to the specimen plane by the magnification factor of the microscope. This is usually the objective power multiplied by any auxiliary magnification that is in place. The eyepieces normally do not contribute to this calculation. Resolution According to the Rayleigh criterion, one can just resolve (identify as separate) two point objects in a microscope image if they are separated by the distance δ = 0.61λ/NA, where NA is the numerical aperture of the objective and λ is the illumination wavelength (Castleman, 1996). The camera should be able to reproduce detail to this degree. Linearity The degree of linearity of the relationship between the input light intensity and the output signal amplitude is another important factor. Although the eye is not particularly critical in this department, the validity of subsequent pro-
cessing can be jeopardized by a nonlinear camera. Noise Finally, one of the most important characteristics of a camera is its noise level. If a uniformly gray image is presented to a camera, its output will show variations in gray level, even though the input brightness is constant across the image. Such noise introduced by the camera is a source of image degradation, and should be small relative to the contrast of the specimen. Requirements Whether or not a particular camera is adequate depends on the specific task at hand. In some applications, digitizing images with relatively few lines, pixels per line, or gray levels or with appreciable noise and nonlinearity may be sufficient. Image cytometry, however, normally requires a high-quality camera that is capable of sensing large images with many gray levels, good linearity, and a low noise level.
TYPES OF CAMERAS Historically, imaging tubes, such as the vidicon and its relatives, were the backbone of image cytometry. Currently, however, solidstate cameras generally offer more flexibility, better performance, and lower cost.
Tube-Type Cameras Vidicon construction Figure 2.3.2 illustrates the construction of the vidicon, a common type of television image-sensing tube. It is a cylindrical glass envelope containing an electron gun at one end and a target and faceplate at the other. The tube is surrounded by a yoke containing electromagnetic focus and beam deflection coils. The faceplate is coated on the inside with a thin layer of photoconductor over a thin transparent metal film, forming the target. Adjacent to the target (to the left in Fig. 2.3.2) is a positively charged fine wire screen called the mesh. A smaller positive charge is applied to the target. Vidicon operation A stream of electrons is projected from the electron gun, focused to a small spot on the target by the focus field, and steered across the target in a scanning pattern by the time-varying deflection field. Electrons decelerate after passing the mesh, and reach the target with approximately zero velocity. The moving electron
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deflection coils electron gun
anode target faceplate
glass envelope electron beam
Figure 2.3.2 Vidicon camera tube construction.
beam deposits a layer of electrons on the inner surface of the photoconductor to balance the positive charge on the metal coating on the opposite side. Light striking an area of the photoconductor causes electrons to flow through, locally depleting the surface charge layer at that point. The optical image formed on the target then causes the photoconductor to leak electrons until an identical electron image is formed on the back of the target. Electrons will be present in dark areas and absent in light areas of the image. As the electron beam scans the target, it replaces the lost electrons, restoring a uniform surface charge. As the electrons are replaced, a current flows in the external circuit of the target. This current is proportional to the number of electrons required to restore the charge and therefore to the light intensity at that point. Current variations in the target circuit produce the video signal. The electron beam repeatedly scans the surface of the target, replacing the charge that bleeds away. The vidicon target is thus an integrating sensor, with the period of integration equal to the scanning frame rate. The vidicon family The photoconductor target of a standard vidicon is made of selenium photoconductor material. Relatives of the vidicon, with similarsounding names, differ mainly in the composition of the photoconductive target, and each excels in certain imaging characteristics.
CCD Cameras
Cameras
Silicon light sensors Pure silicon can be grown in large crystals in which each atom is covalently bonded to its six neighbors in a three-dimensional rectangu-
lar lattice structure. An incident photon can break one of these bonds, freeing an electron. A thin metal layer deposited on the surface of the silicon and charged with a positive voltage creates a potential well that collects and holds the electrons thus freed. Each potential well corresponds to one pixel in an array of sensors. A potential well can hold about 106 electrons on typical chips. Thermal energy also causes random bond breakage, creating thermal electrons that are indistinguishable from photoelectrons. This gives rise to dark current, current produced in the absence of light. Dark current is temperature sensitive, doubling for each 6°C increase in temperature. At the long integration times that are required for image sensing at low light levels, the wells can fill with thermal electrons before filling with photoelectrons. Cooling is often employed to reduce dark current and thereby extend the usable integration time. CCD construction CCD chips are manufactured on a light-sensitive crystalline silicon chip, as discussed above (Janesick and Elliot, 1992). A rectangular array of photodetector sites (potential wells) is built into the silicon substrate. Photoelectrons produced in the silicon are attracted to and held in the nearest potential well. By controlling the electrode voltages, they can be shifted as a charge packet from well to well until they reach an external terminal. CCD operation There are three architectures that can be employed for reading the accumulated charge out of CCD image sensor arrays. These are (1) full-frame architecture, (2) frame-transfer architecture, and (3) interline-transfer architecture (Fig. 2.3.3).
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line shift
image array full-frame array
line shift
line shift
masked storage array
serial register
serial register
serial register
pixel shift full-frame CCD
pixel shift frame-transfer CCD
pixel shift interline transfer CCD
Figure 2.3.3 CCD chip operation using full-frame, frame-transfer, or interline transfer architecture.
Full-frame CCD. Following exposure, a full-frame CCD is shuttered to keep it in the dark during the readout process. It then shifts the charge image out of the bottom row of sensor wells, one pixel at a time. After the bottom row is empty, the charge in all rows is shifted down one row, and the bottom row is again shifted out. This process repeats until all rows have been shifted down and out. The device is then ready to integrate another image. Frame-transfer CCD. A frame-transfer CCD chip has a doubly long sensor array. The top half senses the image in the standard manner, while the storage array on the bottom is protected from incident light by an opaque mask. At the end of the integration period, the charge image that has accumulated in the sensing array is shifted rapidly, row by row, into the storage array. From there it is shifted out pixel by pixel in the standard manner, while the sensing array integrates the next image. Like interline transfer, this technique employs simultaneous integration and readout, making video-rate image sensing possible. Interline-transfer CCD. In an interline transfer CCD every second column of sensors is covered by an opaque mask. These columns of masked wells are used only in the readout process. After exposure, the charge packet in each exposed well is shifted into the adjacent masked well. This transfer requires very little time because all charge packets shift at once. While the exposed wells are accumulating the next image, the charge packets in the masked columns are being shifted out in the same way as in full-frame CCD. In an interline transfer sensor, the number of pixels per line is half the actual number of wells per row on the chip. No more than 50% of the chip area is light-sensi-
tive, because the masked columns cover half its surface. CCD performance Available in a variety of configurations, CCDs give rise to compact and rugged solidstate cameras for both television and image digitizing applications. They are free of geometric distortion and exhibit highly linear response to light. CCDs are therefore emerging as the device of choice for image cytometry. CCDs can be scanned at television rates (30 frames/sec) or much more slowly. Because they can integrate for periods of seconds to hours to capture low-light-level images, they are often used in fluorescence microscopy. Integration times longer than a few seconds require cooling the chip well below room temperature to reduce dark current, which would otherwise fill the wells with thermal electrons before photoelectrons had a chance to build up. Because of imperfections in the crystal lattice, dark current varies significantly from one pixel to the next, particularly in less expensive chips. In long-exposure images, this leaves a “starfield” of fixed-pattern noise due to the few pixels with abnormally high dark current. Because this pattern is stationary, it can be recorded and subtracted out, provided the offending pixels are not allowed to saturate. Defects in the crystal lattice can cause “dead pixels,” which will not hold or shift electrons. This can wipe out all or part of a column of pixels. CCD sensors are graded on the number of such defects, and the higher-grade chips are more expensive. Readout noise is random noise generated by the on-chip electronics. It ranges from a few to many electrons per pixel depending on chip
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row scan
column scan
video signal
Figure 2.3.4 CID chip operation.
Cameras
design, and gets worse as the charge is read out at a faster rate. It is usually the dominant noise factor under short-exposure, low-light conditions where the dark current and photon noise components are small. Photon noise results from the quantum nature of light. The actual number of photons striking any particular pixel in any one exposure will be random. In general the photon noise component is the square root of the number of electrons that accumulate in a well (i.e., photoelectrons plus thermal electrons). It is usually the dominant noise source under high-exposure or high–dark current conditions. The charge developed at a particular pixel may be shifted as many as two thousand times, depending upon its location in the array. The charge transfer efficiency must be extremely high or significant numbers of photoelectrons will be lost in the readout process. Often, half or more of the available area of the sensor is covered by opaque charge-transfer circuitry, leaving gaps between the pixels and reducing the fill factor below the ideal of 100%. The chip can be coated with a thin layer of lenslets, each of which focuses the incoming light it receives onto the sensitive areas of one pixel. Overexposure of a CCD sensor can cause
blooming of the image as excess photoelectrons spread to adjacent pixels. Spectral sensitivity may also be a significant issue. Silicon sensors become less sensitive at the deep blue and ultraviolet end of the wavelength spectrum. This can be overcome by a lumigen coating, which absorbs the shortwavelength photons and then reemits the energy as longer-wavelength photons that the silicon can see. Dynamic range characterizes the performance of the chip at high light levels, where the wells can be filled with photoelectrons (rather than dark current) during a relatively short exposure. It is computed as pixel well capacity divided by readout noise level, both measured in electrons, and is usually expressed in dB. This parameter is independent of exposure conditions (light level and exposure time). The signal-to-noise ratio (SNR) can be computed as the number of photoelectrons received by a well divided by the total (photon plus readout) noise level; that is, SNR = (F × QE × te)/(Np2 + Nr2)1/2, where F is the incident photon flux, QE the quantum efficiency, te the exposure time, Nr the readout noise level, and Np = [(F × QE + DC)te]1/2 the photon noise that results from the statistical nature of light. The
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SNR is quite dependent on light level and exposure time.
CID Cameras Charge injection device (CID) sensors (Williams and Carta, 1989; Kaplan, 1990) employ the photoelectronic properties of silicon as do CCDs, but they use a different method of readout. CID construction At each pixel site, the CID has two adjacent electrodes (Fig. 2.3.4) that are insulated from the silicon surface by a thin metal-oxide layer. Each pixel is connected to all the pixels in its column by one electrode, and to all the pixels in its row by the other electrode. Thus, a single pixel can be addressed by its row and column address. If all rows and columns of electrodes are held at a positive voltage, the entire chip accumulates a photoelectron image. CID operation When one electrode is driven to 0 V, the accumulated photoelectrons will shift under the second electrode, creating a current pulse in the external circuitry. The size of this current pulse reflects the amount of accumulated photoelectronic charge. Because the accumulated photoelectrons remain in the well after the shift, this is a nondestructive type of image readout: the pixel can be read repeatedly by shifting the charge back and forth between the electrodes. When both electrodes are driven to zero, the accumulated photoelectrons are injected into the underlying substrate, producing a current pulse in the external circuitry. The size of the pulse again reflects the amount of accumulated charge, but this process leaves the well empty. This destructive readout mode is used to prepare the chip for integrating another image. The circuitry on the chip controls the voltages on the row and column electrodes to effect image integration and destructive and nondestructive readout. Because the CID can address individual pixels in any order, subimages of any size can be read out at any speed. Nondestructive readout allows one to watch the image accumulate on the chip, which is useful when the length of the required integration period is unknown. CID performance CIDs are largely immune to blooming (charge spreading to adjacent pixels) and to radiation damage. Also, with nondestructive readout, the control program can monitor the
filling of the wells and selectively flush individual pixels that become full before the integration period is over. Because CIDs do not shift charge packets across the array, charge transfer efficiency is not a concern. Unlike the CCD, a small defect in the crystal lattice affects only one pixel. Also, essentially the entire surface area is light sensitive, leaving virtually no gaps between pixels. Even so, CIDs are considerably less light sensitive than similar CCDs.
APS Cameras An emerging new solid-state sensing technology is active pixel sensor (APS) cameras (Janesick and Elliot, 1992; Fossum, 1993, 1995). Like CCD cameras, these are fabricated on a silicon chip, but they use complementary metal-oxide-semiconductor (CMOS) integrated circuit technology. This allows the chip designers to embed on the sensor chip itself processing circuitry that normally exists elsewhere in the camera. Indeed, much of the circuitry that traditionally resides on various circuit boards in the camera can be fabricated directly on the image-sensor chip. Experimental APS camera chips have been developed with amplifiers at each pixel and with special noise-reducing readout circuitry and analog-to-digital (A-to-D) converters on the chip. Some APS chips contain circuitry that allows them to read out a rectangular subimage continuously and nondestructively. APS technology promises to reduce the cost of cameras in the future and perhaps to improve their performance as well.
IMAGE DIGITIZATION Before an image can be processed by computer, it must be converted into an array of numbers. This must be done in such a way that it does not destroy or significantly degrade the specimen content of interest. Both the number of gray levels in the grayscale and the number of pixels per row and column must be adequate for the tasks at hand. Binning is the technique of combining adjacent pixels in a sensor array to form larger pixels. For example, using 2 × 2 binning on a 1024 × 1024 sensor array with 6 × 6-µm pixels would produce a 512 × 512 image where the pixels were effectively 12 × 12 µm.
Noise Sources The time-varying electrical signal emerging from the image sensor is sampled and quantized by an analog-to-digital converter (ADC) cir-
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Table 2.3.1
Front-Illuminated CCD Chips from Kodak
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Microlensing Dynamic range
KAF0400
KAF1000
KAF1300L
KAF1400
KAF1600
KAF4200
KAF6300
9×9 80,000 768 × 512 6.9 × 4.6 0.36
24 × 24 630,000 1,024 × 1,024 24.6 × 24.6 0.40
16 × 16 140,000 1,280 × 1,024 20.5 × 16.4 0.32
6.8 × 6.8 45,000 1,317 × 1,035 9.0 × 7.0 0.40
9×9 80,000 1,536 × 1,024 13.8 × 9.2 0.39
9×9 80,000 2,032 × 2,044 18.4 × 18.4 0.38
9×9 85,000 3,072 × 2,048 27.65 × 18.5 0.38
19 at 1 Mhz 18 at 1 Mhz
14 at 2 MHz
13 at 500 kHz 13 at 500 kHz 12 at 500 kHz 22 at 5 MHz
0.44
14.4
7.2
0.31
0.31
0.90
7.2
Full frame 100% No 72 dB
Full frame 100% No 82 dB
Full frame 100% No 80 dB
Full frame 100% No 72 dB
Full frame 100% No 72 dB
Full frame 100% No 76 dB
Full frame 100% No 72 dB
cuit. If B is the number of bits used in the quantization, the grayscale goes from zero to 2B − 1. Quantization can be viewed as a source of noise, because it alters the gray level at each pixel by a small random amount. The signalto-noise ratio (SNR) for quantization is the (full-scale) signal amplitude divided by the quantization noise level. For images with a Gaussian distribution of gray levels, the SNR, measured in decibels (dB), is 20 × log10[2B/σn] = 6B + 11, where B is the number of bits used in the quantization and σn is the standard deviation of the resulting quantization noise. Each 20 dB of SNR represents a factor of ten in the ratio. The commonly used eight-bit grayscale (B = 8, 2B = 256 gray levels, SNR = 59 dB = 891) is adequate for many image cytometry applications. Normally this quantization noise level (± 0.11% of full scale) is tolerable, but one should verify this and use ten or more bits of grayscale resolution if that is required by the application. The camera introduces other random noise components as well. Readout noise, introduced by the circuitry on the CCD chip, and photon noise, which results from the statistical nature of light, are discussed above (see CCD performance). In general, the different random noise sources combine in such a way that the overall noise level is the square root of the sum of the squares of their individual amplitudes.
Spatial Resolution Cameras
The well-known Shannon sampling theorem states that one can reconstruct, by proper
interpolation, a sinusoidal signal from equally spaced sample points if there are no fewer than two sample points per cycle of the sine wave (Castleman, 1996). If the sampling is done more sparsely, one can encounter the phenomenon of aliasing, which introduces Moiré patterns into the image. A microscope objective cannot pass image detail at frequencies higher than the optical cutoff frequency of fc = 2NA/λ, where NA is its numerical aperture and λ is the illumination wavelength (Castleman, 1996). Thus, aliasing can be avoided completely if the pixel spacing at the specimen is no larger than λ/4 NA. This is 1⁄8 µm for an objective with NA = 1 operating in green (λ = 500) light. For applications in which the specimen does not contain detail at the resolution limit of the objective lens, larger pixel spacing will suffice. However, even smaller pixel spacing may be required for accurate measurement of objects in the image or for optimal display of the image. In these cases, reduced pixel spacing can be achieved by interpolation of the image after it is digitized (Castleman, 1996).
AVAILABLE CCD CHIPS AND CAMERAS An impressive array of solid-state cameras, incorporating a variety of different CCD chips, is commercially available. These cover a wide range of cost and performance. In this section we tabulate some of the ones that are potentially most useful for cytometry. The list is by no means exhaustive, and the CCD camera situation is subject to rapid change.
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The performance of a particular CCD camera depends on two major design factors: the choice of the CCD sensor itself, and the design of the supporting electronics in the camera. A poor quality chip in a well-designed camera, and a good chip embedded in poorly matched circuitry, will be equally disappointing. The circuitry in a well-designed camera will exploit the best characteristics of the sensor chip. Overall camera performance cannot exceed the limitations of either the chip or the electronics. Thus a particular CCD camera must be evaluated as a complete system. With that point made, we herein tabulate chip and camera characteristics separately for the sake of brevity.
CCD Chips Tables 2.3.1, 2.3.2, 2.3.3, 2.3.4, 2.3.5, 2.3.6, 2.3.7, and 2.3.8 have been compiled from manufacturers’ data. Achieving a consistent set of specifications is difficult because each chip maker chooses to specify chip characteristics differently. We include the commonly used
Table 2.3.2
CCD Cameras Table 2.3.9 presents specifications on several instrumentation quality CCD cameras. The terms used in the table are defined above (see Image Digitization). Many of the relevant specifications are not available from all manufacturers in a consistent form. Where entries are not shown, reliable data were unavailable at the time of publication. The very important sensitivity parameter, for example, is specified in so
Front-Illuminated Chips from Scientific Imaging Technologies, Inc.
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Dynamic range
Table 2.3.3
mechanical specifications and show the electrical parameters in common units. Well capacity and RMS readout noise are given in electrons, and dark current in electrons per second for a single pixel at 0°C. Dark current doubles for each 6°C increase in temperature, and vice versa. Dynamic range, computed as well capacity divided by readout noise, also appears in the tables. Signal-to-noise ratio is quite dependent on exposure conditions (light level and exposure time), and thus is not listed in the tables.
SI502AF
STAE01AF, SIA003AF
SIA002AB
24 × 24 325,000 1,100 × 165 26.4 × 4 0.30 20 at 1 Mhz 160 Full frame 100% 84 dB
24 × 24 325,000 1,024 × 1,024 24.6 × 24.6 0.30 20 at 1 Mhz 353 Full frame 100% 84 dB
15 × 15 100,000 4,096 × 2,048 30.7 × 30.7 0.30 7 at 45 kHz 10 Frame transfer 100% 83 dB
Front-Illuminated Chips from Thomson-CSF
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Dynamic range
TH7895M
TH7896M
TH7887
19 × 19 375,000 512 × 512 9.7 × 9.7 0.30 25 at 500 kHz 72 Full frame 100% 84 dB
19 × 19 375,000 1,024 × 1,024 19.5 × 19.5 0.35 25 at 500 kHz 90 Full frame 100% 84 dB
14 × 14 250,000 1,024 × 1,024 14.3 × 14.3 0.10
Frame transfer 100% 80 dB
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overview of their cameras and the CCD chips used.
many different ways by different manufacturers that comparison is impossible, and it is thus conspicuously absent from this table. Price ranges are specified as “low” ($20,000). Photometrics cameras have three gain settings. Specifications are given here for the 1× gain setting, which matches the full scale range of the analog-to-digital converter (ADC) to the well capacity of a single pixel. The 4× gain mode, where one-quarter of full well capacity saturates the ADC, achieves greater sensitivity for use at low light levels. The 0.5× gain mode, when used with binning, increases the effective well size to improve the SNR at high light levels. Photometrics product literature (http://www.photomet.com) provides useful specifications and other helpful information about CCD cameras. Princeton Instruments’ catalog, available in Adobe Acrobat format (http://www.prinst. com/imagprod.htm), provides a comprehensive Table 2.3.4
Each image cytometry application deserves its own analysis of camera and digitizing requirements. When it is possible to select a specific camera, camera characteristics should be evaluated in light of the requirements of the planned experiments. In any case, the camera should be well matched to the problem and to the other system components. Although an inadequate camera can forestall success, camera overkill can waste resources that might be better applied elsewhere. Modern cameras are quite good in performance and reasonable in cost compared to those of the past, and will undoubtedly continue to improve. In addition, some of the image detail lost to camera-induced noise, distortion, and lack of resolution can be recovered with digital image processing.
Front-Illuminated Chips from EEV
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Dynamic range
Table 2.3.5
Cameras
CONCLUSION
EEV CCD37
EEV 02-06
EEV 05-20
15 × 15 165,000 512 × 512 7.7 × 7.7 0.26 23 at 2 Mhz 1,078 Frame transfer 100% 77 dB
22 × 22 500,000 288 × 384 12.7 × 8.4 0.32 28 at 1 Mhz 2,580 Frame transfer 100% 85 dB
22.5 × 22.5 500,000 576 × 770 12.9 × 17.3 0.30 22 at 500 kHz 2,580 Frame transfer 100% 87 dB
Front-Illuminated Chips from Texas Instruments and Loral
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Microlensing Dynamic range
T.I. TC-255
Loral CCD442A
10 × 10 50,000 320 × 240 3.2 × 2.4 0.4 28 97 Full frame 100% No 70 dB
15 × 15 150,000 2,048 × 2,048 30.7 × 30.7 9 at 100 kHz 102
84 dB
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Table 2.3.6
Front-Illuminated Chips from EG&G Reticon
Well size (µm) Frame size (pixels) Chip size (mm) Type of transfer Fill factor Microlensing Dynamic range
Table 2.3.7
RL2005PAQ-11
RL2010PAQ-11
RL2020PAQ-11
14 × 14 512 × 512 7.2 × 7.2 Frame transfer 100% No 48 dB
14 × 14 1,024 × 1,024 14.3 × 14.3 Frame transfer 100% No 48 dB
14 × 14 2,048 × 2,048 28.6 × 28.6 Frame transfer 100% No 48 dB
Back-Illuminated Chips from Scientific Imaging Technologies, Inc.
Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Microlensing Dynamic range
SIAE01AB, SI502AB SIA003AB
SIA002AB
24 × 24 325,000 1,100 × 165 26.4 × 4 0.8 20 at 1 Mhz 64 Full frame 100% No 84 dB
15 × 15 100,000 4,096 × 2,048 30.7 × 30.7 0.8 7 at 45 kHz 10 Frame transfer 100% No 83 dB
24 × 24 325,000 1,024 × 1,024 24.6 × 24.6 0.8 20 at 1 Mhz 353 Full frame 100% No 84 dB
Table 2.3.8 Back-Illuminated Chip from Princeton Instruments
P.I. 1000PB Well size (µm) Well capacity (e−) Frame size (pixels) Chip size (mm) Quantum efficiency at 550 nm Readout noise (e− rms) Dark current (e−/pixel/sec at 0°C) Type of transfer Fill factor Microlensing Dynamic range
15 × 15 60,000 1,000 × 800 15 × 12 0.67 20 at 500 kHz 97 Full frame 100% No 70 dB
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Table 2.3.9
CCD Cameras
Photometrics SenSys 0400/1400/1600 Chip KAF0400/KAF1400/KAF1600 Binning options Arbitrary M × N Cooling method 10°C, thermoelectric A-to-D conversion 12 bits Frame readout time at rate 0.41/1.39/1.6 sec at 1 MHz Dynamic range 72.5/67/72 dB Price Mid-range Web site http://www.photomet.com Photometrics Quantix Chip Binning options Cooling method A-to-D conversion Frame readout time at rate Dynamic range Price
KAF1400 Arbitrary M × N −25°C, forced air; −35°C, liquid circulation 12 bits 0.3 sec at 5 MHz 67 dB High-range
Photometrics S300:1400/4200/502B/003B/003F/7895/7896 Chip KAF1400/KAF4200/SI502AB/SI003AB/SI003AF/TH7895M/ TH7896M Binning options Arbitrary M × N −25°C, forced air; −40°C, liquid circulation; −90°C, −110°C, LN2 Cooling method A-to-D conversion 12, 14, or 16 bits (chip dependent) Frame readout time at rate 2.8/8.7 sec at 500 kHz/1.4/5.4/5.4/1.4/5.5/5.5 sec at 200 kHz Dynamic range 70/77/89/86/86/87/88 dB Price Mid- to high-range Photometrics PXL:37/1000/1300L/1400/6300/003F/003B Chip CCD37/KAF1000/1300L/1400/6300/SI003AF/SI003AB Binning options Arbitrary M × N −25°C, liquid circulation Cooling method A-to-D conversion 12, 14, or 16 bits (chip dependent) Frame readout time at rate 0.16/0.58/0.74/2.6/3.9 sec at 2 mHz/1.2/1.4 sec at 1 MHz Dynamic range 77/82/82/75/75/79/76 dB Price High-range Princeton Instruments CCD-576E/CCD-770E/CCD-1242E Chip EEV 02-06/EEV 05-20/EEV 05-30 Binning options Flexible −35° to −130°C, various methods Cooling method A-to-D conversion 12, 14, or 16 bits Frame readout time at rate 0.286 sec at 1 MHz/1.8/1.5 sec at 500 kHz Dynamic range 85/87/86 dB Price Mid- to high-range Web site http://www.prinst.com/imagprod.htm continued
Cameras
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Table 2.3.9
CCD Cameras, continued
Princeton Instruments CCD-512SF/CCD-1024SF/CCD-512SB/CCD-1024SB Chip SITe SI502FA/SIA003AF/SITe SI502AB/SIA003AB Binning options Flexible −30° to −140°C, various methods Cooling method A-to-D conversion 14 or 16 bits Frame readout time at rate 0.29/1.2/0.5 sec at 1 MHz/2.32 sec at 500 kHz Dynamic range 84/82/84/88 dB Price Mid- to high-range Princeton Instruments CCD-768K/CCD-1280K/CCD-1317K/CCD-1536K/CCD-2033K Chip KAF0400/KAF1300L/KAF1400/KAF1600/KAF4200/KAF6300 Binning options Flexible −35° to −60°C, various methods Cooling method A-to-D conversion 12 or 14 bits Frame readout time at rate 0.41/1.4/1.4/1.8/4.2/6.3 sec at 1 MHz Dynamic range 72/84/73/77/77/72 dB Price Mid- to high-range Princeton Instruments CCD-1000PB Chip P.I. 1000PB Binning options Flexible −35° to −55°C, various methods Cooling method A-to-D conversion 14 or 16 bits Dynamic range 70 dB Price Mid- to high-range PixelVision SV512/SV10K/SV165/SV20K Chip SITe SI502AB(F)/SIA003AB(F)/STAE01AB(F)/Loral CCD442A Binning options Yes Cooling method 40° or 70°C below ambient A-to-D conversion 16 bits Readout rate 100 kHz to 1 MHz Dynamic range 84 dB at 1 MHz Price Mid- to high-range Web site http://www.site-inc.com Hamamatsu C4880-10/20/30/31/40 Chip T.I. TC-215/S5466/SITe SI502A/SI003A/ICX-074 Binning options Yes Cooling method 5° to −120°C, using air, water, LN2 A-to-D conversion 12, 14, or 16 bits Price Mid- to high-range Web site http://www.hamamatsu.com Richter Enterprises Silicon Mountain Design 1M60-20 Chip Thompson TH7887 Binning options 2×2 A-to-D conversion 12 bits Readout rate 20 MHz Web site http://www.smd.com continued Image Cytometry Instrumentation
2.3.13 Current Protocols in Cytometry
Supplement 8
Table 2.3.9
CCD Cameras, continued
Spectra Source Orbis Chip Binning options Cooling A-to-D conversion Price Web site
Various EEV, Texas Instruments, Kodak, and SITe Yes −30° to −120°C 12 or 16 bits Mid- to high-range (some low-priced models) http://www.optics.org/spectrasource/producttoc.html
Capella Cooled CCD Chip Cooling A-to-D conversion Readout rate
Kodak KAF0400, Kodak KAF1300, EEV CCD37 Pelter, LN2 12 or 14 bits 500 kHz to 8 MHz
Diagnostic Instruments Inc. Spot (color) Chip Kodak KAF1400 A-to-D conversion 8 bits per color, liquid-crystal filter Web site http://www.diaginc.com/spotspec.htm Micro Photonics UltraPix Astrocam Chip Kodak KAF0400/KAF1600 Binning Yes Frame readout time 5 sec/20 sec Web site http://www.microphotonics.com/ccdupix.html EG&G Reticon LD2000 Chip Binning Readout rate A-to-D conversion
RL2005PAQ-11/RL2010PAQ-11/RL2020PAQ-11 Yes 33 MHz 10 bits
Integrated Scientific Imaging Chip KAF0400/KAF1600 Cooling 45°C below ambient A-to-D conversion 14 bits Note Integrated with filter wheel Optic PixCel 225 Chip Binning Cooling A-to-D conversion Web site
T.I. TC-255 2×2 35°C below ambient 14 or 16 bits http://www.optecinc.com/pixcel.htm
Xillix Microimager Chip Binning Readout rate A-to-D conversion Price Web site
Kodak KAF-1400 2×2 500 kHz to 8 MHz, up to 5 frames/sec 10 or 12 bits Low- to mid-range http://www.xillix.com/microimagerspecs.html
Cameras
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Current Protocols in Cytometry
LITERATURE CITED Castleman, K.R. 1996. Digital Image Processing. Prentice Hall, Englewood Cliffs, N.J. Fink, D.G. (ed.). 1957. Television Engineering Handbook. McGraw-Hill, New York. Fink, D.G. and Christiansen, D. 1989. Electronics Engineers Handbook. McGraw-Hill, New York. Fossum, E. R. 1993. Active pixel sensors: Are CCDs dinosaurs? Proc. SPIE 1900:2-14. Fossum, E.R. 1995. CMOS image sensors: Electronic camera on a chip. IEEE International Electron Devices Meeting Technical Digest, Dec. 10-13, 1995, Washington, D.C.
Hutson, G., Shepherd, P., and Brice, J. 1990. Colour Television Theory: System Principles, Engineering Practice and Applied Technology. McGrawHill, New York. Janesick, J. and Elliot, T. 1992. History and advancements of large area array scientific CCD imagers. Astronom. Soc. Pacif. Conf. Ser. 23:1-67. Kaplan, H. 1990. New jobs for charge-transfer devices. Photonics Spectra (Nov.). Williams B. and Carta, D. 1989. CID cameras: More than an alternative to CCDs. Adv. Imaging (Jan.).
Contributed by Kenneth R. Castleman Perceptive Scientific Instruments, Inc. League City, Texas
Image Cytometry Instrumentation
2.3.15 Current Protocols in Cytometry
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Optical Filtering Systems for Wavelength Selection in Light Microscopy The recent renaissance of light microscopy is based on a number of exciting technological developments that enable microscopic imaging of biological cells and tissues at greatly improved spatial, temporal, and spectral resolution (Taylor et al., 1997). Numerous disciplines (physics, chemistry, molecular biology, engineering, computer science) contribute to this enhanced performance of microscopic imaging. Fluorescence-based detection is one of the foremost methods, given its sensitivity and specificity. Its high quality is made possible by the ability to detect the emission corresponding to a single molecular species, based on selective labeling and equally selective optical detection, with simultaneously high spatial and wavelength resolution. This need has led to dramatic improvements in filter system design and implementation. With the concomitant development of sensitive fluorescent dyes (at multiple excitation wavelengths), multicolor imaging has become a powerful tool for analyzing structure and function of cells and tissues (Farkas, 2000). This unit reviews some of the main principles and developments in optical filtering, beginning with optical interference filters and colored glass filters that are the basis of color selection for the majority of commercial microscope systems. Table 2.4.1 gives a summary of the relative merits of the various waveTable 2.4.1
length selection techniques that will be discussed.
OPTICAL INTERFERENCE FILTERS AND COLORED GLASS FILTERS Optical interference filters and colored glass filters have gained wide acceptance over other devices for selecting wavelengths of light; they provide a small and relatively inexpensive way to achieve good wavelength selectivity. There are several types of optical filters that can be obtained. Long-pass filters have the light transmission spectra shown in Figure 2.4.1. They are specified according to their “cut-on” wavelength, which is located at half the maximal transmission of the filter. Short-pass filters transmit at short wavelengths. Transmission spectra for filters can be obtained on most scanning absorption spectrometers, although specialized instruments are required to obtain quantitative measurements of extreme levels of light blocking that can be incorporated into precision optical filters. Band-pass filters (Fig. 2.4.2) transmit light in a narrow range of wavelengths and are classified according to the width of the pass band at half-maximal transmission and the center wavelength of the pass region. Some types of filters have multiple pass bands.
Summary of the Relative Merits of the Various Wavelength Selection Techniques
Method/Tool
Main strengths
Main weaknesses
Filters
Excellent rejection Widely available Variable center wavelength
Fixed spectral characteristics Low
LCTF
UNIT 2.4
Reduced throughput Reduced rejection AOTF Variable center wavelength, Reduced throughput band-pass, and intensity Some image blur Fastest switching Reduced rejection Long acquisition time FTIS Good spectral resolution across large region High memory and processing requirements Grating imaging Only one exposure required High CCD requirements High memory and systems for all information processing requirements Gives spectral data across large wavelength region Prism imaging Gives spectral data across Requires scanning to get systems large wavelength region full x, y, and λ data
Relative cost
Moderate High
High
Moderate
Moderate
Image Cytometry Instrumentation Contributed by Alan S. Waggoner, Elliot S. Wachman, and Daniel L. Farkas Current Protocols in Cytometry (2001) 2.4.1-2.4.11 Copyright © 2001 by John Wiley & Sons, Inc.
2.4.1 Supplement 15
0
100 long-pass filter
75
25 50
light
25
filter
100 0
0 100 75
75
dichroic filter (also long-pass)
25
filter 50 25 0 400
long
light short 500
600 700 Wavelength
Reflectance (%)
Transmittance (%)
50
50 75 100
Figure 2.4.1 Transmission spectra for a long-pass interference filter. Maximum filter transmittance is nearly 100%. In the top panel, the wavelength at half-maximal transmission is ∼580 nm, so this filter is classified as a 580-nm long-pass interference filter. Manufacturers may add coatings to the filter that block the appearance of transmission in the wavelength range shorter than 450 nm. The bottom panel illustrates the transmission spectrum shift when the 580-nm long-pass filter of the top panel is rotated 45°. In the latter orientation the filter can be used as a 520-nm long-pass dichroic filter for color separation.
Transmittance (%)
100
pass-band center
pass-band width (at half-maximal transmittance)
0 540 560 580 600 620 640 660 680 Wavelength (nm)
Optical Filtering Systems for Wavelength Selection in Light Microscopy
Figure 2.4.2 Transmission spectrum of a band-pass interference filter. The center wavelength (top arrow) of this filter is ∼610 nm and the bandwidth at 50% maximal transmittance (between the two lower arrows) is ∼60 nm. Both specifications are required to define the filter, e.g., a 610/60 band-pass interference filter. The transmittance scale used in this figure does not show how effectively this filter blocks light outside the pass band. A logarithmic scale of 6 to 10 decades is required to define blocking ability for top-performance filters. An optical density scale (which is logarithmic) of 6 to 10 decades will also suffice (see for example Fig. 2.4.3).
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Dichroic filters are designed to steer light of different colors along separate paths. Usually one color is separated and sent at 90° to the path of the remaining light, as illustrated in Figure 2.4.1. Such “beam-splitters” are used for color separation and are ordinarily included in a standard epifluorescence microscope filter set. Depending on filter design, either short or long wavelengths can be reflected. Dichroic filters are basically long-pass, short-pass, or wideband-pass interference filters which are used in a 45° orientation and which have transmission specifications designated for the 45° orientation. The center wavelength and width of the pass region change with angle, as explained later.
CONSTRUCTION OF OPTICAL FILTER COMPONENTS Optical filters are fabricated in basically two ways: (1) by including light-absorbing molecules in gelatin or glass (colored-glass filters) to filter out certain colors of light, and (2) by generating interference effects that block the passage of certain wavelengths. The latter are called interference filters. Colored glass and gelatin filters have the advantage of being inexpensive and having strong light blocking in certain spectral regions. The transition from blocking to maximal transmission in a colored-glass filter is more gradual than for the band-pass interference filters and the latter have gradually taken over for applications that require highest optical selectivity. Interference filters also predominate in multicolor detection applications because they can be made with sharper transitions between transmitting and blocking regions. Another disadvantage of colored glass filters is that they often are weakly fluorescent; as a result, scattered excitation light may produce an unwanted fluorescence signal in the detection channel. Interference filters are prepared by creating partially reflective “cavities” of a thickness 1⁄2 the wavelength of light to be transmitted by the filter. The cavities are formed by layering dielectric materials on glass or quartz in a vacuum-deposition chamber. The incoming light forms an “in-phase” standing wave between the two partially reflective walls of the cavities and is passed efficiently through the filter. The slightly shorter or longer wavelengths, on the other hand, generate “out-of-phase” interference between the walls of the cavities and are therefore not transmitted, but reflected. Filters must be properly oriented in the light path, or the wavelength selectivity is altered. If
the filter is tipped from an orientation perpendicular to incoming light, a somewhat longer wavelength (different color) will be passed through the cavity because the cavity is thicker in the direction of incoming light. For this reason it is important to keep filters perpendicular to the beam of light (or 45° for dichroics). Figure 2.4.3 shows how the near-band light transmission is reduced and the pass-band walls become steeper as additional cavities are added. Multi-cavity filters are ideal for transmitting the fluorescent light in a defined spectral region without passing light from the excitation source or from other fluorophores. Thus, these highperformance multi-cavity filters are of great importance in multiparameter flow and image cytometry. However, filters with more cavities are more expensive. The following are features found in a good interference filter: 1. Good quality control of band-pass width, band center, and maximal transmission. 2. A set of spectral curves delivered with each filter. 3. High blocking (at least 6 orders of magnitude) in the blocking region. 4. Blocking into the UV and IR. 5. Freedom from pinhole defects in layer material. 6. Nonfluorescent coatings (including the glass, glues, blocking agents, and antireflective coatings). 7. Resistance to degradation by high-intensity light and humidity. The filter should be examined every few months for pinhole defects and color change. 8. Low registration shift: for imaging, the image should not shift on the camera as filter sets are changed. 9. Resistance to damage in handling. 10. Clear labeling that contains pass-band width, central wavelength, and product number information. 11. Arrow indicating proper insertion direction in optical path (usually pointing in the same direction as the incoming light).
INTEGRATING OPTICAL FILTER COMPONENTS INTO THE MICROSCOPE SYSTEM Optimal use of optical filters requires a clear understanding of the entire excitation-detection system (Galbraith et al., 1989). All fluorescence microscopes use some form of the epifluorescence filter system shown in Figure 2.4.4. The light source is usually an arc lamp (mercury or xenon) or a laser. Because lasers
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0
1
Optical density
2 2-cavity 3 4 3-cavity 5 6 6-cavity 400
450
500 Wavelength
4-cavity 550
600
Figure 2.4.3 Steeper walls and greater blocking outside the pass band are obtained by increasing the number of cavities of the filter. Note that the optical density (vertical axis) is logarithmic. Blocking can be achieved to greater than one photon per million.
emit at just a few wavelengths with very narrow wavelength bands, the excitation filter requirements for laser scanning microscopes are less stringent. Otherwise, considerations for filter set design are similar for arc-lamp- and laserbased microscopes. Each filter set must provide the optimal excitation wavelength band to reach the sample and allow the optimal emission wavelength band from the sample to reach the imaging camera (or eye). At the same time, the set should minimize light reaching the camera that is scattered excitation light, autofluorescence, or fluorescence from other excited fluorescent probes on the sample.
Single Fluorophore Imaging
Optical Filtering Systems for Wavelength Selection in Light Microscopy
Epifluorescence filter sets for single fluorophore detection are the simplest filter arrangement to set up. Still, there are several decisions that have to be made. First, how broad should the excitation filter bandwidth be and where should it be centered? If the excitation source is a laser, this decision is easy; a narrow bandpass filter centered at the laser wavelength should be used. For a mercury arc lamp, it is best to center the filter over one of the strong emission lines. If the absorption spectrum of the dye lies between mercury emission lines, the bandwidth of the filter should be wider to pass more photons within the absorption band of the dye. This is also the case for xenon arc
lamps, which have very little structure in their emission spectrum. It should be remembered that a neutral-density filter could be included in the excitation path to reduce illumination intensity if photobleaching is a problem. Generally, it is best to collect as much emission signal as possible, and often long-pass colored-glass filters are used as emission filters. For more challenging experiments where there is fluorescence background in the same wavelength region as the emission signal from the probe or label, it is better to take a smaller slice of the emission spectrum centered around the emission maximum of the dye by using a bandpass interference filter. Rejection of scattered excitation light becomes a challenge when capturing fluorescence from dyes (sometimes called fluorophores or fluorochromes) that emit close to the wavelength of excitation. Yet many of the best fluorophores—such as fluorescein, rhodamine, the BODIPYs, Cy2, Cy3, Cy5, and the Alexa dyes—have small Stokes shifts, i.e., separation between the longest-wavelength absorption band and the shortest-wavelength emission band (Fig. 2.4.5). In order to optimally excite and collect fluorescence from such fluorophores it is necessary for the excitation and emission filter pass-band regions to be positioned close to one another; however, the pass bands must not overlap or else scattered light will
2.4.4 Supplement 15
Current Protocols in Cytometry
Detector
excitation filter
emission (or barrier) filter
dichroic filter
Figure 2.4.4 Typical epifluorescence filter set used for detection of fluorescence with a microscope or other detection instrument. Excitation light from an arc, tungsten, or laser light source (solid line) is passed through an excitation filter which isolates the particular band of light for excitation. The excitation light is reflected 90° by a long-pass dichroic filter (with a cutoff halfway between the absorption maximum and the emission maximum of the fluorescent dye) and passes through a lens to illuminate the sample on a microscope slide. Fluorescence, which occurs at a longer wavelength, is collected by the lens and passes through the dichroic filter to the “barrier” or “emission filter.” The emission filter selects a band of fluorescence to be passed on to the detector.
dominate the signal received at the imaging camera, eye, or photomultiplier tube.
Multicolor-Multiprobe Imaging When the goal is to detect fluorescence signals from several different dyes with different excitation wavelengths, an efficient way to discriminate their signals is to use a separate epifluorescence filter set (as shown in Fig. 2.4.4) for each dye. This is a common procedure in microscopy. The image of each different color probe is acquired using a different filter set. Newer microscopes have multiple filter sets, sometimes under computer control. Multicolor fluorescence microscopy requires careful consideration of the spectral properties of the fluorescent probes. This has become a major challenge in light of the large number of new fluorescent labels and probes with different wavelengths that are designed for multiparameter analysis of cells. Composite images with five fluorescent probes have been published by DeBiasio et al. (1987). The problem is that even
the fluorescent dyes with the narrowest absorption and emission spectra have long tails (on the blue side of the absorption spectrum and the red side of the emission spectrum). Thus, each excitation wavelength used may excite more than one fluorophore, and each fluorophore may produce a signal that appears in fluorescence channels for longer-wavelength dyes. To choose filters with minimal fluorescence signal overlap, the first thing needed is a set of accurate emission spectra of the dyes that are used as probes or labels. The dyes must be in the same microenvironment as the biological experiment, or a similar one at least, because most dyes show wavelength shifts when they are transferred to solvents of different polarity. Once accurate emission spectra are obtained, the investigator should try to visualize what optimal bandwidths and central wavelengths should be. Usually the filter manufacturers will help with custom filter development, if they are provided with the absorption and emission spectra of the fluorophores used.
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Absorbance, fluorescence, or transmittance
400
450
500
550
600
Wavelength (nm) Figure 2.4.5 Absorption and fluorescence (emission) spectra of fluorescein. Superimposed on the spectra are transmission spectra (dotted lines) of excitation and emission band-pass filters of an epifluorescence filter set that would be used to generate and detect the fluorescein emission. The transmission spectrum of the dichroic filter of this set is not shown, but it would be a long-pass filter with a half-maximal transmission centered between the absorption and fluorescence peaks of this dye. It appears that the pass band of this emission filter could be moved to a shorter wavelength to capture a greater part of the maximum fluorescence of the dye, or the excitation filter could be moved to a slightly longer wavelength to provide more excitation. This may be true, but it would be necessary to determine more closely the amount of overlap of the excitation and emission pass bands before moving the pass band so that the amount of scattered light reaching the detector is not increased substantially. A sensitive spectrophotometer (absorption/transmission) would be used for this determination.
Optical Filtering Systems for Wavelength Selection in Light Microscopy
Quantitatively designing the bandwidths and central wavelengths of all the epifluorescence excitation and emission filters to be used in a three- to six-fluorophore experiment requires accurate absorption and emission spectra of the fluorophores (in a molecular environment that simulates that of the sample), information on the distribution of excitation wavelength of the excitation source (if an arc lamp), and the wavelength profile of detector sensitivity. A computer program can be used to optimize the filter parameters for epifluorescence microscopy (Galbraith et al., 1989). Designing emission and dichroic filters for laser scanning microscopes is also a challenge and has been extensively discussed by Brelje et al. (1993). Again, filter manufactures and microscope companies should be consulted for help ( e.g., C hr oma Technology Corp.;
http://www.chroma.com; Omega Technology, Inc.; http://www.omegafilters.com). When constructing filter sets for multicolor imaging, the thickness and flatness of all of the dichroic and emission filters, as well as their orientation within the filter holder of the microscope, must be carefully controlled in order to minimize registration shifts of the different color images. Alternatively, the registration shifts can be measured with a sample that produces a pattern in each of the color channels. Correction factors can then be applied by software methods to register other acquired images (Galbraith and Farkas, 1993).
Single-Laser-Line Excitation with Multiple Fluorescence Signals: Laser Scanning Microscopes For multiple fluorescence signals, the balancing trick is to situate the emission band-pass
2.4.6 Supplement 15
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Fluorescence/Transmittance
488 nm exc.
450
500
550
600
650
Wavelength (nm) Figure 2.4.6 Fluorescence spectra (solid) of fluorescein (green fluorescence, 525-nm peak) and R-PE (orange fluorescence, 575-nm peak) superimposed on the transmission spectra (dotted) of the band-pass filters used to collect the fluorescence signals. Notice the spillover of the fluorescein fluorescence into the orange R-PE detection channel. The intensity of spillover is some fraction of the intensity in the main detection channel, so that the spillover can be subtracted by electronic or software methods. This process is called compensation (UNIT 1.14).
filters over fluorescence peaks of the dyes in such a way as to optimize light collection for each dye in its designated detection channel, while at the same time minimizing the detection of fluorescence signals from other dyes that spill over into that detection channel (Fig. 2.4.6). When it is not possible to exclude spillover signals with the filter sets, there are methods of removing spillover signals by electronic or software compensation, in which a portion of signal from one detector is subtracted from the signal from another detector. This is routinely done in two-color flow cytometry experiments where the fluorescein signal spills into the phycoerythrin channel (Fig. 2.4.6), but it can also be done with laser scanning microscopes.
Multi-Band-Pass Excitation with Multi-Band-Pass Emission A unique method for simultaneously allowing human (or color camera) visualization of two, three, or even four different color fluorophores became available with the development
of multi-band-pass filters (Pinkel et al., 1989). For example, a three-color multi-band-pass excitation filter contains three pass bands optimized for excitation at the absorption peaks of the three dyes for which the set has been designed (Fig. 2.4.7). Similarly, there is a tripleband-pass emission filter for collecting the three fluorescence signals. The three bands of the multi-band-pass emission filter lie between the excitation bands and are offset to longer wavelengths. The dichroic filter is also a tripleband-pass filter set at 45° and is designed to split each of the excitation-emission bands. These remarkable filters have become especially useful for imaging chromosome-specific probes and paints (Pinkel et al., 1989).
Multi-Band-Pass Emission with Excitation Changer In this hybrid system, the emission filter and dichroic filter are triple-band-pass filters, but the excitation filters are single-band-pass filters and are independently moved into position. With this setup, there is no possibility of regis-
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Transmittance (%)
100
0 350
400
450
500
550
600
650
Wavelength Figure 2.4.7 Transmission spectrum of a triple-band-pass excitation filter (dotted) and a tripleband-pass emission filter (solid). Notice that the excitation and emission pass bands are located to excite and collect the fluorescence from three dyes (such as DAPI, fluorescein, and Texas Red) without scattered light reaching the eye or imaging camera.
tration shifts, which can occur when some epifluorescence filter sets are exchanged. A highresolution black-and-white imaging camera is used in this mode. Wavelength discrimination is not as good as with independent epifluorescence filter sets but is better than with dual multi-band-pass filter sets.
OTHER METHODS OF WAVELENGTH SELECTION While optical filters provide multi-wavelength capabilities, a number of more advanced methods additionally allow for a continuous scan of the wavelength domain, thus enabling spectral imaging microscopy (Farkas, 2000).
Liquid Crystal Tunable Filters
Optical Filtering Systems for Wavelength Selection in Light Microscopy
LCTFs consist of a number of liquid crystal layers, each of which passes a number of different frequencies; stacking them results in a single dominant transmission band, along with much smaller side-bands (and in some cases, unfortunately, additional major transmission
bands far from the desired spectral region). This concept is based on the Lyot birefringent interferometer. In addition to a computer, a small controller box is needed to drive an LCTF assembly. LCTFs can be switched from wavelength to wavelength in ∼50 msec, and are optically well behaved in that they do not seem to induce image distortion or shift. Each filter assembly can span ∼1 octave of wavelength (e.g., 400 to 800 nm) and their useful range can extend into the near IR. They can be introduced into either the illumination (excitation) or emission pathways, or both. Throughput is a problem, however, in that half the light corresponding to one polarization state is lost automatically, and peak transmission of the other half is limited. Overall throughput of a device approximating the bandwidth of the interference filter is ∼10%. In addition, focal-length variations can be introduced into the optical system as the LCTF is tuned from wavelength to wavelength. Finally, out-of-band rejection is not sufficient to prevent excitation light from leaking
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Current Protocols in Cytometry
into the emission channel without the use of a dichroic mirror or cross-polarization (Hoyt, 1996). Apart from these limitations, LCTFs are useful when multi-spectral microscopy is required, since their electronic tunability provides great flexibility in choice of center wavelengths for illumination and/or fluorescence excitation.
Acousto-Optic Tunable Filters In an AOTF, a radio frequency (RF) electronic signal is applied to an acoustic transducer attached to one face of the crystal. This generates a traveling acoustic wave in the crystal at the frequency of the applied RF signal. The resulting modulation of the crystal’s index of refraction acts as a sinusoidal phase grating for incoming light. Hence, when broad-band light is incident on an AOTF the narrow-band filtered light exiting the AOTF is angularly deflected away from the incident beam, as it would be by a conventional diffraction grating. The central wavelength of this filtered beam is determined by the acoustic frequency of the AOTF; this wavelength can be changed in less than 50 µsec to any other wavelength—several orders of magnitude faster than with other spectral filtering technologies. AOTFs have the ability to vary not only the wavelength, but also the bandwidth and the intensity of the transmitted light. Thus, experiments involving luminescence lifetimes or very rapid acquisition of multiple wavelengths are possible using this technology. Over the past few years, considerable progress has been made (Farkas et al., 1998) in overcoming the difficulties of using AOTFs for high-resolution imaging: (1) as with LCTFs, throughput is reduced since only one polarization state is typically used; (2) intrinsic image blur is present; and (3) out-of-band rejection is no greater than 10−2 to 10−3. Approaches and solutions to some of these problems have been described previously (Wachman et al., 1997), including use of both polarization beams to increase throughput, use of long-path-length imaging AOTF crystals to reduce blur, and transducer apodization (in the emission path) to improve out-of-band rejection. Nevertheless, additional optics (such as dichroics or rejection filters) are typically used to achieve adequate rejection for fluorescence measurements. Like LCTFs, AOTFs can be used for excitation and emission and give the benefit of continuous tunability. In addition, however, they have the capability for variable bandwidths,
which may be useful in multi-fluorophore applications. Finally, AOTFs are the technology of choice if sub-millisecond wavelength switching speed is required.
Fourier Transform Methods Fourier transform interference spectroscopy (FTIS) is a technique most widely used for generating spectral information in the infrared region. This method takes advantage of the principle that when light is allowed to interfere with itself at a number of optical path lengths, the resulting interferogram reflects its spectral constitution. Monochromatic light generates a pure cosine wave; multispectral light, which consists of a mixture of wavelengths, will generate interferograms that can be modeled as a sum of their respective cosine waves. An inverse Fourier transform of such an interferogram will regenerate the presence and intensities of all the contributing wavelengths. The classical Michelson interferometer, which uses two separate light paths to generate an interference pattern, is usable in the infrared region of the optical spectrum, because mechanical tolerances at these wavelengths are not relatively large. However, in the visible region, the required optical pathlength differences are much smaller, and vibration and mechanical imprecision become limiting. The Sagnac interferometer design, which directs the interfering light beams in opposite directions around a common path, overcomes many of these problems, and has been successfully adopted for use for visible light imaging spectroscopy by Applied Spectral Imaging (Garini et al., 1996). FTIS systems can be used to provide very rapid acquisition of emission spectra and have been used in experimental multicolor fluorescence flow cytometry and image cytometry (Buican, 1990; Cabib et al., 1996; Farkas et al., 1996). The optimal use of such a device occurs when several fluorophores can be simultaneously excited with a single laser wavelength but fluoresce at different wavelengths, as is true with fluorescein and phycoerythrin. In this case a long-pass filter can be used to remove laser scattering and the FTIS system can acquire the emission spectra. It is much more challenging to use an FTIS system when the fluorophores all absorb in different regions of the spectrum. In such a case, a double- or triple-dichroic filter cube (e.g., Fig. 2.4.7) is required in the optical path to ensure that each excitation band is excluded from the interferometer. Thus, instead of being able to collect light from a broad spectral region (and
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thereby able to acquire detailed spectral shape information), the instrument is presented with light from several relatively narrow spectral regions inside of which it may be difficult to get enough data to distinguish spectrally similar dyes. Significant advances have been made in this area. It has been shown, for example, that six fluorophores with different spectra can be resolved by analyzing wavelength shifts that can be observed in the three pass-band regions of a triple-band-pass filter. A commercially available instrument (Applied Spectral Imaging) has been used with five fluorescent dyes (Garini et al., 1996) to resolve all chromosomes in human metaphase spreads that have been hybridized with chromosome paints (Schröck et al., 1996). Such instrumentation is also useful for multicolor immunofluorescence imaging (Farkas et al., 1998). There are advantages and drawbacks to Fourier transform techniques. As with tunable filters, and in contrast to fixed interference filters, wavelength ranges can be adjusted to match the spectral properties of the image. Compared to monochromator- or prism-based devices, FT spectroscopy enjoys a throughput advantage, as it does not require a narrow-slit aperture that would reduce the signal-to-noise ratio. Finally, FITS systems provide good spectral resolution (5 mW). In addition, commercial systems exist which can switch wavelengths in 180°C in a sealed tube
continued
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Table 3.4.1
Commonly Used Hazardous Chemicalsa, continued
Chemical
Hazards
Remarksb
Hydrogen peroxide (30%)
Carcinogenic, corrosive, mutagenic, oxidizer
Hydroxylamine
Corrosive, flammable, mutagenic, toxic Carcinogenic Corrosive, toxic Carcinogenic, mutagenic, toxic Carcinogenic, mutagenic Carcinogenic, toxic Stench, toxic Teratogenic, toxic Teratogenic, toxic Carcinogenic, mutagenic, teratogenic, toxic Mutagenic, toxic Carcinogenic, toxic Teratogenic, toxic Mutagenic
Avoid bringing into contact with organic materials, which may form explosive peroxides; may decompose violently in contact with metals, salts, or oxidizable materials; see Basic Protocol 7 Explodes in air at >70°C
3-β-Indoleacrylic acid (IAA) Iodine Iodoacetamide Janus green B Lead compounds 2-Mercaptoethanol (2-ME) Mercury compounds Methionine sulfoximine (MSX) Methotrexate (amethopterin) Methylene blue Methyl methanesulfonate (MMS) Mycophenolic acid (MPA) Neutral red Nigrosin, water soluble Nitric acid, concentrated Nitroblue tetrazolium (NBT) Orcein, synthetic Oxonols Paraformaldehyde Phenol
See Basic Protocol 8 See Basic Protocol 2
See Basic Protocol 9
See Basic Protocol 2 See Basic Protocol 5 See Basic Protocol 2 See Basic Protocol 2
Corrosive, oxidizer, teratogenic, toxic Toxic See Basic Protocol 2
Phenylmethylsulfonyl fluoride (PMSF) Phorbol 12-myristate 13-acetate (PMA) Phycoerythrins (PE) Piperidine Potassium hydroxide, concentrated
Toxic Toxic Carcinogenic, corrosive, teratogenic, toxic Enzyme inhibitor Carcinogenic, toxic Toxic Flammable liquid, teratogenic, toxic Corrosive, toxic
Propane sultone Propidium iodide (PI) Pyridine Rhodamine and derivatives Rose Bengal Safranine O
Carcinogenic, toxic Mutagenic Flammable liquid, toxic Toxic Carcinogenic, teratogenic Mutagenic
Readily absorbed through the skin See Basic Protocol 11
Produces a highly exothermic reaction when solid is added to water See Basic Protocol 5 See Basic Protocol 2 or 6
See Basic Protocol 2 See Basic Protocol 2
continued
3.4.4 Supplement 20
Current Protocols in Cytometry
Table 3.4.1
Commonly Used Hazardous Chemicalsa, continued
Chemical
Hazards
Remarksb
Sodium azide
Carcinogenic, toxic
Adding acid liberates explosive volatile, toxic hydrazoic acid; can form explosive heavy metal azides, e.g., with plumbing fixtures—do not discharge down drain; see Basic Protocol 10
Sodium deoxycholate (Na-DOC) Sodium dodecyl sulfate (sodium lauryl sulfate, SDS) Sodium hydroxide, concentrated
Carcinogenic, teratogenic, toxic Sensitizing, toxic
Sodium nitrite Sulfuric acid, concentrated
SYTO dyes Tetramethylammonium chloride (TMAC) N,N,N′,N′-Tetramethyl-ethylenediamine (TEMED) Texas Red (sulforhodamine 101, acid chloride) Toluene Toluidine blue O Nα-p-Tosyl-L-lysine chloromethyl ketone (TLCK) N-p-Tosyl-L-phenylalanine chloromethyl ketone (TPCK) Trichloroacetic acid (TCA) Triethanolamine acetate (TEA) Trifluoroacetic acid (TFA) Trimethyl phosphate (TMP) Trypan blue Xylenes
Corrosive, toxic
A highly exothermic reaction ensues when the solid is added to water
Carcinogenic Corrosive, oxidizer, teratogenic, toxic Reaction with water is very exothermic; always add concentrated sulfuric acid to water, never water to acid Toxic Toxic Corrosive, flammable liquid, toxic Toxic Flammable liquid, teratogenic, toxic Mutagenic, toxic Toxic, enzyme inhibitor
See Basic Protocol 2 See Basic Protocol 11
Toxic, mutagenic, enzyme inhibitor
See Basic Protocol 11
Carcinogenic, corrosive, teratogenic, toxic Carcinogenic, toxic Corrosive, toxic Carcinogenic, mutagenic, teratogenic May explode on distillation Carcinogenic, mutagenic, teratogenic See Basic Protocol 2 Flammable liquid, teratogenic, toxic
aFor extensive information on the hazards of these and other chemicals as well as cautionary details, see Bretherick (1986), O’Neil (2001), Furr (2000),
Lewis (1999), Lunn and Sansone (1994a), and Bretherick et al. (1999). bCAUTION: These chemicals should be handled only in a chemical fume hood by knowledgeable workers equipped with eye protection, lab coat, and
gloves. The laboratory should be equipped with a safety shower and eye wash. Additional protective equipment may be required.
No gloves resist all chemicals, and no gloves resist any chemicals indefinitely. Disposable gloves labeled “exam” or “examination” are primarily for protection from biological materials (e.g., viruses, bacteria, feces, blood). They are not designed for and usually have not been tested for resistance to chemicals. Disposable gloves generally offer extremely marginal protection from chemical hazards in most cases and should be removed immediately upon contamination before the chemical can pass through. If possible, design handling procedures to eliminate or reduce potential for contamination. Never assume that disposable gloves will offer the same protection or even have the same properties as
Safety Procedures and Quality Control
3.4.5 Current Protocols in Cytometry
Supplement 20
Table 3.4.2
Examples of Chemical Incompatibility
Chemical
Incompatible with
Acetic acid
Aldehydes, bases, carbonates, chromic acid, ethylene glycol, hydroxides, hydroxyl compounds, metals, nitric acid, oxidizers, perchloric acid, peroxides, phosphates, permanganates, xylene Acids, amines, concentrated nitric and sulfuric acid mixtures, oxidizers, plastics Copper, halogens, mercury, oxidizers, potassium, silver Acids, aldehydes, carbon dioxide, carbon tetrachloride or other chlorinated hydrocarbons, halogens, ketones, plastics, sulfur, water Acids, aldehydes, amides, calcium hypochlorite, hydrofluoric acid, halogens, heavy metals, mercury, oxidizers, plastics, sulfur Acids, alkalis, chlorates, chloride salts, flammable and combustible materials, metals, organic materials, phosphorus, reducing agents, sulfur, urea Acids, aluminum, dibenzoyl peroxide, oxidizers, plastics Any reducing agent Acids, heavy metals, oxidizers Acetaldehyde, alcohols, alkalis, amines, ammonia, combustible materials, ethylene, fluorine, hydrogen, ketones (e.g., acetone, carbonyls), metals, petroleum gases, sodium carbide, sulfur Acids, ethanol, fluorine, organic materials, water Alkali metals, calcium hypochlorite, halogens, oxidizers Sodium Acids, ammonium salts, finely divided organic or combustible materials, powdered metals, sulfur Acetylene or other hydrocarbons, alcohols, ammonia, benzene, butadiene, butane, combustible materials, ethylene, flammable compounds (e.g., hydrazine), hydrogen, hydrogen peroxide, iodine, metals, methane, nitrogen, oxygen, propane (or other petroleum gases), sodium carbide, sodium hydroxide Ammonia, hydrogen, hydrogen sulfide, mercury, methane, organic materials, phosphine, phosphorus, potassium hydroxide, sulfur Acetic acid, acetone, alcohols, alkalis, ammonia, bases, benzene, camphor, flammable liquids, glycerin (glycerol), hydrocarbons, metals, naphthalene, organic materials, phosphorus, plastics Acetylene, calcium, hydrocarbons, hydrogen peroxide, oxidizers Acids (organic or inorganic) Acids, alkaloids, aluminum, iodine, oxidizers, strong bases Ammonium nitrate, chromic acid, halogens, hydrogen peroxide, nitric acid, oxidizing agents in general, oxygen, sodium peroxide All other chemicals
Acetone Acetylene Alkali metals, alkaline earth metals Ammonia (anhydrous)
Ammonium nitrate
Aniline Arsenical materials Azides Bromine
Calcium oxide Carbon (activated) Carbon tetrachloride Chlorates Chlorine
Chlorine dioxide
Chromic acid, chromic oxide
Copper Cumene hydroperoxide Cyanides Flammable liquids
Fluorine Safe Use of Hazardous Chemicals
continued
3.4.6 Supplement 20
Current Protocols in Cytometry
Table 3.4.2
Examples of Chemical Incompatibility, continued
Chemical
Incompatible with
Hydrocarbons (liquid or gas) Hydrocyanic acid Hydrofluoric acid
See flammable liquids
Hydrogen peroxide Hydrogen sulfide Hydroperoxide Hypochlorites Iodine Mercury Nitric acid Nitrites Nitroparaffins Oxalic acid Oxygen
Perchloric acid Peroxides, organic Phosphorus (white) Potassium chlorate
Potassium perchlorate Potassium permanganate Selenides and tellurides Silver Sodium Sodium nitrate Sodium peroxide
Sulfides Sulfuric acid
Alkali, nitric acid Ammonia, metals, organic materials, plastics, silica (glass, including fiberglass), sodium All organics, most metals or their salts, nitric acid, phosphorus, sodium, sulfuric acid Acetylaldehyde, fuming nitric acid, metals, oxidizers, sodium, strong bases Reducing agents Acids, activated carbon Acetylaldehyde, acetylene, ammonia, hydrogen, metals, sodium Acetylene, aluminum, amines, ammonia, calcium, fulminic acid, lithium, oxidizers, sodium Acids, nitrites, metals, most organics, plastics, sodium, sulfur, sulfuric acid Acids Amines, inorganic bases Mercury, oxidizers, silver, sodium chlorite All flammable and combustible materials, ammonia, carbon monoxide, grease, metals, oil, phosphorus, polymers All organics, bismuth and alloys, dehydrating agents, grease, hydrogen halides, iodides, paper, wood Acids (organic or mineral), avoid friction, store cold Air, alkalis, oxygen, reducing agents Acids, ammonia, combustible materials, fluorine, hydrocarbons, metals, organic materials, reducing agents, sugars Alcohols, combustible materials, fluorine, hydrazine, metals, organic matter, reducing agents, sulfuric acid Benzaldehyde, ethylene glycol, glycerin, sulfuric acid Reducing agents Acetylene, ammonium compounds, fulminic acid, oxalic acid, ozonides, peroxyformic acid, tartaric acid Acids, carbon dioxide, carbon tetrachloride, hydrazine, metals, oxidizers, water Acetic anhydride, acids, metals, organic matter, peroxyformic acid, reducing agents Acetic anhydride, benzaldehyde, benzene, carbon disulfide, ethyl acetate, ethyl or methyl alcohol, ethylene glycol, furfural, glacial acetic acid, glycerin, hydrogen sulfide, metals, methyl acetate, oxidizers, peroxyformic acid, phosphorus, reducing agents, sugars, water Acids Alcohols, bases, chlorates, perchlorates, permanganates of potassium, lithium, sodium, magnesium, calcium Safety Procedures and Quality Control
3.4.7 Current Protocols in Cytometry
Supplement 20
Table 3.4.3
Safe Use of Hazardous Chemicals
Chemical Resistance of Commonly Used Glovesa,b
Chemical
Neoprene Latex gloves gloves
Butyl gloves
Nitrile gloves
*Acetaldehyde Acetic acid *Acetone Ammonium hydroxide *Amyl acetate Aniline *Benzaldehyde *Benzene Butyl acetate Butyl alcohol Carbon disulfide *Carbon tetrachloride *Chlorobenzene *Chloroform Chloronaphthalene Chromic acid (50%) Cyclohexanol *Dibutyl phthalate Diisobutyl ketone Dimethylformamide Dioctyl phthalate Epoxy resins, dry *Ethyl acetate Ethyl alcohol *Ethyl ether *Ethylene dichloride Ethylene glycol Formaldehyde Formic acid Freon 11, 12, 21, 22 *Furfural Glycerin Hexane Hydrochloric acid Hydrofluoric acid (48%) Hydrogen peroxide (30%) Ketones Lactic acid (85%) Linseed oil Methyl alcohol Methylamine Methyl bromide *Methyl ethyl ketone *Methyl isobutylketone Methyl methacrylate Monoethanolamine
VG VG G VG F G F P G VG F F F G F F G G P F G VG G VG VG F VG VG VG G G VG F VG VG G G VG VG VG F G G F G VG
VG VG VG VG F F G P F VG F P F P F F G G G G F VG G VG VG F VG VG VG F G VG P G G G VG VG F VG G G VG VG VG VG
G VG P VG P P G F P VG F G P E F F VG G P G VG VG F VG G P VG VG VG G G VG G G G G P VG VG VG G F P P F VG
G VG VG VG P F F P F VG F P P P P P F P F F P VG F VG G P VG VG VG P G VG P G G G VG VG P VG F F G F G G
continued
3.4.8 Supplement 20
Current Protocols in Cytometry
Table 3.4.3
Chemical Resistance of Commonly Used Glovesa,b, continued
Chemical
Neoprene Latex gloves gloves
Butyl gloves
Nitrile gloves
Morpholine Naphthalene Naphthas, aliphatic Naphthas, aromatic *Nitric acid Nitric acid, red and white fuming Nitropropane (95.5%) Oleic acid Oxalic acid Palmitic acid Perchloric acid (60%) Perchloroethylene Phenol Phosphoric acid Potassium hydroxide Propyl acetate i-Propyl alcohol n-Propyl alcohol Sodium hydroxide Styrene (100%) Sulfuric acid Tetrahydrofuran *Toluene Toluene diisocyanate *Trichloroethylene Triethanolamine Tung oil Turpentine *Xylene
VG G VG G G P F VG VG VG VG F VG VG VG G VG VG VG P G P F F F VG VG G P
VG F F P F P F G VG VG G P G VG VG G VG VG VG P G F P G P G F F P
G G VG G F P F VG VG VG G G F VG VG F VG VG VG F G F F F G VG VG VG F
VG F F P F P P F VG VG F P F G VG F VG VG VG P G F P G F G P F P
aPerformance varies with glove thickness and duration of contact. An asterisk indicates limited use. Abbreviations: VG,
very good; G, good; F, fair; P, poor (do not use). bAdapted from the July 8, 1998, version of the DOE OSH Technical Reference Chapter 5 (APPENDIX C at
http://tis.eh.doe.gov/docs/osh_tr/ch5c.html). For more information also see Forsberg and Keith (1999).
nondisposables. Select gloves carefully and always look for some evidence that they will protect against the materials being used. Inspect all gloves before every use for possible holes, tears, or weak areas. Never reuse disposable gloves. Clean reusable gloves after each use and dry carefully inside and out. Observe all common-sense precautions—e.g., do not pipet by mouth, keep unauthorized persons away from hazardous chemicals, do not eat or drink in the lab, wear proper clothing in the lab (sandals, open-toed shoes, and shorts are not appropriate). Order hazardous chemicals only in quantities that are likely to be used in a reasonable time. Buying large quantities at a lower unit cost is no bargain if someone (perhaps you) has to pay to dispose of surplus quantities. Substitute alcohol-filled thermometers for mercury-filled thermometers, which are a hazardous chemical spill waiting to happen.
Safety Procedures and Quality Control
3.4.9 Current Protocols in Cytometry
Supplement 20
Although any number of chemicals commonly used in laboratories are toxic if used improperly, the toxic properties of a number of reagents require special mention. Chemicals that exhibit carcinogenic, corrosive, flammable, lachrymatory, mutagenic, oxidizing, teratogenic, toxic, or other hazardous properties are listed in Table 3.4.1. Chemicals listed as carcinogenic range from those accepted by expert review groups as causing cancer in humans to those for which only minimal evidence of carcinogenicity exists. No effort has been made to differentiate the carcinogenic potential of the compounds in Table 3.4.1. Oxidizers may react violently with oxidizable material (e.g., hydrocarbons, wood, and cellulose). Before using any of these chemicals, thoroughly investigate all its characteristics. Material Safety Data Sheets are readily available; they list some hazards but vary widely in quality. A number of texts describing hazardous properties are listed at the end of this unit (see Literature Cited). In particular, Sax’s Dangerous Properties of Industrial Materials, 10th ed. (Lewis, 1999) and the Handbook of Reactive Chemical Hazards, 6th ed. (Bretherick et al., 1999) give comprehensive listings of known hazardous properties; however, these texts list only the known properties. Many chemicals, especially fluorochromes, have been tested only partially or not at all. Prudence dictates that, unless there is good reason for believing otherwise, all chemicals should be regarded as volatile, highly toxic, flammable human carcinogens and should be handled with great care. Waste should be segregated according to institutional requirements, for example, into solid, aqueous, nonchlorinated organic, and chlorinated organic material, and should always be disposed of in accordance with all applicable federal, state, and local regulations. Extensive information and cautionary details along with techniques for the disposal of chemicals in laboratories have been published (Bretherick, 1986; Lunn and Sansone, 1994a; O’Neil, 2001; Furr, 2000). Some commonly used disposal procedures are outlined in Basic Protocols 1 to 11. Incorporation of these procedures into laboratory protocols can help to minimize waste disposal problems. Alternate Protocols 1 to 7 describe decontamination methods for some of the chemicals. Support Protocols 1 to 9 describe analytical techniques that are used to verify that reagents have been decontaminated; with modification, these assays may also be used to determine the concentration of a particular chemical. DISPOSAL METHODS A number of procedures for the disposal of hazardous chemicals are available; protocols for the disposal and decontamination of some hazardous chemicals commonly encountered in cytometry laboratories are listed in Table 3.4.4. These procedures are necessarily brief; for full details consult the original references or a collection of these procedures (see Lunn and Sansone, 1994a). CAUTION: These disposal methods should be carried out only in a chemical fume hood by workers equipped with eye protection, a lab coat, and gloves. Additional protective equipment may be necessary. BASIC PROTOCOL 1
Safe Use of Hazardous Chemicals
DISPOSAL OF BENZIDINE AND DIAMINOBENZIDINE Benzidine and diaminobenzidine can be degraded by oxidation with potassium permanganate (Castegnaro et al., 1985; Lunn and Sansone, 1991a). This protocol presents a method for decontamination of benzidine and diaminobenzidine in bulk. This method can also be adapted to the decontamination of benzidine and diaminobenzidine spills (see Alternate Protocol 1). These compounds can also be removed from solution using horseradish peroxidase in the presence of hydrogen peroxide (see Alternate Protocol 2). Destruction and decontamination are >99%. Support Protocol 1 is used to test for the presence of benzidine and diaminobenzidine.
3.4.10 Supplement 20
Current Protocols in Cytometry
Table 3.4.4
Protocols for Disposal of Some Hazardous Chemicals
Protocol
Disposal method for
Basic Protocol 1 Alternate Protocol 1 Alternate Protocol 2 Support Protocol 1
Benzidine and diaminobenzidine Spills of benzidine and diaminobenzidine Aqueous solutions of benzidine and diaminobenzidine Analysis for benzidine and diaminobenzidine
Basic Protocol 2 Alternate Protocol 3 Support Protocol 2
Biological stains Large volumes of dilute biological stains Analysis for biological stains
Basic Protocol 3
Silanes
Basic Protocol 4 Support Protocol 3
Cyanide and cyanogen bromide Analysis for cyanide
Basic Protocol 5
Dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, 1,3-propane sultone Analysis for dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, 1,3-propane sultone
Support Protocol 4 Basic Protocol 6 Alternate Protocol 4 Alternate Protocol 5 Alternate Protocol 6 Support Protocol 5
Ethidium bromide and propidium iodide Equipment contaminated with ethidium bromide Ethidium bromide in isopropanol containing cesium chloride Ethidium bromide in alcohols Analysis for ethidium bromide and propidium iodide
Basic Protocol 7
Hydrogen peroxide
Basic Protocol 8
Iodine
Basic Protocol 9 Alternate Protocol 7 Support Protocol 6
Mercury compounds Waste water containing mercury Analysis for mercury
Basic Protocol 10 Support Protocol 7 Support Protocol 8
Sodium azide Analysis for sodium azide Analysis for nitrite
Basic Protocol 11 Support Protocol 9
Enzyme inhibitors Analysis for enzyme inhibitors
Materials Benzidine or diaminobenzidine tetrahydrochloride dihydrate 0.1 M HCl (for benzidine) 0.2 M potassium permanganate: prepare immediately before use 2 M sulfuric acid Sodium metabisulfite 10 M potassium hydroxide (KOH) Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1) 1. For each 9 mg benzidine, add 10 ml of 0.1 M HCl or for each 9 mg diaminobenzidine tetrahydrochloride dihydrate, add 10 ml water. Stir the solution until the aromatic amine has completely dissolved.
Safety Procedures and Quality Control
3.4.11 Current Protocols in Cytometry
Supplement 20
2. For each 10 ml of solution, add 5 ml freshly prepared 0.2 M potassium permanganate and 5 ml of 2 M sulfuric acid. Allow the mixture to stand for ≥10 hr. 3. Add sodium metabisulfite until the solution is decolorized. 4. Add 10 M KOH to make the solution strongly basic, pH >12. CAUTION: This reaction is exothermic.
5. Dilute with 5 vol water and pass through filter paper to remove manganese compounds. 6. Test the filtrate for the presence of aromatic amines (i.e., benzidine or diaminobenzidine; see Support Protocol 1). 7. Neutralize the filtrate with acid and discard. ALTERNATE PROTOCOL 1
DECONTAMINATION OF SPILLS INVOLVING BENZIDINE AND DIAMINOBENZIDINE Additional Materials (also see Basic Protocol 1) Glacial acetic acid 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid: prepare immediately before use Absorbent material (e.g., paper towels, Kimwipes) High-efficiency particulate air (HEPA) vacuum (Fisher) Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1) CAUTION: This procedure may damage painted surfaces and Formica. 1. Remove as much of the spill as possible using absorbent material and high-efficiency particulate air (HEPA) vacuuming. 2. Wet the surface with glacial acetic acid until all the amines are dissolved, then add an excess of freshly prepared 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid to the spill area. Allow the mixture to stand ≥10 hr. 3. Ventilate the area and decolorize with sodium metabisulfite. 4. Mop up the liquid with paper towels. Squeeze the solution out of the towels and collect in a suitable container. Discard towels as hazardous solid waste. 5. Add 10 M KOH to make the solution strongly basic, pH ≥12. CAUTION: This reaction is exothermic.
6. Dilute with 5 vol water and filter through filter paper to remove manganese compounds. 7. Test the filtrate for the presence of aromatic amines (i.e., benzidine or diaminobenzidine; see Support Protocol 1). 8. Neutralize the filtrate with acid and discard it. 9. Verify complete decontamination by wiping the surface with a paper towel moistened with water and squeezing the liquid out of the towel. Test the liquid for the presence of benzidine or diaminobenzidine (see Support Protocol 1). Repeat steps 1 to 9 as necessary.
Safe Use of Hazardous Chemicals
3.4.12 Supplement 20
Current Protocols in Cytometry
DECONTAMINATION OF AQUEOUS SOLUTIONS OF BENZIDINE AND DIAMINOBENZIDINE The enzyme horseradish peroxidase catalyzes the oxidation of the amine to a radical which diffuses into solution and polymerizes. The polymers are insoluble and fall out of solution.
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) Aqueous solution of benzidine or diaminobenzidine 1 N HCl or NaOH 3% (v/v) hydrogen peroxide Horseradish peroxidase (see recipe) 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid 5% (w/v) ascorbic acid Porous glass filter or Sorvall GLC-1 centrifuge or equivalent Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1) 1. Adjust the pH of the aqueous benzidine or diaminobenzidine solution to 5 to 7 with 1 N HCl or NaOH as required and dilute so the concentration of aromatic amines is ≤100 mg/liter. 2. For each liter of solution, add 3 ml of 3% hydrogen peroxide and 300 U horseradish peroxidase. Let the mixture stand 3 hr. 3. Remove the precipitate by filtering the solution through a porous glass filter or by centrifuging 5 min at room temperature in a benchtop centrifuge to pellet the precipitate. The precipitate is mutagenic and should be treated as hazardous waste.
4. Immerse the porous glass filter in 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid. Clean the filter in a conventional fashion and discard potassium permanganate/sulfuric acid solution as described for benzidine and diaminobenzidine (see Basic Protocol 1). 5. For each liter of filtrate, add 100 ml of 5% ascorbic acid. 6. Test the filtrate for the presence of aromatic amines (see Support Protocol 1). 7. Discard the decontaminated filtrate. ANALYTICAL PROCEDURES TO DETECT BENZIDINE AND DIAMINOBENZIDINE Reversed-phase HPLC (Snyder et al., 1997) is used to test for the presence of aromatic amines. The limit of detection is 1 µg/ml for benzidine and 0.25 µg/ml for diaminobenzidine.
SUPPORT PROTOCOL 1
Materials Decontaminated aromatic amine solution 10:30:20 (v/v/v) acetonitrile/methanol/1.5 mM potassium phosphate buffer (1.5 mM K2HPO4/1.5 mM KH2PO4) (benzidine) or 75:25 (v/v) methanol/1.5 mM potassium phosphate buffer (diaminobenzidine) 250-mm × 4.6-mm-i.d. Microsorb C-8 reversed-phase HPLC column (Varian) or equivalent Additional reagents and equipment for reversed-phase liquid chromatography (Snyder et al., 1997)
Safety Procedures and Quality Control
3.4.13 Current Protocols in Cytometry
Supplement 20
Analyze the decontaminated aromatic amine solution by reversed-phase HPLC using a 250-mm × 4.6-mm-i.d. Microsorb C-8 column or equivalent. To detect benzidine, elute with 10:30:20 (v/v/v) acetonitrile/methanol/1.5 mM potassium phosphate buffer at a flow rate of 1.5 ml/min and UV detection at 285 nm. To detect diaminobenzidine, elute with 75:25 (v/v) methanol/1.5 mM potassium phosphate buffer at a flow rate of 1 ml/min and UV detection at 300 nm. BASIC PROTOCOL 2
DISPOSAL OF BIOLOGICAL STAINS Biological stains (Table 3.4.5), as well as ethidium bromide and propidium iodide, can be removed from solution using the polymeric resin Amberlite XAD-16. The decontaminated solution may be disposed of as nonhazardous aqueous waste and the resin as hazardous solid waste. The volume of contaminated resin generated is much smaller than the original volume of the solution of biological stain, so the waste disposal problem is greatly reduced. The final concentration of any remaining stain should be less than the limit of detection (see Support Protocol 2 and Table 3.4.5). In each case decontamination should be >99%. This protocol describes a method for batch decontamination in which the resin is stirred in the solution to be decontaminated and removed by filtration at the end of the reaction time. Large volumes of biological stain can be decontaminated using a column (see Alternate Protocol 3). For full details refer to the original literature (Lunn and Sansone, 1991b) or a compilation (Lunn and Sansone, 1994a).
Table 3.4.5
Safe Use of Hazardous Chemicals
Decontamination of Biological Stains
Compound
Time required for complete decontamination
Volume of solution (ml) decontaminated per gram resin
Acridine orange Alcian blue 8GX Alizarin red S Azure A Azure B Brilliant blue R Congo red Coomassie brilliant blue G Cresyl violet acetate Crystal violet Eosin B Erythrosin B Ethidium bromide Janus green B Methylene blue Neutral red Nigrosin Orcein Propidium iodide Rose Bengal Safranine O Toluidine blue O Trypan blue
18 hr 10 min 18 hr 10 min 10 min 2 hr 2 hr 2 hr 2 hr 30 min 30 min 18 hr 4 hr 30 min 30 min 10 min 2 hr 2 hr 2 hr 3 hr 1 hr 30 min 2 hr
20 500 5 80 80 80 40 80 40 200 40 10 20 80 80 500 80 200 20 20 20 80 40
3.4.14 Supplement 20
Current Protocols in Cytometry
Materials Amberlite XAD-16 resin (Supelco) 100 µg/ml biological stain in water Additional reagents and equipment for testing for the presence of biological stain (see Support Protocol 2) For batch decontamination of 20 ml stain 1a. Add 1 g Amberlite XAD-16 to 20 ml of 100 µg/ml biological stain in water. For aqueous solutions having stain concentrations other than 100 ìg/ml, use proportionately greater or lesser amounts of resin to achieve complete decontamination. For solutions of erythrosin B, use 2 g Amberlite XAD-16 for 20 ml stain.
2a. Stir the mixture for at least the time indicated in Table 3.4.5. For batch decontamination of larger volumes of stain 1b. Add 1 g Amberlite XAD-16 to the volume of a 100 µg/ml biological stain in water indicated in Table 3.4.5. 2b. Stir the mixture for at least 18 hr. 3. Remove the resin by filtration through filter paper. 4. Test the filtrate for the presence of the biological stain (see Support Protocol 2). 5. Discard the resin as hazardous solid waste and the decontaminated filtrate as liquid waste. CONTINUOUS-FLOW DECONTAMINATION OF AQUEOUS SOLUTIONS OF BIOLOGICAL STAINS
ALTERNATE PROTOCOL 3
For treating large volumes of dilute aqueous solutions of biological stains (Table 3.4.5), it is possible to put the resin in a column and run the contaminated solution through the column in a continuous-flow system (Lunn et al., 1994). Limited grinding of the Amberlite XAD-16 resin increases its efficiency. Additional Materials (also see Basic Protocol 2) 25 µg/ml biological stain in water Methanol (optional) 300-mm × 11-mm-i.d. glass chromatography column fitted with threaded adapters and flow-regulating valves at top and bottom nut and insert connectors, and insertion tool (Ace Glass) or 300-mm × 15-mm-i.d. glass chromatography column (Spectrum 124010, Fisher) Glass wool 1.5-mm-i.d. × 0.3-mm-wall Teflon tubing Waring blender (optional) Rubber stopper fitted over a pencil QG 20 lab pump (Fluid Metering) Additional reagents and equipment for testing for the presence of biological stain (see Support Protocol 2)
Safety Procedures and Quality Control
3.4.15 Current Protocols in Cytometry
Supplement 20
Table 3.4.6 Breakthrough Volumes for Continuous-Flow Decontamination of Biological Stains
Compound Acridine orange Alizarin red S Azure A Azure B Cresyl violet acetate Crystal violet Ethidium bromide Janus green B Methylene blue Neutral red Safranine O Toluidine blue O
Breakthrough volume (ml) Limit of detection
1 ppm
5 ppm
465 120 615 630 706 1020 260 170 420 >2480 365 353
>990 150 810 882 >1396 >1630 312 650 645 >2480 438 494
>990 240 >975 >1209 >1396 >1630 416 >870 1050 >2480 584 606
Using a slurry of Amberlite XAD-16 1a. Prepare a 300-mm × 11-mm-i.d. glass chromatography column. To prevent clogging of the column outlet, place a small plug of glass wool at the bottom of the chromatography column. Connect 1.5-mm-i.d. × 0.3-mm wall Teflon tubing to the adapters using nut and insert connectors. Attach the tubing using an insertion tool. 2a. Mix 10 g Amberlite XAD-16 and 25 ml water in a beaker and stir 5 min to wet the resin. Using a finely ground Amberlite XAD-16 slurry 1b. Prepare a 300-mm × 15-mm-i.d. glass chromatography column. To prevent clogging of the column outlet, place a small plug of glass wool at the bottom of the chromatography column. 2b. Grind 20 g Amberlite XAD-16 with 200 ml water for exactly 10 sec in a Waring blender. 3. Pour the resin slurry into the column through a funnel. As the resin settles, tap the column with a rubber stopper fitted over a pencil to encourage even packing. Attach a QG 20 lab pump. 4. Pump the 25-µg/ml biological stain solution through the column at 2 ml/min. Alternatively, gravity flow coupled with periodic adjustment of the flow-regulating valve can be used.
5. Check the effluent from the column for the presence of biological stain (see Support Protocol 2). Stop the pump when stain is detected. Table 3.4.6 lists breakthrough volumes at different detection levels for a number of biological stains.
6. Discard the decontaminated effluent and the contaminated resin appropriately. 7. Many biological stains can be washed off the resin with methanol so the resin can be reused. Discard the methanol solution of stain as hazardous organic liquid waste. Safe Use of Hazardous Chemicals
3.4.16 Supplement 20
Current Protocols in Cytometry
Table 3.4.7
Methods for Detecting Biological Stainsa
Compound
Reagentb
Procedure Wavelength(s)(nm)
Limit of detection (ppm)
Acridine orange Alcian blue 8GX Alizarin red S Azure A Azure B Brilliant blue R Congo red Coomassie brilliant blue G Cresyl violet acetate Crystal violet Eosin B Erythrosin B Ethidium bromide Janus green B Methylene blue Neutral red Nigrosin Orcein Propidium iodide Rose Bengal Safranine O Toluidine blue O Trypan blue
DNA solution
F A A A A A A A
ex 492, em 528 615 556 633 648 585 497 610
0.0032 0.9 0.46 0.15 0.13 1.0 0.25 1.7
F A A F F A A A A A F F F A A
ex 588, em 618 588 514 ex 488, em 556 ex 540, em 590 660 661 540 570 579 ex 350, em 600 ex 520, em 576 ex 460, em 582 626 607
0.021 0.1 0.21 0.025 0.05 0.6 0.13 0.6 0.8 1.15 0.1 0.04 0.03 0.2 0.22
1 M KOH
pH 5 buffer
pH 5 buffer 1 M KOH DNA solution
aAbbreviations: A, absorbance; em, emission; ex, excitation; F, fluorescence bSee Support Protocol 2
ANALYTICAL PROCEDURES TO DETECT BIOLOGICAL STAIN Depending on the biological stain, the filtrate or eluate from the decontamination procedure can be analyzed using either UV absorption (A) or fluorescence detection (F).
SUPPORT PROTOCOL 2
Materials Filtrate or eluate from biological stain decontamination (see Basic Protocol 2 or Alternate Protocol 3) pH 5 buffer (see recipe) 1 M KOH solution 20 µg/ml calf thymus DNA in TBE electrophoresis buffer, pH 8.1 (APPENDIX 2A) Test the filtrate or eluate from the biological stain decontamination procedure using the appropriate method as indicated in Table 3.4.7. Traces of acid or base on the resin may induce color changes in the stain. Accordingly, with cresyl violet acetate or neutral red, mix aliquots of the filtrate with 1 vol pH 5 buffer before analyzing. With alizarin red S and orcein, mix aliquots of the filtrate with 1 vol of 1 M KOH solution before analyzing. Increase the fluorescence of solutions of acridine orange, ethidium bromide, and propidium iodide by mixing an aliquot of the filtrate with an equal volume of 20 ìg/ml calf thymus DNA in TBE electrophoresis buffer, pH 8.1. Let the solution stand 15 min before measuring the fluorescence.
Safety Procedures and Quality Control
3.4.17 Current Protocols in Cytometry
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BASIC PROTOCOL 3
DISPOSAL OF CHLOROTRIMETHYLSILANE AND DICHLORODIMETHYLSILANE Silane-containing compounds are hydrolyzed to hydrochloric acid and polymeric siliconcontaining material (Patnode and Wilcock, 1946). 1. Hydrolyze silane-containing compounds by cautiously adding 5 ml silane to 100 ml vigorously stirred water in a flask. Allow the resulting suspension to settle. 2. Remove any insoluble material by filtration and discard it with the solid or liquid hazardous waste. 3. Neutralize the aqueous layer with base and discard it.
BASIC PROTOCOL 4
DISPOSAL OF CYANIDES AND CYANOGEN BROMIDE Inorganic cyanides (e.g., NaCN) and cyanogen bromide (CNBr) are oxidized by sodium hypochlorite (NaOCl; e.g., Clorox) in basic solution to the much less toxic cyanate ion (Lunn and Sansone, 1985a). Destruction is >99.7%. Materials Cyanide (e.g., NaCN) or cyanogen bromide (CNBr) 1 M NaOH 5.25% (v/v) sodium hypochlorite (NaOCl; i.e., standard household bleach) Additional reagents and equipment for testing for the presence of cyanide (see Support Protocol 3) 1. Dissolve cyanide in water to give a concentration ≤25 mg/ml or dissolve CNBr in water to give a concentration ≤60 mg/ml. If necessary, dilute aqueous solutions so the concentration of NaCN or CNBr does not exceed the limit.
2. Mix 1 vol NaCN or CNBr solution with 1 vol 1 M NaOH and 2 vol fresh 5.25% NaOCl. Stir the mixture 3 hr. IMPORTANT NOTE: With age bleach may become ineffective; use of fresh bleach is strongly recommended.
3. Test the reaction mixture for the presence of cyanide (see Support Protocol 3). 4. Neutralize the reaction mixture and discard it. SUPPORT PROTOCOL 3
ANALYTICAL PROCEDURE TO DETECT CYANIDE This protocol is used to detect cyanide or cyanogen bromide at ≥3 µg/ml. Materials Cyanide or cyanogen bromide decontamination reaction mixture (see Basic Protocol 4) Phosphate buffer (see recipe) 10 mg/ml sodium ascorbate in water: prepare fresh daily 100 mg/liter NaCN in water: prepare fresh weekly 10 mg/ml chloramine-T in water: prepare fresh daily Cyanide detection reagent (see recipe) Sorvall GLC-1 centrifuge or equivalent
Safe Use of Hazardous Chemicals
1. If necessary to remove suspended solids, centrifuge two 1-ml aliquots of the cyanide or cyanogen bromide decontamination reaction mixture 5 min in a benchtop centrifuge, room temperature. Add each supernatant to 4 ml phosphate buffer in separate tubes.
3.4.18 Supplement 20
Current Protocols in Cytometry
2. If an orange or yellow color appears, add 10 mg/ml freshly prepared sodium ascorbate dropwise until the mixture is colorless, but do not add more than 2 ml. 3. Add 200 µl of 100 mg/liter NaCN to one reaction mixture (control solution). 4. Add 1 ml freshly prepared 10 mg/ml chloramine-T to each tube. Shake the tubes and let them stand 1 to 2 min. 5. Add 1 ml cyanide detection reagent, shake, and let stand 5 min. A blue color indicates the presence of cyanide. If destruction has been complete and the analytical procedure has been carried out correctly, the treated reaction mixture should be colorless and the control solution, which contains NaCN, should be blue.
6. Centrifuge tubes 5 min, room temperature, if necessary to remove suspended solids. Measure the absorbance at 605 nm with appropriate standards and blanks. DISPOSAL OF DIMETHYL SULFATE, DIETHYL SULFATE, METHYL METHANESULFONATE, ETHYL METHANESULFONATE, DIEPOXYBUTANE, AND 1,3-PROPANE SULTONE
BASIC PROTOCOL 5
Dimethyl sulfate is hydrolyzed by base to methanol and methyl hydrogen sulfate (Lunn and Sansone, 1985b). Subsequent hydrolysis of methyl hydrogen sulfate to methanol and sulfuric acid is slow. Methyl hydrogen sulfate is nonmutagenic and a very poor alkylating agent. The other compounds can be hydrolyzed with base in a similar fashion (Lunn and Sansone, 1990a). Destruction is >99%. A method to verify that decontamination is complete is also provided (see Support Protocol 4). NOTE: The reaction times given in the protocol should give good results; however, reaction time may be affected by such factors as the size and shape of the flask and the rate of stirring. The presence of two phases indicates that the reaction is not complete, and stirring should be continued until the reaction mixture is homogeneous. Materials Dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone 5 M NaOH Additional reagents and equipment for testing for the presence of dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone (see Support Protocol 4) For bulk quantities of dimethyl sulfate 1a. Add 100 ml dimethyl sulfate to 1 liter of 5 M NaOH. Stir the reaction mixture. 2a. Fifteen minutes after all the dimethyl sulfate has gone into solution, neutralize the reaction mixture with acid. For bulk quantities of diethyl sulfate 1b. Add 100 ml diethyl sulfate to 1 liter of 5 M NaOH. Stir the reaction mixture 24 hr. 2b. Neutralize the reaction mixture with acid.
Safety Procedures and Quality Control
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For bulk quantities of methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, and 1,3-propane sultone 1c. Add 1 ml methyl methanesulfonate, ethyl methanesulfonate, or diepoxybutane, or 1 g of 1,3-propane sultone to 10 ml of 5 M NaOH. Stir the reaction mixture 1 hr for 1,3-propane sultone, 2 hr for methyl methanesulfonate, 22 hr for diepoxybutane, or 24 hr for ethyl methanesulfonate. 2c. Neutralize the reaction mixture with acid. 3. Test the reaction mixture for the presence of the original compound (see Support Protocol 4). 4. Discard the decontaminated reaction mix. SUPPORT PROTOCOL 4
ANALYTICAL PROCEDURE TO DETECT THE PRESENCE OF DIMETHYL SULFATE, DIETHYL SULFATE, METHYL METHANESULFONATE, ETHYL METHANESULFONATE, DIEPOXYBUTANE, AND 1,3-PROPANE SULTONE This protocol is used to verify decontamination of solutions containing dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone. The detection limit for this assay is 40 µg/ml for dimethyl sulfate, 108 µg/ml for diethyl sulfate, 84 µg/ml for methyl methanesulfonate, 1.1 µg/ml for ethyl methanesulfonate, 360 µg/ml for diepoxybutane, and 264 µg/ml for 1,3-propane sultone. Materials Reaction mixture containing dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone 98:2 (v/v) 2-methoxyethanol/acetic acid 5% (w/v) 4-(4-nitrobenzyl)pyridine in 2-methoxyethanol Piperidine 2-Methoxyethanol 1. Dilute an aliquot of the reaction mixture with 4 vol water. 2. Add 100 µl diluted reaction mixture to 1 ml of 98:2 (v/v) 2-methoxyethanol/acetic acid. Swirl to mix. 3. Add 1 ml of 5% (w/v) 4-(4-nitrobenzyl)pyridine in 2-methoxyethanol. Heat 10 min at 100°C, then cool 5 min in ice. 4. Add 0.5 ml piperidine and 2 ml of 2-methoxyethanol. 5. Measure the absorbance of the violet reaction mixture at 560 nm against an appropriate blank. The absorbance of a decontaminated solution should be 0.000.
BASIC PROTOCOL 6
Safe Use of Hazardous Chemicals
DISPOSAL OF ETHIDIUM BROMIDE AND PROPIDIUM IODIDE Ethidium bromide and propidium iodide in water and buffer solutions may be degraded by reaction with sodium nitrite and hypophosphorous acid in aqueous solution (Lunn and Sansone, 1987); destruction is >99.87%. This reaction may also be used to decontaminate equipment contaminated with ethidium bromide (see Alternate Protocol 4; Lunn and Sansone, 1989) and to degrade ethidium bromide in organic solvents (see Alternate Protocol 5 and Alternate Protocol 6; Lunn and Sansone, 1990b). Ethidium bromide and propidium iodide may also be removed from solution by adsorption onto Amberlite XAD-16 resin (see Basic Protocol 2).
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Materials Ethidium bromide– or propidium iodide–containing solution in water, buffer, or 1 g/ml cesium chloride 5% (v/v) hypophosphorous acid: prepare fresh daily by diluting commercial 50% reagent 1/10 0.5 M sodium nitrite: prepare fresh daily Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide or propidium iodide (see Support Protocol 5) 1. If necessary, dilute the ethidium bromide– or propidium iodide–containing solution so the concentration of ethidium bromide or propidium iodide is ≤0.5 mg/ml. 2. For each 100 ml solution, add 20 ml of 5% hypophosphorous acid solution and 12 ml of 0.5 M sodium nitrite. Stir briefly and let stand 20 hr. 3. Neutralize the reaction mixture by adding sodium bicarbonate until the evolution of gas ceases. 4. Test the reaction mixture for the presence of ethidium bromide or propidium iodide (see Support Protocol 5). 5. Discard the decontaminated reaction mixture. DECONTAMINATION OF EQUIPMENT CONTAMINATED WITH ETHIDIUM BROMIDE
ALTERNATE PROTOCOL 4
Glass, stainless steel, Formica, floor tile, and the filters of transilluminators have been successfully decontaminated using this protocol. No change in the optical properties of the transilluminator filter could be detected, even after a number of decontamination cycles. Materials Equipment contaminated with ethidium bromide Decontamination solution (see recipe) Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide (see Support Protocol 5) 1. Wash the equipment contaminated with ethidium bromide once with a paper towel soaked in decontamination solution. The pH of the decontamination solution is 1.8. If this would be too corrosive for the surface to be decontaminated, wash with a paper towel soaked in water instead.
2. Wash the surface five times with paper towels soaked in water using a fresh towel each time. 3. Soak all the towels 1 hr in decontamination solution. 4. Neutralize the decontamination solution by adding sodium bicarbonate until the evolution of gas ceases. 5. Test the decontamination solution for the presence of ethidium bromide (see Support Protocol 5). 6. Discard the decontamination solution and the paper towels as nonhazardous liquid and solid wastes.
Safety Procedures and Quality Control
3.4.21 Current Protocols in Cytometry
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ALTERNATE PROTOCOL 5
DECONTAMINATION OF ETHIDIUM BROMIDE IN ISOPROPANOL SATURATED WITH CESIUM CHLORIDE Materials Ethidium bromide in isopropanol saturated with cesium chloride Decontamination solution (see recipe) Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide (see Support Protocol 5) 1. If necessary, dilute the ethidium bromide in isopropanol saturated with cesium chloride so the concentration of ethidium bromide is ≤1 mg/ml. 2. For each volume of ethidium bromide solution, add 4 vol decontamination solution. Stir the reaction mixture 20 hr. 3. Neutralize the reaction mixture by adding sodium bicarbonate until the evolution of gas ceases. 4. Test the reaction mixture for the presence of ethidium bromide (see Support Protocol 5). 5. Discard the decontaminated solution.
ALTERNATE PROTOCOL 6
DECONTAMINATION OF ETHIDIUM BROMIDE IN ISOAMYL ALCOHOL AND 1-BUTANOL Materials Ethidium bromide in isoamyl alcohol or 1-butanol Decontamination solution (see recipe) Activated charcoal Sodium bicarbonate Separatory funnel Additional reagents and equipment for testing for the presence of ethidium bromide 1. If necessary, dilute the ethidium bromide in isoamyl alcohol or 1-butanol so the concentration is ≤1 mg/ml final. 2. For each volume of ethidium bromide solution, add 4 vol decontamination solution. Stir the two-phase reaction mixture rapidly for 72 hr. 3. For each 100 ml of reaction mixture, add 2 g activated charcoal. Stir another 30 min. 4. Filter the reaction mixture. 5. Neutralize the filtrate by adding sodium bicarbonate until the evolution of gas ceases. Separate the layers using a separatory funnel. More alcohol may tend to separate from the aqueous layer on standing.
6. Test the alcohol and aqueous layers for the presence of ethidium bromide. 7. Discard the alcohol and aqueous layers appropriately. Discard the activated charcoal as solid waste. The aqueous layer contains 4.6% 1-butanol or 2.3% isoamyl alcohol.
Safe Use of Hazardous Chemicals
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Current Protocols in Cytometry
ANALYTICAL PROCEDURE TO DETECT ETHIDIUM BROMIDE OR PROPIDIUM IODIDE This protocol is used to verify that solutions no longer contain ethidium bromide or propidium iodide. The limits of detection are 0.05 parts per million (ppm) for ethidium bromide and 0.1 ppm for propidium iodide.
SUPPORT PROTOCOL 5
Materials Reaction mixture containing ethidium bromide or propidium iodide TBE buffer, pH 8.1 (APPENDIX 2A) 20 µg/ml calf thymus DNA in TBE buffer, pH 8.1 1. Mix 100 µl reaction mixture containing ethidium bromide or propidium iodide with 900 µl TBE buffer, pH 8.1. 2. Add 1 ml of 20 µg/ml calf thymus DNA in TBE, pH 8.1. Prepare a blank solution (100 µl water + 900 µl TBE + 1 ml of 20 µg/ml calf thymus DNA) and control solutions containing known quantities of ethidium bromide or propidium iodide. Let the mixtures stand 15 min. 3. To detect ethidium bromide, measure the fluorescence with an excitation wavelength of 540 nm and an emission wavelength of 590 nm. To detect propidium iodide, measure the fluorescence with an excitation wavelength of 350 nm and an emission wavelength of 600 nm. If a spectrophotofluorometer is not available, fluorescence of ethidium bromide can be qualitatively determined using a hand-held UV lamp on the long-wavelength setting (Lunn and Sansone, 1991c).
DISPOSAL OF HYDROGEN PEROXIDE Hydrogen peroxide can be reduced with sodium metabisulfite (Lunn and Sansone, 1994b).
BASIC PROTOCOL 7
Materials 30% hydrogen peroxide 10% (w/v) sodium metabisulfite 10% (w/v) potassium iodide 1 M HCl 1% (w/v) starch indicator solution 1. Add 5 ml of 30% hydrogen peroxide to 100 ml of 10% sodium metabisulfite. Stir the mixture at room temperature until the temperature starts to drop, indicating that the reaction is over. 2. Test for the presence of hydrogen peroxide by adding a few drops of the reaction mixture to an equal volume of 10% potassium iodide. Add a few drops of 1 M HCl to acidify the reaction mixture, then add a drop of 1% starch indicator solution. A deep blue color indicates the presence of excess oxidant. If necessary, add more 10% sodium metabisulfite until the starch test is negative.
3. Discard the decontaminated mixture.
Safety Procedures and Quality Control
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Supplement 20
BASIC PROTOCOL 8
REDUCTION OF IODINE Iodine is reduced with sodium metabisulfite to iodide (Lunn and Sansone, 1994b). Materials Iodine 10% (w/v) sodium metabisulfite 1 M HCl 1% (w/v) starch indicator solution 1. Add 5 g iodine to 100 ml of 10% sodium metabisulfite. Stir the mixture until the iodine has completely dissolved. 2. Acidify a few drops of the reaction mixture by adding a few drops of 1 M HCl. Add 1 drop of 1% starch indicator solution. A deep blue color indicates the presence of iodine. If reduction is not complete, add more sodium metabisulfite solution.
3. Dispose of the decontaminated solution. BASIC PROTOCOL 9
DISPOSAL OF MERCURY COMPOUNDS Solutions of mercuric acetate can be decontaminated using Dowex 50X8-100, a strongly acidic gel-type ion-exchange resin with a sulfonic acid functionality. Solutions of mercuric chloride can be decontaminated using Amberlite IRA-400(Cl), a strongly basic gel-type ion-exchange resin with a quaternary ammonium functionality. The final concentration of mercury is 98.3%). The exact reaction conditions depend on the solvent (see Table 3.4.8). The solutions that were decontaminated are representative of those described in the literature.
BASIC PROTOCOL 11
Materials Solutions of APMSF, AEBSF, PMSF, DFP, TLCK, or TPCK in buffer, DMSO, isopropanol, or water 1 M NaOH Glacial acetic acid Additional reagents and equipment for testing for the presence of the enzyme inhibitors (see Support Protocol 9) 1. If necessary, dilute the solutions with the same solvent so that the concentrations given in Table 3.4.8 are not exceeded. Bulk quantities of AEBSF, PMSF, and TPCK may be dissolved in isopropanol and bulk quantities of APMSF and TLCK may be dissolved in water at the concentrations shown in Table 3.4.8. Bulk quantities of DFP (a liquid) may be mixed directly with 1 M NaOH, taking care to make sure that all the DFP has mixed thoroughly, in the ratio shown in Table 3.4.8 (e.g., 40 ìl DFP with 1 ml of 1 M NaOH).
2. Add 1 M NaOH so that the ratio of solution to 1 M NaOH is that listed in Table 3.4.8. 3. Shake to ensure complete mixing, check that the solution is strongly basic (pH ≥12), and allow to stand for the time given in Table 3.4.8. 4. Neutralize the reaction mixture with acetic acid and test for the presence of residual enzyme inhibitor (see Support Protocol 9). 5. Discard the decontaminated reaction mixture.
Safety Procedures and Quality Control
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Supplement 20
Table 3.4.9 HPLC Conditions for Enzyme Inhibitors
Compound
Mobile phase
Detector
Retention time
Limit of detection
AEBSF
40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 50:50 (v/v) acetonitrile:water 40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 48:52 (v/v) acetonitrile:10 mM pH 7 phosphate buffer
UV 225 nm
9.5 min
0.1 µg/ml
UV 232 nm
7.7 min
0.5 µg/ml
UV 220 nm
8 min
0.9 µg/ml
UV 228 nm
9.5 min
0.37 µg/ml
UV 228 nm
10.5 min
2 µg/ml
APMSF
PMSF TLCK
TPCK
SUPPORT PROTOCOL 9
ANALYTICAL PROCEDURES TO DETECT ENZYME INHIBITORS DFP can be determined using a complex procedure involving the inhibition of chymotrypsin activity. For more information, refer to Lunn and Sansone (1994d). A gas chromatographic method has also been described by Degenhardt-Langelaan and Kientz (1996). AEBSF, APMSF, PMSF, TLCK, and TPCK may be determined by reversed-phase HPLC (Snyder et al., 1997). The chromatographic conditions and limits of detection are shown in Table 3.4.9 (Lunn and Sansone, 1994c). Materials Decontaminated enzyme inhibitor solutions Acetonitrile (HPLC grade) Water (HPLC grade) 0.1% (v/v) trifluoroacetic acid in water 10 mM phosphate buffer, pH 7 250-mm × 4.6 mm-i.d. Microsorb C-8 reversed-phase HPLC column (Varian) or equivalent Additional reagents and equipment for reversed-phase liquid chromatography (Snyder et al., 1997) Analyze the decontaminated enzyme inhibitor solutions by reversed-phase HPLC using a 250-mm × 4.6-mm-i.d. Microsorb C-8 reversed-phase column, or equivalent, using the conditions shown in Table 3.4.9. In each case, the injection volume was 20 µl, the separation occurred at ambient temperature, and the flow rate was 1 ml/min. Check the analytical procedures by spiking an aliquot of the acidified reaction mixture with a small quantity of a dilute solution of the compound of interest.
Safe Use of Hazardous Chemicals
3.4.30 Supplement 20
Current Protocols in Cytometry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Cyanide detection reagent Stir 3.0 g barbituric acid in 10 ml water. Add 15 ml of 4-methylpyridine and 3 ml concentrated HCl while continuing to stir. Cool and dilute to 50 ml with water. Store at room temperature. CAUTION: This reaction is exothermic.
Decontamination solution Dissolve 4.2 g sodium nitrite (0.2 M final) and 20 ml hypophosphorous acid (3.3% w/v final) in 300 ml water. Prepare fresh. Horseradish peroxidase Dissolve hydrogen-peroxide oxidoreductase (EC 1.11.1.7 [Type II]; specific activity 150 to 200 purpurogallin U/mg, Sigma) in 1 g/liter sodium acetate to give 30 U/ml. Prepare fresh daily. For small-scale reactions, a more dilute solution can be used to avoid working with inconveniently small volumes.
pH 5 buffer 2.04 g potassium hydrogen phthalate (0.05 M final) 38 ml 0.1 M potassium hydroxide (15 mM) H2O to 200 ml Store at room temperature Phosphate buffer 13.6 g monobasic potassium phosphate (KH2PO4; 0.1 M final) 0.28 g dibasic sodium phosphate (Na2HPO4; 2 mM final) 3.0 g potassium bromide (KBr; 25 mM final) 1 liter H2O Store at room temperature Potassium bromide is necessary to make the assay for cyanide work correctly.
Sodium azide indicator solution 0.1 g bromocresol purple (0.4% final) 18.5 ml 0.01 M potassium hydroxide (KOH; 7.4 mM final) H2O to 25 ml Store at room temperature Sulfanilic acid solution Dissolve 0.5 g sulfanilic acid in 150 ml of 15% (v/v) aqueous acetic acid. Use immediately. LITERATURE CITED Bretherick, L. (ed.) 1986. Hazards in the Chemical Laboratory, 4th ed. Royal Society of Chemistry, London. Bretherick, L., Urben, P.G., and Pitt, M.J. 1999. Bretherick’s Handbook of Reactive Chemical Hazards, 6th ed. Butterworth-Heinemann, London. Castegnaro, M., Barek, J., Dennis, J., Ellen, G., Klibanov, M., Lafontaine, M., Mitchum, R., van Roosmalen, P., Sansone, E.B., Sternson, L.A., and Vahl, M. (eds.) 1985. Laboratory Decontamination and Destruction of Carcinogens in Laboratory Wastes: Some Aromatic Amines and 4-Nitrobiphenyl. IARC Scientific Publications No. 64. International Agency for Research on Cancer, Lyon, France. Cunniff, P. (ed.) 1995. Official Methods of Analysis of the Association of Official Analytical Chemists, 16th ed., Ch. 4, p. 14. Association of Official Analytical Chemists, Arlington, Va.
Safety Procedures and Quality Control
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Degenhardt-Langelaan, C.E.A.M. and Kientz, C.E. 1996. Capillary gas chromatographic analysis of nerve agents using large volume injections. J. Chromatogr. A.723:210-214. Forsberg, K. and Keith, L.H. 1999. Chemical Protective Clothing Performance Index Book, 2nd ed. John Wiley & Sons, New York. Furr, A.K. (ed.) 2000. CRC Handbook of Laboratory Safety, 5th ed. CRC Press, Boca Raton, Fla. Lewis, R.J. Sr. 1999. Sax’s Dangerous Properties of Industrial Materials, 10th ed. John Wiley & Sons, New York. Lunn, G. and Sansone, E.B. 1985a. Destruction of cyanogen bromide and inorganic cyanides. Anal. Biochem. 147:245-250. Lunn, G. and Sansone, E.B. 1985b. Validation of techniques for the destruction of dimethyl sulfate. Am. Ind. Hyg. Assoc. J. 46:111-114. Lunn, G. and Sansone, E.B. 1987. Ethidium bromide: Destruction and decontamination of solutions. Anal. Biochem. 162:453-458. Lunn, G. and Sansone, E.B. 1989. Decontamination of ethidium bromide spills. Appl. Ind. Hyg. 4:234-237. Lunn, G. and Sansone, E.B. 1990a. Validated methods for degrading hazardous chemicals: Some alkylating agents and other compounds. J. Chem. Educ. 67:A249-A251. Lunn, G. and Sansone, E.B. 1990b. Degradation of ethidium bromide in alcohols. BioTechniques 8:372-373. Lunn, G. and Sansone, E.B. 1991a. The safe disposal of diaminobenzidine. Appl. Occup. Environ. Hyg. 6:49-53. Lunn, G. and Sansone, E.B. 1991b. Decontamination of aqueous solutions of biological stains. Biotech. Histochem. 66:307-315. Lunn, G. and Sansone, E.B. 1991c. Decontamination of ethidium bromide spills-author’s response. Appl. Occup. Environ. Hyg. 6:644-645. Lunn, G. and Sansone, E.B. 1994a. Destruction of Hazardous Chemicals in the Laboratory, 2nd ed. John Wiley & Sons, New York. Lunn, G. and Sansone, E.B. 1994b. Safe disposal of highly reactive chemicals. J. Chem. Educ. 71:972-976. Lunn, G. and Sansone, E.B. 1994c. Degradation and disposal of some enzyme inhibitors. Scientific note. Appl. Biochem. Biotechnol. 48:57-59. Lunn, G. and Sansone, E.B. 1994d. Safe disposal of diisopropyl fluorophosphate (DFP). Appl. Biochem. Biotechnol. 49:165-171. Lunn, G., Klausmeyer, P.K., and Sansone, E.B. 1994. Removal of biological stains from aqueous solution using a flow-through decontamination procedure. Biotech. Histochem. 69:45-54. Manufacturing Chemists Association. 1973. Laboratory Waste Disposal Manual. p. 136. Manufacturing Chemists Association, Washington, D.C. Mason, K.G. 1967. Hydrogen azide. In Mellor’s Comprehensive Treatise on Inorganic and Theoretical Chemistry, Vol. VIII (Suppl. II) pp. l-15. John Wiley & Sons, New York. National Research Council. 1983. Prudent Practices for Disposal of Chemicals from Laboratories, p. 88. National Academy Press, Washington, D.C. O’Neil, M.J. (ed.) 2001. The Merck Index, 13th ed. Merck & Co., Whitehouse Station, N.J. Patnode, W. and Wilcock, D.F. 1946. Methylpolysiloxanes. J. Am. Chem. Soc. 68:358-363. Shirakashi, T., Nakayama, K., Kakii, K., and Kuriyama, M. 1986. Removal of mercury from laboratory waste water with iron powder. Chem. Abstr. 105:213690y. Snyder, L.R., Kirkland, J.J., and Glajch, J.L. 1997. Practical HPLC Method Development, 2nd ed. John Wiley & Sons, New York.
KEY REFERENCES The following are good general references for laboratory safety. American Chemical Society, Committee on Chemical Safety. 1995. Safety in Academic Chemistry Laboratories, 6th ed. American Chemical Society, Washington, D.C. Castegnaro, M. and Sansone, E.B. 1986. Chemical Carcinogens. Springer-Verlag, New York. DiBerardinis, L.J., First, M.W., Gatwood, G.T., and Seth, A.K. 2001. Guidelines for Laboratory Design, Health and Safety Considerations, 3rd ed. John Wiley & Sons, New York. Safe Use of Hazardous Chemicals
Fleming, D.D., Richardson, J.H., Tulis, J.J., and Vesley, D. 1995. Laboratory Safety, Principles and Practices, 2nd ed. American Society for Microbiology, Washington, D.C. Freeman, N.T. and Whitehead, J. 1982. Introduction to Safety in the Chemical Laboratory. Academic Press, San Diego.
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Fuscaldo, A.A., Erlick, B.J., and Hindman, B. (eds.) 1980. Laboratory Safety, Theory and Practice. Academic Press, San Diego. Lees, R. and Smith, A.F. (eds.) 1984. Design, Construction, and Refurbishment of Laboratories. Ellis Horwood, Chichester, United Kingdom. Montesano, R., Bartsch, H., Boyland, E., Della Porta, G., Fishbein, L., Griesemer, R.A., Swan, A.B., and Tomatis, L. (eds.) 1979. Handling Chemical Carcinogens in the Laboratory, Problems of Safety. IARC Scientific Publications No. 33. International Agency for Research on Cancer, Lyon, France. National Research Council. 1995. Prudent Practices in the Laboratory: Handling and Disposal of Chemicals. National Academy Press, Washington, D.C. Occupational Health and Safety. 1993. National Safety Council, Chicago. Pal, S.B. (ed.) 1991. Handbook of Laboratory Health and Safety Measures, 2nd ed. Kluwer Academic Publishers, Hingham, Mass. Rosenlund, S.J. 1987. The Chemical Laboratory: Its Design and Operation: A Practical Guide for Planners of Industrial, Medical, or Educational Facilities. Noyes Publishers, Park Ridge, N.J. Young, J.A. (ed.) 1991. Improving Safety in the Chemical Laboratory: A Practical Guide, 2nd ed. John Wiley & Sons, New York.
INTERNET RESOURCES http://www.ilpi.com/msds/index.html Where to find MSDSs on the internet. Contains links to general sites, government and nonprofit sites, chemical manufacturers and suppliers, pesticides, and miscellaneous sites. http://www.OSHA.gov OSHA web site. http://www.osha-slc.gov/OshStd_data/1910_1450.html Text of OSHA Standard 29 CFR 1910.1450: Occupational Exposure to Hazardous Chemicals in Laboratories. http://www.osha-slc.gov/OshStd_data/1910_1000_TABLE_Z-1.html Tables of permissible exposure limits (PELs) for air contaminants. http://www.osha-slc.gov/OshStd_data/1910_1000_TABLE_Z-2.html Tables of PELs for toxic and hazardous substances. http://hazard.com/msds/index.php Main site for Vermont SIRI. One of the best general sites to start a search. Browse manufacturers alphabetically (for sheets not in the SIRI collection) or do a keyword search in the SIRI MSDS database. Lots of additional safety links and information. http://siri.uvm.edu/msds Alternate site for Vermont SIRI. http://tis.eh.doe.gov/docs/osh_tr/ch5.html DOE OSH technical reference chapter on personal protective equipment.
Contributed by George Lunn Baltimore, Maryland Gretchen Lawler Purdue University West Lafayette, Indiana
Safety Procedures and Quality Control
3.4.33 Current Protocols in Cytometry
Supplement 20
Method for Visualizing Aerosol Contamination in Flow Sorters Deflection-droplet cell sorters are an important part of scientific research because of their capability to separate different populations of cells based on size and granularity as well as on extrinsic fluorescent properties. In the operation of a cell sorter, a cell sample is hydrodynamically focused within a pressurized flowing stream of sheath fluid. Under normal circumstances, i.e., without external forces, the stream emerging from an orifice is hydrodynamically unstable, meaning that the droplet breakoff point is unpredictable and, therefore, not stable.
UNIT 3.5
BASIC PROTOCOL
Stabilization of the droplet breakoff in time and space is achieved by applying vibration to the stream with a steady frequency (by means of a piezoelectric crystal transducer), thereby fixing the droplet breakoff point. When the desired criteria for sorting are met, droplets are charged positively or negatively, and then passed through an electrostatic field. The highly positive or negative deflection plates attract negative or positive droplets, respectively, causing a separation in the stream. Aerosols are fine mists or sprays that can contain minute particles. During routine operation of a flow cytometer, microdroplets and droplets of varying sizes (3 to 7 µm, 40 to 200 µm, respectively) are formed when the pressurized sheath fluid leaves the flow-cell orifice, resulting in the generation of aerosols. In addition, splashing of side streams into receptacles, as well as center waste-stream splashing, can also form aerosols. Since the normal operation of cell sorters generates aerosols, the potential for aerosol creation is even greater in failure mode due to clogs or air in the fluidics. Usually, aerosols are contained within the collection chamber (Ferbas et al., 1995). However, this potential for aerosol generation should be a concern for sorter operators as well as for other individuals present in the facility who can potentially be exposed to contamination during sorting of biohazardous samples. Most commercially available sorters have some means of reducing the potential for exposure to biohazardous aerosols; however, none is designed to be 100% effective in eliminating aerosols. By using established precautionary guidelines and regularly testing the efficiency of an instrument’s capability to reduce aerosols, sorter operators can protect themselves as well as others around them. This unit provides a visual method to examine aerosol containment using a modified, commercially available product called Glo Germ. Glo Germ was developed for teaching aseptic techniques in hospitals, industry, restaurants, and schools and is visualized with UV or black light. Glo Germ is available in three forms: a white powder, an orange oil-based suspension of a melamine copolymer resin, or the orange melamine copolymer resin in dry form. The white powder is not used in this protocol because it fluoresces blue under black-light illumination, making it difficult to differentiate from ever-present lint. The orange oil-based suspension was successfully used prior to the availability of the dry form (Oberyszyn and Robertson, 2001). The newly available dry orange resin is described in this unit. The reported size of the Glo Germ particles (≤5 µm) is similar to that of yeast and bacteria, making it a comparable indicator. The Glo Germ suspension is run as a normal sample and sorted, thereby mimicking a biological sample sort. Placing slides in and around the sort collection area and examining the slides for orange fluorescent particles either by eye or under a fluorescent microscope allows for the detection of aerosols. Safety Procedures and Quality Control Contributed by Andrew S. Oberyszyn Current Protocols in Cytometry (2002) 3.5.1-3.5.7 Copyright © 2002 by John Wiley & Sons, Inc.
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The preparation of the suspension of resin is extremely straightforward, inexpensive, and quick. The visualization of orange fluorescent particles under a microscope is very easy due to their very intense fluorescence. Materials Glo Germ melamine copolymer resin (Glo Germ) 95% and 100% ethanol PBS without calcium or magnesium (PBS (−)) containing 10% filtered fetal bovine serum (FBS; APPENDIX 2A or Life Technologies) 100% acetone 15-ml conical tubes (Falcon) Deflected-droplet cell sorter to be tested Black light source (e.g., GE Bright Stik) Microscope slides Kimwipes (or any low-lint/lint-free paper) Fluorescent microscope (with DAPI or FITC exciter filter) 12 × 75–mm culture tubes (Falcon) Tubing (∼1.3-mm i.d.) Compressed air with thin extension tube Prepare Glo Germ sample suspension 1. Weigh out 0.5 g dry orange Glo Germ and place in a 15-ml conical tube containing 10 ml 95% or 100% ethanol. It is best to prepare the Glo Germ suspension as far away from the cell sorter as possible to prevent potential spreading of resin during preparation, preferably in another room. Without the ethanol suspension step, the Glo Germ will not suspend in the PBS (−)/10% FBS.
2. Vortex tube well to completely suspend resin. 3. Centrifuge 15-ml conical tube 5 min at 2000 × g, 25°C. 4. Remove supernatant and resuspend pellet in 10 ml PBS (−)/10% FBS. Vortex sample periodically to keep resin in suspension. The Glo Germ resin concentration is ∼5%. Higher resin concentration can cause clumping and a decline in sample flow rate during sorting. If clumping does occur, a final concentration of 1% can also be used successfully. The FBS is included to mimic a real sort sample.
Prepare cell sorter for sorting 5. Use standard manufacturer-suggested method for setting up the cell sorter (e.g., determining frequency, drive, and delay). Make sure the cell sorter is working optimally to be able to examine normal sort conditions.
6. With as many lights off as possible, examine the cell sorter area with a black light source. Wearing gloves, clean off any lint or fluorescent particles with 100% ethanol to make sure the area is “spotless.” Lint will glow light blue when excited by a black light source. It is important to eliminate as much nonspecific fluorescence as possible prior to starting the aerosolization evaluation. Methods for Visualizing Aerosol Contamination in Flow Sorters
7. Wearing gloves, meticulously clean several microscope slides by rinsing with 100% acetone. Air dry the slides and wipe clean with a Kimwipe. Rinse the slides with 100% ethanol and wipe dry with a Kimwipe.
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E A
B D C
Figure 3.5.1 Examples of positions of slides for examination of aerosol extent for a Beckman Coulter Elite ESP; (A,B,C,D) Slides located outside sort chamber on workstation; (E,F) Slides located inside sort chamber.
Acetone essentially “melts” any Glo Germ resin that may be on the slide. Ethanol cleans the slides further to make sure nothing fluorescent is present on the slides.
8. Examine slides using a fluorescent microscope to make sure that no fluorescence is present. It is imperative that the slides do not contain any fluorescence. Lint and dust will fluoresce and can give false positive results.
9. Label the microscope slides and place in and around the sort collection area in various locations (Fig. 3.5.1). Locations should include: directly inside the collection chamber, directly outside the collection chamber door, directly below sample chamber, near the computer keyboard, and so on. Make a note of slide location to be able to identify region of aerosol deposition.
10. Place collection tubes into cell sorter sample holders inside the collection chamber. Collection tubes should mimic a normal sorter collection tube (i.e., medium or PBS, with FBS).
Perform sorting in regular mode 11. Using forward scatter (FS) versus side scatter (SS), set sort regions. Since Glo Germ particles do not show distinct fluorescent peaks, FS versus SS is sufficient for gate setting. The regions selected should include large and small particle sizes.
12. Sort the Glo Germ resin for 1 to 2 hr at rates ranging from the lowest to the highest regularly used on the instrument. Keep collection chamber closed during sorting except for replacement of collection tubes. If the sorter is equipped with a biohazard filter, make sure it is on. This protocol was developed on a Beckman-Coulter Elite fitted with the high-speed ESP upgrade. Rates examined were 1000 to 10,000 events/sec. For regular-mode sorting, it is best to follow normal sorting procedure. This will allow one to determine if precautions regularly used are sufficient.
Safety Procedures and Quality Control
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13. Turn off as many lights as possible, visualize the area in and around the sort collection chamber with a black light source, and look for any orange fluorescence. If aerosolization is extensive, fluorescent regions or particles may be seen with the naked eye.
14. Carefully remove slides and proceed with microscopic visualization. Perform sorting in mock-failure mode 15. Repeat steps 6 to 10. If normal-mode sorting was performed prior to mock-failure mode, cleaning is imperative due to the fact that orange fluorescence will be seen near the waste drain as well as the location where the center stream falls inside the collection chamber.
16. Connect tubing to the thin extension tube from the compressed air and attach behind and below the deflection plate, making sure to point the end of the tubing toward the center stream (Fig. 3.5.2). 17. Sort the Glo Germ resin as described in step 12. Keep collection chamber open during sorting. If the sorter is equipped with a biohazard filter, make sure it is off. Since this is a failure-mode run, the principle is to try to achieve a “worst-case” scenario. To examine actual containment of aerosols, a “best-case” scenario should also be tested where all safety measures available are in place (i.e., biohazard filter on, collection chamber door closed, etc.) with the addition of the failure mode. What would be expected in a situation like this is that aerosols should be contained within the sorting collection chamber. This will determine if the employed safety measures are sufficient.
18. During sorting, spray a 10- to 30-sec burst of compressed air, deflecting the center stream to mimic a clogged flow cell.
center stream
compressed air
deflection plate
sort streams tubing
Methods for Visualizing Aerosol Contamination in Flow Sorters
“mock” clog
Figure 3.5.2 Tubing attached to compressed air was positioned below the deflection plates, behind the center waste stream. A 10-sec burst was used to simulate a failure mode.
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Alternatively, with the deflection voltage on, turn off the drive, which gives extensive fanning of the sorting streams.
19. Turn off as many lights as possible, visualize the area in and around the sort collection chamber with a black light source, and look for any orange fluorescence. 20. Carefully remove slides and proceed with microscopic visualization. Visualize aerosolization extent by fluorescent microscopy 21. Place slides on the stage of a fluorescent microscope. Examine areas of the slides looking for any orange fluorescence. A DAPI-exciter filter (405 nm) gives the brightest orange fluorescence; however a FITCexciter filter (490 nm) gives a very similar intensity. A Texas Red–exciter filter (570 nm) has also been used, although the orange fluorescence is noticeably less intense. In many instances, particles can be visualized with bright-field illumination; however, very small particles are more easily visualized by their intense orange fluorescence.
COMMENTARY Background Information A major advantage of sorting flow cytometers is the ability to separate cell populations based on cell size, cell density, or fluorescence. With increased analysis and separation of viable biological specimens comes an increase in the hazard of sorter operator exposure to unknown as well as known pathogens, such as hepatitis and HIV. Additionally, hazards can include commonly used staining reagents that can be toxic if inhaled. The normal working of the cell sorter forms droplets generated by pressurized fluid coming out of a vibrating nozzle. If desired sorting criteria are met, droplets containing the desired cells are electrostatically charged positively and/or negatively and are deflected into receptacles. Due to sample pressurization and electrostatic charges, aerosols of small droplets and microdroplets (size range: 40 to 200 µm and 3 to 7 µm, respectively) are formed, causing potential hazard to the operator. Secondary aerosols can also be formed when the sorting streams splash into receptacles (Merril, 1981; Bakker, 1992). Failure modes of the cell sorter such as a clogged tip, air in the fluidics system, or other instrumental malfunction can cause increased production of aerosols. The potential safety of the sorter operator, as well as of individuals in the sorter area, is further compromised due to the fact that aerosols have been shown to be of importance in the spread of infectious diseases (Almeida et al., 1971; Sattar and Ijaz, 1987; Schoenbaum et al., 1990; Ijaz et al., 1994). As a result, the International Society for Analytical Cytology (ISAC) Biohazard Working Group (Schmid et al., 1997) has developed guidelines as precautions for cell sorter operators. It is important to
remember that sorters used for analysis and not sorting of biohazardous samples still generate aerosols and should also be tested. The generally large size of cell sorters makes it impossible for them to be placed inside a biological hood. It is also not practical to create a “room-sized” biological hood to house them. Most current models of commercially available flow cytometers have designs that reduce the formation of aerosols and their subsequent release outside the sorting collection chamber (Ferbas et al., 1995). Custom-designed modifications to contain aerosols have been developed (Bakker, 1992; Giorgi, 1994). Methods for assessing the efficiency of aerosol containment during the process of sorting biohazardous samples have been developed as well (Anderson, 1958; Merril, 1981; UNIT 3.3). The manufacturer of the Glo Germ resin reports that the particle sizes are ≤5 µm. However, examining the mixture of Glo Germ resin and Flow-Check beads (10-µm nominal diameter; Beckman Coulter) in Figure 3.5.3 reveals that this is not completely correct. Regardless of this discrepancy, particles as small as ∼1 µm can be easily visualized under 20× magnification. Current methods for testing of aerosol containment during cell sorting include monitoring gravitational deposition of droplets (Merril, 1981; UNIT 3.3), and the use of air samplers (Anderson, 1958; UNIT 3.3). There are several advantages to using Glo Germ. Since there is no need for knowledge of microbiology, there is less potential for error compared to titration of T4 bacteriophage and agar plate handling (UNIT 3.3). Preparation takes 3 when 1 µg/100 µl is required, the antibody is unsuitable for use. For some cell types, there is a continuum from positive to negative cells without a clearcut distinction. Their epitopes are especially difficult to titer, resulting in a plateau effect or no peak in the signal-to-noise ratio. Identifying antibodies with the greatest possible S/N assists in distinguishing positive and negative cells. Antibodies that never show a plateau in MCF or a peak S/N should not be used, and an alternate supplier found.
Time Considerations The titering procedures for epitopes or antibodies should take 0.01% final concentration) also interferes with accurate readings (Cesarone et al., 1979). The pH of the assay solution is critical to sensitivity and should be ∼7.4 (Stout and Becker, 1982; Labarca and Paigen, 1980). At a pH 8.0 the background becomes much higher and there is a concomitant loss of fluorescence enhancement. High-quality double-stranded DNA is recommended. With very small fragments of DNA, the Hoechst 33258 dye binds to doublestranded DNA only. Thus, the assay will not work with single-stranded oligomers. Linear and circular DNA give approximately the same levels of fluorescence (Daxhelet et al., 1989). When preparing DNA standards, an attempt should be made to equalize the GC content of the standard DNA and that of the sample DNA. In most situations, salmon sperm or calf thymus DNA is suitable. An extensive list of estimated GC content for various organisms is available (Marmur and Doty, 1962). Eukaryotic cells vary somewhat in GC content but are generally in the range of 39% to 46%. Within this range, the fluorescence per microgram of DNA does not vary substantially. In contrast, the GC content of prokaryotes can vary from 26% to 77%, causing considerable variation in the fluorescence signal. In these situations, the sample DNA should be first quantitated via transmission spectroscopy and compared to a readily
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Relative flourescence units
1000 900 800 700 600 500 400 300 200 100 0
A
H33258 Ex: 365 nm r 2: = 0.9999 Ex: 455 nm
60 50 40 30 20 10 0
H33258
0
10
20
ethidium bromide Ex: 546 nm r 2: = 0.9993 Ex: 590 nm
50
30
B
40 30 20 10 0
0
100
200
300
400
DNA (ng/ml)
500
Figure 4.5.1 Fluorochrome Hoechst 33258 (H33258) (A) and ethidium bromide (B) DNA concentration standard curves. Assays were performed as described in alternate protocols, at indicated excitation and emission wavelengths. The concentrations of H33258 and ethidium bromide were 0.1 and 5 µg/ml, respectively. Assays contained the indicated concentrations of calf thymus DNA standards suspended in a final volume of 2.0 ml. Inset shows low DNA concentration curve for the H33258 assay. Note that, under these conditions, H33258 produces ∼20 times more relative fluorescence units than ethidium bromide. A Shimadzu RF-5000 scanning fluorescence spectrometer was used for both assays.
available standard (e.g., calf thymus DNA). Future measurements would then use calf thymus as a standard, but with a correction factor for difference in fluorescence yield between the two DNA types. For further troubleshooting information, see Van Lancker and Gheyssens (1986), in which the effects of interfering substances on the Hoechst 33258 assay (and several other assays) are compared. In the ethidium bromide assay, singlestranded DNA gives approximately half the signal as double-stranded calf thymus DNA. Ribosomal RNA also gives about half the fluorescent signal as double-stranded DNA, and RNase and DNase both severely decrease the signal. Closed circular DNA also binds less ethidium bromide than nicked or linear DNA. Further critical parameters of the ethidium bromide assay are described by Le Pecq (1971).
(r2) of 0.98 to 0.99 (Fig. 4.5.1). Table 4.5.3 provides a comparison of the sensitivities and specificities of the three assays.
Anticipated Results
Literature Cited
The detection limit of absorption spectroscopy will depend on the sensitivity of the spectrophotometer and any UV-absorbing contaminants that might be present. The lower limit is generally ∼0.5 to 1 µg nucleic acid. Typical values for a highly purified sample of DNA are shown in Table 4.5.2. For the Hoechst 33258 and ethidium bromide assays, a plot of relative fluorescence units or estimated concentration (y axis) versus actual concentration (x axis) typically produces a linear regression with a correlation coefficient
Applied Biosystems. 1989. User Bulletin Issue 11, Model No. 370. Applied Biosystems, Foster City, Calif.
Time Considerations The three assays described can be performed in a short period of time. In a well-planned series of assays, 50 samples can be prepared and read comfortably in 1 hr. Although some error might be introduced, DNA samples can be sequentially added to the same cuvette containing working dye solution. The increase in fluorescence with each sample is noted and subtracted from the previous reading to give relative fluorescence or concentration of the new sample, eliminating the need to change solutions for each sample. Be certain that the final amount of DNA does not exceed the linear portion of the assay.
Cesarone, C.F., Bolognesi, C., and Santi, L. 1979. Improved microfluorometric DNA determination in biological material using 33258 Hoechst. Anal. Biochem. 100:188-197. Daxhelet, G.A., Coene, M.M., Hoet, P.P., and Cocito, C.G. 1989. Spectrofluorometry of dyes with DNAs of different base composition and conformation. Anal. Biochem. 179:401-403. Molecular and Cellular Probes
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Labarca, C. and Paigen, K. 1980. A simple, rapid, and sensitive DNA assay procedure. Anal. Biochem. 102:344-352. Le Pecq, J.-B. 1971. Use of ethidium bromide for separation and determination of nucleic acids of various conformational forms and measurement of their associated enzymes. In Methods of Biochemical Analysis, Vol. 20 (D. Glick, ed.) pp. 41-86. John Wiley & Sons, New York. Marmur, J. and Doty, P. 1962. Determination of the base composition of deoxyribonucleic acid from its thermal denaturation temperature. J. Mol. Biol. 5:109-118. Portugal, J. and Waring, M.J. 1988. Assignment of DNA binding sites for 4′,6-diamidine-2- phenylindole and bisbenzimide (Hoechst 33258): A comparative footprinting study. Biochem. Biophys. Acta 949:158-168. Stout, D.L. and Becker, F.F. 1982. Fluorometric quantitation of single-stranded DNA: A method applicable to the technique of alkaline elution. Anal. Biochem. 127:302-307.
Van Lancker, M. and Gheyssens, L.C. 1986. A comparison of four frequently used assays for quantitative determination of DNA. Anal. Lett. 19:615-623. Wallace, R.B. and Miyada C.G. 1987. Oligonucleotide probes for the screening of recombinant DNA libraries. In Methods of Enzymology, Vol. 152: Guide to Molecular Cloning Techniques (S.L. Berger and A.R. Kimmel, eds.) pp. 432442. Academic Press, San Diego.
Key References Labarca and Paigen, 1980. See above. Contains a detailed description of the Hoechst 33258 fluorometric DNA assay.
Contributed by Sean R. Gallagher Motorola Corporation Tempe, Arizona
Spectroscopic Analysis Using DNA and RNA Fluorescent Probes
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CHAPTER 5 Specimen Handling, Storage, and Preparation INTRODUCTION
T
his chapter describes well-established, quality-controlled methods and procedures for the handling, storage, and preparation of subcellular components (such as chromosomes), and tissue for flow cytometric assay. Numerous assays can be performed with samples prepared following these fundamental techniques.
Common to all flow cytometric procedures is the requirement for a single-cell, singlenucleus, or single-chromosome preparation. Although human blood is obtained conveniently in cell suspension, specific procedures must still be followed, as discussed in UNIT 5.1, to ensure the quality of the resulting single-cell preparation, which can be affected by a variety of parameters. Conditions for obtaining, transporting, and storing material prior to assay should prevent or minimize artefactual debris, degradation of protein, RNA, or DNA, and loss of functional activity. In many cases it is also desirable to obtain specific blood cell fractions desired for analysis in enriched or purified form; procedures for obtaining such fractions are also detailed in UNIT 5.1. The same critical parameters that apply to blood samples are equally important in the preparation of other human tissues. Unlike blood cells, such tissues are not initially obtained initially as single-cell suspensions but must be prepared as such. The cell parameters or components to be measured (e.g., protein, DNA, or RNA) and the localization of the markers (e.g., cell surface, cytoplasmic, or nuclear) will influence the choice of dissociation technique as well as the optimal methods for collection, transport, and storage of specimens. Several methods to accommodate flow cytometric analysis of both whole cells and nuclei are described in UNIT 5.2. Flow cytometric analysis and sorting of either animal or plant chromosomes is an invaluable method of obtaining DNA to be used for physical gene mapping, isolation of molecular markers, or construction of chromosome-specific DNA libraries, among other applications. Varying sample handling, storage, and preparation conditions are needed depending on the cell type used as the chromosome source. UNIT 5.3 details procedures for accumulation of plant cells in metaphase, for preparation of plant chromosome suspensions, and for flow cytometric analysis and sorting; methods for further processing of sorted plant chromosomes are also provided. Specific applications, such as those described in other chapters of this manual, may require a unique method that works optimally with that application and may not be included in this chapter. These applications may require a different fixative, medium, or time frame for preparation. Therefore, users are advised to review their specific flow cytometry applications prior to the preparation of cells. Alberto Orfao Specimen Handling, Storage, and Preparation Contributed by Alberto Orfao Current Protocols in Cytometry (1999) 5.0.1 Copyright © 1999 by John Wiley & Sons, Inc.
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UNIT 5.1
Human peripheral blood samples are probably the most common specimens submitted to the flow cytometry laboratory for analysis. How specimens are obtained, handled, and stored is intimately related to the reason for obtaining them. Although the majority of specimens are for lymphocyte immunophenotyping, peripheral blood is also commonly obtained for the analysis of platelets, neutrophil function, lymphocyte mitogenesis or blastogenesis, and intracellular antigen expression, and for reticulocyte enumeration. Many of these assays impose specific requirements for specimen acquisition, transport, and storage. Moreover, although flow cytometry can be used to analyze heterogeneous cell populations, it is often desirable to enrich, purify, or separate cell populations prior to performing flow cytometry. Preparation of cells for flow cytometry will vary depending on which cell lineage is to be examined and what assay is to be performed on the cells. Therefore, it is not possible to present a single, generic protocol for acquisition and transportation of all specimens. However, as a general principle, cells should be examined as soon as possible after the specimen is obtained and in as close to the native state as possible (i.e., with the least amount of processing possible), and certain general procedures for obtaining blood samples apply. 1. Before taking the sample, read thoroughly the procedure for the specific assay to be performed. Contact the venipuncturist with any information concerning the volume of blood required for the assay and the anticoagulant to be used, as well as any other pertinent information. 2. Procure blood samples from the venipuncture team as quickly as possible. Maintain blood sample at room temperature unless the specific protocol dictates otherwise. In general, avoid subjecting specimens to extremes of temperature or holding them for prolonged periods of time prior to processing. 3. Add tissue culture medium, anticoagulant, preservatives, or other additives as directed by the specific protocol. 4. Be sure to label all specimens properly with type of specimen, time and date of collection, patient name (if appropriate), and test to be performed. Also note the type of anticoagulant used if not evident from the color of the stopper or another indicator. 5. Store the specimen as appropriate for the assay to be performed. If no specific additives are indicated, maintain blood sample aseptically at room temperature until needed; this temperature is usually acceptable for short-term storage, although this may not be universally true. Gentle rocking may help in preserving cellular aggregation. (Table 5.1.1 provides general recommendations for anticoagulants and storage times for blood samples obtained for a variety of common assays.) This unit presents protocols for separating white cells by lysing erythrocytes (see Basic Protocol 1), isolating mononuclear cells using trilineage separation (see Basic Protocol 2), enrichment of lymphocytes by adherence (see Support Protocol), and assorted methods of isolating or enriching specific cell populations, including peripheral blood monocytes (see Basic Protocol 3), neutrophils (see Basic Protocol 4), T lymphocytes (see Basic Protocols 5 and 7), B lymphocytes (see Basic Protocols 6, 7, and 8), NK cells (see Basic Protocol 8), stem cells (see Basic Protocol 9), and platelets (see Basic Protocol 10). Specimen Handling, Storage, and Preparation Contributed by J. Philip McCoy, Jr. Current Protocols in Cytometry (1998) 5.1.1–5.1.13 Copyright © 1998 by John Wiley & Sons, Inc.
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Table 5.1.1 Recommended Anticoagulants and Storage Times for Commonly Performed Assays
Assay
Anticoagulant
Time limitation
Lymphocyte immunophenotyping Myeloid immunophenotyping Neutrophil function Platelet activation Platelet markers Reticulocyte enumeration DNA analysis
Sodium heparin or EDTA
Store ≤72 hr
EDTA
Use immediately
Sodium heparin or EDTA EDTA EDTA EDTA Sodium heparin or EDTA
Use immediately Use immediately Use immediately Store ≤72 hr at 4°C Use immediate for cell-cycle analysis; store ≤72 hr for ploidy analysis
CAUTION: When working with human blood, cells, or infectious agents, biosafety practices should be followed; see Critical Parameters for further information. NOTE: All solutions and equipment coming in contact with cells must be sterile, and proper sterile techniques should be used accordingly. BASIC PROTOCOL 1
PREPARATION OF WHITE CELL SUSPENSION BY LYSIS OF ERYTHROCYTES Although erythrocytes can be separated from mononuclear cells by density-gradient separation (see Basic Protocol 2), many laboratories prefer lysis methods to eliminate erythrocytes from various specimens. Lysis is much quicker than gradient separation and in general leaves the remaining white cell populations relatively unperturbed. Blood samples may be treated with any anticoagulant. This procedure, in which erythrocytes are lysed with ammonium chloride, may be used for unstained blood or blood that has already been incubated with monoclonal antibodies. In general, this method will not affect the pattern of staining observed for most lymphoid markers. The viability of white blood cells subjected to this treatment is good. Commercial reagents such as FACSLyse, ImmunoLyse, and Optilyse can be used in place of ammonium chloride to lyse erythrocytes. However, as FACSLyse contains a fixative, staining for cell surface markers on leukocytes should be performed prior to lysis of the erythrocytes with this reagent. Stain whole blood with antibodies, then lyse erythrocytes according to manufacturer’s instructions. Optilyse may be used without subsequent washing of the leukocytes. Materials Whole blood sample 1× ammonium chloride lysing solution (prepare fresh from 10× stock; see APPENDIX 2A) or FACSLyse (Becton Dickinson Immunocytometry), ImmunoLyse (Coulter), or Optilyse (Immunotech) Phosphate-buffered saline (PBS; APPENDIX 2A) 1. Place 200 µl whole blood sample in a centrifuge tube and add 3 ml fresh 1× ammonium chloride lysing solution. Incubate 10 min at room temperature.
Handling, Storage, and Preparation of Human Blood Cells
2. Centrifuge 5 min at 300 × g, room temperature (22° to 25°C). 3. Discard supernatant. Resuspend cells in 2 ml PBS. Centrifuge 5 min at 300 × g, room temperature.
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4. Discard supernatant and repeat wash with 2 ml PBS. 5. Resuspend cells as necessary for specified assay. Use immediately or store as indicated for assay. ISOLATION OF MONONUCLEAR CELLS BY DENSITY GRADIENT SEPARATION
BASIC PROTOCOL 2
Trilineage separation methods (Boyum, 1968, 1977) are used when purification of cell populations is required rather than simple removal of erythroid contaminants. Densitygradient separation techniques may not yield as many cells as the simple lysis methods, but they have other distinct advantages. Separating cells by Ficoll-Hypaque centrifugation often decreases the cytometry time for acquisition and removal of nonviable cells. Materials 1.077 g/ml Ficoll-Hypaque (Pharmacia Biotech) or Histopaque-1077 (Sigma) Anticoagulated blood in heparin or EDTA Phosphate-buffered saline (PBS; APPENDIX 2A) Tissue culture medium (optional; APPENDIX 3B) 15- and 50-ml conical centrifuge tubes Centrifuge 1. With a sterile pipet, place the Ficoll-Hypaque solution into a 50-ml conical centrifuge tube, using 2 ml Ficoll-Hypaque per ml blood. Volume of Ficoll-Hypaque may vary with brand used. Consult manufacturer’s recommendations.
2. Mix anticoagulated blood with an equal volume of PBS. 3. Slowly layer the diluted blood over the Ficoll-Hypaque solution by gently pipetting the diluted blood down the side of the tube containing the Ficoll-Hypaque. 4. Centrifuge 40 min at 400 × g, 22°C, with no brake. 5. Using a sterile Pasteur pipet, carefully remove the mononuclear cells, located at the interface between the plasma (upper layer) and the Ficoll-Hypaque (bottom). 6. Transfer the aspirated mononuclear cells to a 15-ml conical tube. Add 10 ml PBS or tissue culture medium and mix thoroughly. Centrifuge 10 min at 400 × g, 4°C. 7. Discard the supernatant and repeat wash with PBS or tissue culture medium as needed. ENRICHMENT OF LYMPHOCYTES FROM MONONUCLEAR CELL PREPARATIONS BY DEPLETION OF MONOCYTOID CELLS THROUGH ADHERENCE TO PLASTIC
SUPPORT PROTOCOL
The density-gradient procedure described in Basic Protocol 2 yields a preparation of mononuclear cells that may include monocytoid cells. For most individuals, this protocol provides a specimen with more lymphoid than monocytoid cells. However, in some instances, even further purity of the lymphocytes is desired. This protocol enriches the lymphocyte population by depleting monocytoid cells from the mononuclear cell preparation using adherence to a plastic tissue culture flask. Additional Materials (also see Basic Protocol 2) Mononuclear cell preparation (see Basic Protocol 2) Tissue culture medium (APPENDIX 3B) containing ≥10% FBS (APPENDIX 2A)
Specimen Handling, Storage, and Preparation
5.1.3 Current Protocols in Cytometry
75-cm2 or 150-cm2 plastic tissue culture flask Humidified 37°C, 5% CO2 incubator Additional reagents and equipment for assessing cell viability (UNIT 9.2) 1. Suspend mononuclear cells in tissue culture medium to a final concentration of 1–2 × 106 cells/ml. 2. Transfer cell suspension to tissue culture flask. An appropriate volume for a 150-cm2 flask is 50 ml.
3. Incubate cells 1 hr in a humidified 37°C, 5% CO2 incubator. 4. Gently pour supernatant containing nonadherent lymphoid cells into a 50-ml centrifuge tube. Rinse flask gently with 10 ml tissue culture medium and add to centrifuge tube. 5. Check cell suspension manually or with a hematology analyzer to determine percentage of monocytoid cells remaining in the specimen. If additional depletion of monocytoid cells is desired, repeat steps 1 to 4. 6. Assess viability of lymphocytes using trypan blue exclusion (APPENDIX 3B) or flow cytometry (UNIT 9.2). BASIC PROTOCOL 3
ISOLATION OF PERIPHERAL BLOOD MONOCYTES BY GRADIENT CENTRIFUGATION WITH FICOLL-HYPAQUE FOLLOWED BY PERCOLL Peripheral blood monocytes are isolated from blood by centrifugation through a FicollHypaque or Histopaque gradient (Denholm and Wolber, 1991) followed by Percoll. Materials Venous blood in sodium citrate Phosphate-buffered saline (APPENDIX 2A) 1.077 g/ml Ficoll-Hypaque (Pharmacia Biotech) or Histopaque-1077 (Sigma) 10× Hanks balanced salt solution (HBSS; APPENDIX 2A) Percoll, specific gravity 1.130 g/ml (Sigma) Tissue culture medium (optional; APPENDIX 3B) 50-ml conical centrifuge tubes 10 × 15–cm round bottom polypropylene tubes silanized with Surfasil 1. Mix 30 to 40 ml of blood with an equal volume of PBS. Layer diluted blood over Ficoll-Hypaque solution, using 3 parts diluted blood to 2 parts Ficoll-Hypaque. Centrifuge 30 min at 500 × g, 25°C. 2. Using a sterile Pasteur pipet, carefully collect mononuclear cells, located at the interface between the plasma (upper layer) and Ficoll-Hypaque solution (bottom of tube). 3. Transfer cells to a 50-ml centrifuge tube. Add 10 ml PBS. Centrifuge 10 min at 400 × g, 4°C. 4. Discard supernatant and resuspend cells in 4 ml PBS.
Handling, Storage, and Preparation of Human Blood Cells
5. Mix 1.65 ml of 10× Hanks balanced salt solution with 10 ml Percoll. Adjust pH to 7.0 with 0.1 N HCl (~30 µl should be sufficient). 6. Mix 8 ml Percoll solution with 4 ml mononuclear cells in a silanized 10 × 1.5–cm round-bottom polypropylene tube. Mix thoroughly by inverting tube three or four times. Centrifuge 25 min at 370 × g, 25°C.
5.1.4 Current Protocols in Cytometry
7. Aspirate monocytes by gentle pipetting into a clean test tube. Add PBS or tissue culture medium as needed. After centrifugation, monocytes appear as a cloudy layer in the top 5 mm of the gradient.
8. Wash the cells in PBS or tissue culture medium. ISOLATION OF NEUTROPHILS BY PERCOLL GRADIENT CENTRIFUGATION
BASIC PROTOCOL 4
In general, neutrophils require more prompt and delicate handling than lymphoid cells. Specimens more than a few hours old are not optimal for use in neutrophil isolation procedures. Materials Blood sample in acid citrate dextrose, formula A 0.6% (w/v) dextran in 0.9% (w/v) NaCl 0.9% NaCl 1.10 g/ml, 1.095 g/ml, and 1.085 g/ml Percoll in phosphate-buffered saline (PBS; APPENDIX 2A) 15-ml polypropylene tubes 1. Mix 45 ml of blood with 10 ml of 6% dextran in 0.9% NaCl. Allow to stand for 1 hr at 25°C. 2. Remove leukocyte-rich supernatant and centrifuge 10 min at 400 × g, 4°C. 3. Discard supernatant and resuspend cells in 0.9% NaCl to a final concentration of 5 × 107 cells/ml. 4. Prepare a discontinuous Percoll gradient: transfer 4 ml of 1.10 g/ml Percoll in PBS to a 15-ml polypropylene tube; then overlay this with 3 ml of 1.095 g/ml Percoll and finally with 3 ml of 1.085 g/ml Percoll. 5. Layer 5 ml of cell suspension over the Percoll gradient. Centrifuge 30 min at 700 × g, 25°C, in a swinging-bucket rotor with no braking. 6. Sample will contain three bands and a cell pellet. Using sterile polypropylene pipets, collect the two lower bands, which contain the enriched neutrophil fraction. Wash in 0.9% NaCl. ISOLATION OF T LYMPHOCYTES BY ERYTHROCYTE ROSETTING Erythrocyte rosetting (Madsen et al., 1979; Indiveri et al., 1980) is one of several methods used to isolate T lymphocytes. Other methods include panning (see Basic Protocol 7) and separation using magnetic beads coated with appropriate antibodies (see Basic Protocol 6). E-rosetting, a classical method for isolating T cells, involves separating T cells from other mononuclear cells by exploiting the unique ability of the lymphocytes to bind to and form rosettes with sheep red blood cells. The method remains very useful and inexpensive today. Materials Sheep red blood cells in Alsevers solution Phosphate-buffered saline, isotonic, pH 7.0 (PBS; APPENDIX 2A) 2-aminoethylisothiouronium hydrobromide (AET) solution: 0.143 M AET in H2O, pH 9.0 (adjust pH with 4 N NaOH), prepared fresh for each use Tissue culture medium (APPENDIX 3B)
BASIC PROTOCOL 5
Specimen Handling, Storage, and Preparation
5.1.5 Current Protocols in Cytometry
FBS (heat inactivated; APPENDIX 2A) 1–2 × 107 cell/ml mononuclear cell preparation (see Basic Protocol 2) prepared fresh in tissue culture medium 1.077 g/ml Ficoll-Hypaque (Pharmacia Biotech) or Histopaque-1077 (Sigma) 10× ammonium chloride lysing solution (APPENDIX 2A; prepare fresh) 15-ml polystyrene round-bottom centrifuge tubes 15- and 50-ml conical centrifuge tubes 1. Place 6 ml sheep red blood cells in a 50-ml conical centrifuge tube. Add 30 ml isotonic PBS at room temperature. Centrifuge 10 min at 400 × g, 4°C. Discard supernatant and repeat wash two more times. 2. Transfer 2 ml of washed sheep erythrocytes to a 50-ml conical centrifuge tube. Add 10 ml AET solution. Vortex the mixture gently and incubate 15 min at 37°C, mixing gently every 5 min. 3. Wash the AET-treated cells three times with 30 ml of 4°C PBS as in step 1. Resuspend cells in 50 ml tissue culture medium. 4. Place 1 ml mononuclear cells in a 15-ml round bottom centrifuge tube. Add 2 ml AET-treated sheep erythrocytes and 2 ml heat-inactivated FBS. Mix well and incubate 10 min at 37°C. 5. Centrifuge 10 min at 200 × g, 4°C. Remove the supernatant and incubate pellet 60 min at 4°C. 6. Gently resuspend pelleted mixture with a pipet and slowly layer over an equal volume of Ficoll-Hypaque solution in a 15-ml conical centrifuge tube. Centrifuge 40 min at 400 × g, room temperature. T cells will pellet at the bottom of the centrifuge tube.
7. Remove T cells with a pipet. To lyse erythrocytes, add 15 ml ammonium chloride lysing solution to pellet and let stand 10 min (until solution becomes transparent), then transfer to 50-ml conical tube containing tissue culture medium. Centrifuge 10 min at 400 × g, 4°C. 8. Discard supernatant and resuspend T cells in tissue culture medium at a concentration appropriate for their desired use. BASIC PROTOCOL 6
Handling, Storage, and Preparation of Human Blood Cells
ISOLATION OF B LYMPHOCYTES BY MAGNETIC BEAD–ANTIBODY SEPARATION Lymphoid subsets can be isolated by either positive or negative selection. Positive selection is more direct, but in some instances the cells of interest may suffer from being manipulated. In those instances, negative selection can be applied by using antibodies that will bind all lymphoid subsets other than the one of interest. The positive selection procedure that follows (based on procedures developed by Rasmussen et al., 1992, and Molday et al., 1977) employs magnetic beads coated with anti-CD19 antibodies to bind B cells. Materials 1% FBS/PBS: phosphate-buffered saline (PBS; APPENDIX 2A) with 1% (v/v) FBS (heat inactivated; APPENDIX 2A), 4°C Heparinized blood Tissue culture medium with 10% FBS (APPENDIX 2A) Detach-a-bead (Dynal) or other anti-mouse Fab antibody (optional)
5.1.6 Current Protocols in Cytometry
Anti-CD19-coated magnetic beads (Dynal) Magnetic separation device (e.g., Dynal) 75-cm2 tissue culture flask 15-ml polypropylene test tube Humidified 37°C, 5% CO2 incubator Bind B cells with anti-CD19-coated magnetic beads 1. Thoroughly mix 100 µl of resuspended beads with 10 ml of 4°C 1% FBS/PBS in a 15-ml polypropylene test tube. Place tube in magnetic separation device for 5 to 10 min to draw beads to the side of the tube. Gently remove the PBS by pipetting or aspiration. 2. Repeat suspension and separation as in step 1. Resuspend cells in 2 ml of 4°C 1% FBS/PBS. 3. Mix 10 ml heparinized blood with 30 ml of 4°C 1% FBS/PBS. Place diluted blood in tissue culture flask or polypropylene centrifuge tube. Add washed magnetic beads to the flask and incubate 30 min at 4°C with gentle rocking. 4. Separate the bead-cell complexes using a magnet or by placing tube in magnetic separation device for 5 to 10 min. For this separation, cells may be transferred to a different vessel if necessary.
5. Aspirate and discard fluid from the tube or separation vessel. Remove vessel from magnetic separation device. 6. Resuspend bead-cell complexes in 10 ml of 4°C 1% FBS/PBS. Incubate 30 min at 4°C with gentle rocking. 7. Repeat separation and washing with 1% FBS/PBS as many times as desired. Dissociate B cells from beads by any of several methods, including the following: Capping: 8a. Suspend bead-cell complexes in 10 ml complete tissue culture medium with 10% FBS in a tissue culture flask and incubate 12 to 24 hr at 37°C in a humidified 37°C, 5% CO2 incubator. 9a. Expose flask to a magnet or place in magnetic separation device for 5 to 10 min. Exposure to anti-mouse antibodies (Detach-a-bead): 8b. Suspend bead-cell complexes in 100 to 200 µl complete tissue culture medium with 10% FBS in a polypropylene test tube. Add 10 to 20 µl Detach-a-bead or an appropriate volume of other anti-mouse Fab antibody. Incubate 1 hr at 25°C with gentle rocking. 9b. Add 5 to 10 ml tissue culture medium with 10% FBS, mix thoroughly, and expose tube to a magnet or place in magnetic separation device for 5 to 10 min. 10. Remove and save suspension, which contains unbound B cells. Repeat capping or exposure to anti-mouse antibodies as desired to remove more beads. 11. Centrifuge 10 min at 200 × g, 4°C. Resuspend cells in tissue culture medium. Specimen Handling, Storage, and Preparation
5.1.7 Current Protocols in Cytometry
BASIC PROTOCOL 7
ISOLATION OF T OR B LYMPHOCYTES BY PANNING T or B lymphocyte subsets may be purified through the use of panning techniques (Wysocki and Sato, 1978). These are very similar to the magnetic bead separation technique described in Basic Protocol 6, except the antibodies are used to adhere the cells of interest to a plastic vessel while the unwanted cells are washed away. Yield will vary with cell type to be isolated and the antibody used. Materials Highly specific anti-mouse IgG (Dako) 0.05 M Tris⋅Cl, pH 9.5 Specific mouse monoclonal antibodies (e.g., CD19 for B lymphocytes and CD3 for mature T lymphocytes) 10% sodium azide (optional) Phosphate-buffered saline, pH 7.2 (PBS; APPENDIX 2A) Mononuclear cell preparation (see Basic Protocol 2) 5% and 1% FBS/PBS: PBS with 5% or 1% (v/v) FBS (heat inactivated; APPENDIX 2A), 4°C Complete tissue culture medium (APPENDIX 3B) 100 × 15 mm–petri dish, bacteriological grade 15-ml polystyrene centrifuge tubes Additional reagents and equipment for counting cells and assessing cell viability by trypan blue exclusion (APPENDIX 3B) or using propidium iodide (UNIT 9.2) 1. Dilute anti-mouse IgG to ∼10 µg/ml in 0.05 M Tris⋅Cl, pH 9.5. Place 10 ml in a petri dish and incubate 1 hr at room temperature. 2. Add 2 × 107 mononuclear cells to a 15-ml polystyrene tube. Add specific monoclonal antibody and, optionally (depending on what the cells will be used for), 0.01% sodium azide. Incubate 15 min to 1 hr, at room temperature if sodium azide is used or at 4°C without azide. The amount of antibody to be used will be variable. Roughly the same concentration which yields the brightest staining by immunofluorescence or immunoperoxidase methods is generally appropriate.
3. While cells are incubating, remove unbound IgG from petri dish using a sterile pipet. Add 10 ml of 5% FBS/PBS to plate, swirl, and remove. Repeat wash with a second 10 ml of 5% FBS/PBS. 4. Wash the antibody-treated cells from step 2 by adding 10 ml of 5% FBS/PBS, centrifuging 10 min at 400 × g, and removing supernatant. Repeat wash, then resuspend the cells in 3 ml of 5% FBS/PBS. 5. Remove FBS/PBS from coated petri dish and pour cells onto dish. Incubate 1 hr at 4°C on a flat surface. Swirl cells once midway through the incubation to ensure even coating of dish. 6. Remove nonadherent cells by gently pipetting supernatant from the dish. Wash cells gently as in step 3 with 5 to 10 ml of 1% FBS/PBS. Repeat wash. 7. Remove adherent cells by adding 5 to 10 ml of 1% FBS/PBS to the dish and gently scraping with a pipet bulb or tissue culture scraper. Alternatively, harvest cells by aggressive pipetting. Handling, Storage, and Preparation of Human Blood Cells
8. Wash cells (as in step 4) in 1% FBS/PBS, count (APPENDIX 3A), and resuspend in complete tissue culture medium. 9. Check viability using trypan blue exclusion (APPENDIX (UNIT 9.2).
3B)
or propidium iodide
5.1.8 Current Protocols in Cytometry
ENRICHMENT OF B OR NK CELLS BY REMOVAL OF T LYMPHOCYTES THROUGH COMPLEMENT-MEDIATED LYSIS
BASIC PROTOCOL 8
This protocol presents a relatively simple procedure for using negative selection to deplete T cells from mononuclear cell preparations when B cells or NK cells are the population of interest. T cells are lysed with antibody and complement. The subtleties of this procedure revolve around each lot of reagent used. The complement must not be cytotoxic to any of the cells in the absence of specific antibody and the amount of complement and antibody to be used must be determined by titration. Materials Rabbit complement (Harlan Bioproducts for Science) in tissue culture medium Mononuclear cell preparation (see Basic Protocol 2) Anti–T cell antibodies: e.g., anti-CD2, CD3, CD5, or CD7 15-ml conical centrifuge tube 1% FBS/PBS: phosphate-buffered saline (PBS; APPENDIX 2A) with 1% (v/v) FBS (heat inactivated; APPENDIX 2A) 1. Place 107 mononuclear cells in a 15-ml conical centrifuge tube. Centrifuge 10 min at 400 × g, 10°C. 2. Remove supernatant and resuspend cells in an appropriate dilution of antibody. Incubate 30 min at 25°C. 3. Centrifuge 10 min at 400 × g, 10°C. 4. Remove the supernatant and resuspend cells in the appropriate concentration of complement in tissue culture medium. Incubate 45 to 60 min at 25°C. The concentration of complement that gives maximum antibody-specific cytotoxicity is generally 20% to 50% (depending on the source). This can be determined by titration: simultaneously perform the procedure with different dilutions of complement, and then use the concentration that gives minimal nonspecific cytotoxicity and maximum antibody-specific cytotoxicity.
5. Wash the enriched cells from step 4 with 10 ml of 1% FBS/PBS, centrifuging 10 min at 400 × g, 4°. ISOLATION OF PERIPHERAL BLOOD STEM CELLS USING AN IMMUNOSORBENT COLUMN
BASIC PROTOCOL 9
The method for isolating peripheral blood stem cells may vary depending on how one defines stem cells. In this protocol, CD34+ cells are considered to be synonymous with stem cells. If stem cells are defined by additional markers or by antigen density, this method may be used as an enrichment technique. Materials CellPro Ceperate LC Kit containing: Avidin column PBS P30 gel 1% and 5% BSA in PBS Precolumn gel Primary antibody (CD34) Secondary biotinylated antibody Sample/wash chamber Sample stand
Specimen Handling, Storage, and Preparation
5.1.9 Current Protocols in Cytometry
Cell preparation (see Basic Protocol 2) Peristaltic pump 15-ml centrifuge tubes 1. Mount sample/wash chamber assembly onto the sample stand. Remove assembly from its pouch and mount on the sample stand as directed in manufacturer’s instructions. Remove cap at the bottom of the three-way valve. 2. Remove the avidin column from its foil pouch, remove luer lock cap from top of column, and attach top of the column to bottom of the three-way valve on sample assembly. Tighten the luer nut at the connection. 3. Insert tubing from the bottom of avidin column into the peristaltic pump. Cut end of tubing and place tubing into a 15-ml centrifuge. 4. Prime the sample/wash chamber and column with PBS to remove any bubbles that may interfere with cell separation. Open sample chamber and turn three-way valve to permit flow between the sample chamber and column. Squeeze the chamber to remove air, and, while squeezing, add 5 ml PBS to sample chamber. Release chamber and turn the three-way valve to allow PBS to flow from the sample chamber to the wash chamber. Turn the three-way valve to permit PBS to flow from the sample chamber into the column. Add 10 ml PBS to wash chamber and 5 ml PBS to sample chamber. Add 0.5 ml P30 gel to sample chamber and allow it to settle onto filter near the bottom of the chamber. 5. Wash P30 gel and column with 5 ml PBS, then wash with 5 ml of 5% BSA/PBS using the peristaltic pump. Stop when the fluid reaches the top of the P30 gel in the sample chamber. 6. Wash cells in PBS with 1% BSA and suspend at a density of 108 cells/ml in 1% BSA/PBS. Add the appropriate amount of CD34 antibody to cells, mix, and incubate 25 min at 25°C. 7. Wash cells twice with 1% BSA/PBS. Resuspend cells to a density of 108 cells/ml in 1% BSA/PBS and add the secondary biotinylated antibody. Mix and incubate 25 min at 25°C. 8. Wash cells in 5% BSA/PBS and resuspend at a density of 108 cells/ml in 5% BSA/PBS. 9. Load labeled cells onto the column by layering cells onto the P30 gel in the sample chamber. Pump cells through the column at a flow rate of 0.75 ml/min. When the last of the cell suspension reaches the top of the P30 gel, add 2 ml of 5% BSA/PBS to the sample chamber and pump the remaining cells into the column. When sample chamber wash is nearing the top of the P30 gel, turn the three-way valve to allow 4 ml of PBS from the wash chamber to be pumped through the column. Close the clamp below the wash chamber. 10. Collect the adsorbed CD34+ stem cells. Cut the tubing 10 to 15 cm below the column, but before the pump. Place the end of the tubing into a new 15-ml centrifuge tube. Open the clamp below the wash chamber to permit PBS to start flowing through the column and squeeze the column ten times. Allow column to drain into the collection tube. Allow all of the PBS in the wash chamber to drain into the column. Handling, Storage, and Preparation of Human Blood Cells
The centrifuge tube contains the enriched CD34+ stem cells. These may be assayed as desired. If the cells are not used immediately, they should be transferred to another medium for optimal storage.
5.1.10 Current Protocols in Cytometry
PREPARATION OF PLATELET-ENRICHED PLASMA For many platelet assays, the platelets do not need to be purified by density-gradient separation. Platelet-enriched plasma, prepared by enrichment of platelets from peripheral blood (Ault, 1988), is often an acceptable specimen.
BASIC PROTOCOL 10
Materials Peripheral blood in EDTA or appropriate anticoagulant Tyrode’s buffer (see recipe) 15-ml conical centrifuge tube 1. Centrifuge 7 ml blood (in collection tube) 10 min at 200 × g, 25°C. 2. With a sterile pipet, transfer the plasma layer to a 15-ml conical centrifuge tube. Centrifuge 10 min at 1600 × g, 25°C. 3. Remove and discard supernatant. Resuspend pellet containing platelets in Tyrode’s buffer or a buffer containing EDTA. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Tyrode’s buffer 137 mM NaCl 12 mM NaHCO3 5.5 mM glucose 2 mM KCl 1 mM MgCl2 0.3 mM Na2HPO4 Adjust pH to 7.4 with HCl COMMENTARY Background Information Flow cytometry is a relatively unique technology in that cells need not be purified or separated for the study of a particular subpopulation or clone within a more heterogenous population of cells. Multiparameter analyses and electronic gating permit flow cytometry studies to be performed without purification of the subpopulations. It is even theoretically possible to analyze peripheral blood leukocytes without prior lysis of erythrocytes using multiparameter methods (Terstappen et al., 1991). Indeed, “live” gating techniques where data are gathered during acquisition and prior to analysis my be viewed as “electronic separation” of cell populations, permitting analysis of small populations of cells without their physical separation. Additionally, flow cytometry provides an excellent means of isolating specific cell populations with high purity, if high yield is not an overriding factor. Although flow cytometry may be used to analyze cells in heterogeneous populations, it
is quite often desirable to enrich, purify, or in some way separate populations of cells prior to flow cytometric analysis. The most common example of this is the removal of erythrocytes from peripheral blood prior to analysis of leukocytes. Though not absolutely required, the ease with which erythrocyte removal can be performed, the minimal impact this has on most assays, and the extent of the cytometric problems this avoids makes this removal desirable. Separation or enrichment of cell populations prior to flow cytometric sorting studies may also be well worth the effort. The throughput and, ultimately, the yield of a desired population of cells may be greatly enhanced by enriching for the target cells in the starting population. The techniques described in this unit can be used in many different ways. Protocols using antibodies for separation can be applied to a variety of cell types if the appropriate antibodies and cell surface antigens are available. Several of the methods, including the immunosor-
Specimen Handling, Storage, and Preparation
5.1.11 Current Protocols in Cytometry
bent columns and the magnetic bead separation techniques, can be used in either a positive- or a negative-selection mode. Again, the manner in which these techniques are used depends on the cell surface antigens and their corresponding antibodies, as well as any effect these may have on cell function.
Critical Parameters
Handling, Storage, and Preparation of Human Blood Cells
Numerous points must be considered in using preenriching or separation techniques prior to flow cytometric studies. In addition to yield and purity, possible alterations in cell function must be considered when cells are enriched for subsequent functional studies. Viability and processing time are other issues that may affect the separation or enrichment methods used, or whether any precytometric method should be used at all. Precytometric enrichment or separation techniques are not without ancillary effects, both beneficial and detrimental. For example, erythrocyte lysis techniques are quite effective in removing mature erythroid cells from blood, but may leave numerous immature, nucleated erythroid cells in the specimen if they were present in the original blood sample. These may be detected in the lymphoid gate by light scatter and may confuse the determination of how lymphoid subsets are distributed (Slade et al., 1988). On the other hand, a separation technique such as Ficoll-Hypaque not only removes granulocytes from the mononuclear cells, it has the added advantage of removing many dead lymphoid cells from the specimen as well. If cell selection methods are used prior to flow cytometric immunophenotyping, care should be taken that these methods do not affect the staining or light-scatter attributes of the cells of interest. For example, use of antibodies in these methods may interfere with subsequent immunofluorescence staining. Alternatively, granulocytes, for example, may have significantly different scatter properties after cell separation procedures. After any cell separation or preparation technique, particularly if cell surface antigens are to be examined, it is highly recommended that cellular viability be determined. Cells should also be thoroughly washed in isotonic buffer to assure the cells are as free of contaminants as possible and in a native condition. The latter may be of particular concern when staining cells with lectins. Lectins bind carbohydrate structures on the cells, and this may be hindered by residual sugars from the medium or from density gradients.
One last note concerning the handling and preparation of human specimens for flow cytometric analyses: All human-derived material should be considered potentially infectious and should be handled with appropriate caution! The precise precautions taken—e.g., whether specimens are prepared under a laminar flow hood—may vary somewhat from laboratory to laboratory. Certain universal precautions, such as wearing of latex gloves, disinfecting of countertops and equipment with bleach solutions, and the use of face shields should always be followed. No specimen from a human, even a “normal control,” should ever be considered “safe.” Fixation of cells with formalin or paraformaldehyde after staining reduces the infectious potential of human specimens considerably, yet prudent precautions are still advised even when working with fixed specimens.
Troubleshooting In attempting to troubleshoot the procedures presented in this unit, it is critical that an initial assessment be made of the specimen (cells) to be purified or enriched. The percentage of cells of interest in the initial specimen, and the absolute number of cells of interest should be determined prior to any of the procedures. Similarly, the types and numbers of unwanted cells contaminating the initial specimen should be determined. In general, this may best be accomplished by flow cytometric immunopyhenotyping together with manual or automatic cell counts. There is usually a trade-off between obtaining maximum purity and maximum yield in isolating cells of interest. This should be borne in mind when separating cells and whether purification (maximum purity) or enrichment (maximum yield) is the ultimate goal. This goal is obviously determined by the final use of the cells and the influence of the remaining, contaminating cells in preparation in the final application. Unwanted contaminants in the final cell preparation can be the result of any of a number of factors. Assuming protocols are followed precisely and the beginning specimen was typical, a common reason for failure in purification or enrichment of cells is reagent failure. This may be due to substandard lots of reagents, improper storage, or the use of inappropriate or expired reagents. If possible, all reagents should be checked (at least grossly) prior to use. Such checks might include examining the reagent bottles for obvious contamination, re-
5.1.12 Current Protocols in Cytometry
agent expiration date, and any unusual discoloration or precipitate. It is also crucial in most instances to assess the viability of separated cells. This may be easily done by any number of methods (e.g., see APPENDIX 3B & UNIT 5.2). Additionally, viability should be determined prior to separation procedures. Low initial viability may greatly alter purification, even if viable cells are not needed in the final specimen. Low viability in the final specimen may be due to the presence of cytotoxic agents, such as preservatives, in one of the reagents. A reagent to be considered is the fetal bovine serum, which may not have been properly heat inactivated. Alternatively, poor viability may be due to overly harsh treatment of the cells (e.g., centrifugation).
Anticipated Results The yield and purity from each of these methods is different. As stated above, before using any of these separation procedures is used, a decision must be made whether this is to be a purification or enrichment process. The resulting yields must be anticipated and evaluated in this light.
Time Considerations The length of time required to separate cells by these procedures ranges from several minutes to many hours. With most procedures, optimal results will be obtained if cells are processed as soon as possible after collection from the donor. This will minimize problems with viability and any possible alteration of the cell surface antigens.
Literature Cited Ault, K.S. 1988. Flow cytometric measurement of platelet-associated immunoglobulin. Pathol. Immunopathol. Res. 7:395-408. Boyum, A. 1968. Isolation of mononuclear cells and granulocytes from human blood. Scand. J. Clin. Lab. Invest. Suppl. 21:77-89. Boyum, A. 1977. Separation of lymphocytes, lymphocyte subgroups and monocytes: A review. Lymphology 10:71-76. Denholm, E.M. and Wolber, F.M. 1991. A simple method for the purification of human peripheral blood monocytes. A substitute for SepracellMN. J. Immunol. Methods 144:247-251. Indiveri, F., Huddlestone, J., Pellegrino, M., and Ferrone, S. 1980. Isolation of human T lymphocytes: Comparison between wool filtration and rosetting with neuraminidase (VCN) and 2-aminoethylisothiouronium bromide (AET)–treated sheep red blood cells. J. Immunol. Methods 34:107-115.
Madsen, M., Johnsen, H.E., and Kissmeyer-Nielsen, F. 1979. Separation of human T and B lymphocytes using ET-treated sheep red blood cells. Transplant. Proc. 11:1381-1382. Molday, R.S., Yen, S.P., and Rembaum, A. 1977. Application of magnetic microspheres in labelling and separation of cells. Nature 268:437-438. Rasmussen, A.M., Smeland, E.B., Erikstein, B.K., Caignault, L., and Funderud, S. 1992. A new method for detachment of Dynabeads from positively selected B lymphocytes. J. Immunol. Methods 146:195-202. Slade, H.B., Greenwood, J.H., Hudson, J.L., Beekman, R.H., Riedy, M.C., and Schwartz, S.A. 1988. Lymphocyte phenotyping of infants with congenital heart disease: Comparison of cell preparation techniques. Diagn. Clin. Immunol. 5:249-255. Terstappen, L.W., Johnson, D., Mickaels, R.A., Chen, J., Olds, G., Hawkins, J.T., Loken, M.R., and Levin, J. 1991. Multidimensional flow cytometric blood cells differentiation without erythrocyte lysis. Blood Cells 17:585-602. Wysocki, L.J. and Sato, V.L. 1978. Panning for lymphocytes: A method for cell separation. Proc. Natl. Acad. Sci. U.S.A. 75:2844-2848.
Key References NCCLS. 1989. Clinical Applications of Flow Cytometry: Quality Assurance and Immunophenotyping of Peripheral Blood Lymphocytes. 1989. NCCLS Doc. H42-P, Vol. 9, No.13. Provides general guidelines for performance of flow cytometric immunophenotyping. NCCLS. 1993. Clinical Applications of Flow Cytometry: Quality Assurance and Immunophenotyping of Leukemic Cells; Proposed Guidelines. 1993. NCCLS Doc. H43-P, Vol. 13, No. 23. Provides more detailed guidelines for flow cytometric immunophenotyping of leukemias and lymphomas. McCoy, J.P., Carey, J.L., and Krause, J.R. 1990. Quality control in flow cytometry for diagnostic pathology: I. Cell surface phenotyping and general laboratory procedures. Am. J. Pathol. 93(Suppl. 1):S27-S37. Discusses quality control issues in the performance of low cytometric immunophenotyping. Stewart, C.C. 1990. Cell preparation for the identification of leukocytes. Methods Cell Biol. 33:411-426. Details sample preparation methods for immunophenotyping studies.
Contributed by J. Philip McCoy, Jr. Cooper Hospital/University Medical Center Camden, New Jersey
Specimen Handling, Storage, and Preparation
5.1.13 Current Protocols in Cytometry
Handling, Storage, and Preparation of Human Tissues
UNIT 5.2
In contrast to blood cells, human tissue for use in flow cytometry must first be prepared as an adequate single-cell suspension. Selection of the appropriate methods for specimen collection, transport, storage, and tissue dissociation depend on the cell parameters being measured (e.g., protein, DNA, or RNA) and the localization of the markers (e.g., cell surface, cytoplasm, or nucleus). This unit includes a general method for collecting and transporting tissue samples to the flow cytometry laboratory (see Basic Protocol 1) and preparing a tissue imprint to assess the sample (see Support Protocol 1). Single-cell suspensions of whole cells can be prepared from fresh tissue using mechanical disaggregation (see Basic Protocol 2) or enzymatic disaggregation (see Alternate Protocol 1). Single-cell suspensions of whole cells can also be prepared from fine-needle aspitates (see Basic Protocol 3) or pleural effusions, abdominal fluids, or other fluids (see Alternate Protocol 2). Once a single-cell suspension is obtained, the cells can be fixed (see Support Protocol 2), used for cytospin preparations (see Support Protocol 3), or frozen for future use (see Support Protocol 4). Intact nuclei can be prepared for flow cytometry from fresh or frozen tissue (see Basic Protocol 4) or from paraffin-embedded tissue (see Alternate Protocol 3). Debris can be removed from nuclear preparations, from frozen or paraffinembedded samples, by a sucrose step gradient (see Support Protocol 5). CAUTION: Handle all standards, controls, and patient samples as if they are capable of transmitting hepatitis and AIDS. Refer to Universal Precaution Procedures for correct handling procedures. COLLECTION AND TRANSPORT OF SAMPLES TO THE FLOW CYTOMETRY LABORATORY
BASIC PROTOCOL 1
Most often human tissues are received from a hospital pathology laboratory or surgical suite. Before using tissue for any assay, the investigator should ensure that appropriate tissue has already been obtained for diagnosis by the pathologist and that, if needed, appropriate consent forms have been signed. The collaborating pathologist or tissue procurement person should be aware of the needs of the study so that representative tissue is appropriately collected and transported, especially if the use for the tissue requires special handling (see Table 5.2.1). For example, if RNA must remain intact, the tissue should be frozen immediately, but freezing is not a requirement for DNA ploidy and S-phase determinations. For most protein and DNA flow cytometry assays, tissue can be collected at room temperature and placed in a clean (sterile, if possible) vessel containing growth medium. The sample is placed on ice and transported to the laboratory. For Table 5.2.1
Sample Collection and Transport Conditions
To study
Collect tissue
Transport tissue
DNA (e.g., ploidy, % S) Nuclear proteins
Fresh Fresha
At 4°C At 4°Ca
Cell-surface proteins RNA
Fresha Freeze immediately
At 4°Ca In liquid nitrogen or isopentene
aDepending on the nature of the protein—e.g., enzymes—the conditions for transport may vary and may require
freezing; the investigator will need to identify optimal conditions depending on the specific protein being assayed.
Contributed by Lynn G. Dressler and Dan Visscher Current Protocols in Cytometry (1997) 5.2.1-5.2.15 Copyright © 1997 by John Wiley & Sons, Inc.
Specimen Handling, Storage, and Preparation
5.2.1 Supplement 1
efficiency, refrigerated prefilled containers of growth medium can be kept at the sample collection site. Cells in suspension as aspirates or fluid samples can simply be obtained in a sterile container, bag, or syringe and transported on ice to the laboratory. Once the tissue is received, it is essential to include a quality assurance check to ensure that appropriate tissue (normal or malignant) is obtained; this can be accomplished in a variety of methods including analyzing tissue imprints (see Support Protocol 1), frozen sections, or cytospin preparations (see Support Protocol 3). Materials Growth medium (e.g., MEM or RPMI 1640) containing 5% (v/v) heat-inactivated FBS (APPENDIX 2A) or Hanks’ balanced salt solution (HBSS; APPENDIX 2A) Liquid nitrogen or isopentane bath Sample containers (e.g., 50-ml conical centrifuge tubes or cryovials), sterile Curved and straight forceps, sterile Cryovials Additional reagents and equipment for preparing a tissue imprint (see Support Protocol 1) To collect samples for DNA or protein assays 1a. Add 25 ml growth medium containing 5% FBS to a 50-ml conical centrifuge tube. Keep tube on ice. Alternatively, use tubes that have been filled with growth medium and refrigerated. Sterility is preferable but not essential.
2a. Have the pathologist or appropriate person procure a representative piece of tissue and trim away fat and necrotic areas. If possible, prepare a tissue imprint (see Support Protocol 1). 3a. Using sterile forceps, place sample in conical tube on ice. Label tube with appropriate identification information, date, and time of collection. Transport tube to lab on ice. 4a. Keep sample at 4°C and dissociate as soon as possible, preferably within 2 hr. Depending on the assay, samples may be stored overnight in the refrigerator, if necessary, before dissociation (see analysis protocols for details).
To collect samples for RNA assays or those which are susceptible to protein or nucleic acid degradation 1b. Have the pathologist or appropriate person procure a representative piece of tissue, and trim away fat and necrotic areas. Prepare a tissue imprint (see Support Protocol 1). Sample may need to be cut into multiple pieces depending on size of tissue and investigator need.
2b. Place tissue in a dry cryovial, marked with appropriate identification information and date of collection. Using long forceps, place vial in liquid nitrogen or isopentane bath for 30 to 60 seconds. 3b. Transfer vial to bucket containing dry ice or keep in liquid nitrogen or isopentane bath and transport to lab. Keep sample frozen during transport. Handling, Storage, and Preparation of Human Tissues
4b. Store sample at −70°C or lower.
5.2.2 Supplement 1
Current Protocols in Cytometry
PREPARING A TISSUE IMPRINT Tissue imprints are used to verify that the correct tissue has been obtained and to provide a preliminary evaluation of cellularity. The tissue sample is touched to a slide, and the slide is fixed and stained.
SUPPORT PROTOCOL 1
Materials Tissue specimen, freshly collected 95% ethanol Forceps Glass slides Coplin jar Papanicolaou staining set or hematoxylin-eosin staining series 1. Gently hold the tissue specimen with forceps. Touch the freshly cut surface of the tissue with a glass slide. 2. Fix the slide immediately by immersing it in a Coplin jar containing 95% ethanol ≥30 sec. These preparations will air dry almost immediately because of their relatively low cellularity, so it is essential that slides be fixed immediately to prevent drying artifacts.
3. Stain slide with Papanicolaou or hematoxlyin-eosin, add a coverslip, and examine for presence of appropriate cells and cellularity (e.g., for cancer specimens, ensure that ≥15% of cells are malignant). MECHANICAL DISAGGREGATION OF WHOLE CELLS FROM FRESH HUMAN TISSUE
BASIC PROTOCOL 2
Flow cytometric analysis of fresh solid tumors requires the recovery of sufficient numbers of intact, viable tumor cells, as a single-cell suspension, either by mechanical or enzymatic disaggregation (see Alternate Protocol 1). Mechanical disaggregation involves the dispersion of cells by scraping and mincing the tumor with a scalpel. The resulting cell suspension is filtered, counted, and fixed (see Support Protocol 2) prior to flow cytometry (Hitchcock, et al., 1996). Materials Tumor tissue, freshly obtained (see Basic Protocol 1) Growth medium (e.g., RPMI 1640 medium, Life Technologies) FBS, heat-inactivated (Life Technologies or APPENDIX 2A) 0.4% (w/v) trypan blue (Sigma) Gauze, sterile No. 22 scalpel blades and handle 100-mm petri dishes 80-µm metal-mesh sieve (EC Apparatus) 15- and 50-ml conical plastic centrifuge tubes IEC CL centrifuge and swinging bucket rotor (or equivalent) Additional reagents and equipment for counting cells (APPENDIX 3A) and assessing viability (APPENDIX 3B), and fixing cell suspension (see Support Protocol 2) NOTE: This protocol should be carried out in biological safety cabinet.
Specimen Handling, Storage, and Preparation
5.2.3 Current Protocols in Cytometry
1. Remove fresh tumor tissue from medium and blot on sterile gauze. Although it is optimal to dissociate cells as soon as possible, for most DNA flow cytometry protocols, samples may be stored at 4°C up to 24 hr without deleterious effects.
2. Using a no. 22 scalpel blade, trim fat and areas of necrosis. Weigh remaining tissue. 3. Place tissue in a petri dish containing 5 to 10 ml growth medium. Using forceps and a no. 22 scalpel blade, gently scrape the tissue until the solution becomes turbid. Then bisect the specimen to provide more surface area and repeat. Continue to scrape and mince the tissue until no more cells are released. 4. Pour the solution through an 80-µm metal-mesh sieve into a 15-ml conical centrifuge tube. Centrifuge 10 min at 250 × g (4000 rpm in swinging bucket rotor), room temperature. 5. Remove supernatant and resuspend pellet in 1 ml growth medium plus 1 ml heat-inactivated FBS. 6. Combine 10 µl of cell suspension with 10 µl of 0.4% trypan blue. Using a hemacytometer, count the cells (APPENDIX 3A) and assess viability (APPENDIX 3B). CAUTION: Trypan blue is a potential carcinogen and should be handled carefully. Dead cells take up trypan blue.
7. Fix cell suspension (see Support Protocol 2). If the antigen to be assayed is destroyed by fixation, the cells do not have to be fixed, except to prepare cytospin slides. ALTERNATE PROTOCOL 1
ENZYMATIC DISAGGREGATION OF WHOLE CELLS FROM FRESH HUMAN TISSUE Enzymatic disaggregation of solid tumors uses trypsin to disrupt intercellular adhesions and collagenase Type II to digest stromal components. DNase I is also added to ensure that ploidy is determined only from populations of intact cells. Enzymatic disaggregation can be used alone or in combination with mechanical methods and it is recommended for complete dissociation of tumors of squamous origin. Additional Materials (also see Basic Protocol 2) Trypsin Collagenase type II DNAse I Additional reagents and equipment for counting cells (APPENDIX 3A) and assessing viability (APPENDIX 3B), and fixing cell suspension (see Support Protocol 2) 1. In a 50-ml centrifuge tube, prepare a 10-ml enzyme cocktail containing 2.5 mg/ml trypsin, 0.5 mg/ml collagenase Type II, and 20 µg/ml DNase I in growth medium. Warm 15 min at 37°C. 2. Place fresh tumor tissue or mechanically disaggregated tissue (Basic Protocol 2, step 5) into the enzyme cocktail. Incubate 1 hr at 37°C with gentle agitation.
Handling, Storage, and Preparation of Human Tissues
3. Pour the solution through an 80-µm metal-mesh sieve into a 15-ml centrifuge tube. Add 2 ml heat-inactivated FBS to stop the reaction. Centrifuge 10 min at 250 × g (4000 rpm in swinging bucket rotor), room temperature. 4. Discard the supernatant and resuspend the pellet in growth medium.
5.2.4 Current Protocols in Cytometry
5. Combine 10 µl of cell suspension with 10 µl of 0.4% trypan blue. Using a hemacytometer, count the cells (APPENDIX 3A) and assess viability (APPENDIX 3B). CAUTION: Trypan blue is a potential carcinogen and should be handled carefully. Dead cells take up trypan blue.
6. Fix cell suspension (see Support Protocol 2). If the antigen to be assayed is destroyed by fixation, the cells do not have to be fixed, except to prepare cytospin slides.
PREPARATION OF CELLS FROM FINE-NEEDLE ASPIRATES Frequently human cells are obtained as a single-cell suspension in fine-needle aspirates, bone marrow aspirates, or pleural, ascitic, or other fluid samples (see Alternate Protocol 2). No dissociation is required for these samples. If the sample contains red blood cells, hemolyzing reagent should be added to lyse them.
BASIC PROTOCOL 3
Materials Needle or bone marrow aspirate in a syringe or sterile container Growth medium (e.g., MEM, HBSS, or RPMI 1640) containing 5% (v/v) heat-inactivated FBS (Life Technologies or APPENDIX 2A) Hemolyzing reagent: 3.6% (w/v) NaCl in H2O, cold Heparin 15-ml or 50-ml conical centrifuge tubes Beckman RJ-6 centrifuge and TH-4 rotor (or equivalent) Forceps, sterile Additional reagents and equipment for counting cells (APPENDIX 3A) 1. Collect needle or bone marrow aspirate in sterile syringe or container. Transport to lab on ice and store at 4°C. Cells should be processed (washed and centrifuged) within 2 hr for DNA flow cytometry.
2. Add contents of syringe and needle or container to a 15- or 50-ml conical tube containing 5 to 10 ml growth medium with 5% FBS. If possible, add needle as well to obtain additional cells. Keep tube on ice. 3. Remove needle from tube with sterile forceps. Cap tube and invert several times to mix contents. 4. Optional: If the suspension is bloody, centrifuge 5 min at 125 × g (800 rpm in TH-4 rotor), 4°C. Resuspend pellet in 12 ml cold water and add 4 ml of cold hemolyzing reagent. Invert tube several times to mix contents and incubate a total of ∼20 sec. Centrifuge 5 min at 125 × g, 4°C. If the pellet is still bloody, repeat. A 20-sec incubation should be sufficient to lyse red blood cells.
5. Centrifuge 5 min at 125 × g (800 rpm in TH-4 rotor), 4°C, to pellet cells. 6. Resuspend cells in 1 to 2 ml fresh growth medium. Count a sample of the suspension in a hemacytometer (see APPENDIX 3A). Adjust cell concentration to 1 to 3 × 106 cells/ml. Sample can be either used for flow cytometry or frozen and stored for later use (see Support Protocol).
Specimen Handling, Storage, and Preparation
5.2.5 Current Protocols in Cytometry
ALTERNATE PROTOCOL 2
PREPARATION OF CELLS FROM PLEURAL EFFUSIONS, ABDOMINAL FLUIDS, AND OTHER FLUIDS Large volumes of fluid (pleural, abdominal, or other fluid) need to be centrifuged and the cell pellets washed and pooled for use. It is important to add hemolyzing agent if the sample contains red blood cells. Additional Materials (also see Basic Protocol 3) Pleural effusion, abdominal fluid, or other fluid 25-ml pipet and bulb, sterile 15- or 50-ml centrifuge tubes or 250-ml centrifuge bottles, sterile Additional reagents and equipment for counting cells (APPENDIX 3B) 1. Using a 25-ml pipet and bulb or a Pasteur pipet, transfer pleural effusion, abdominal fluid, or other fluid sample to 15-ml or 50-ml sterile centrifuge tubes or 250-ml sterile centrifuge containers (depending on total volume). Centrifuge 5 min at 150 × g (1200 rpm in TH-4 rotor), 4°C. 2. Discard supernatant and pool sample pellets. 3. Optional: If the suspension is bloody, centrifuge 5 min at 125 × g (800 rpm in TH-4 rotor), 4°C. Resuspend pellet in 12 ml cold water and add 4 ml of cold hemolyzing reagent. Invert tube several times to mix contents and incubate a total of ∼20 sec. Centrifuge 5 min at 125 × g, 4°C. If the pellet is still bloody, repeat. A 20-sec incubation should be sufficient to lyse red blood cells.
4. Discard supernatant and resuspend pellets in an appropriate volume (for cell counts) of growth medium containing 5% FBS. Count cells in hemacytometer (APPENDIX 3A). Adjust cell concentration to 1 to 2 × 106 cells/ml for immediate assay or 3 to 5 × 106 cells/ml for freezing (see Support Protocol 4) and future use. SUPPORT PROTOCOL 2
ETHANOL FIXATION OF CELL SUSPENSIONS Fixation of disaggregated solid tumor single-cell suspensions is accomplished by adding 70% ethanol drop-wise while vortexing. Fixed specimens can be stored at 4°C for up to 5 days without compromising nuclear integrity. This method is sufficient for most flow cytometry DNA ploidy analysis and can be used in conjunction with simultaneous immunofluorescence for a variety of intracellular antigens including intermediate filaments and/or leukocyte common antigen. Materials Whole cell suspension (see Basic Protocol 2 or 3 or Alternate Protocol 1 or 2) 70% ethanol, 4°C 15-ml centrifuge tube 1. Transfer 2 ml resuspended single-cell suspension into a 15-ml centrifuge tube. While vortexing, add 6 ml cold 70% ethanol drop-wise. This cell/70% ethanol ratio results in a final ethanol concentration of 50%. If the sample is >2 ml, use multiple tubes or larger tubes with the same cell/ethanol ratio.
2. Cap centrifuge tube and store at 4°C for a minimum of 30 min to a maximum of 5 days. Handling, Storage, and Preparation of Human Tissues
3. Use fixed cells for flow cytometry or to prepare cytospin slides (see Support Protocol 3).
5.2.6 Current Protocols in Cytometry
PREPARATION OF CYTOSPIN SLIDES FROM CELL SUSPENSIONS Cytospin slides prepared from dissociated solid tumor cell suspensions can be used as an important quality control for flow cytometry (Dressler and Bartow, 1989). In addition to reflecting tumor cell versus non-malignant cell representation, cytospin slides also provide an assessment of specimen quality in terms of relative amounts of debris and cellular damage. Prepared cytospin slides are stained using either Papanicolaou or Leishman stains.
SUPPORT PROTOCOL 3
Materials PBS (see APPENDIX 2) Ethanol-fixed single-cell suspension (see Support Protocol 2) 0.1% (w/v) bovine serum albumin (BSA, Baxter) in PBS Leishman or Papanicolaou stain (Sigma) 12 × 75-mm tube Glass slides Cytocentrifuge (Shandon), slide chambers, filter cards Dilute cells 1. Pipet 500 µl PBS into a 12 × 75–mm tube. 2. Add 50 µl ethanol-fixed single-cell suspension at an adjusted concentration of 1 × 106 cells/ml. Mix gently. Assemble cytospin apparatus 3. Label glass slide with specimen number, date, and other identification information. 4. Assemble the slide, filter card, and cytospin chamber, carefully aligning the hole in the filter card with the opening of the chamber. 5. Insert the slide/chamber unit into a slot in the centrifuge head. Balance the cytocentrifuge. Prepare slides 6. Add 300 µl diluted cell suspension to the chamber. 7. Add 50 µl 0.1% mg/ml BSA to maintain cellular integrity during cytocentrifugation. 8. Close and lock the cover of the cytocentrifuge. Centrifuge slide/chamber units 5 min at 1000 rpm. 9. When the cytocentrifuge cover unlocks, carefully remove the slide/chamber unit. 10. Discard the filter card and allow the preparation to air dry. Wash the chamber before reusing. 11. Stain slides with Leishman or Papanicolaou stain according to manufacturer’s instructions.
Specimen Handling, Storage, and Preparation
5.2.7 Current Protocols in Cytometry
Supplement 6
SUPPORT PROTOCOL 4
FREEZING DISSOCIATED SINGLE CELLS Single whole cell suspensions prepared from tumor tissue can be frozen and stored for future use. Pelleted whole cells are resuspended in cryopreservation solution and frozen at −70°C. Materials Whole cell suspension (see Basic Protocol 2 or 3 or Alternate Protocol 1 or 2), unfixed Growth medium (e.g., RPMI, MEM, Medium 199) containing 5% (v/v) heat-inactivated FBS (APPENDIX 2A) Cryopreservation solution (see recipe) 1.25-ml cryovials 12 × 75–mm polystyrene tubes Beckman RJ-6 centrifuge and TH-4 rotor 1. Transfer 1.0 ml whole-cell suspension at a concentration of 3 to 5 × 106 cells/ml in growth medium containing 5% FBS into 12 × 75–mm polypropylene tube. Centrifuge 5 min at 150 × g (1200 rpm in TH-4 rotor), 4°C. 2. Decant supernatant and quickly resuspend cells in 1.0 ml cryopreservation solution. 3. Immediately transfer contents of tube into a labeled 1.25-ml cryovial and place in −70°C freezer. Samples can be stored for 4 to 6 months. The resuspension and transfer should be done carefully but quickly to minimize contact of cells with DMSO in warm cryopreservation solution.
BASIC PROTOCOL 4
ISOLATION OF NUCLEI FROM FRESH OR FROZEN TISSUE Nuclei can be prepared from fresh or frozen tissue by mechanical or enzymatic procedures. Some investigators use a combination of both. The goal is to obtain nuclei that are relatively free of any contaminating cytoplasmic debris and yet retain an intact nuclear membrane. The simple technique can be applied to fresh or frozen tissue. Nuclei are isolated from single-cell suspensions of whole cells prepared from tumor tissue. Alternatively, nuclei can be prepared from paraffin-embedded tissues (see Alternate Protocol 3). For assays that require intact whole cells see Basic Protocol 2. Materials Tumor tissue, fresh or frozen Growth medium (e.g., MEM or RPMI 1640) containing 5% (v/v) heat-inactivated FBS (APPENDIX 2A), 4°C Sample container, sterile and dry Petri dish, disposable Curved and straight forceps, sterile Scalpel and no. 20 blades 50- to 70-µm nylon mesh (Tetko) 15-ml conical centrifuge tube Beckman RJ-6 centrifuge and TH-4 rotor (or equivalent) Additional reagents and equipment for counting cells (APPENDIX 3A)
Handling, Storage, and Preparation of Human Tissues
1a. For fresh tumor tissue: Obtain tumor tissue in a dry sterile container and put on ice or at 4°C. 1b. For frozen tumor tissue: Remove vial or container from −70°C freezer and place on ice. Incubate on ice ∼8 to 10 min until the tissue reaches ∼4°C.
5.2.8 Supplement 6
Current Protocols in Cytometry
2. Using sterile forceps, transfer the tissue to a disposable petri dish on ice. Add 5 to 10 ml cold growth medium containing 5% FBS. Keep the tissue on ice or at 4°C. 3. Cut the tissue into small pieces using a scalpel with a no. 20 blade. 4. Hold the tissue in the medium using a pair of forceps. Using moderate pressure, scrape the cut surface of the tissue with a glass slide held at a 45° angle until only connective tissue remains (1 to 3 min). The medium will turn cloudy as cells are released from the tissue.
5. Using a Pasteur pipet, collect the cells and medium and filter through a 50- to 70-µm nylon mesh into a 15-ml conical centrifuge tube. 6. Centrifuge 5 min at 150 × g (1200 rpm in TH-4 rotor), 4°C, to gently pellet the cells. 7. Resuspend the pellet in 2 ml growth medium containing 5% FBS. Centrifuge 5 min at 150 × g, 4°C. If the sample is small or there is minimal cell yield, this step can be eliminated to reduce cell loss.
8. Resuspend the pellet in 1 ml growth medium containing 5% FBS. Count the cells (APPENDIX 3A) and adjust the concentration to 1 to 2 × 106 cells/ml for flow cytometry or 3 to 5 × 106 cells/ml for freezing and storage. These cells will lyse and release nuclei when they are added to the staining solution for flow cytometry.
ISOLATION OF NUCLEI FROM PARAFFIN-EMBEDDED TISSUE Nuclei can also be isolated from tumor tissue in formalin-fixed paraffin blocks for flow cytometric determination of DNA. Sections cut from the block are first dewaxed with xylene and rehydrated with ethanol and water. The sample is then digested with pepsin to disaggregate the tissue and isolate the nuclei. Isolated nuclei can be stained for flow cytometric analysis of DNA content (Dressler and Bartow, 1989).
ALTERNATE PROTOCOL 3
Materials Formalin-fixed paraffin-embedded tissue Hematoxylin-eosin or Diff-Qwik stain Xylene or Histoclear Ethanol series: 100%, 95%, 70% and 50% Pepsin solution: 0.5% (w/v) bovine pancreatic pepsin in 0.9% (w/v) NaCl Growth medium (e.g., MEM, RPMI, HBSS) containing 5% (v/v) heat-inactivated FBS (APPENDIX 2A) Propidium iodide buffer (see recipe) Microtome (adjustable, to cut 50-µm and 4-µm sections) Glass slides 13 × 100–mm glass screw top tubes 50-µm nylon mesh (Tetko4) Additional reagents and equipment for counting cells (APPENDIX 3A) Cut sections 1. Deface block of formalin-fixed paraffin-embedded tissue. Mount block on a microtome and cut one 4-µm section. Prepare a slide with this section, label it “top,” and include appropriate identifying information. Set aside for routine staining (step 4). The block should be examined by a pathologist who should mark the block for areas of malignant and nonmalignant tissue. If distinct areas are marked, first make a razor cut
Specimen Handling, Storage, and Preparation
5.2.9 Current Protocols in Cytometry
Supplement 6
along the demarcation line so malignant and nonmalignant sections for each area can be processed separately.
2. Cut three 50-µm sections from the same block. Let each section curl up. Using forceps, carefully transfer each thick section to a glass 13 × 100–mm screw top tube. Label tube appropriately with solvent pen. 3. Cut one 4-µm section from the same block. Prepare a slide, label it “bottom,” and include other identifying information. Set aside for routine staining (step 4). 4. Dewax, rehydrate, and stain the “top” and “bottom” slides using hematoxylin-eosin or Diff-Qwik stain according to manufacturer’s instructions. 5. Evaluate top and bottom sections to ensure that representative tissue is present on both sections before processing and analyzing material from thick sections. Keep stained sections on file for future reference. Dewax and rehydrate thick sections 6. Dewax thick sections using two changes of xylene (or Histoclear): add 10 ml xylene to each tube and let sit 10 min at room temperature (without vortexing). Carefully remove xylene with Pasteur pipet, replace with 10 ml fresh xylene, and incubate 10 min. CAUTION: Perform steps 6 and 7 in a fume hood. CAUTION: Collect used xylene in a discard beaker and transfer to a xylene waste container. Never pour xylene down the sink!
7. Remove xylene and rehydrate by incubating 10 min in 10 ml of each of the following (do not vortex): 100% ethanol, two 10-min changes 95% ethanol 70% ethanol 50% ethanol With each successive ethanol step, the tissue becomes more fragile and will break up into pieces. Be careful to remove only the alcohol solution without losing the tissue pieces.
8. After the final ethanol step, remove ethanol and add 10 ml distilled water to each tube, cap tube, and let stand overnight at room temperature. This step should allow any remaining alcohol trapped in the tissue to be released. Sections can stay in distilled water up to 10 days without adverse effects, but must stay ≥12 to 24 hr for optimal results.
Disaggregate tissue and isolate nuclei 9. After ≥12 to 24 hr, remove distilled water from each tube and add 10 ml fresh distilled water. Let stand 10 min at room temperature. During this 10-min incubation, take pepsin out of freezer and warm to room temperature. Add 10 ml fresh distilled water to each tissue tube. 10. Prepare 1.5 ml pepsin solution per tube. Warm solution 5 min at 37°C before adding to tubes. 11. Remove distilled water from tube and place tube in a rack in a 37°C water bath. 12. Add 1.5 ml pepsin solution to each tube and vortex 10 seconds. Handling, Storage, and Preparation of Human Tissues
13. Incubate tube 30 min in 37°C water bath, vortexing each tube at 5-min intervals. If cells stick to side of tube, wash them down with a Pasteur pipet.
5.2.10 Supplement 6
Current Protocols in Cytometry
The solution in the tube should turn cloudy as cells are released from tissue.
14. After the incubation, filter digested suspension through 50-µm nylon mesh. Wear gloves. If the filtered suspension contains considerable debris, remove the debris on a sucrose step gradient (see Support Protocol 5).
15. Centrifuge 5 min at 150 × g (1200 rpm in TH-4 rotor), 4°C, to obtain a cell pellet. Resuspend in 1.0 to 2.0 ml growth medium containing 5% FBS. Count cells and nuclei in a hemacytometer (APPENDIX 3A) and adjust to a concentration of 1 to 2 × 106 cells and nuclei/ml in propidium iodide buffer. Long-term storage of unstained nuclei is not recommended. Once isolated, nuclei should remain in growth medium at 4°C and, for optimal results, should be processed within 2 hr. Isolated, unstained nuclei can be held overnight at 4°C and stained within 24 hr; however, the risk of losing select populations (e.g., aneuploid nuclei) increases the longer nuclei are held. Time course experiments have shown that propidium iodide–stained nuclei can be held for longer periods of time, up to 24 to 72 hr; however, the risk of selective nuclear loss remains, usually of deteriorating aneuploid nuclei, which appear to have more fragile nuclear membranes. Experimental studies have shown a decrease in the aneuploid population as a function of storage time; this is somewhat variable from sample to sample. Therefore, for optimal results, use nuclei for flow cytometry within 2 hr of preparation.
REMOVING DEBRIS FROM SUSPENSIONS OF NUCLEI Sometimes filtered nuclear suspensions prepared from fresh, frozen, or paraffin-embedded tissue may contain considerable debris or increased amounts of fibrous connective tissue. In these cases, subcellular debris can be removed by centrifuging the suspension on a sucrose step gradient.
SUPPORT PROTOCOL 5
Materials 1.75 M sucrose in PBS (APPENDIX 2A) 1.5 M sucrose in PBS (APPENDIX 2A) Nuclear suspension from fresh, frozen, or paraffin-embedded tissue, filtered (see Basic Protocol 4 or Alternate Protocol 3) Growth medium containing 5% (v/v) heat-inactivated FBS (APPENDIX 2A) 12 × 75–mm centrifuge tube Beckman RJ-6 centrifuge and TH-4 rotor (or equivalent) 1. Add 800 µl of 1.75 M sucrose to a 12 × 75–mm centrifuge tube. 2. Carefully layer 800 µl of 1.5 M sucrose on top of the 1.75 M sucrose. 3. Carefully add 1.5 ml filtered nuclear suspension on top of the sucrose cushions. 4. Centrifuge 40 min at 250 × g (4000 rpm in TH-4 rotor), 4°C, to remove subcellular debris. 5. Discard supernatant. Resuspend the nuclear pellet in 1 ml growth medium containing 5% FBS.
Specimen Handling, Storage, and Preparation
5.2.11 Current Protocols in Cytometry
Supplement 1
REAGENTS AND SOLUTIONS Use sterile deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see SUPPLIERS APPENDIX.
Cryopreservation solution 1.765 g NaCl (0.3 M final) 15 ml dimethyl sulfoxide (DMSO; 15% final) H2O to 100 ml Store at 4°C Propidium iodide buffer 250 ml H2O 1.211 g Tris base (0.04 M final) 0.2541 g MgCl2⋅6H2O (5 mM final) 75 µl 0.3% (v/v) Nonidet P-40 (NP-40) 0.0125 g propidium iodide (0.05 mg/ml final) Adjust pH to near 7.0 with concentrated HCl Adjust pH to exactly 7.0 with 1 M HCl Store up to 3 weeks at 4°C in a foil-wrapped container COMMENTARY Background Information
Handling, Storage, and Preparation of Human Tissues
A fundamental requirement of flow cytometry is a sample prepared as a single-cell or single-nucleus suspension. It is imperative that the user understand the nature of the parameter being measured before choosing a procedure for sample collection, transport, and preparation (see Table 5.2.1). For example, paraffin block material can be used to measure nuclear parameters (e.g., DNA ploidy, S phase, and nuclear proteins); this fixed material cannot be used for measuring membrane proteins because the fixation and dissociation process destroys cell membrane integrity. Analysis of membrane proteins requires fresh tissue or dissociated cells from optimally fixed fresh tissue. Therefore, the user needs to understand the specific flow cytometric application and identify optimal conditions for assay, including transport and storage. Dissociation of intact cells can be accomplished by both mechanical (see Basic Protocol 2) and enzymatic (see Alternate Protocol 1) disaggregation techniques. The success of any disaggregation procedure can be evaluated by several parameters including cell yield, cell viability, maintenance and integrity of aneuploid populations, and histogram quality (coefficient of variation and baseline debris; Visscher and Crissman, 1984). The ability to successfully address these issues varies with disaggregation method as well as with tumor type. Mechanical disaggregation involves scraping and mincing to disperse cells from solid
tumors. Although cell yield and viability can vary, mechanical disaggregation has been demonstrated to be superior to enzymatic digestion for aneuploid cell recovery in breast (McDivitt et al., 1994), colon (Crissman et al., 1988), prostate (König et al., 1993), and ovarian tumors (Costa et al., 1987). Neoplasms of squamous origin or differentiation, however, require enzymatic digestion for their complete dissociation. Cell yield and viability and aneuploid cell recovery are all increased when compared to mechanical methods for these tumors (Bijman et al., 1985; Ensley et al., 1988). There is no universal disaggregation method, so any technique should be optimized to obtain maximum recovery of target cell populations comprising a solid tumor and individualized for tumor type. The most frequent use of nuclei prepared from formalin-fixed paraffin-embedded tissue is for measurement of DNA ploidy and S phase by DNA flow cytometry. David Hedley (Hedley et al., 1983; Hedley, 1994) revolutionized DNA flow cytometry with a procedure that can be applied to tissue fixed in paraffin blocks. Since then, many variations of this technique have been developed to improve the quality of the DNA histogram obtained. Basic to all procedures is the isolation of nuclei from fixed tissue by proteolytic dissolution of the disulfide linkages formed during formalin fixation. Regardless of the technique used, the user must first select an adequately fixed, representative block of tissue that contains minimal necrosis and, if possible, minimal fibrosis. This first step
5.2.12 Supplement 1
Current Protocols in Cytometry
of quality assurance is essential to any DNA flow cytometric assay (Dressler, 1993). Optimally, for flow cytometry, material should be fixed in buffered formalin for ∼24 hr prior to paraffin processing and embedding. However, the investigator usually has little control over the initial tissue processing because this is usually done by the hospital or clinical laboratory following a routine histologic procedure. Finished blocks should never be exposed to extremes of temperature; they should be transported neither frozen, nor under conditions that cause paraffin to melt (>80°F). To ensure that tissue processed for flow cytometry is representative, thin 4- to 5-µm sections immediately preceding and after the thick sections cut for assay should be obtained and stained for histopathologic evaluation. The number of thick sections required for assay depends on the cellularity of the lesion. For most solid tumors, three 50-µm sections should provide a sufficient number of nuclei for flow cytometry. For lymph nodes (either as controls or from lymphoma cases), one to two 50-µm sections are usually sufficient. Although some investigators have used 30-µm sections, most have found that a thicker cut, at 50-µm, reduces debris caused by cutting through nuclei and yields high-quality histograms. It is imperative that a pathologist review the corresponding hematoxylin-eosin stained slides, not only to ensure representative tissue on the section, but also to determine if selective areas of the block should be prepared for flow cytometry. For example, if tumor tissue is localized in one portion of the block and nonmalignant tissue in another, each portion should be processed separately. The pathologist can mark the stained slide (and the block, if possible) with a line or semicircle demarcating the malignant versus nonmalignant areas for cutting by the histotechnician. This is an excellent way to obtain malignant and nonmalignant areas and to provide a nonmalignant diploid control (Dressler and Bartow, 1989; Dressler and Seamer, 1994).
Critical Parameters and Troubleshooting Three main critical parameters need to be addressed: tissue representation, disaggregation, and cell yield (Dressler and Seamer, 1994; Shankey, et al., 1993). To a great extent, representation of appropriate cells (neoplastic or nonneoplastic) in a dissociation protocol reflects adequacy of sampling, although a variety of biologic factors may also interfere with tumor representation. These include admixture of
neoplastic and benign tissue in some lesions as well as admixture of preinvasive and invasive tissue in other lesions. In general, sampling is optimized if tissue is obtained from the areas of the “invasion front” (i.e., the advancing edge) of the tumor. For this reason, consistent sampling of fresh, frozen, or paraffin-embedded tissue requires an experienced technologist or pathologist. For some tumor systems, such as prostate carcinoma, it may be difficult to distinguish benign from malignant tissue. Further, for fresh or frozen tissue, the decision to sample tissue requires diagnostic evaluation in order to assess proximity to margins and neoplasm size. Experienced diagnostic evaluation is essential in any experimental protocol that employs unfixed human tissue samples. Fine needle aspiration will often selectively withdraw aneuploid cells, as the intercelluar bridges of aneuploid cells are often less intact than those of diploid cells. Disaggregation protocols sometimes result in excessive debris, usually observed as baseline elevation of the DNA histogram or as increased coefficient of variation. In general, fresh samples have less debris than fixed specimens, but this is not always the case. Fine needle aspirates may also contain excess debris. Debris may reflect inadequate filtering of the cellular suspension, presence of abundant mucin within the neoplasm, tissue autolysis prior to dissociation, or sampling from necrotic or hemorraghic areas with low cellularity. In paraffin-embedded material, debris may also result from slicing through nuclei and cells, especially in thin sections. Debris can be minimized by using a double-layer sucrose cushion (see Support Protocol 5); this technique can be applied to whole cells or nuclei, fresh or frozen, in addition to paraffin-embedded tissue. Refrigeration of tissue prior to dissociation is suggested to avoid tissue autolysis. Use of an isotonic growth medium also optimizes cellular preservation. Addition of 5% heat-inactivated fetal bovine serum (FBS) to the medium appears to enhance this preservation. For fresh or frozen samples, tissue scraping to release cells may be insufficiently vigorous to yield cells or so vigorous that cells are damaged. It is useful to examine the scalpel blade after scraping the tissue to look for the presence of adherent, slightly turbid fluid on the blade surface. Minimal observable fluid means the dissociation was probably not sufficiently vigorous, and a cytospin preparation will reveal large clusters or aggregates of cells. If there are fragments of tissue, then dissociation has likely been too
Specimen Handling, Storage, and Preparation
5.2.13 Current Protocols in Cytometry
vigorous and cell viability will decline noticeably. In either case, overall cell yield decreases. The third critical parameter is cell yield. Optimal specimens have high cellularity, so after processing, cell yield is sufficient and also representative of the original specimen. An inherent limitation to dissociation of fresh tissue specimens (as compared to dissociation of formalin-fixed paraffin-embedded tissue), is that histologic features of the tissue are not necessarily known at the time of sampling. Although a tissue imprint (see Support Protocol 1) is suggested for quality assurance of representative tissue, the cell yield of dissociation will still be a function of the volume of tissue dissociated, the inherent cellularity of the tissue or neoplasm, the ratio of neoplastic cells to benign stromal or inflammatory cells, and factors intrinsic to the dissociation protocol. It should be noted that there is considerable intercase variability in these parameters, even for a neoplasm of a given type. For example, some breast carcinomas are characterized by extensive areas of “desmoplastic” (scar-like) stromal reaction which contains relatively few neoplastic cells. Another type of breast carcinoma (medullary carcinoma) is characterized by a high percentage of tumor-infiltrating lymphocytes.
Anticipated Results
Typically, cell yield is ∼107 cells per milligram of tissue. Optimally, 58% of the dissociated cells are intact, as estimated by viability studies for fresh tissue dissociation. Approximately 20% to 80% of the cells from a tumor slice will be neoplastic, depending on the degree of host cell contribution. An evaluation of the degree of benign and/or nonneoplastic cell admixture as well as assessment of neoplastic cell morphology and representation is critical. As noted, tissue imprints, cytospin preparations, and paraffin block sections stained with Papanicolau or hemotoxylin-eosin are necessary for evaluation. The quality of the results obtained from these different dissociation techniques will largely depend on the quality and cellularity of the initial sample obtained for assay.
Time Considerations
Handling, Storage, and Preparation of Human Tissues
Disaggregation of solid tumors from fresh or frozen specimens, fixation, and cytospin preparations can be performed in a few hours. Formalin-fixed, paraffin-embedded tissue samples require 2 days for preparation: 2 to 3 hr are required on the first day for processing thin
sections and rehydration, and ∼3 hr on the second day for isolation of nuclei from the hydrated tissue sections (depending on the total number of samples that are processed). Twelve to eighteen samples is a reasonable number for manual preparation by one technician. For procuring fresh samples, the investigator should establish a coordinated protocol with the cooperating pathologist and/or tissue procurement person. The time required for tissue procurement is usually 5% dead cells (i.e., 90% for lymphocytes. Large lymphocytes, which are most important in disease, are frequently excluded from the analysis. Note that the large cells with low SS to the right of the dense cluster have been included in R9 because the lymphocyte gate (R9) circumscribes only the dense lymphocyte cluster. These cells are very important and should always be included even though they often reside under the monocyte cluster. R9, rather than R1, is used as the gate region because most analysis software applications assign colors in a hierarchical fashion. If R1 were the gate, it would not be possible to assign colors to the various cell populations resolved by the Boolean logical gates shown in Table 6.2.1. It is desirable to use multiparameter software that allows at least nine regions, as shown in Figures 6.2.1 and 6.2.2 and Table 6.2.1. Regions 1 to 8 are used for the quadrant markers and regions 9 and higher are used for gating. Some software applications attach a color based on a Venn spectrum to the gate logic, in a hierarchical order instead of allowing userselected colors for each population. In viewing multiparameter data, this software restriction is often undesirable because the color mixtures can duplicate the background color and the population will completely disappear from the view. The hierarchical nature associated with gating means the gate must always be last. Because of hierarchical color assignment, if the gate region is first, it will not be possible to assign colors to any other region. Thus, it must be last so that its color does not dominate. Color assignments for quadrant markers combined by Boolean equations must be assigned by the user and not the software.
Anticipated Results The most important single potential problem with results stems from the quality of the antibody. Refer to UNIT 4.1 for a more complete discussion of these problems. The objective of these protocols is to resolve epitope-positive cells from negative ones. To do so, there must be enough epitopes, and the antibody’s fluorochrome must be bright enough for the investigator’s instrument to measure. One may not be able to do anything about the number of epitopes but the fluorochrome and the instrument
Phenotypic Analysis
6.2.17 Current Protocols in Cytometry
are within the investigator’s power. Often, stream-in-air detection is less sensitive than stream-in-liquid (flow-cell cuvette) detection. Thus, the instrument plays a role in sensitivity. Because PE-containing fluorochromes are five to seven times brighter than the others, a strategy to use them for conjugation to antibodies that will be used to detect the cells with the least number of epitopes can markedly improve results. Finally, Fc-receptor binding, dead cells, and nonspecific binding affect both the result and its interpretation. Fc-receptor binding can be eliminated by appropriate blocking; dead cells can often, but not always be removed from analysis by appropriate gating. Nonspecific binding problems can be reduced by washing. When a cell population is poorly resolved by one antibody, it can often be well resolved by combining it with a second to pull the population away from other cells. Even if both antibodies are directed toward antigens in low abundance, their combined effect can be rewarding. This is one of the best reasons for performing multicolor flow cytometry. Investigators have been taught to expect that immunophenotyping by flow cytometry will resolve positive and negative cells. Clearly, if one desires to have a CD4 count or a CD34 count, this is what to expect. But the pathologist does not expect this result when using immunohistochemistry to identify cells in a tissue. When cell populations are to be identified, it is possible to combine antibodies that resolve different cells and their pattern in bivariate plots can be used to identify them. In the case of pathology, their pattern is the expected result, and deviations from the known pattern may indicate abnormal processes such as malignancy.
Time Considerations One should always set up the necessary tubes and reagents prior to getting cells. If 96 tubes (one full standard rack) are used as an example, this preparation should take ∼30 min. The time to prepare cells depends on where they come from; nevertheless, they should be obtained just before they are to be stained, when-
ever possible. Counting, washing, and blocking takes at most 15 min. Adding cells to the antibody cocktails takes another 5 min. For each block of steps, 20 min is required. Thus—in a procedure where staining with an unconjugated antibody (20 min) and subsequent staining with secondary antibody (20 min) is combined with a block (10 min) prior to adding a biotinylated antibody combined with directly conjugated antibodies (20 min) and finally adding fluorochrome-conjugated antibody or antibodies (20 min)—up to 90 min may be necessary for staining. For acquiring data on the instrument, the authors usually require 1 min per tube. This will vary depending on cell concentration and the number of events acquired. It is difficult to evaluate analysis time, because this depends on how much prior knowledge of the samples is available. If it is a routine evaluation where the analysis protocol is known, it may require only 1 to 2 min per sample. Thus, 96 samples can take as little as 4 hr to process from start to finish.
Literature Cited Riedy, M.C., Muirhead, K.A., Jensen, C.P., and Stewart, C.C. 1991. The use of a photolabeling technique to identify nonviable cells in fixed homogenous or heterologous cell populations. Cytometry 12:133-139.
Key References Stewart, C.C. and Stewart, S.J. 1994. Cell preparation for the identification of leukocytes. In Methods In Cell Biology (Z. Darzynkiewicz, J. Robinson, and H. Crissman, eds.) pp. 39-60. Academic Press, New York. Provides additional information on preparing cells. Stewart, C.C. and Stewart, S.J. 1994. Multiparameter analysis of leukocytes by flow cytometry. In Methods in Cell Biology (Z. Darzynkiewicz, J. Robinson, and H. Crissman, eds.) pp. 61-79. Academic Press, New York. Provides additional information on instrument setup, data acquisition, and data analysis.
Contributed by C.C. Stewart and S.J. Stewart Roswell Park Cancer Institute Buffalo, New York
Immunophenotyping
6.2.18 Current Protocols in Cytometry
High-Sensitivity Immunofluorescence/ Flow Cytometry: Detection of Cytokine Receptors and Other Low-Abundance Membrane Molecules
UNIT 6.3
Immunofluorescence-based flow cytometry, in common with other analytical techniques, has limited sensitivity. Thus, it is important to understand that when it is said that T cells do not express CD32, it is only known with certainty that CD32 is undetectable on T cells with the methods used. The detection limit, or sensitivity, depends on the reagents, staining, and instrument parameters. For the most commonly used procedures and reagents, the sensitivity is ∼2000 molecules of target antigen per cell (this parameter is different from the widely used “molecules of equivalent soluble fluorescein,” MESF). This level of sensitivity is adequate for most purposes. Molecules such as CD3, CD4, and CD8 are expressed at ∼20,000 to 100,000 molecules/cell on the appropriate T cell subsets. Most of the widely used leukocyte markers are expressed at similar levels on the cells of interest (Table 6.3.1). Molecules expressed at much lower concentrations are, however, biologically significant. Growth factors can activate cells through receptors expressed at very low concentrations (e.g., 1000 molecules or less; Table 6.3.1). The same may be true for molecules involved in intercellular interactions, such as B7 (CD80 and CD86) and its ligand CTLA-4 (CD152), receptors for Ig, or adhesion and homing molecules. For the example above, CD32 Table 6.3.1 Typical Concentration of a Selection of Widely Used Immunological Markers, Compared with Cytokine Receptors
Marker
Cell typea
Concentration (molecules/cell)
Reference
Commonly used markers CD2
T cells (blood)
40,000
Martin et al. (1983)
CD3
T cells (blood)
57,000
Bikoue et al. (1996)
CD4
T cell subset (blood)
47,000
Bikoue et al. (1996)
CD8
T cell subset (blood)
145,000
Bikoue et al. (1996)
CD5
T cells (blood)
50,000
Bikoue et al. (1996)
CD19
B cells (blood)
27,000
Bikoue et al. (1996)
CD45
Lymphocytes
sIg
Chronic lymphocytic leukemia
217,000 6,500-22,500
Bikoue et al. (1996) Dighiero et al. (1980)
Cytokine receptors CD121a
Lymphocytes
0.15
5
546 546 552
6
1 month).
8. Add 50 µl ice-cold pretitrated PE-SA to the cells and mix gently. Incubate and wash twice as above (see steps 3 to 5). Sigma PE-SA can be used at a 1/20 dilution, but a preliminary titration (see Support Protocol 1) is necessary for each new batch or after prolonged storage (>1 month).
9. Optional (for whole-blood samples): To lyse erythrocytes, add 2 ml lysing solution, vortex briefly, and allow to stand 10 min at room temperature in the dark. Centrifuge 5 min at 200 × g, and remove supernatant. Wash once with 1 ml DPBS/azide. 10. Resuspend cells thoroughly in 100 to 200 µl ice-cold DPBS/azide. If analysis is to be delayed until another day, resuspend in 200 µl flow cytometry fixative (Lanier and Warner, 1981) and store up to 7 days at 4◦ C.
Phenotypic Analysis
6.3.5 Current Protocols in Cytometry
Supplement 30
Analyze cells 11. Analyze by flow cytometry (see Chapter 1). Select cell population of interest by gating on forward and side scatter, and print out fluorescence-intensity histograms (see Fig. 6.3.1 for examples). ALTERNATE PROTOCOL
MULTICOLOR STAINING AND ANALYSIS In many situations, the expression of markers in a mixed cell population must be analyzed. While the cell subset of interest can first be purified and then examined by single-color analysis, a more elegant solution is to use multicolor analysis. One or more colors are used on cell-subset markers to identify the cell populations, and a different color is used to measure the expression of the marker of interest. The major factor in planning such a study is to reserve the highest-sensitivity label (phycoerythrin) for the low-abundance marker. A number of directly labeled cell-specific reagents are commercially available. Fluorescein- or PE/cyanine 5 (Cy5)–conjugated monoclonal antibodies with specificities for lymphocyte subsets (e.g., anti-CD3, anti-CD4, anti-CD8, anti-CD19) and other cell populations are available in good quality from various suppliers (e.g., Caltag Labs, BD/PharMingen, Beckman Coulter, Sigma). Becton Dickinson Immunocytometry supplies subset markers labeled with peridin chlorophyll protein (PerCP). PerCP and PE/Cy5 energy transfer reagents (e.g., Tricolor from Caltag Labs, CyChrome from BD/PharMingen, PE/Cy5 from Beckman Coulter, or Quantum Red from Sigma) are better than PE/Texas Red combinations, because there is less spectral overlap with PE. Nevertheless, electronic compensation for fluorescence spectral overlap must be set carefully to avoid reducing sensitivity (UNIT 6.14). Recently, allophycocyanin (APC)–MAb reagents have also become available from several manufacturers. These can be used once the flow cytometer is equipped with a red laser. The range of antibodies conjugated to the Alexa dyes is at present limited, but increasing.
Additional Materials (also see Basic Protocol) Normal mouse serum or 1 mg/ml normal mouse Ig (Sigma) in DPBS/azide (store refrigerated), ice cold Pretitrated, direct fluorochrome-conjugated, cell-specific MAb(s) (see introduction; see Support Protocol 1 for titration) 1. Follow steps 1 to 7 of Basic Protocol. 2. Add 50 µl ice-cold normal mouse serum or 50 µl of 1 mg/ml normal mouse Ig and incubate 10 min on melting ice. The purpose of this step is to block sites on the biotinylated anti–mouse Ig that may bind subsequent monoclonal antibodies.
3. Add 5 µl ice-cold pretitrated, direct fluorochrome-conjugated, cell-specific MAb(s) and 50 µl PE-SA to the cells and mix gently. Incubate and wash twice (see Basic Protocol, steps 3 to 5). Cell-specific MAbs and PE-SA can be added together. Sigma PE-SA can be used at a 1/20 dilution, but a preliminary titration (see Support Protocol 1) is necessary for each new batch or after prolonged storage (>1 month). Streptavidin labeled with PE/Cy5 can be used in place of PE-SA (see Strategic Planning; see Critical Parameters). High-Sensitivity Detection of Low-Abundance Membrane Molecules
4. Perform steps 9 (optional) and 10 (see Basic Protocol). 5. Analyze by flow cytometry (see Chapter 1). Select cell population of interest by gating on forward and side scatter, and print out two-color dot diagrams. Alternatively, gate
6.3.6 Supplement 30
Current Protocols in Cytometry
Figure 6.3.3 Two-color staining and analysis. High sensitivity staining of Fas (CD95) and analysis of expression on B cells (CD19), T cells (CD3), and T cell subpopulations (CD4 and CD8). Neg, negative control.
on dual scatter and one or two fluorescence parameters, and print out fluorescence intensity histograms (see Fig. 6.3.3 for examples).
TITRATION AND QUALITY CONTROL OF MONOCLONAL ANTIBODIES AND DETECTION REAGENTS
SUPPORT PROTOCOL 1
The primary monoclonal antibody (MAb) and the detection reagents must be titrated to determine optimal concentrations. This is done by performing staining and analysis with varied concentrations of MAb and detection reagents. Thus, reagents and specific methods are as described (see Basic Protocol). Modifications for titration are described below. Note that the use of an isotype-matched negative control is an essential but not necessarily adequate control, because one antibody can give higher nonspecific binding than another for reasons that are difficult to control, such as aggregation or partial denaturation during purification or storage. Phenotypic Analysis
6.3.7 Current Protocols in Cytometry
Supplement 30
MAb Titration Prepare cells (see Basic Protocol, step 1). Make dilutions of primary monoclonal antibody in DPBS/azide. As a guide, start with ∼5 µg/ml if the antibody concentration is known. For unpurified ascitic fluid, a dilution of 1/100 is a reasonable starting point. When using culture supernatant as antibody, undiluted supernatant is likely to be adequate. Use three to five doubling dilutions to cover the likely range of antibody concentrations. Add cell suspension to tubes (see Basic Protocol, step 3) before adding MAb, then add 50 µl MAb/tube directly to the cell suspension, mix gently, and proceed with the incubation and remaining procedure (steps 3 to 11). Select the optimal concentration on the basis of the best separation between positive and negative populations when the antibody gives a bimodal distribution (Figs. 6.3.1 and 6.3.2). When the antibody does not yield a bimodal distribution, select on the basis of the best discrimination between the antibody and an equivalent concentration of an isotype-matched negative control antibody.
Titration of Detection Reagents To test and titrate detection reagents, use three sets of tubes: a known strong antibody that will resolve positive and negative populations (e.g., CD3), a known antibody that produces low-level staining with a bimodal distribution (e.g., CD25), and a known negative control. While the test can be done as a checkerboard titration to determine optimal concentrations of both the biotinylated anti–mouse Ig and the PE-SA, it is more efficient to vary one reagent at a time. Prepare cells and perform MAb labeling (see Basic Protocol, steps 1 to 6). Make dilutions of biotinylated anti–mouse Ig and PE-SA in DPBS/azide, using the manufacturer’s recommendation as a starting point and adding 2-fold higher and lower concentrations. Use these dilutions to perform indirect labeling and analysis (see Basic Protocol, steps 7 to 11). A typical set of results is shown for human peripheral blood lymphocytes (PBLs) in Figure 6.3.1. This type of analysis provides an important quality control tool, to compare reagent batches, to check reagent activity after storage, and to check instrument performance. The CD3 pattern gives a measurement of overall sensitivity (the separation between the positive and negative peaks), while comparison of CD25 with the negative control provides confirmation that the assay is operating at the necessary level of sensitivity. The CD25 pattern should give a clear shoulder of positive cells. SUPPORT PROTOCOL 2
High-Sensitivity Detection of Low-Abundance Membrane Molecules
CELL SORTING WITH HIGH SENSITIVITY Staining protocols for high-sensitivity analysis can be translated directly to sorting applications, and modern sorters are capable of sensitivity equivalent to that of analytical cytometers. In principle, therefore, there are no difficulties associated with translating a high-sensitivity analytical application to preparative use. In practice, there are a number of variables that can cause difficulty. The first is the concentration of reagents used in staining. Because sorting applications normally involve staining a relatively large number of cells (typically 107 or more, compared to 0.5 × 106 generally used for analysis) there is a temptation to avoid using the same ratio of reagent to cells, to economize on antibodies. However, because this can lead to lower sensitivity, it is important to titrate the reagents to determine the optimum concentrations (see Support Protocol 1). A second potential source of reduced sensitivity derives from the relatively long sorting times, compared with analysis. Antibody may dissociate with time, particularly at higher temperatures, and it may be essential to cool the sample tube during sorting. Stream-in-air optical systems sacrifice some signal due to the air interfaces, and quartz cuvettes are better for sensitivity. Quartz cuvettes are used almost universally for analytical
6.3.8 Supplement 30
Current Protocols in Cytometry
instruments but not on all sorters, and stream-in-air optics are universally used for highspeed sorting. The best approach is to optimize sensitivity on an analyzer, then translate the application as far as possible to the sorter. If sensitivity is not adequate, contact the manufacturer of the sorter for advice—the major companies are aware of the sensitivity issue and can provide help with instrument optimization.
COMMENTARY Background Information Cytokine receptors are complex structures composed of two or more protein chains. One protein confers cytokine-binding specificity while a second protein, which may be shared by several other cytokines, increases binding affinity and mediates signal transduction (Taga and Kishimoto, 1992; Noguchi et al., 1993). Cytokine receptors can be expressed on activated cells at very high levels (>30,000 molecules/cell for CD25), but they are usually expressed at much lower concentrations (100 to 500 molecules) in cells taken directly from blood (Zola et al., 1990, 1995; Zola, 1995). Monoclonal antibodies are available against many cytokine receptor proteins (Schlossman et al., 1995; Zola, 1995; Kishimoto et al., 1997; Mason et al., 2002). Receptor expression may be analyzed using the labeled cytokine as ligand. This approach may give a different result from analysis using monoclonal antibody, since the cytokine will usually bind to the multichain receptor complex, while the antibody detects individual proteins. Other molecules that may be expressed and functional at low concentrations include Fc receptors (Mantzioris et al., 1993) and costimulatory molecules such as B7 and CTLA-4. The technique described in this protocol is also useful for detection of transfectants expressing low levels of transgene, and for detection of low-affinity binding.
Critical Parameters Maintain cells and reagents cold throughout the procedure to reduce loss of antigen and dissociation of the antibody-antigen complex. All reagents should be stored refrigerated, and the reaction tubes should be in contact with melting ice. Centrifugation should be carried out at 4◦ C. Blocking of Fc receptors (see Basic Protocol step 6), however, is performed at room temperature to increase the rate of binding. After centrifugation remove supernatant thoroughly, carefully, and reproducibly. If the amount of supernatant left behind after washing varies from tube to tube, the next
reagent will be diluted to a variable degree, introducing variability in staining. Resuspend cells after each centrifugation, to ensure a single-cell suspension. Careful use of a vortex mixer is recommended. Excessive mixing may denature protein reagents. Although the recommended protocols specify two washes at each stage, it may be worthwhile experimenting with fewer washes, particularly with low-affinity antibodies. Sensitivity As has been emphasized (see Strategic Planning), phycoerythrin (PE) is the fluorochrome that gives the highest sensitivity. The use of a three-step staining protocol provides amplification by increasing the number of fluorochrome molecules per antigen epitope. Different staining reagents that are equivalent in principle may differ greatly in sensitivity, and reagents may lose activity on storage. Titration and quality control are therefore essential. Instrument parameters can also have a major influence on sensitivity (see Strategic Planning). Sensitivity is not improved simply by increasing the signal strength, as can be demonstrated very simply by turning up the photomultiplier voltage. Nonspecific (background) staining levels must be minimized to achieve sensitivity. The major sources of background staining are autofluorescence, cross-reactivity of anti-Ig reagents with cell membrane Ig, and binding of antibodies to Fc receptors or other membrane components. Strategies for reducing Fc-mediated staining have been described in the protocols. Cross-reactivity of anti–mouse Ig with human Ig may be removed by absorption on immobilized human Ig. Include controls to determine the major contributors in a particular situation, and then take steps to minimize the major contributors. Effect of fluorescence compensation on sensitivity Electronic compensation must be carried out carefully (UNITS 1.3 & 1.14). Some loss of sensitivity is generally experienced in multicolor compared with single-color analysis. The
Phenotypic Analysis
6.3.9 Current Protocols in Cytometry
Supplement 30
emission spectrum of fluorescein has a substantial “tail” that overlaps the spectral window used to collect PE signals. Electronic correction for this involves some reduction in the PE signal; if the compensation is perfect the reduction will exactly match the signal from fluorescein and will not reduce the genuine PE signal, but in practice it is difficult to achieve this precisely. In principle, digital fluorescence compensation, available on some instruments, should be more precise. The spectral overlap between PE and PE/Texas Red is also considerable, but that between PE and PE/Cy5 is small. This combination should therefore be chosen where possible, but PE/Cy5 conjugates need to be checked to see if they emit in the PE spectral window, which can happen if there is
High-Sensitivity Detection of Low-Abundance Membrane Molecules
any free PE. To avoid spectral overlap, use the 488-nm laser to excite FITC while using a 532-nm dye-pumped solid-state (DPSS) laser to excite PE. Although each dye will be excited by both lasers, the excitation of FITC at 532 nm will be very low. The signals have to be measured separately and then combined for correlated dual-color analysis, which can be achieved by separating the laser/stream intersection points and using gated amplifiers. Simultaneous analysis of two low-abundance molecules If two low-abundance molecules are to be measured together, use PE and PE/Cy5 as the fluorochromes, because they give the strongest signals and require minimal compensation (but
Figure 6.3.4 Three-color staining and analysis, including two high-sensitivity colors. Cells were stained directly with FITC-conjugated anti-CD4 antibody and PE-conjugated anti-Fas (CD95) antibody (DX-2; BD/PharMingen), and indirectly with anti-CD25 detected with biotinylated horse anti-mouse (Vector Labs) and streptavidin-PE/Cy5 (Sigma). The bottom panel shows CD4 cells with a major population coexpressing CD25 and Fas (results from Ms. Natasha Wuttke).
6.3.10 Supplement 30
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see note about checking PE emission from PE/Cy5, in above discussion on effect of fluorescence compensation). Alternatively, combine PE with one of the Alexa dyes, or use two different Alexa conjugates. The biotin-avidin amplification system can be used for only one of the markers, and is best reserved for the weaker one. A second indirect system can be used, such as that based on digoxigenin, but the availability of reagents usually means that one of the markers will have to be detected with a direct conjugate. In this situation it is worth exploring alternative reagents within the same specificity; for example, Becton Dickinson Immunocytometry’s CD25-PE conjugate and BD/PharMingen’s CD95-PE conjugate give exceptionally good staining.
Troubleshooting Loss of sensitivity is probably due to deterioration of one of the reagents, and they should be retitrated (see Support Protocol 1). A high background signal, usually in the form of a “tail” to the negative population in the nega-
tive control, is probably due to Fc-mediated uptake or cross-reactivity (see Critical Parameters). Identify the cause of the problem and remedy as described in the protocols. Not infrequently, a batch of negative control antibody will start showing nonspecific binding that had not been seen previously. This may result from aggregation or other denaturation and is best remedied by using a new batch.
Anticipated Results Cytokine receptors are still commonly referred to in the literature as absent from cells that have not been activated in vitro. As a result, many immunologists would anticipate negative results with cells taken directly from tissue. However, as may be seen in Figures 6.3.1, 6.3.2, 6.3.3, and 6.3.4 and in a number of publications (Zola and Flego, 1992; Zola et al., 1993a,b), optimization for low-abundance antigens has shown that a number of receptors clearly stain subsets of lymphocytes. Furthermore, the level of staining may have clinical significance (Zola et al.,
Figure 6.3.5 Application of high-sensitivity fluorescence outside the cytokine receptors area: detection of low-avidity, single-chain antibody fragment. Rat thymocyte population was stained with negative control antibody (top panels), scFv (single chain variable fragments) derived from the anti-CD4 antibody (middle panels), and with anti-CD4 antibody (lower panels). While the highsensitivity method did not provide any advantage for staining with whole antibody, it showed clear staining with scFv, which could not be discerned clearly using conventional staining reagents (results from Dr. Michael Thiel). MFI, mean fluorescence intensity.
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1994; Borvak et al., 1995). The value of this method is not restricted to cytokine receptors. It has been used to demonstrate CD32 on T cells (Mantzioris et al., 1993), to detect lowlevel expression of transfected genes, and to detect binding of relatively low-affinity, genetically engineered antibody fragments (Nicholson et al., 1997; Fig. 6.3.5). The measures described above to reduce nonspecific staining usually work well for T cells and less well for B cells, while monocytes tend to produce some nonspecific staining that is difficult to remove entirely, resulting in a small “tail” of positive cells in the negative control.
Time Considerations Allow 1 hr for cell preparation. During cell preparation, label reaction tubes, place in rack in melting ice, and aliquot monoclonal antibodies ready for use. Allow 2.5 hr for the performance of the test, once the cells are ready. The whole procedure, starting from a blood or tissue sample, therefore takes 3.5 hr, excluding flow cytometry (see Chapter 5 for time considerations). The three protocols (single-color, multicolor, and titration) are roughly similar with respect to time. The size of the test (number of samples) will affect the time and may lead to significant variations in incubation periods. The author generally limits a test to 50 to 60 samples. The protocols are all readily performed by one person.
Literature Cited Ben Aribia, M.H., Moire, N., Metivier, D., Vaquero, C., Lantz, O., Olive, D., Charpentier, B., and Senik, A. 1989. IL-2 receptors on circulating natural killer cells and T lymphocytes. Similarity in number and affinity but difference in transmission of the proliferation signal. J. Immunol. 142:490-499. Bikoue, A., George, F., Poncelet, P., Mutin, M., Janossy, G., and Sampol, J. 1996. Quantitative analysis of leukocyte membrane antigen expression: Normal adult values. Cytometry 26:137147. Borvak, J., Chou, C.-S., Bell, K., van Dyke, G., Zola, H., Ramilo, O., and Vitetta, E.S. 1995. Expression of CD25 defines peripheral blood mononuclear cells with productive versus latent HIV infection. J. Immunol. 155:3196-3204.
High-Sensitivity Detection of Low-Abundance Membrane Molecules
Dighiero, G., Bodega, E., Mayzner, R., and Binet, J.L. 1980. Individual cell-by-cell quantitation of lymphocyte surface membrane Ig in normal and CLL lymphocytes and during ontogeny of mouse B lymphocytes by immunoperoxidase assay. Blood 55:93-100. Dower, S.K., Kronheim, S.R., March, C.J., Conlon, P.J., Hopp, T.P., Gillis, S., and Urdal, D.L. 1985.
Detection and characterization of high affinity plasma membrane receptors for human interleukin 1. J. Exp. Med. 162:501-515. Kishimoto, T. 1989. The biology of interleukin-6. Blood 74:1-10. Kishimoto, T., Kikutani, H., von dem Borne, A.E.G., Goyert, S.M., Mason, D.Y., Miyasaka, M., Moretta, L., Okumura, K., Shaw, S., Springer, T.A., Sugamura, K., and Zola, H. 1997. Leucocyte Typing VI: White cell differentiation antigens. Garland Publishing, New York. Lanier, L.L. and Warner, N.L. 1981. Paraformaldehyde fixation of hematopoietic cells for quantitative flow cytometry (FACS) analysis. J. Immunol. Methods 47:25-30. Le Mauff, B., Gascan, H., Olive, D., Moreau, J.F., Mawas, C., Soulillou, J.P., and Jacques, Y. 1987. Parameters of interaction of a novel monoclonal antibody (33B3.1) with the human IL2-receptors: Interrelationship between 33B3.1, anti-Tac, and IL2 binding sites. Hum. Immunol. 19:53-68. Lidke, D.S., Nagy, P., Heintzmann, R., Arndt-Jovin, D., Post, J.N., Grecco, H.E., Jares-Erijman, E.A., and Jovin, T.M. 2004. Quantum dot ligands provide new insights into erbB/HER receptormediated transduction. Nat. Biotech. 22:198203. Lowenthal, J.W., Castle, B.E., Christiansen, J., Schreurs, J., Rennick, D., Arai, N., Hoy, P., Takebe, Y., and Howard, M. 1988. Expression of high affinity receptors for murine interleukin 4 (BSF-1) on hemopoietic and nonhemopoietic cells. J. Immunol. 140:456-464. Mantzioris, B.X., Berger, M.F., Sewell, W., and Zola, H. 1993. Expression of the Fc receptor for IgG (FcgammaRII/CDw32) by human circulating T and B lymphocytes. J. Immunol. 150:51755184. Martin, P.J., Longton, G., Ledbetter, J.A., Newman, W., Braun, M.P., Beatty, P.G., and Hansen, J.A. 1983. Identification and functional characterization of two distinct epitopes on the human T cell surface protein Tp50. J. Immunol. 131:180185. Mason, D.Y., Andr´e, P., Bensussan, A., Buckley, C., Civin, C., Clark, E.A., de Haas, M., Goyert, S., Hadam, M., Hart, D., Horejs´ı, V., Meuer, S., Morrissey, J., Schwartz-Albiez, R., Shaw, S., Simmons, D., Uguccioni, M., van der Schoot, E., Vivier, E., and Zola, H. 2002. CD antigens 2001. Blood. 99:3877-80. Mavrangelos, C., Swart, B., Nobbs, S., Nicholson, I.C., Macardle, P.J., and Zola, H. 2004. Detection of low-abundance membrane markers by immunofluorescence: A comparison of alternative high-sensitivity methods and reagents. J. Immunol. Methods 289:169-78. Nicholson, I.C., Lenton, K.A., Little, D.J., DeCorso, T., Lee, F.T., Scott, A.M., Zola, H., and Hohmann, A.W. 1997. Construction and characterization of a functional single chain Fv fragment for immunotherapy of B lineage leukemia and lymphoma. Mol. Immunol. 34:11571165.
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Noguchi, M., Nakamura, Y., Russell, S.M., Ziegler, S.F., Tsang, M., Cao, X., and Leonard, W.J. 1993. Interleukin-2 receptor gamma chain: A functional component of the interleukin-7 receptor. Science 262:1877-1880. Schlossman, S.F., Boumsell, L., Gilks, W., Harlan, J.M., Kishimoto, T., Morimoto, C., Ritz, J., Shaw, S., Silverstein, R., Springer, T.A., Tedder, T.F., and Todd, R.F. 1995. Leucocyte Typing V: White cell differentiation antigens. Oxford University Press, Oxford. Taga, T. and Kishimoto, T. 1992. Cytokine receptors and signal transduction. FASEB J. 6:3387-3396. Zola, H. 1995. Use of flow cytometry to detect cytokine receptors. In Current Protocols in Immunology (R. Coico, J.E. Coligan, E.M. Shevach, D.H. Margulies, W. Strober, and A.M. Kruisbeek, eds.) pp. 6.21.1-6.21.24. John Wiley & Sons, New York. Zola, H. and Flego, L. 1992. Expression of interleukin-6 receptor on blood lymphocytes without in vitro activation. Immunology 76:338340. Zola, H., Flego, L., and Weedon, H. 1993a. Expression of membrane receptor for tumor necrosis factor on human blood lymphocytes. Immunol. Cell Biol. 71:281-288.
Zola, H., Neoh, S.H., Mantzioris, B.X., Webster, J., and Loughnan, M.S. 1990. Detection by immunofluorescence of surface molecules present in low copy numbers. High sensitivity staining and calibration of flow cytometer. J. Immunol. Methods 135:247-255. Zola, H., Siderius, N., Flego, L., Beckman Coulter, I.G.R., and Seshadri, R. 1994. Cytokine receptor expression in leukemic cells. Leuk. Res. 18:6573. Zola, H., Fusco, M., Macardle, P.J., Flego, L., and Roberton, D. 1995. Expression of cytokine receptors by human cord blood lymphocytes— comparison with adult lymphocytes. Pediatr. Res. 38:397-403.
Key Reference Zola et al., 1990. See above. This paper describes the high-sensitivity immunofluorescence procedure in detail and shows typical results.
Contributed by Heddy Zola Child Health Research Institute North Adelaide, Australia
Zola, H., Flego, L., and Weedon, H. 1993b. Expression of IL-4 receptor on human T and B lymphocytes. Cell Immunol. 150:149-158.
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Enumeration of CD34+ Hematopoietic Stem and Progenitor Cells
UNIT 6.4
Hematopoietic progenitor cells (HPCs) can be mobilized from the bone marrow into the peripheral blood by cytotoxic drugs, cytokines, or combinations of the two. In the majority of cases, this will allow these primitive cells to be collected by apheresis in sufficient quantities for transplantation procedures. CD34 is the first documented cell-surface antigen whose expression within the hematopoietic system is restricted to stem and progenitor cells of all lineages. Thus, CD34+ HPCs can restore multilineage hematopoiesis in myelo-ablated patients. Flow cytometric enumeration of CD34+ cells per kilogram of recipient body weight has been shown to be the most useful indicator of the hematopoietic reconstitutive capacity of peripheral blood stem cell (PBSC) transplants. As flow cytometry can produce results within an hour, serial assessments of CD34+ cell concentrations in peripheral blood also enable the optimal timing of the apheresis sessions, ensuring the harvest of sufficient numbers of CD34+ cells. As CD34+ cells occur only in low frequencies in peripheral blood and apheresis products following their mobilization (i.e., typically between 0.1% and 2%), accurate enumeration of CD34+ cells represents rare event analysis and it is not uncommon for the number of specifically stained CD34+ cells to be outnumbered by nonspecifically stained events. The Basic Protocol is a modified version of the protocol developed by Sutherland et al. (1994) that was subsequently incorporated into a set of clinical guidelines for the International Society for Hematotherapy and Graft Engineering (ISHAGE; Sutherland et al., 1996a). It features counterstaining of CD34 by the CD45 monoclonal antibody (MAb), allowing the identification of leukocytes (CD45+) and the verification of “true” CD34+ cells as being dim for CD45 fluorescence and having low side scatter (CD45dim, SSlow). This flexible approach to CD34+ cell enumeration can be applied to the widest range of specimens, including those most commonly used for transplantation (e.g., apheresis products, bone marrow, and cord blood). The addition of predefined numbers of counting beads to the cell suspension yields the concentration of CD34+ cells per unit of sample volume (i.e., the absolute CD34+ cell count) from a single flow-cytometric assessment (single-platform technique). Dead cells can be excluded from analysis using the DNA stain 7-aminoactinomycin D (7-AAD). Support Protocol 1 allows the simultaneous enumeration of both CD34+ cells and CD3+ T cells. An ability to accurately enumerate the latter can be particularly useful in the allogeneic transplant setting, where reducing the number of CD3+ cells in the graft has been shown to result in decreased severity of graft-versus-host disease (GVHD). In this modification, CD3 is added (in place of 7-AAD) as a third antibody conjugate. The modified assay can also be performed in the presence of counting beads, allowing the generation of absolute CD3+ and CD34+ cell counts from a single assay tube. Support Protocol 2 addresses the further immunophenotypic characterization of CD34+ cells. This assay is of use in studies to identify more precisely which CD34+ cell subsets may have the highest relevance for prediction of engraftment. ENUMERATION OF ABSOLUTE NUMBERS OF CD34+ CELLS Counterstaining CD34 with CD45 MAb allows the elimination of debris and nonspecifically stained events from the analysis, as well as the generation of a reliable denominator against which to measure CD34+ cells (i.e., leukocytes). Importantly, this approach also allows the discrimination of HPCs (which express relatively low levels of CD45 on their surface) from lymphocytes and monocytes (which express high levels). Just as lymphoContributed by D. Robert Sutherland, Michael Keeney, and Jan W. Gratama Current Protocols in Cytometry (2003) 6.4.1-6.4.23 Copyright © 2003 by John Wiley & Sons, Inc.
BASIC PROTOCOL
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cytes, monocytes, and granulocytes form discrete clusters on bivariate plots of CD45 versus side scatter (SS), so do nonmalignant CD34+ hematopoietic stem and precursor cells. Thus, in the Basic Protocol presented here, a multiparameter flow methodology is applied that utilizes the maximum information available of four parameters—i.e., forward scatter (FS), side scatter (SS), CD34 expression, and intensity of CD45 expression. These four parameters are combined in a sequential Boolean gating strategy (UNIT 10.4) that can be used on a variety of clinical samples such as peripheral blood, apheresis products, purified CD34+ cell suspensions, cord blood, and bone marrow specimens. To enumerate absolute numbers of CD34+ cells (i.e., the concentration per unit of volume) by a single flow cytometric measurement (single-platform assay), an accurately measured volume of sample is pipetted into a tube and the same accurately measured volume of counting beads is added. This procedure establishes a ratio of counting beads to volume of sample. The known number of added counting beads allows direct calculation of the absolute CD34+ cell count. To avoid the loss of cell, and/or counting beads, lyse/no-wash sample processing is employed. These modifications convert the ISHAGE Protocol into a single-platform assay (Keeney et al., 1998a). Optional exclusion of dead cells from CD34+ cell enumeration can be achieved by inclusion of the DNA G-C intercalating dye 7-AAD, which allows discrimination between viable and nonviable (CD34+) cells. This modification is simple to apply and may be of value in all sample types. It is highly recommended for apheresis samples over 4 hr old, all “fresh” cord blood and marrow samples, or any samples that have been manipulated in any manner, and is essential for the accurate analysis of post-thawed samples, regardless of source (see below). 7-AAD is excited at 488 nm and has maximum emission at 660 nm. Consequently, the dye cannot be used in single-laser systems with other fluorochromes that emit at >600 nm, such as PerCP or PE-Cy5. See UNIT 9.2 for a discussion on compensation for spectral overlap between PE and 7-AAD. This procedure requires reverse pipetting of the sample and counting beads (see Critical Parameters and Troubleshooting). Additionally, this protocol uses terminology and gating parameters that are unique to specific instruments, i.e., Beckman Coulter XL and BD Biosciences FACS series instruments (see Fig. 6.4.1 and Fig. 6.4.2, respectively). However, the procedure can certainly be adapted for other flow cytometers. Materials Sample of interest: peripheral blood, apheresis, cord blood, bone marrow, selected CD34+ cells, or cryopreserved and thawed samples Phycoerythrin-conjugated CD34 monoclonal antibody (CD34-PE MAb), appropriately titered (UNIT 4.1) Fluorescein isothiocyanate–conjugated CD45 monoclonal antibody (CD45-FITC MAb), appropriately titered (UNIT 4.1) 1× ammonium chloride lysing solution (APPENDIX 2A) Phosphate-buffered saline (PBS; APPENDIX 2A) supplemented with 1% human serum albumen (or similar; PBS/HSA) Flow-Count counting beads (Beckman Coulter) 12 × 75–mm polypropylene tubes Flow cytometer with at least three fluorescence detectors and appropriate filter sets for detection of FITC, PE, and if required, 7-AAD Enumeration of CD34+ Hematopoietic Stem and Progenitor Cells
Additional reagents and equipment for assessing leukocyte count (APPENDIX 3A)
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Figure 6.4.1 Example of the Basic Protocol combined with optional dead-cell exclusion for a Beckman Coulter XL flow cytometer equipped with four fluorescence detectors. Data acquisition was done using System 2 Software and analysis using Expo software. Data are shown from a prediluted apheresis sample containing 16% dead cells (i.e., 7-AAD+) and 300 viable CD34+ cells/µl. In this example, 180 viable CD34+ cells were gated in region D and 6754 singlet beads were gated in region H. The assayed Flow-Count bead concentration was 1046/µl and the cell dilution factor was 1/10.
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Figure 6.4.2 Example of the Basic Protocol combined with optional dead-cell exclusion for a BD Biosciences FACScan flow cytometer. Data were acquired and analyzed using CellQuest version 3.3 software. Data are shown from apheresis collection, stored overnight at 4°C after dilution with autologous plasma. Sample (diluted 1/10 before staining) contained ∼11% dead cells (i.e., 7-AAD+ gated in R8) and 117 viable CD34+ cells/µl. In this example, 456 viable CD34+ cells were identified in gate 4 and 4079 singlet beads were enumerated in gate 7. The assayed bead concentration was 1046/µl.
Enumeration of CD34+ Hematopoietic Stem and Progenitor Cells
Prepare sample 1. Assess the leukocyte count (APPENDIX 3A) of blood and apheresis samples to determine whether or not dilution of the sample is necessary. Dilute the sample with PBS/HSA so that the leukocyte count is ≤20 × 109/liter in a volume of 0.5 ml. 2. For each sample, label three 12 × 75–mm polypropylene tubes: two for CD34 and one for PBS.
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3. Mix diluted sample well and add 100 µl to each of the tubes labeled CD34. Add 2 ml PBS to the PBS tube. 4. Add 10 µl CD45-FITC MAb and 10 µl CD34-PE MAb to each of the two CD34 tubes. Gently mix. 5. Incubate tubes 15 min at room temperature, protected from light. 6. Add 2 ml of 1× ammonium chloride lysing solution at room temperature to the CD34 tubes. Vortex gently. Incubate 10 to 15 min at room temperature, protected from light. Incubation for 10 min is usually sufficient to lyse erythrocytes in peripheral blood samples and apheresis products. Incubation for 15 min is recommended for cord blood samples because of the higher concentration and greater resistance to lysis of the erythrocytes. If erythrocytes are only a minor population (e.g., apheresis products or enriched CD34+ cells), lysis is not necessary and samples can simply be resuspended in 2 ml PBS/HSA. Loss of counting beads may occur when the analysis is performed on undiluted apheresis samples using polystyrene tubes in the absence of protein (Brando et al., 2001). Optional (for dead cell exclusion): Before adding lysing solution, prepare a fresh 100 ìg/ml working solution of 7-AAD by diluting a 1 mg/ml stock solution (UNIT 9.2) with PBS. Add 20 ìl working solution to 2 ml ammonium chloride lysing solution and then add this 7-AAD/lysing solution to each tube. Vortex gently and incubate 15 min at room temperature protected from light.
CAUTION: 7-AAD is a suspected carcinogen and should be handled with care. In particular, wear gloves when weighing out the dye and do not inhale dust. 7. Immediately prior to use, resuspend Flow-Count beads gently but thoroughly. This can be achieved manually by end-over-end rotation to avoid the generation of air bubbles. Do not vortex. Add 100 µl Flow-Count beads to the CD34 tubes with the same pipet used for sample dispensing and gently mix to distribute the beads evenly. If necessary Flow-Count beads should be properly resuspended before use by vortexing, especially if they have been sitting for 12 hr or more. This is best performed at least 2 hr prior to use so that any air bubbles inadvertently generated during this process will not be present when beads are pipetted into the sample. Samples without 7-AAD can be analyzed immediately or kept on melting ice for a maximum of 1 hr. Samples should be gently mixed immediately prior to analysis. Samples with 7-AAD should be analyzed immediately.
Set up instrument 8. Position FS, SS, and fluorescence (FL) windows of analysis and adjust electronic correction for spectral overlap according to standard procedures for lyse/no-wash immunophenotyping assays (UNIT 1.3). Flow-Count beads are detected on FACScan and FACSCalibur instruments at lower FS levels than lymphocytes; thus, the FS threshold must be lowered so that singlet beads are not excluded from acquisition (see Fig. 6.4.2 histogram 7).
9. Create a total of seven or eight bivariate histograms (dot plots) as follows: a. b. c. d. e.
Histogram 1: CD45-FITC (green fluorescence) versus SS Histogram 2: CD34-PE (orange-red fluorescence) versus SS Histogram 3: CD45-FITC (green fluorescence) versus SS Histogram 4: FS versus SS Histogram 5: CD45-FITC (green fluorescence) versus CD34-PE (orange-red fluorescence)
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f. Histogram 6: FS versus SS g. Histogram 7: Time versus counting-bead fluorescence (red fluorescence; for Beckman Coulter XL) or time versus FS (for BD Biosciences FACS) Optional (for dead cell exclusion): For Beckman Coulter XL instruments, acquire histogram 8 as 7-AAD (far-red fluorescence) versus SS. On Beckman Coulter XL instruments with three fluorescence detectors, replace the 620-nm red fluorescence band-pass filter with a 675-nm band-pass filter for optimal detection of 7-AAD. For BD Biosciences FACS instruments, acquire histogram 8 as 7-AAD (far-red fluorescence) versus SS.
Create gating regions In the following steps, specific terminology for regions and gates are given parenthetically for different instruments. For example, region A is for Beckman Coulter XL instruments, and region R1 and gate G1 (see below) are for BD Biosciences FACS instruments. See Figures 6.4.1 and 6.4.2. 10. Histogram 1 (leukocyte gate): Display all events. Draw a rectangular gate (A; R1) to include all CD45dim to CD45bright events and to exclude debris, platelets, and unlysed erythrocytes, which are all CD45neg. For Beckman Coulter XL instruments: By ensuring that the right edge of region A does not extend further than the brightest CD45+ cells, the Flow-Count beads (in the top right-hand corner of histogram 1) can be excluded from region A. For BD Biosciences FACS instruments: The counting beads are found in the brightest green, orange-red, and far-red channels and also exhibit very high side scatter. Since the gate statistics (Fig. 6.4.2) are obtained from events gated in R1 (displayed on histogram 2), ensure that all the CD45+ events as well as all the counting beads are gated in R1 by including the highest green fluorescence and SS channels.
11. Histogram 2 (total CD34+ gate): Display events from the leukocyte gate (i.e., gated on A; gate G1 = R1). Draw an amorphous polygon (nonrectangular) region (B; R2) to include all CD34+ events. 12. Histogram 3 (CD34+ blast gate): Display events that fulfill the criteria of both above gates (i.e., regions A and B, gated on AB; regions R1 and R2, G2 = R2 and G1). Draw an amorphous polygon region (C; R3) to include only those events that form a cluster with low to intermediate SS and CD45dim expression. 13. Histogram 4 (lymph-blast gate): Display events that fulfill the criteria of all three above gates (i.e., regions A, B, and C, gated on ABC; regions R1, R2, and R3, G3 = R3 and G2). Draw an amorphous polygon region (D; R4) to include only those events that form a cluster with low to intermediate SS and low to high FS. On BD Biosciences FACS instruments, set logical gate G4 = R4 and G3. The lymph-blast gate serves to exclude platelets and debris that may show weak nonspecific binding of CD34 and CD45 MAbs. Its lower boundaries are verified in histogram 6 (see below).
14. Histogram 5: Display ungated data. Draw a quad-stat region in order to establish the lower limit of CD45 expression by the CD34+ events. On histogram 5, draw a small rectangular region H (R6) to include the brightest events that fall in the highest green and orange-red channels. Although not readily visible, all the Flow-Count beads (e.g., singlets and aggregates) are contained in this region. Enumeration of CD34+ Hematopoietic Stem and Progenitor Cells
15. Histogram 6 (duplicate lymph-blast gate): On histogram 1, draw an amorphous polygon region (E; R5) on the lymphocytes (CD45bright, SSlow) to create a lymphocyte
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gate. Display the events from this region in histogram 6 (i.e., gated on E; G5 = R5). Then: a. For Beckman Coulter XL: Draw an amorphous polygon region F in histogram 6 that is similar to region D in histogram 4, creating the duplicate lymph-blast gate. Adjust region F on histogram 6 so that lymphocytes from region E are just included. Once region F has been optimized, place region D in histogram 4 similarly. b. For BD Biosciences FACS: Copy region R4 from histogram 4 into histogram 6 (creating the duplicate lymph-blast gate), and adjust the position of the duplicate so that lymphocytes from region R5 are just included. The original region R4 on histogram 4 will automatically move to the same position on histogram 4. This step establishes the minimum FS and SS range for the lymph-blast gate (F; R4) in histogram 6.
16. Histogram 7 (bead gate); gated on H (R6): a. For Beckman Coulter XL: Set a rectangular region (G) to include only single-bead events as per the manufacturer’s recommendations. Adjust counting-bead (red fluorescence) high voltage to ensure that both singlet and doublet bead populations are visible. On the Beckman Coulter XL-MCL, define region G as a CAL region to allow automatic calculation of absolute numbers of CD34+ cells. b. For BD Biosciences FACS: Display events from R6 on histogram 7 (time versus FS). If beads are not visible, lower FS threshold until singlet beads are clearly visible. Set region R7 to include only singlet beads. Set logical gate G7 = R6 and R7. Optional (for dead cell exclusion): On histogram 8 for Beckman Coulter XL, draw a rectangular region (J) to include only living (i.e., 7-AAD−) cells, and display listmode data from region J only in histograms 2 to 6. A summary of the resulting logical gates is shown in Table 6.4.1. For FACS, draw a rectangular region (R8) on histogram 8 that includes the dead cells (7-AAD+) but excludes the counting beads (that are present in the highest SS and far-red fluorescence channels). Exclude dead cells from further analysis by setting logical gates as in Table 6.4.1.
Table 6.4.1 Summary of Instrument-Specific Logical Gates Using Dead Cell Exclusion
Histogram 1 2 3 4 5 6 7 8
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BD Biosciences FACS
Gated on J Gated on AJ Gated on ABJ Gated on ABCJ Ungated Gated on EJ Gated on H Ungated
G8 = not R8 G1 = (not R8) and R1 G2 = R2 and G1 G3 = R3 and G2 Ungated G5 = (not R8) and R5 G6 = R6 Ungated Phenotypic Analysis
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Acquire data NOTE: Data analysis is performed using System 2/Expo Software for Beckman Coulter XL instruments or using CellQuest Software for BD Biosciences FACS series instruments. 17. Acquire a minimum of 100 CD34+ events (region D; G4 = R4 and G3) from the first CD34 tube. If a sensitivity level of 0.1% is sufficient, stop at 50,000 CD45+ events (region A; R1). 18. Repeat step 17 for the second CD34 tube. The total number of events to be acquired is dependent on the desired sensitivity (e.g., a level of 0.1% implies the detection of 100 CD34+ cells among a total of 100,000 CD45+ cells) in the two tubes analyzed.
19. Run the PBS tube for 1 min between two patient specimens to flush fluids and prevent specimen carryover. Flushing the system with PBS is not required between duplicates, but should be done before another pair of duplicate samples is run.
Analyze data 20. If the total number of events in region D (G4 = R4 and G3) of the two CD34 tubes is ≥100, calculate the absolute number of CD34+ cells/µl as follows (for instruments that do not calculate the absolute CD34+ cell number automatically): CD34 + cells / µl =
no. CD34 + × B × DF no. beads
where the number of CD34+ cells is determined from region D (G4 = R4 and G3), the number of singlet beads is determined from region G (G7), B is bead concentration (specified per lot), and DF is the sample dilution factor. Average the results of the two tubes. 21. If it is necessary to know the % CD34+ cells in a sample (i.e., as a fraction of leukocytes) or if the total number of events in region D (G4 = R4 and G3) of the two CD34 tubes is 95%) should be used. It has been shown that samples which are not immediately processed after collection are not in optimal condition for calculating the absolute numbers of a given cell population, since they contain varying numbers of dead cells; therefore, samples are ideally processed within 3 hr after being obtained in order to perform reliable identification, enumeration, and phenotypic characterization of antigen-presenting cells, including DCs. Sample storage, handling, and preparation are also important variables that might affect the accuracy of DC enumeration. Accordingly, the accuracy of pipetting, which should be performed by reverse pipetting (UNIT 6.4), is of critical importance in single-platform assays, where the result is directly affected by the amount of both sample and counting beads added. Likewise, the inclusion of washing steps is not recommended when microbeads are used
Phenotypic Analysis
6.9.11 Current Protocols in Cytometry
Supplement 17
for enumerating DCs, as washing could result in selective loss of either cells or counting beads. Regarding immunophenotypical studies for the characterization of each DC subset, the time from sample collection to its technical processing may not be so limiting as for DC enumeration; however, it should be noted that at present, information in the literature in this respect is scanty. Thus, processing of samples as soon as possible is recommended for the phenotypic characterization of DC subsets.
Phenotypic Analysis of Human PB Dendritic Cells
Choice of mAbs and fluorochromes Currently, no specific markers exist that can identify all human DC subsets. Thus, it is recommended to use a mixture of mAbs directed against T and B lymphocytes, NK cells, and monocytes in order to be able to select for the minor populations of antigen-presenting cells present in the sample, including DCs, on the basis of their negativity for these markers; therefore, all the mAbs included in the cocktail must be conjugated with the same fluorochrome molecule. In the authors’ experience, the best combination of such mAbs for normal human PB includes: CD3 (to exclude all mature T cells), CD19 (for B lymphocytes), CD56 (for NK cells; alternatively, CD16 could be used but would prevent the identification of CD16+ antigen-presenting cells), and CD14 (mature monocytes can be excluded by their strong positivity for this antigen). In principle, all these mAbs should be FITC-conjugated, since PE is better used for either specific identification of CD16 reactivity or the expression of other molecules of interest. Positive identification of peripheral blood DCs requires the presence of anti-HLA-DR, and CD33 allows for the specific discrimination between the different subsets of DCs and the CD16+/HLA-DR+/lineage− antigen-presenting cells. Additionally, it should be noted that the identification and enumeration of DCs in PB samples that contain abnormally increased numbers of other HLA-DR+ cells (e.g., malignant blast cells, CD34+ hemopoietic progenitors) may need special consideration in order to exclude these events from the DC region (i.e., inclusion of CD34 mAb in the exclusion cocktail). If only a three-color instrument is available, the study of DCs would then be, in principle, limited to their identification and enumeration, since at present the fourth color would be necessary for their characterization.
Number of events to acquire To obtain statistically reliable DC subset counts by flow cytometry, it is crucial to acquire a large number of events included in the fraction in which DCs are located. As the different DC subsets are considered to be rare events, in the protocols described above for the enumeration of peripheral blood DC subsets, including the CD16+ subset of antigen-presenting cells, the authors recommend acquiring and storing information on a minimum of 3000 to 5000 total DCs/CD16+ antigen-presenting cells. Since these cells represent ∼1% of all nucleated cells in normal human peripheral blood, it is necessary to acquire information on at least 3–5 × 105 events, corresponding to the whole peripheral blood cellularity. Although information can be stored for all events acquired, the authors recommend use of a two-step acquisition procedure, information from the second step being stored exclusively through a live gate on HLADR+ events, to prevent computer-memory problems with an over-large listmode file.
Anticipated Results Enumeration of DC subpopulations present in normal human peripheral blood samples The overall percentage of PB DCs and CD16+ antigen-presenting cells from normal human individuals is ∼1% of all nucleated cells (1.03% ± 0.33%), ranging between 0.46% and 1.99%. The most frequent cell subset is CD16+, representing 0.72% ± 0.35% of all nucleated cells. From the other two subsets, CD16− /CD33strong+ DCs represent ∼0.2% (0.19% ± 0.08%) of all PB nucleated cells and the CD16− /CD123strong+ DC sub po pu lation ∼1.0% (0.11% ± 0.04%; Almeida et al., 2001). In absolute numbers the distribution of these cell subsets is as follows: 39.9 ± 17.4 CD16+ antigen-presenting cells/µl, 10.5 ± 3.9 CD16− /CD33strong+ DCs/µl, and 6.3 ± 2.6 CD16− /CD123strong+ DCs/µl. Immunophenotypic characteristics of DC subpopulations present in normal human peripheral blood samples The technical procedure described above represents a sensitive and specific method for the systematic study of the immunophenotypic characteristics of the different DC subsets. As an example, the authors have seen that the three different DC subsets can be distinguished by their distinct reactivity for Fcγ and complement receptors, certain adhesion and costimulatory molecules, in addition to their different pattern
6.9.12 Supplement 17
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of expression of CD33 and HLA-DR (Almeida et al., 2001). In principle, any molecule of interest present either on the surface or inside the DCs can be specifically studied and precisely measured using the four-color flow cytometric protocols described in this unit.
Time Considerations
Lane, P.J. and Brocker, T. 1999. Developmental regulation of dendritic cell function. Curr. Opin. Immunol. 11:308-313. O’Doherty, U., Peng, M., Gezelter, S., Swiggard, W.J., Betjes, M., Bhardwaj, M., and Steinman, R.M. 1994. Human blood contains two subsets of dendritic cells, one immunologically mature and the other immature. Immunology 82:487493.
Cell preparation takes 45 to 60 min. Acquisition on the flow cytometer will take 10 to 15 min per sample. The Alternate Protocol will take 1 hr total.
Olweus, J., BitMansour, A., Warnke, R., Thompson, P.A., Carballido, J., Picker, L.J., and Lund-Johansen, F. 1997. Dendritic cell ontogeny: A human dendritic cell lineage of myeloid origin. Proc. Natl. Acad. Sci. U.S.A. 94:12551-12556.
Literature Cited
Orfao, A., Escribano, L., Villarrubia, J., Velasco, J.L., Cerveró, C., Ciudad, J., Navarro, J.L., and San Miguel, J.L. 1996. Flow cytometric analysis of mast cells from normal and pathologic human bone marrow samples. Identification and enumeration. Am. J. Pathol. 149:1493-1499.
Almeida, J., Bueno, C., Algueró, M.C., Sanchez, M.L., Cañizo, M.C., Fernandez, M.E., Vaquero, J.M., Escribano, L., Laso, F.J., San Miguel, J.F., and Orfao, A. 1999a. Extensive characterization of the immunophenotype and the pattern of cytokine production by distinct subpopulations of normal human peripheral blood dendritic cells. Clin. Exp. Immunol. 118:392-401. Almeida, J., Orfao, A., Ocqueteau, M., Mateo, G., Corral, M., Caballero, M.D., Blade, J., Moro, M.J., Hernandez, J., and San Miguel, J.F. 1999b. High sensitive immunophenotyping and DNA ploidy studies for the investigation of minimal residual disease in multiple myeloma. Br. J. Haematol. 107:121-131. Almeida, J., Bueno, C., Algueró, M.C., Sanchez, M.L., Escribano, L., Diaz-Agustin, B., Vaquero, J.M., Laso, F.J., San Miguel, J.F., and Orfau, A. 2001. Comparative analysis of the morphological, cytochemical, immunophenotypical and functional characteristics of normal human per ip h er al bloo d lineag e−/CD16+/HLADR+/CD14−/low cells, CD14+ monocytes and CD16− dendritic cells. Clin. Immunol. In press. Arpinati, M., Green, C.L., Heimfeld, S., Heuser, J.E., and Anasetti, C. 2000. Granulocyte-colony stimulating factor mobilizes T helper 2-inducing dendritic cells. Blood 95:2484-2490. Banchereau, J. and Steinman, R.M. 1998. Dendritic cells and the control of immunity. Nature 392:245-252. Bernhard, H., Disis, M.L., Heimfeld, S., Hand, S., Gralow, J.R., and Cheever, M.A. 1995. Generation of immunostimulatory dendritic cells from human CD34+ hematopoietic progenitor cells of the bone marrow and peripheral blood. Cancer Res. 55:1099-1104. Dzionek, A., Fuchs, A., Schmidt, P., Cremer, S., Zysk, M., Miltenyi, S., Buck, D.W., and Schmitz, J. 2000. BDCA-2, BDCA-3, and BDCA-4: Three markers for distinct subsets of dendritic cells in human peripheral blood. J. Immunol. 165:6037-6046.
Pickl, W.F., Majdic, O., Kohl, P., Stöckl, J., Riedl, E., Scheinecker, C., Bello-Fernández, C., and Knapp, W. 1996. Molecular and functional characteristics of dendritic cells generated from highly purified CD14+ peripheral blood monocytes. J. Immunol. 157:3850-3859. Rissoan, M.-C., Soumelis, V., Kadowaki, N., Grouard, G., Briere, F., Waal Malefyt, R., and Liu, Y.-J. 1999. Reciprocal control of T helper cell and dendritic cell differentiation. Science 283:1183-1186. Schäkel, K., Mayer, E., Federle, C., Schmitz, M., Riethmüller, G., and Rieber, P. 1998. A novel dendritic cell population in human blood: Onestep immunomagnetic isolation by a specific mAb (M-DC8) and in vitro priming of cytotoxic T lymphocytes. Eur. J. Immunol. 28:4084-4093. Steinman, R.M. 1996. Dendritic cells and immunebased therapies. Exp. Hematol. 24:859-862. Thomas, R. and Lipsky, P.E. 1994. Human peripheral blood dendritic cell subsets. Isolation and characterization of precursor and mature antigen-presenting cells. J. Immunol. 153:40164028. Williams, L.A., Egner, W., and Hart, D.N. 1994. Isolation and function of dendritic cells. Int. Rev. Cytol. 153:41-103. Young, J.W. and Steinman, R.M. 1996. The hematopoietic development of dendritic cells: A distinct pathway for myeloid differentiation. Stem Cells 14:376-387.
Contributed by Julia Almeida and Clara Bueno Universidad de Salamanca Salamanca, Spain
Hart, D.N.J. 1997. Dendritic cells: Unique leukocyte populations which control the primary immune response. Blood 90:3245-3287. Phenotypic Analysis
6.9.13 Current Protocols in Cytometry
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Immunophenotypic Analysis of Platelets
UNIT 6.10
With an average diameter of 3 µm, platelets are the smallest circulating cellular component in peripheral blood. The primary role of circulating platelets is to maintain hemostasis. The evaluation of platelets by flow cytometry has proven beneficial in the investigation of many disease states, including inherited defects such as Bernard-Soulier syndrome, Glanzmann thrombasthenia, and storage pool disease (Michelson et al., 2001). Flow cytometric techniques have been used in blood bank applications such as quality control of platelet concentrates, immunophenotyping of platelet surface receptor polymorphisms, platelet crossmatching, and detection of feto-maternal anti-platelet antibodies. Platelet hyporeactivity may result in potentially life-threatening bleeding including intracranial hemorrhage, while platelet hyperreactivity may result in intravascular thrombosis resulting in potentially life-threatening acute myocardial infarction or stroke. Consequently, antiplatelet therapies designed to reduce platelet responsiveness in vivo are now common practice in clinical cardiovascular medicine. Perhaps most common is the use of flow cytometry to study the role of platelet function and platelet activation in cardiovascular disease. A more complete list of the applications of flow cytometry to the study of platelets is shown in Table 6.10.1 and discussed in Michelson et al. (2001). Resting platelets constitutively express many surface glycoproteins that are easily identified by flow cytometry. Upon platelet activation, many surface receptors are modulated in both copy number and conformation, while others, absent from the resting platelet surface, are newly expressed. This unit describes several strategies to evaluate platelet function by evaluating surface receptor expression on resting and activated platelets using flow cytometry. Three methods are described here in detail: determination of resting platelet surface receptor expression (see Basic Protocol 1 and Alternate Protocol); determination of platelet activation using P-selectin (CD62P) expression (see Basic Protocol 2), which reflects platelet α-granule release (McEver, 2001), or PAC1 binding, which detects the activated conformation of glycoprotein (GP) IIb-IIIa (integrin αIIbβ3; Shattil et al., 1985); and determination of procoagulant platelets and platelet-derived microparticles using annexin V binding or monoclonal antibodies specific for coagulation factors V/Va or X/Xa (see Basic Protocol 3; Furman et al., 2000). The methods described here are performed using the more physiologically relevant milieu of whole blood, which has the following advantages over platelet-rich plasma or washed platelet systems: (1) red cells and leukocytes are present, both of which affect platelet activation; (2) minimal sample manipulation minimizes artifactual in vitro activation and potential loss of platelet subpopulations; (3) both the activation state of circulating platelets and the reactivity of circulating platelets can be determined; (4) only minuscule volumes (∼5 µl) of blood are required, making whole-blood flow cytometry particularly advantageous for neonatal studies; and (5) platelets of patients with profound thrombocytopenia can also be accurately analyzed. This unit does not specifically address platelet-associated IgG, heparin-induced thrombocytopenia, monitoring of GPIIb-IIIa receptor antagonists, reticulated platelets (see UNIT 7.10), leukocyte-platelet aggregate formation, or platelet counting. For a review of platelet-associated analysis techniques not covered by this unit see Schmitz et al. (1998).
Phenotypic Analysis Contributed by Lori A. Krueger, Marc R. Barnard, A.L. Frelinger III, Mark I. Furman, and Alan D. Michelson
6.10.1
Current Protocols in Cytometry (2002) 6.10.1-6.10.17 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 19
Table 6.10.1 Applications of Flow Cytometry to the Study of Plateletsa
Measurement of platelet activationb Activation-dependent monoclonal antibodies/reagents Modulation of constitutively expressed surface receptors Procoagulant platelet-derived microparticles Leukocyte-platelet aggregates Platelet-platelet aggregates Diagnosis of specific disorders Bernard-Soulier syndrome Glanzmann thrombasthenia Storage pool disease Heparin-induced thrombocytopenia Immune thrombocytopenias Monitoring of antiplatelet agents GPIIb-IIIa antagonists Thienopyridines Monitoring of thrombopoiesis Reticulated platelets Blood bank applications Quality control of platelet concentrates Identification of leukocyte contamination in platelet concentrates Immunophenotyping of platelet HPA-1a Detection of maternal and fetal anti-HPA-1a antibodies Platelet cross-matching Platelet counting Research applications Platelet survival, tracking, and function in vivo Platelet recruitment Bacteria-platelet interactions Calcium flux Cytoskeletal rearrangement Fluorescence resonance energy transfer Signal transduction aTable order reflects the most commonly studied, and relevant applications. bIncludes circulating activated platelets, platelet hyperreactivity, or platelet hyporeactivity.
STRATEGIC PLANNING Blood Collection Careful experimental planning is required for accurate and consistent results when immunophenotyping platelets by flow cytometry. To minimize ex vivo platelet activation, blood samples should be processed within ∼30 min after drawing blood for many assays. The act of drawing blood is itself a potential source of artifactual platelet activation; therefore, the following recommendations are suggested (Michelson et al., 2001): Use a light tourniquet or none at all Use a 21-G (or larger bore) needle Ensure a smooth draw (i.e., good flow) Discard the first 2 ml of blood drawn Immunophenotypic Analysis of Platelets
Adhering to these recommendations will minimize tissue thromboplastin contamination of blood samples and red cell hemolysis that could lead to artifactual platelet activation.
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Table 6.10.2
Anticoagulants Used in the Study of Platelets
Anticoagulant
Mechanism of action
Weak Ca2+ chelator Chelates Ca2+ and increases intracellular cAMP, keeping platelets “quiet” Activated coagulation factor XII inhibitor Strong Ca2+ chelator, dissociates GPIIb-IIIa complex Combines with anti-thrombin III to inhibit thrombin activity Hirudin Direct thrombin inhibitor D-Phenylalanyl-L-prolyl-L-arginine Direct thrombin inhibitor chloromethyl ketone (P-PACK) Sodium citrate Weak Ca2+ chelator
Acid citrate dextrose (ACD) Citrate theophylline adenosine dipyridimole (CTAD) Corn trypsin inhibitor EDTAa Heparina
aThese anticoagulants should be avoided for evaluation of platelet function studies by flow cytometry (see Strategic
Planning).
Each laboratory should determine whether their method of collection, including the drawing of samples through angioplasty and other catheters, results in artifactual in vitro platelet activation, as determined by the binding of activation-dependent monoclonal antibodies. Choice of Anticoagulant Although sodium citrate (a weak calcium chelator) is the most common anticoagulant used for platelet studies, others have been successfully used. EDTA (a strong calcium chelator) should be avoided, because it causes dissociation of the integrin αIIbβ3 (GPIIbIIIa) complex. Heparin should also be avoided because it binds to, and may activate, platelets. Nonchelating anticoagulants such as P-PACK (a direct thrombin inhibitor) may be preferable for the monitoring of GPIIb-IIIa antagonist therapy. The anticoagulants listed in Table 6.10.2 have reportedly been used in studies of platelets. Sample Handling The length of time between sample draw and sample preparation should be minimized to reduce spontaneous platelet activation. The blood should be properly mixed with anticoagulant, avoiding unnecessary agitation prior to testing. The whole-blood samples should be collected and maintained in a container with nonwettable surfaces such as siliconized glass or polypropylene. Experimental Design Many platelet surface receptors are modulated during platelet activation (see Table 6.10.3). For example, GPIb-IX-V may be cleaved and/or internalized to the surface-connected canalicular system upon platelet activation (Michelson et al., 1996a). This must be taken into consideration in experimental design. For example, if using a GPIbα-specific antibody (i.e., anti-CD42b) as a platelet identifier, then an adjustment in the instrument threshold (or discriminator) may be necessary when evaluating activated versus resting platelets. Alternatively, if the reduction in platelet GPIbα is being used as an indicator of platelet activation, then the maximal change in receptor expression will be observed only if platelets are labeled with the GPIbα-specific antibody after platelet activation has taken place. If a directly-conjugated GPIb-IX-specific test monoclonal antibody is added to the platelets prior to activation, then the activation-induced redistribution of GPIb-IX to the surface-connected canalicular system will not result in a significant decrease in platelet fluorescence, because fluorescence will be detected irrespective of whether the conjugated antibody is on the surface of or within the platelet; therefore, in flow cytometric
Phenotypic Analysis
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Table 6.10.3 Activation-Dependent Changes in Platelet Surface Labeling of Monoclonal Antibodies and Annexin Va
Activation-dependent platelet surface change Changes in surface receptor expression CD36 GPIb-IX GPIIb-IIIa Conformational changes in GPIIb-IIIa (integrin αIIbβ3) Ligand-induced binding sites (LIBS) PAC1 Receptor-induced binding sites on fibrinogen (RIBS) Development of a procoagulant surface Factor VIII binding Factor V/Va binding Factor X/Xa binding Phosphatidylserine expression (detected by annexin V) Exposure of granule membrane proteins CD40L (or CD154) CD63 (lysosomes) LAMP-1 (lysosomes) LAMP-2 (lysosomes) Lectin-like oxidized LDL receptor-1 (LOX-1) P-selectin (CD62P, α-granules) Platelet surface binding of secreted platelet proteins Multimerin Thrombospondin
Resting platelet
Activated platelet
+ ++ ++
++ + +++
− − −
+++ +++ +++
− − − −
+++ +++ +++ +++
− − − − − −
+ ++ ++ ++ + +++
− −
+ +
aAnnexin V is a 35 to 36 kDa protein that binds to phosphatidylserine in the presence of Ca2+.
assays examining platelet surface GPIb-IX modulation, GPIb-IX-specific antibodies that are directly conjugated must be added to the assay after the addition of the agonist. BASIC PROTOCOL 1
Immunophenotypic Analysis of Platelets
IMMUNOPHENOTYPING OF PLATELET SURFACE RECEPTORS This procedure may be used to qualitatively and/or quantitatively evaluate a platelet surface receptor such as GPIIb-IIIa (reduced or absent in Glanzmann thrombasthenia) or GPIb-IX-V (reduced or absent in Bernard-Soulier syndrome). Additionally, this protocol may be used to evaluate the in vivo activation status of circulating platelets. The optimal final concentration of platelet-specific monoclonal antibody reagents must be determined by titration. At a minimum, two antibodies, each conjugated to a different fluorochrome (e.g., fluorescein and phycoerythrin), are used. The fluorescent conjugate of the platelet identifier determines the thresholding parameter used in the flow cytometric analysis. The choice of the platelet identifier is determined by the analysis being performed. For example, anti-CD42a or anti-CD42b (GPIX- and GPIbα-specific, respectively) are used as platelet identifiers when measuring surface expression of GPIIb-IIIa (CD41 and CD61) in the investigation of Glanzmann thrombasthenia, an inherited deficiency of GPIIb-IIIa. Anti-CD41 or anti-CD61 may be used as platelet identifiers when investigating Bernard-Soulier syndrome, an inherited deficiency of GPIb-IX-V (CD42b, CD42a, and CD42d). Anti-CD41, -CD61, -CD42a, or -CD42b may be used as platelet identifiers when evaluating activation-dependent receptors such as P-selectin
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(recognized by anti-CD62P) and PAC1; however, surface expression of GPIIb-IIIa and GPIb-IX is modulated upon platelet activation, which must be considered when selecting a platelet identifier. Monoclonal antibody reagents are prepared in modified HEPES/Tyrode’s (HT) buffer. Materials Whole blood (WB) containing anticoagulant (see Strategic Planning) or isolated platelets (see Support Protocol) Modified HT buffer (see recipe) Platelet-specific antibody cocktail titrated in modified HT buffer (minimum two antibody specificities each conjugated to a different fluorochrome): Specific platelet identifier: monoclonal anti-CD41, -CD61, -CD42a, or -CD42b Marker of platelet activation: monoclonal anti-CD62P or PAC1 Negative control: antibody isotype-, concentration-, fluorochrome-, and F:P ratio-matched to the activation marker, or blocking agent inhibiting platelet-specific marker binding 1% formalin fixative (see recipe) 1. Within 30 min of blood draw, dilute whole blood (WB) containing anticoagulant 1/10 in modified HT buffer (e.g., 10 µl WB and 90 µl HT). In the diagnosis of Glanzmann thrombasthenia and Bernard-Soulier syndrome it is wise to use blood from a normal healthy donor analyzed in parallel with patient blood to differentiate normal and abnormal expression of GPIIb-IIIa or GPIb-IX, respectively.
2. Immediately mix diluted whole blood (dWB) with appropriately titrated platelet-specific antibody cocktail (e.g., 10 µl dWB and 30 µl antibody cocktail). Include a negative control if a platelet activation marker is used. Incubate at room temperature (e.g., 20 min). Individual laboratories must determine the optimal antibody concentration and incubation time.
3. Fix labeled cells with 10× to 20× total assay volume (e.g., 400 to 800 µl) of 1% formalin solution. Let fixation proceed 15 min at room temperature, then place tubes at 4°C until analysis. Stability of fixed preparations must be determined by individual laboratories.
4. Analyze by flow cytometry using the following setup: a. Use the platelet identifier as the thresholding (or discriminating) parameter (Fig. 6.10.1A). b. Collect light-scatter parameters in logarithmic mode. (Heterogeneity in platelet size and platelet morphology contributes to a relatively broad distribution of light scattering detectable by flow cytometry.) c. Analyze diluted fixed sample using low flow rate (e.g., 150 to 250 platelet events per second) to minimize, as far as possible, coincidence between two or more events. d. Identify single platelet events by their characteristic light scatter and positive labeling with platelet-specific identifier (Fig. 6.10.1B). e. Measure the platelet surface receptor or receptors of interest from gated events (Fig. 6.10.1C). Platelets are smaller than lymphocytes and erythrocytes; therefore, high-voltage settings used for leukocyte and erythrocyte studies may not be appropriate for platelet analysis.
Phenotypic Analysis
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100
103
102
103
104 102
101
Forward scatter 101
B
100
104 103 102 101 100
Platelet identifier fluorescence
A
104
100
101
0
101
102
103
P-selectin fluorescence
104
103
104
103
104
102
103
D
101 100
20
30
Forward scatter
104
C
100
102
Side scatter
10
Count
40 50 60
Side scatter
100
101
102
FX/Xa fluorescence
Figure 6.10.1 Evaluation of platelet activation by whole-blood flow cytometry. (A) Platelets labeled with specific platelet identifier. The instrument threshold (or discriminator) is set on the fluorescence parameter corresponding to the conjugate of the platelet identifier. (B) Characteristic light-scatter profile of a platelet population in diluted whole blood. Data are collected and displayed using logarithmic-orthogonal and logarithmic-forward light scatter. Single platelet events are identified by their characteristic light-scatter properties and positive labeling with a platelet-specific monoclonal antibody reagent. (C) Single-parameter fluorescence histogram of the platelet activation marker P-selectin (CD62P). The positive analysis region is determined by the negative isotype control (thin solid line). Events displayed are generated from gated events in both panel A and panel B. (D) Determination of procoagulant platelets and platelet-derived microparticles. Two-parameter histogram displaying Factor X/Xa binding versus forward-angle light scatter. The analysis region for platelets capable of binding coagulation protein–-activated factors V and X or expressing phosphatidylserine is established on this two-parameter histogram.
ALTERNATE PROTOCOL
IMMUNOPHENOTYPING OF PLATELET SURFACE RECEPTORS Whole blood may be fixed first and subsequently labeled with platelet-specific reagents. Many platelet surface receptors are well preserved and recognized by specific monoclonal antibodies after formalin fixation; however, labeling intensity may diminish with prolonged fixation. Antibody concentrations required for optimal labeling may be different for fixed versus unfixed cells; therefore, optimal antibody concentrations should be verified by titration using fixed cells. Some antigen epitopes such as that recognized by PAC1 may not label after formalin fixation.
Immunophenotypic Analysis of Platelets
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Additional Materials (also see Basic Protocol 1) 2% formalin fixative (see recipe) 1. Within 30 min of blood draw, dilute WB 1:1 with 2% formalin fixative. (i.e., 100 µl WB and 100 µl of 2% formalin). Fix 15 to 60 min at room temperature. Store samples at 4°C. Antibody labeling may decrease in intensity with prolonged fixation; therefore, the length of time that fixed samples can be stored prior to labeling must be determined by individual laboratories for each antibody clone being used.
2. Dilute fixed WB 1:10 in modified HT buffer. Dilution can be performed prior to storage at 4°C.
3. Mix dWB with appropriately titrated antibody cocktail (e.g., 20 µl dWB and 20 µl antibody cocktail) and incubate at room temperature (e.g., 20 min). Individual laboratories must determine the optimal antibody concentration and incubation time.
4. Dilute labeled cells with 10× to 20× total assay volume (e.g., 400 to 800 µl) of 1% formalin solution. Place tubes at 4°C until analysis. Stability of fixed preparations must be determined by individual laboratories.
5. Analyze by flow cytometry as described above (see Basic Protocol 1, step 4) DETERMINATION OF PLATELET ACTIVATION USING P-SELECTIN OR PAC1 EXPRESSION
BASIC PROTOCOL 2
This procedure may be used to evaluate platelet reactivity in response to agonist. Materials Whole blood (WB) containing anticoagulant (see Strategic Planning) or isolated platelets (see Support Protocol) Modified HT buffer (see recipe) Platelet-specific antibody cocktail titrated in modified HT buffer (minimum two antibody specificities each conjugated to a different fluorochrome): Specific platelet identifier: e.g., monoclonal anti-CD41, -CD61, -CD42a, -CD42b Specific marker of platelet activation: e.g., monoclonal anti-CD62P or -PAC1 Platelet agonist (see recipe): e.g., ADP, epinephrine, human α-thrombin, thrombin receptor-activating peptide (TRAP) Negative controls: antibody isotype-, concentration-, fluorochrome-, and F:P ratio-matched to the specific activation marker, or blocking agent that inhibits the platelet-specific marker binding 10 mM GPRP (see recipe) 1% formalin fixative (see recipe) 1. Within 30 min of blood draw, dilute WB 1:10 in modified HT buffer (i.e., 10 µl WB and 90 µl HT). 2. Immediately mix dWB with appropriately titrated platelet-specific antibody cocktail and with platelet agonist (e.g., 10 µl dWB, 20 µl antibody cocktail, and 10 µl platelet agonist). Include a negative control if a platelet activation marker is used. Incubate at room temperature (e.g., 20 min). If thrombin is being used as the platelet agonist, add GPRP to a final concentration of 2.5 µm to prevent fibrin polymerization and clot formation (Michelson, 1994).
Phenotypic Analysis
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The fluorescent conjugate of the platelet identifier determines the thresholding parameter used in the flow cytometric analysis. Surface expression of GPIIb-IIIa and GPIb-IX are modulated upon platelet activation, which must be considered when selecting a platelet identifier, establishing instrument thresholds, and gating platelet populations. Individual laboratories must determine the optimal antibody concentration and incubation time.
3. Fix labeled cells with 10× to 20× total assay volume (e.g., 400 to 800 µl) of 1% formalin fixative. Let fixation proceed 15 min at room temperature, then place tubes at 4°C until analysis. Stability of fixed preparations must be determined by individual laboratories.
4. Analyze by flow cytometry as described above (see Basic Protocol 1, step 4). BASIC PROTOCOL 3
DETERMINATION OF PROCOAGULANT PLATELETS USING ANNEXIN V BINDING OR MONOCLONAL ANTIBODIES SPECIFIC FOR COAGULATION FACTOR V/Va OR X/Xa This procedure may be used to evaluate procoagulant platelets and the ability of platelets to generate procoagulant microparticles in response to an agonist (Furman et al., 2000). Materials Whole blood (WB) containing anticoagulant (see Strategic Planning) or isolated platelets (see Support Protocol) Modified HT buffer (see recipe) containing 5 mM GPRP (see recipe) Coagulation factor V/Va or X/Xa Platelet agonist (see recipe) supplemented with 6 mM CaCl2: e.g., collagen, combined thrombin/collagen mixture, or calcium ionophore A23183 Modified HT buffer Platelet-specific antibody cocktail titrated in modified HT buffer—minimum two antibodies, or one identifier and annexin V—each conjugated to a different fluorochrome: Specific platelet identifier: e.g., anti-CD41, anti-CD61, anti-CD42a, or anti-CD42b Marker of platelet procoagulant activity: e.g., annexin V, monoclonal anti-coagulation factor V/Va or X/Xa 1% formalin fixative (see recipe) 1. Within 1 hr of draw, dilute WB 1:10 in modified HT buffer containing 5 mM GPRP (i.e., 10 µl WB and 90 µl HT/GPRP). If coagulation factor V/Va or X/Xa is to be detected, include these factors in the HT/GPRP diluent. Sources of V/Va or X/Xa include autologous platelet-poor plasma or purified coagulation factors. Each individual laboratory must titrate the optimal concentration of autologous plasma or purified coagulation factor to add back for optimal platelet surface detection.
2. Immediately combine dWB with an equal volume of platelet agonist supplemented with 6 mM CaCl2, or modified HT buffer alone—e.g., 15 µl dWB/GPRP and 15 µl agonist or buffer alone (negative control). Incubate 20 min at 37°C. The optimal incubation time must be determined by individual laboratories.
Immunophenotypic Analysis of Platelets
The final concentration of GPRP is 2.5 mM during the assay incubation; therefore, for a 1:1 (dWB:agonist) mix in the assay, the GPRP concentration in the dWB buffer is 2× or 5 mM. Similarly, HT supplemented with 3 mM Ca2+ (final concentration) used alone or as the agonist diluent is prepared 2× or 6 mM.
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The negative control for this assay incorporates HT diluent (rather than HT supplemented with CaCl2) in place of agonist. Annexin V and coagulation factors V/Va and X/Xa will not bind in the absence of Ca2+.
3. Label with titrated platelet-specific antibody cocktail (e.g., 30 µl dWB/GPRP/agonist or buffer, and 10 µl antibody reagent cocktail) 20 min at room temperature. As a minimum, two antibody specificities (or one platelet identifier and annexin V) each conjugated to a different fluorochrome (such as fluorescein and phycoerythrin) are used. The fluorescent conjugate of the platelet identifier determines the thresholding (or discriminating) parameter used in the flow cytometric analysis. Surface expression of GPIIb-IIIa and GPIb-IX is modulated upon platelet activation, which must be considered when selecting a platelet identifier. The optimal final concentration of platelet-specific monoclonal antibody reagents must be determined by titration.
4. Fix labeled cells 15 min with 10× to 20× total assay volume (e.g., 400 to 800 µl) of 1% formalin fixative at room temperature. Place tubes at 4°C until analysis. Stability of fixed preparation must be determined by individual laboratories.
5. Analyze by flow cytometry as described (see Basic Protocol 1, step 4). Procoagulant platelets often display dramatic light-scatter changes requiring adjustments to fluorescence and to light scatter gates. Additionally, the distinction between the procoagulant (positive labeling for annexin V, anti-V/Va, or anti-X/Xa) and nonprocoagulant (negative labeling for annexin V, anti-V/Va, or anti-X/Xa) phenotype may be difficult to define on a single-parameter fluorescence histogram; therefore, a two-parameter histogram displaying fluorescence (annexin V, anti-V/Va, or anti-X/Xa) versus forward-angle light scatter may better define the procoagulant platelet population (Fig. 6.10.1, panel D).
PREPARATION OF ISOLATED PLATELETS Some studies may require the isolation of platelets from other cellular and/or plasma components. Platelets may be separated from other cellular components by centrifugation and further isolated from plasma components by gel filtration or washing.
SUPPORT PROTOCOL
Materials Anticoagulated whole blood collected in 5-ml vacutainer tubes or 15-ml conical tubes if drawn by syringe Sepharose 2B beads Modified HT buffer (see recipe) Citrate wash buffer (see recipe) Benchtop centrifuge with rotors for 5-ml Vacutainer tubes and 15-ml conical tubes Polypropylene or siliconized glass test tube 10-ml syringe column 15-ml conical tubes Preparation of platelet-rich plasma 1. Within 30 min of blood draw, prepare platelet-rich plasma (PRP) by centrifuging anticoagulated whole blood collected in 5-ml vacutainer tubes or 15-ml conical tubes if drawn by syringe in a benchtop centrifuge with an appropriate rotor 10 to 15 min at 150 to 200 × g, room temperature. Remove PRP to a clean polypropylene or siliconized glass test tube, being careful not to disturb the buffy coat and red cell layers (also see UNIT 5.1).
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Isolation of platelets by gel filtration: 2a. Pack a 10-ml syringe column with Sepharose 2B beads. 3a. Equilibrate column with 10 vol deionized water, followed by 3 vol modified HT buffer. 4a. Layer PRP (step 1) on the top of the column. 5a. Let PRP fully enter the column and carefully layer with 30 ml modified HT. 6a. Collect gel-filtered platelets (i.e., eluent). Isolation of platelets by centrifugation: 2b. Fill a 15-ml conical tube containing 1 to 8 ml PRP (step 1) with citrate wash buffer. 3b. Centrifuge in a benchtop centrifuge with an appropriate rotor 10 min at 1,200 × g, at room temperature. 4b. Aspirate the supernatant and gently resuspend platelet pellet. 5b. Repeat wash procedure (steps 2b to 4b). 6b. After final wash, resuspend platelet pellet in modified HT. Use isolated platelets as soon as possible after preparation, regardless of preparation technique. Isolated platelet preparations may be substituted for whole blood in any of the preceding basic protocols.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. Pass all reagents through 0.2- to 0.4-µm filters prior to use. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Citrate wash buffer 11 mM glucose 128 mM NaCl 4.3 mM NaH2PO4 7.5 mM Na2HPO4 4.8 mM sodium citrate 2.4 mM citric acid 0.35% (w/v) BSA Adjust pH to 6.5 with 0.1 M NaOH or 0.1 M HCl Store up to 1 year at −20°C. Bring to room temperature and add 1 mg/ml prostaglandin PGE1 (see recipe) to a final concentration of 50 ng/ml immediately prior to use. Formalin fixative, 1%, 2% Dilute 10% (v/v) ultrapure methanol-free formalin (Polysciences) to 1% or 2% (v/v) in HEPES buffered saline (HBS; see recipe). Store up to 1 month at 4°C. Bring to room temperature prior to use. GPRP, 10 mM Dilute GPRP (Gly-Pro-Arg-Pro) in modified HT buffer (see recipe) to a concentration of 10 mM. Store up to 1 week at 4°C or 1 year at −20°C. Bring to room temperature prior to use. Immunophenotypic Analysis of Platelets
Use for diluting thrombin and in assays containing CaCl2 to prevent fibrin polymerization and clot formation (Michelson, 1994).
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HEPES buffered saline (HBS) 10 mM HEPES 0.15 mM NaCl Adjust pH to 7.4 with 0.1 M NaOH or 0.1 M HCl Store up to 6 months at 4°C Bring to room temperature prior to use Modified HEPES/Tyrode’s (HT) buffer 10 mM HEPES 137 mM NaCl 2.8 mM KCl 1 mM MgCl2 12 mM NaHCO3 0.4 mM Na2HPO4 0.35% (w/v) BSA 5.5 mM glucose Adjust pH to 7.4 with 0.1 M NaOH or 0.1 M HCl Store up to 1 week at 4°C or 1 year at −20°C Bring to room temperature prior to use pH may need readjustment after storage at 4°C.
Prostaglandin PGE1 Prepare 1 mg/ml stock solution in 100% ethanol. Store in small aliquots at −80°C until immediately prior to use. Avoid multiple freeze/thaw cycles. Platelet agonists Prepare all platelet agonist working solutions (concentrations determined empirically) by diluting stocks in modified HT buffer (see recipe), with or without 6 mM CaCl2 as appropriate, just prior to use. Bring to room temperature just before use. Discard left-over working solutions daily. ADP: Adenosine diphosphate (ADP) is typically used at concentrations of 20 µM (maximal platelet activation in diluted whole blood) to submaximal doses of 0.5 µM. Store stock ADP according to manufacturer’s instructions (often frozen at −20°C). Calcium ionophore A23187: Prepare 10 mM stock A23187 in DMSO and store up to 1 year at −80°C. A23187 is typically used at concentrations of 10 to 20 µM (maximal platelet activation in diluted whole blood). Dilute stock at least 1:100 (final DMSO concentration
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8b. If using a 96-well tissue-culture plate, wash samples three times as follows: a. Using a repeat pipettor, pipet 75 µl of 1:10 diluted flow bead wash buffer/water to each well. b. Replace cover and vortex the plate to mix the beads well. c. Add another 75 µl of 1:10 diluted flow bead wash buffer/water to the wells. Do not vortex at this point because of the risk of splashover between wells. d. Replace plate cover and centrifuge 3 min at 900 × g with the brake on. e. Remove the plate from the centrifuge and flick the plate to remove the supernatant. To flick, remove the plate cover and quickly invert the plate over an appropriate receptacle. The inversion should be forceful enough that the supernatant is removed from the well, but not the beads. While still holding the plate upside down, place it on an absorbent pad to wick off the remaining liquid.
f. Replace the cover and gently run the plate across the vortexer in order to loosen the bead pellet. g. Repeat wash two additional times. Stain with secondary antibody 9. Add 20 µl FITC-conjugated anti-human IgG to the dry button. If testing for IgM antibodies use FITC-conjugated anti-human IgM.
10. Vortex samples and incubate 30 min in the dark at room temperature. After 15 min, gently vortex to resuspend beads. 11. Wash samples as in step 8a or 8b. 12a. If using 6 × 50–mm glass tubes, resuspend in 200 µl of 1:10 diluted flow bead wash buffer/water and vortex. 12b. If using a 96-well tissue culture plate, add 75 µl of 1:10 diluted flow bead wash buffer/water, vortex, and add another 75 µl of 1:10 diluted flow bead wash buffer/water. Transfer the volume from the wells to prenumbered 6 × 50–mm glass tubes using the multi-channel pipettor. 13. Run samples immediately on a flow cytometer and analyze, or store ≤24 hr at 4°C. Samples may be transferred to 12 × 75–mm tubes if the cytometer will take nothing else. However, the 6 × 50–ml tubes hold only 200 ìl, a volume that may be too small in a 12 × 75–tube for some instruments. Alternatively, for BD instruments and the Beckman Coulter FC500, the 6 × 50–ml tubes can be fitted into 12 × 75–mm tubes and will work well if care is taken to get the sample probe into the smaller tube when the sample is run. This configuration permits staining of very small volumes of sample and low cell numbers yet maintains reasonable acquisition times. For these assays, particle (cell) to antibody (serum) relationships are critical. Since antibody concentration is unknown, keeping the number of particles constant and minimal maximizes the potential sensitivity.
Acquire and analyze data 14. Set up flow cytometer. The flow cytometer should be calibrated daily using the appropriate control particles for each instrument. Calibration practices should be designed for optimal detection of low fluorescence values, i.e., maximizing signal to noise is essential.
Flow Cytometric Assessment of HLA Alloantibodies
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15. Gate on the bead population in the forward scatter versus side scatter histogram and acquire 15,000-20,000 events. Plot green versus orange (bead) fluorescence (Fig. 6.16.5). Since results for a given sample are a summation of four separate tubes, data analysis is best performed as a separate post-acquisition step. Assign the individual beads as positive or negative based on their shift relative to the negative control markers. Refer to Anticipated Results for more detail.
CELL-BASED FLOW CYTOMETRIC PANEL REACTIVE ANTIBODY (FC-PRA) ASSAY The flow cytometric crossmatch is the most sensitive method for detecting anti-HLA antibodies in the sera of potential allograft recipients. The fact that the FC-PRA is more sensitive than the antiglobulin-enhanced-complement-dependent cytotoxicity (AHGCDC) indicates that situations may arise wherein the antiglobulin-enhanced-complement-dependent cytotoxicity (CDC) crossmatch is negative but the flow cytometric (FC) crossmatch is positive, thereby precluding transplantation in certain instances. In order to better identify and define alloantibodies, the FC-PRA was developed to address routine antibody screening for selected patients. Such patients include new transplant candidates who have a history significant for sensitization (i.e., multiple transfusions or pregnancies) and currently active patients in whom the antibody titer, by AHG-CDC or flow screening beads, has significantly declined.
BASIC PROTOCOL 3
The FC-PRA is performed by using pools of panel cells (7 pools with 4 cells/pool). Patient sera are tested undiluted against the pools and the reactivity patterns are recorded. From the reaction patterns, a percent PRA can be calculated and in some instances specificities can be assigned. The nature of the pools is such that broadly cross-reactive antibodies (CREGs) can be identified rather than multiple unique specificities. The goal of the FC-PRA is to determine the presence or absence of alloantibodies in selected patients and to assign specificity, albeit broadly reactive or CREGs. Cell-based flow PRA is useful in identifying predominantly HLA class I antibodies. As such, only T lymphocyte reactivity is assessed. Materials Sera to be tested Negative controls: normal human and pooled human sera (NHS) Positive control: pooled positive serum titered for the sensitivity of the flow (PPS) Frozen cell pools, frozen using techniques that preserve maximum viability; appropriate concentration is ∼8 × 106/ml RPMI containing 20% FBS Flow wash buffer (FWB, see recipe), ice cold FITC-conjugated goat anti-human IgG [F(ab′)2, Fc specific; Jackson Laboratories] Phycoerythrin (PE)-conjugated anti-human CD3 monoclonal antibody (Becton-Dickinson), diluted according to manufacturer’s specification 1% paraformaldehyde in PBS, pH 7.2 ± 0.2 Airfuge (e.g., Beckman) with microultracentrifuge tubes and protective caps 15-ml conical tubes Colored tape Benchtop centrifuge (e.g., Beckman GP) 6-ml polypropylene tubes (e.g., Falcon) 96-well U-bottom cell culture plate (Corning-Costar) Repeat pipettor (Eppendorf) and 8-channel attachment with appropriate tips Transfer pipet (e.g., Brinkman Transferpette-12) with appropriate tips Absorbent pad
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Flow cytometer with 488-nm excitation and filters for collection of green and orange fluorescence 6 × 50–mm glass tubes (Baxter diSPo culture tubes) Additional reagents and equipment for counting cells (APPENDIX 3A) Prepare serum 1. Remove frozen patient sera (maximum of nine patients) and positive (PPS) and negative (NHS) controls. Allow to thaw completely at room temperature. If serum has not been frozen, flash-freeze 10 min in liquid nitrogen.
2. Vortex all samples. Airfuge all patient sera and controls 10 min at 28 psi. A total volume of 250 ìl per sample is needed. Vortex two microultracentrifuge tubes with 125 ìl of serum in each tube. Thaw frozen cell pools 3. For each pool, label one 15-ml conical tube with colored tape (select a different color for each pool). Record the lot number for each pool. 4. Add 1 ml RPMI containing 20% FBS to each tube. 5. For each pool, obtain the required volume of cells and place on dry ice in a styrofoam container. For a concentration of 8 × 106/ml, one vial of each pool is sufficient. If the cell concentration is less, more than one vial will be needed.
6. Thaw one pool at a time. Follow the laboratory procedure for thawing frozen cells or see APPENDIX 3B. 7. Place the thawed cells into the appropriately labeled 15-ml conical tube containing RPMI with 20% FBS. Slowly add more RPMI with 20% FBS, drop by drop until the tube is filled. 8. Repeat steps 6 and 7 for the remaining pools, one at a time. Wash the cell pools 9. Cap the tubes containing the thawed cells and centrifuge 1 min at 1000 × g, room temperature (2400 rpm in a Beckman-GP centrifuge). 10. Decant the supernatant and resuspend the cells in 10 ml RPMI with 20% FBS. 11. Repeat the centrifugation in step 9 and decant the supernatant. 12. Label 6-ml polypropylene tubes, one for each pool, with the appropriate colored tape. To each tube, add 2 ml FWB. 13. Transfer each cell preparation from the 15-ml conical tube to the appropriately labeled 6-ml polypropylene tube. 14. Check the viability of all pools and perform a cell count (APPENDIX 3A). Cleanup of the flow PRA pool cells is usually not needed, but if viability is 50% of the beads) increase in fluorescence intensity is considered positive.
Flow Cytometric Assessment of HLA Alloantibodies
ple exhibit fluorescence intensity that is significantly above the negative control? Contained within this statement are two difficult determinations. First is establishing a negative control value and range, and second is the determination of what is “significant” fluorescence above the control. Of these, the most critical determination is establishing a negative control. The negative control serum should approximate the background fluorescence of a patient sample that is devoid of HLA antibodies. For this to be true the negative serum must exhibit an average fluorescence. If the control serum underestimates average background fluorescence, then test results may be skewed towards false-positive results. In contrast, overestimating background fluorescence will lead to false-negative results. For these reasons, it is recommended that the control sera be from a pool of individuals and that two control sera be used. Once appropriate negative control sera are identified, calculation of test results can proceed. Determining a positive value is, in theory, very straightforward in that one must decide if the fluorescence value of the test serum is greater than that of the negative control. Such a determination would then indicate that HLA antibodies have bound to the target cell. For this calculation, the simplest approach is to utilize
channel values. The binding of an HLA antibody will produce an increase in fluorescence that will be measured by a channel displacement. If this displacement is significant, the test result will be deemed “positive.” How one decides what is “significant” will be dependent upon the variability of the negative control (e.g., c.v. of the negative control) as well as the sensitivity of the instrument. On average, using a 1024-channel scale, a displacement of ≥40 channels compared to the negative control may be considered as positive. Such a channel displacement, if converted to molecules of equivalent soluble fluorochrome (MESF), represents ∼1000 to 1500 MESF above background. Obviously, because of the inherent biological variability in this assay, a 40-channel displacement can be considered only a guideline. Therefore, displacements of 41 channels would not always be positive, nor would displacements of 39 channels always be negative. For values very close to the cut-point, patient history must be taken into consideration. Other approaches that may be used to represent the change in fluorescence would be (1) recording the actual linear fluorescence value and determining what a “significant” increase in fluorescence should be, and (2) converting fluorescence values to MESF. Within an indi-
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Figure 6.16.6 Class I flow-specific beads (A) and class I single-antigen beads (B) tested with patient sera containing anti-HLA antibodies. In plot A, the second bead from the top is considered positive, although it is not displaced across the line determined by the negative control. In this instance, the background fluorescence of this bead was less than that of the other beads when stained with a negative control serum. To accommodate these differences in background fluorescence, a box was drawn to delineate the position of this bead with the negative control. As shown, 100% of the beads clearly moved out of the box. (B) A positive patient serum with the specificities of anti-A2, A69, and A29. Note that the A26 bead is contained in the box, which has a fluorescence intensity that is less than that of the other beads but consistent with the negative control.
vidual laboratory the method used is not important. The most important requirement is that the laboratory set up its range and cut-point properly and perform appropriate quality control to ensure consistency in results reporting. As with the FCXM, analysis and interpretation of data for HLA detection by microparticle screening should be made by an individual who is appropriately qualified in both clinical histocompatibility and flow cytometry. Analysis of results, while not difficult, does require a good degree of expertise. The interpretation of a positive result is made by observing increases in fluorescence above background (i.e., negative control serum). In addition to a mere shift in fluorescence, the architecture of the histogram plot can also be of importance. Figures 6.16.3 and 6.16.4 show representative examples of results obtained with actual patient serum. Figure 6.16.3 shows class I and class II beads stained with either negative control serum (Fig. 6.16.3A, B) or serum from a patient who possesses both class I and class II antibodies (Fig. 6.16.3C, D). Figure 6.16.4 also shows a third
set of beads that have been coupled with human serum albumin and that serve as a control for anti-bead reactivity. The logical gate for this bead should be R7 = R1 and R6. In this example, the patient possesses antibodies against HLA class II antigens as well as an antibody that reacts with the bead plastic. Thus, the fluorescence increase seen with the class I beads is the result of an anti-bead antibody and not an HLA antibody. Failure to use a control bead may result in the incorrect reporting of HLA antibody. For each sample, both a percent PRA and antibody specificity can be determined from the flow specificity bead assay. After printing the results for a patient sample, assign the individual beads as positive or negative based on their shifts relative to the negative control markers. Any significant shift of the bead population to the right of the negative control marker is positive. A significant shift is considered as >50% of the bead population moving to the right of the negative control marker. If the population remains to the left of the negative control marker, then it is negative. Any population that
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Figure 6.16.7 Sample results from the flow cytometric PRA. Pooled lymphocytes are incubated with patient serum, then stained for IgG binding as well as for identification of CD3-positive cells. (A) Initially, a scatter gate is drawn around the apparent lymphocyte population (R1). (B) Next, a plot of FS versus CD3 is generated. From this plot, the CD3 cells are clearly delineated and an analysis gate is set (R2). A logical gate R3 = R1 and R2 is then generated and single-parameter histograms are displayed from this gate. Representative histograms are shown. (C) Results with a negative control serum. (D) Sample for which 1 cell (25% of the total events) was positive. (E) Sample for which 2 cells (50% of the total events) were positive. Depending upon the makeup of each pool and the distribution of positive and negative results, HLA specificities can be assigned.
Flow Cytometric Assessment of HLA Alloantibodies
straddles the line should be considered indeterminate until specificity is analyzed. Based on antibody assignment, the undetermined bead population may then be reassessed for positive or negative interpretation. The percent PRA may then be calculated based on the number of positive reactions divided by the panel size. Specificity analysis is based on the positive reaction pattern noted for a specific sample. Figure 6.16.5 illustrates an example of negative and positive serum samples. The quadrant marker for the negative control is placed to the right of the negative bead population. Note that the top bead (no. 1) in Figure 6.16.5 is an outlier (i.e., high background), as it appears to straddle the negative control line. To adjust for this difference, a box is drawn around this population to denote the
difference and indicate that when a patient sample is tested, these beads will be considered positive only when they have moved outside the delineated area. This type of adjustment is needed since it is not possible to control the background fluorescence of all bead preparations. Once the negative markers are established, they are not moved during the remainder of the patient analysis since they are used as points of reference for the shift in FITC fluorescence. The positive control shows that all bead populations have shifted to the right of their negative control markers, thus validating the test setup. This evaluation is repeated for each group of beads used. Figure 6.16.6 shows two illustrations of patient samples tested with the class I specificity beads (Fig. 6.16.6A) and the class I single-antigen beads (Fig. 6.16.6B).
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Figure 6.16.7 shows some representative examples of negative and positive results for flow PRA analysis. Once positive reactions have been assigned, the probability of an HLA antibody can be evaluated by knowing the HLA phenotype of the cells included in each of the seven pools.
Time Considerations The time required for any of the above flow cytometric evaluations will vary depending upon the number of samples tested. As a guideline, the flow cytometric crossmatch can be completed in ∼2.5 to 3 hr provided that all samples are ready for testing. Flow cytometric antibody screening can be performed in about the same time frame. For flow cytometric specificity testing, the time required is generally longer due to the increase complexity of set up and analysis. In general, 4 to 6 hr would be required to complete specificity testing of ∼10 serum samples.
Literature Cited Bray, R.A. 1994. Flow cytometric crossmatching in solid organ transplantation. In Methods in Cell Biology: Flow Cytometry, Vol. 41 (Z. Darzynkiewicz, H.A. Crissman, and J.P. Robinson, eds.) pp. 103-119. Academic Press, New York. Bray, R.A. and Gebel, H.M. 2000. Transplantation immunophenotyping. In Immunophenotyping: Cytometric and Cellular Analysis (C. Stewart and J. Nicholson, eds.) pp. 321-332. Academic Press, New York. Bray, R.A., Lebeck, L.L., and Gebel, H.M. 1989. The flow cytometric crossmatch: Dual-color analysis of T and B cells. Transplantation 48:834-840. Bray, R.A., Cook, D.J., and Gebel, H.M. 1997. Flow cytometric detection of HLA alloantibodies using class I coated microparticles. Hum. Immunol. 55:36. Bray, R.A, Sinclair, D.A, Wilmoth-Hosey, L., Lyons, C., Chapman, P., and Holcomb, J. 1998. Significance of the flow cytometric PRA (FCPRA) in the evaluation of patients awaiting renal transplantation. Hum. Immunol. 59:121. Chapman, J.R., Deierhoi, M.H., Carter, N.P., Ting, A., and Morris, P.J. 1985. Analysis of flow cytometry and cytotoxicity crossmatches in renal transplantation. Transplant. Proc. 17:2480. Cook, D.J., Terasaki, P.I., Iwaki, Y., Terashita, G.Y., and Lau, M. 1987. An approach to reducing early kidney transplant failure by flow cytometry crossmatching. Clin. Transplant 1:253. Garovoy, M.R., Rheinschmilt, M.A., Bigos, M., Perkins, H., Colombe, B., Feduska, N., and Salvatierra, O. 1983. Flow cytometry analysis: A high technology crossmatch technique facilitating transplantation. Transplant. Proc. 15:1939.
Gebel, H.M. and Bray, R.A. 2000. Sensitization and sensitivity: Defining the unsensitized patient. Transplantation 69:1370-1374. Gebel, H.M., Bray, R.A., Ruth, J.A., Zibari, G.B., McDonald, J.C., Kahan, B.D., and Kerman, R.H. 2001. Flow PRA to detect clinically relevant HLA antibodies. Transplant. Proc. 33:477. Harris, S.B., Bray, R.A., Josephson, C.D., Hillyer, C.D., and Gebel, H.M. 2003. Presence of HLA antibodies in blood components: An unappreciated risk factor for transplant patients? Am. J. Transplant. 3:558. Karpinski, M., Rush, D.R., Jeffery, J., Exner, M., Regele, H., Dancea, S., Pochinco, D., Birk, P., and Nickerson, P. 2001. Flow cytometric crossmatching in primary renal transplant recipients with a negative anti-human globulin enhanced cytotoxicity crossmatch. J. Am. Soc. Nephrol. 12:2807-2814. Kerman, R., Gebel, H., Bray, R., Garcia, C., Renna, S., Branislav, R., Knight, R., Katz, S., Van Burren, C., and Kahan, B. 2002. HLA antibody and donor reactivity define patients at risk for rejection or graft loss. Amer. J. Transplant. 2:258. Lazda, V.A., Pollak, R., Mozes, M.F., and Jonasson, O. 1988. The relationship between flow cytometer crossmatch results and subsequent rejection episodes in cadaver renal allograft recipients. Transplantation 45:562. Le Bas-Bernardet, S., Hourmant, M., Valentin, N., Paitier, C., Giral-Classe, M., Curry, S., Follea, G., Soulillou, J.P., and Bignon, J.D. 2003. Identification of the antibodies involved in B-cell crossmatch positivity in renal transplantation. Transplantation 75:477-482. Muller-Steinhardt, M., Fricke, L., Kirchner, H., Hoyer, J., and Kluter, H. 2000. Monitoring of anti-HLA class I and II antibodies by flow cytometry in patients after first cadaveric kidney transplantation. Clin. Transplant. 14:85-89. Ogura, K., Terasaki, P.I., Johnson, C., Mendez, R., Rosenthal, J.T., Ettenger, R., Martin, D.C., Dainko, E., Cohen, L., Mackett, T., et al. 1993. The significance of a positive flow cytometric crossmatch test in primary renal transplantation. Transplantation 56:294-298. Pei, R., Wang, G., Tarsitani, C., Rojo, S., Chen, T., Takemura, S., Liu, A., and Lee, J. 1998. Simultaneous HLA class I and class II antibodies screening with flow cytometry. Hum. Immunol. 5:313-322. Pei, R., Lee, J.-H., Shih, N.-J., Chen, M., and Terasaki, P.I. 2003. Single human leukocyte antigen flow cytometry beads for accurate identification of human leukocyte antibody specificities. Transplantation 75:43-49. Piazza, A., Adorno, D., Poggi, E., Borrelli, L., Buonomo, O., Pisani, E., Valeri, M., Torlone, N., Camplone, C., Monaco, P.I., Fraboni, D., and Casciani, C.U. 1998. Flow cytometry crossmatch: A sensitive technique for assessment of acute rejection in renal transplantation. Transplant. Proc. 30:1769-1771.
Phenotypic Analysis
6.16.23 Current Protocols in Cytometry
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Piazza, A., Poggi, E., Borrelli, L., Servetti, S., Monaco, P.L., Buonomo, O., Valeri, M., Torlone, N., Adorno, D., and Casciani, C.U. 2001. Impact of donor-specific antibodies on chronic rejection occurrence and graft loss in renal transplantation: Posttransplant analysis using flow cytometric techniques. Transplantation 71:1106-1112. Scornik, J.C., Brunson, M.E., Schaub, B., Howard, R.J., and Pfaffa, W.W. 1994. The crossmatch in renal transplantation. Evaluation of flow cytometry as a replacement for standard cytotoxicity. Transplantation 57:621-625. Scornik, J.C., Clapp, W., Patton, P.R., Van der Werf, W.J., Hemming, A.W., Reed, A.I., and Howard, R.J. 2001. Outcome of kidney transplants in patients known to be flow cytometry crossmatch positive. Transplantation 71:1098-1102. Talbot, D., Givan, A.L., Shenton, B.K., Stratton, A., Proud, G., and Taylor, R.M.R. 1988. Rapid detection of low levels of donor specific IgG by flow cytometry with single and dual color fluorescence in renal transplantation. J. Immunol. Methods 112:279-283.
Thistlethwaite, J.R., Buckingham, M., Stuart, J.K., Garber, A.O., Mayes, J.T., and Stuart, F.P. 1987. T cell immunofluorescence flow cytometry cross-match results in cadaver donor renal transplantation. Transplant. Proc. 19:722. Utzig, M.J., Blumke, M., Wolff-Vorbeck, G., Lang, H., and Kirste, G. 1997. Flow cytometry crossmatch: A method for predicting graft rejection. Transplantation 63:551-554. Vaidya, S., Cooper, T.Y., Avandsalehi, J., Barnes, T., Brooks, K., Hymel, P., Noor, M., Sellers, R., Thomas, A., Stewart D., Daller, J., Fish, J.C., Gugliuzza, K.K., and Bray, R.A. 2001. Improved flow cytometric detection of HLA alloantibodies using pronase. Transplantation 71:422-428.
Contributed by Robert A. Bray and Howard M. Gebel Emory University Atlanta, Georgia Thomas M. Ellis The Blood Center of Southeastern Wisconsin Milwaukee, Wisconsin
Flow Cytometric Assessment of HLA Alloantibodies
6.16.24 Supplement 27
Current Protocols in Cytometry
Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
UNIT 6.17
Recent advances in analytical cytometry have improved diagnostic tools for the study of erythropoiesis in anemic patients and resolution of the differential diagnosis in diseases of the erythron. This unit presents three applications of red blood cell (RBC) analysis— quantitation of fetal red cells (see Basic Protocol 1), F cell enumeration (see Basic Protocol 2), and F reticulocyte analysis (see Basic Protocol 3)—that improve diagnostic precision, sensitivity, and specificity, and provide better laboratory indicators of therapeutic efficacy in a variety of hematologic and obstetric disorders. Such advances also include the measurement and quantitation of RBC hemoglobins and their relative nucleic acid levels. These advances not only promise to improve diagnostic accuracy and laboratory precision over techniques such as the traditional manual reticulocyte counting method and the Kleihauer-Betke stain method for evaluating fetomaternal hemorrhage (FMH), but also serve as tools for newer assays of anemia diagnosis and improved clinical outcomes. In addition to the primary methods, supporting techniques for preparing spiked controls (see Support Protocol 1), setting up a fetal hemoglobin acquisition protocol (see Support Protocol 2), and assaying reticulocytes using thiazole orange (see Support Protocol 3) are also presented. QUANTITATION OF FETAL RED CELLS BY FLOW CYTOMETRY Flow cytometric methods are more sensitive and precise than routine visual counting and reliably detect low levels of fetal RBCs in circulating maternal blood. The following protocol for the detection of fetal hemoglobin (HbF)–containing RBCs uses a monoclonal antibody conjugated to fluorescein isothiocyanate (FITC), a method for the intracytoplasmic staining of red blood cells, and WinList or any other equivalent Software (Davis et al., 1998; Chen et al., 2000); however, the method can be easily adapted for use with anti-HbF monoclonal antibody reagents conjugated to other fluorochromes and with other data-analysis software.
BASIC PROTOCOL 1
Materials Whole-blood samples anticoagulated with EDTA or other suitable anticoagulant (test within 4 hr of collection or store ≤72 hr at 4°C) Fetaltrol stabilized three-level controls (Caltag Laboratories) or spiked control blood samples (see Support Protocol 1) PBS/0.1% BSA (see recipe) 0.05% glutaraldehyde (see recipe) 0.1% Triton X-100 (see recipe) Anti-HbF antibody dilution (see recipe) 1% formaldehyde fixative (see recipe) 12 × 75–mm disposable polystyrene tubes (Falcon) with rack DAC II automatic cell washer (Baxter) or equivalent (optional) Multiparameter flow cytometer with 488-nm excitation and filters for detection of green and orange fluorescence WinList Software (Verity Software House; http://www.vsh.com) or equivalent Additional reagents and equipment for counting cells (APPENDIX 3A) and acquisition of data (see Support Protocol 2) Phenotypic Analysis Contributed by Bruce H. Davis and Kathleen Thompson Davis Current Protocols in Cytometry (2004) 6.17.1-6.17.16 Copyright © 2004 by John Wiley & Sons, Inc.
6.17.1 Supplement 28
Table 6.17.1
Volume to Fix for a Given RBC Concentration
RBC X (µl) (× 106/µl)
RBC X (µl) (× 106/µl)
RBC X (µl) (× 106/µl)
5.0 4.9 4.8 4.7 4.6 4.5 4.4 4.3 4.2 4.1 4.0
3.9 3.8 3.7 3.6 3.5 3.4 3.3 3.2 3.1 3.0 2.9
2.8 2.7 2.6 2.5 2.4 2.3 2.2 2.1 2.0 1.9 1.8
5.0 5.1 5.2 5.3 5.4 5.6 5.7 5.8 6.0 6.1 6.3
6.4 6.6 6.8 6.9 7.1 7.4 7.6 7.8 8.1 8.3 8.6
8.9 9.3 9.6 10.0 10.4 10.9 11.5 11.9 12.5 12.9 13.5
1. Mix anticoagulated whole blood samples and spiked control blood samples or Fetaltrol stabilized three-level controls by rolling the tubes ten times between the hands. Place tubes on a rocker to continue mixing and allow to reach ambient temperature if chilled. Run all three controls (negative, low positive, and high positive) with each batch of patient specimens.
2. Measure the RBC count for each patient sample and each control (APPENDIX 3A). If the RBC count is >5.0 × 106 cells/µl, dilute 1:2 with PBS/0.1% BSA and count again. 3. Using Table 6.17.1, determine the volume of blood for processing (denoted by X). X contains ∼2.5 × 107 RBCs. This volume of RBCs in the fixation tube will result in ∼5 × 105 cells in the staining tube.
4. Label one 12 × 75–mm disposable polystyrene tube for each control and two for each patient sample. Place in a rack. Patient samples are run in duplicate.
Fix cells 5. Fix X µl whole blood or control in 1 ml cold 0.05% glutaraldehyde 10 min at room temperature. Vortex immediately for 15 sec at high speed. In subsequent steps, when instructed to vortex, do so using these conditions. Note that other fixatives may reduce red cell lysis.
6a. For washing with a cell washer: Load the tubes into a DACII automatic cell washer and select three wash cycles using flow cytometric isotonic sheath fluid. 6b. For washing by centrifugation: Wash cells with 2 ml PBS/BSA and centrifuge 5 min at 200 × g, at room temperature. 7. Resuspend the cell pellet by vortexing. Add 500 µl of 0.1% Triton X-100. Mix by vortexing. Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
8. Incubate 3 to 5 min at room temperature. 9. Load tubes into the DACII automatic cell washer and select one wash cycle (see step 6a). Alternatively, perform a manual wash as described (see step 6b).
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Current Protocols in Cytometry
10. Resuspend the cell pellet by vortexing. Add 500 µl PBS/BSA and mix by vortexing. 11. During the above incubation and washes, prepare tubes for the staining step. Label two 12 × 75–mm tubes for each control and three for each duplicate patient sample. A total of six tubes is labeled for each patient.
Stain cells 12. To each tube add 10 µl anti-HbF antibody dilution and 80 µl PBS/BSA. 13. Add 10 µl (∼1–2 × 105 cells) of the suspensions made in step 10 to the tubes containing Ab. Mix by vortexing. 14. Incubate 15 min in the dark at room temperature. 15. Load the tubes into a DACII automatic cell washer, select two wash cycles, and start the machine (step 6a). Alternatively, perform two manual washes (step 6b). 16. Resuspend the cell pellet by vortexing. Add 500 µl of 0.1% formaldehyde. Vortex to mix. 17. Store tubes up to 24 hours at 4°C until ready for flow cytometric acquisition. Acquire sample data 18. Perform cytometer start up and run daily quality control to verify proper operation. If all instrument checks are satisfactory, launch the acquisition protocol (see Support Protocol 2). Instrument settings must be checked and/or adjusted if there are any changes in the laser. A change in the laser will require new voltage settings for the detectors and PMTs. When selecting new voltage settings, place the negative population fully within the first decade.
19. Run tubes in the following order: Patient sample tubes Blank tube of water or buffer Fetaltrol Level 1 or negative control Fetaltrol Level 2 or low-positive (0.15%) control Fetaltrol Level 3 or high-positive (1.5%) control. If there are specimens from more than one patient, run a blank (water) for 30 sec between patient sets (six tubes) to prevent carryover. It is not necessary to run a blank between the three levels of controls since they are known levels arranged from lowest to highest. Carryover will not affect the results.
20. Collect at least 50,000 ungated events at high flow rate for each specimen. 21. Name and save listmode files. Analyze data 22. Select the HbF macro in WinList software to bring up the analysis template. 23. Gate on RBCs (R1) using the orange fluorescence versus SS parameters to exclude autofluorescent (orange positive) events of intensity greater than the first decade (Fig. 6.17.1A). 24. Using the Fetaltrol Level 3 or high-positive control files, set region markers closely around the fetal RBC peak to define the fetal cell region R4 (Fig. 6.17.1B). Include all 50,000 events by clicking on the Data Source bar.
Phenotypic Analysis
6.17.3 Current Protocols in Cytometry
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B 104
250
103
200
adult cells
150
R2 F cells
Number
Autofluorescence
A
102 R1
fetal cells R4
101
100 100
R3
100 50
101
102 103 Side scatter
104
0 100 101 102 103 104 Green fluorescence (HbF-FITC)
Figure 6.17.1 Analysis of anti-HbF stained Fetaltrol sample with 1.5% fetal cells. Listmode files are collected with parameters of light scatter, green fluorescence (anti-HbF FITC signal), and orange fluorescence (autofluorescence signal). Analysis is performed by gating events to exclude small particles and autofluorescent nucleated cells as shown as the R1 region in panel A with the R1 gated events displayed as a single fluorescence histogram of the HbF distribution as shown in panel B. The distribution of HbF in red cells typically shows three regions of adult red cells lacking HbF content (R2), adult F cells with an intermediate level of HbF (R3), and fetal red cells (if present) with high levels of HbF (R4), as shown in panel B.
25. With the R4 region marker unchanged, open the Fetaltrol Level 2 and Level 1 or low-positive and negative control files. 26. Report the % gate value for R4 as the value representing the percentage of HbF-positive cells. 27. Evaluate the control values. Results must fall within the reference (Fetaltrol) or established laboratory range (in-house controls). If they fall outside, troubleshoot the procedure and repeat the batch. 28. If the controls are within range, open the patient files. The R4 region marker will remain unchanged, but the R1 gate may need to be adjusted. Include all events. 29. Report the % gate value for R4 as the percentage of HbF-positive fetal red blood cells. Normal values for adult blood: HbF (fetal cells) ≤0.07%
Patient results which fall outside the range cannot be reported. BASIC PROTOCOL 2
Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
F-CELL ENUMERATION BY FLOW CYTOMETRY The procedure for sample staining and flow cytometric analysis is identical to that used for fetal red cell enumeration. A no-wash modification of the assay has also been reported (Mundee et al., 2000); however, it is recommended that laboratories first gain experience with the original method, which has been more extensively validated. Refer to the detailed description of the fetal red cell enumeration method (see Basic Protocol 1), paying particular attention to the instrument setup instructions on how to adjust the fluorescence detection settings such that the peak positions of cells lacking HbF and autofluorescent leukocytes are similarly positioned on both fluorescence parameters. This instrument setup then allows use of the autofluorescence signal to determine the thresholds or cursor positions for counting F cells based upon the anti-HbF antibody fluorescence with a high level of precision.
6.17.4 Supplement 28
Current Protocols in Cytometry
Data are analyzed using WinList (Verity Software House) or other equivalent software . The instrument calibration and adjustment for the quantitation of F cells result in the placement of the predominant RBC population fully within the first decade of the log fluorescence scale at an equal amount of green and orange intensity (see Fig. 6.17.2). The color compensation is adjusted so that the autofluorescence signals of reticulocytes, leukocytes, and other nucleated cells in an unstained glutaraldehyde-fixed blood sample fall on the 45° angle or line of equivalence in the green versus orange fluorescence histogram (Fig. 6.17.2A). Data analysis for F-cell enumeration on any sample is performed to initially exclude the autofluorescent nucleated cells, which are primarily leukocytes, from gates set on histograms showing the fluorescent cutoff between F cells and other erythrocytes (Fig. 6.17.2B). The orange fluorescence is used to define the cursor position or threshold for F cells with the FITC-conjugated anti-HbF antibody by finding the channel at which ≥99.8% of the cells are to the left of the cursor or ≤0.02% of the orange fluorescent cells are to the right of the cursor (Fig. 6.17.2C). The same channel position of the cursor from the orange autofluorescence histogram is then used as the cursor position on the green fluorescence histogram, thus defining the F cells as all events to the right of that cutoff position (Fig. 6.17.2D).
C 104
500
103
400 Number
Orange fluorescence
A
102
300 200
101
100
100 100
101 102 103 Green fluorescence
B
0 100
104
D 104
1100 900
HbF-
103 700 leuk
Number
Orange fluorescence
101 102 103 Orange fluorescence
102
F cells
500 300
101
RBC 100
100
101
102 103 Side scatter
104
100
102 103 101 Green fluorescence
Figure 6.17.2 Histogram setup for analysis of F cells as described above in the text. Normal values: F cells in 40 adult normals = 3.8% ± 3.6%.
Phenotypic Analysis
6.17.5 Current Protocols in Cytometry
Supplement 28
SUPPORT PROTOCOL 1
PREPARATION OF SPIKED CONTROL BLOOD SAMPLES It is imperative that multilevel assayed control samples be used as a means of assessing both the procedure and the analysis. Low staining intensity, increased F-cell levels, and improper compensation and gating can seriously impact the validity of the generated results. There are two sources of quality-control samples: Fetaltrol and in-house preparations. Fetaltrol, a stabilized three-level blood control product produced by Trillium Diagnostics (http://www.trilliumdx.com), is a convenient alternative to “home-brew” controls. It contains vials of negative, low-level positive (∼0.15% fetal RBCs), and high-level positive (∼1.5% fetal RBCs) controls, and has a 3-month shelf life. This product is used exactly like whole blood in the protocols. Alternatively, spiked control blood samples may be prepared in house following the procedure outlined here. Additional Materials (also see Basic Protocols 1 and 2) Peripheral blood from a healthy nonpregnant adult Alsever’s solution (see recipe) Umbilical-cord or newborn blood Anticoagulant (e.g., EDTA)-containing vacutainer Additional reagents and equipment for collection of EDTA-anticoagulated blood and determining blood ABO grouping Prepare negative control 1. Using an anticoagulant-containing vacutainer, collect a whole-blood sample from a healthy nonpregnant adult. 2. Take 1 ml of this normal sample and immediately add 1 ml Alsever’s solution. Mix well. 3. Measure and record the new RBC count (APPENDIX 2A). 4. Store ≤1 week at 4°C. Prepare positive controls 5. Obtain whole blood from an umbilical cord or newborn, in an anticoagulant-containing vacutainer. Determine the ABO grouping. 6. Wash cells three times with 5 ml PBS/0.1% BSA. 7. From a healthy nonpregnant adult, obtain a fresh ABO-compatible whole blood sample, anticoagulated with EDTA, with normal hematology CBC parameters. 8. Spike 1 ml adult whole blood with the washed cord cells (see step 4) to make 1.5% and 0.1-0.2% mixtures, which can serve as high and low level fetal RBC controls. 9. Immediately add 1 ml Alsever’s solution to each spiked mixture, mix well, and determine and record the new RBC count. Store ≤2 weeks at 4°C.
SUPPORT PROTOCOL 2
Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
SETTING UP A FETAL HEMOGLOBIN ACQUISITION PROTOCOL The acquisition protocol for the fetal red cell assay requires a multi-parameter instrument with a 488-nm excitation laser and the ability to collect light scatter and at least two fluorescence parameters. This procedure describes the optimal instrument setup to allow for both fetal red cells and adult F cells. Although the procedure uses only a single fluorescently labeled antibody (anti-HbF), the collection of a second fluorescent signal for autofluorescence is important for both exclusion of nucleated cells from analysis and accurate enumeration of F cells (Chen et al., 2000). Once the acquisition protocol is established and stored on the instrument as described below, it does not need to be performed again unless major service alignment adjustments, PMT replacements, or laser replacements are performed.
6.17.6 Supplement 28
Current Protocols in Cytometry
Additional Materials (also see Basic Protocol 1) Blood sample, preferably with high leukocyte count (>25 × 109 cells/liter) Fetaltrol Level 3 or adult blood spiked with washed ABO-compatible cord cells (see Support Protocol 1) 1. Select two blood samples, one preferably with a high WBC and the other either Fetaltrol Level 3, or adult blood spiked with washed ABO-compatible cord cells. 2. Stain as described (see Basic Protocol 1, steps 1 to 17) and run on the flow cytometer. 3. Draw the following histograms: Log FS versus log SS HbF-FITC (green fluorescence) versus log FS Autofluorescence (orange fluorescence) versus log FS Autofluorescence (orange fluorescence) versus HbF-FITC (green fluorescence) Number versus HbF-FITC (green fluorescence). 4. While running the tube containing the stained spiked blood (or Fetaltrol), adjust FS and SS so that the RBC population lies midscale on both axes. 5. Adjust threshold on FS to eliminate unwanted events (platelets, cell debris) with a lower signal than that of the red cell population. 6. Adjust FITC (green fluorescence) and autofluorescence (orange fluorescence) so that the red cell peaks are in the center of the first decade for both parameters and the entire peak can be visualized on the histogram. 7. While running the tube containing the stained sample with the high WBC, adjust orange minus green compensation so that the (stained) RBCs along the green fluorescence axis will fall mostly within the first decade of the orange fluorescence axis, and WBCs will line up on a 45° diagonal. 8. Set acquisition to collect at least 50,000 events in a listmode file with all parameters (FS, SS, green fluorescence, and orange fluorescence). Name and save the instrument settings and the protocol template. F-RETICULOCYTE ANALYSIS Reliable reticulocyte analysis on blood samples is considered to be the least expensive and fastest way to evaluate human bone marrow erythropoietic production. Flow cytometry is presently the preferred and most precise method for reticulocyte measurement (Davis, 2001a,b). Hence the measurement of F-cell production can be achieved by combining the monoclonal HbF assay with the thiazole orange reticulocyte assay. The cells that are the newly released precursors to mature circulating F cells are called F reticulocytes. This measurement of F reticulocytes can be used to specifically monitor the rate of F-cell production, such as in various anemias or myelodysplasia or in evaluation of new therapies in patients with sickle cell disease or other hemoglobinopathies (Nagel et al., 1993; Maier-Redelsperger et al., 1998a,b; Bohmer et al., 2000; Mundee et al., 2001). Materials Whole-blood sample, anticoagulated with EDTA or other suitable anticoagulant (test within 6 hr of collection or store ≤72 hr at 4°C) PE-Cy5 (Tri-color)–conjugated anti-HbF monoclonal antibody (Caltag Laboratories) 1% formaldehyde fixative (see recipe)
BASIC PROTOCOL 3
Phenotypic Analysis
6.17.7 Current Protocols in Cytometry
Supplement 28
Retic-Count kit (thiazole orange; BD Biosciences) Sheath fluid: filtered PBS (0.40 µm pore) Retic-COUNT solution (BD Biosciences) or an alternative thiazol orange solution in filtered PBS Multiparameter flow cytometer with 488-nm excitation and filters for collection of green and red fluorescence Additional reagents and equipment for staining blood samples (see Basic Protocol 1, steps 1 to 17) and thiozole orange reticulocyte analysis (see Support Protocol 3).
A
B 104
Autofluorescence
104
Side scatter
103 102 101
103
leukocytes
R4
102 R2
101
100 100
101 102 103 Forward scatter
104
C
101
102 103 Side scatter
104
D 104 103 102
700 leukocytes
600
R3
Number
Autofluorescence
100
101
500 400 300 200
fetal cells (R4)
100 100 100 101 102 103 104 Green fluorescence (HbF-FITC)
Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
100 101 102 103 104 Green fluorescence (HbF-FITC)
Figure 6.17.3 Representative histograms following optimal setup of protocol for analysis of fetal red cells in blood sample stained with anti-HbF monoclonal antibody. The positioning of the “negative” population of red cells lacking HbF should be within the first decade for both the anti-HbF and autofluorescence signals, and the leukocyte population with high level of autofluorescence should be positioned on a 45° diagonal. Panel A shows the typical light scatter signal of red cells. Panel B shows the higher level of autofluorescence shown by leukocytes (and other nucleated cells) gated in R1 and the red cells lacking significant autofluorescence (large cluster), which is the population gated in R2 for eventual analysis for fetal cells. Panel C demonstrates the optimal instrument setup such that the two fluorescent signals have balanced and equal intensities for the autofluorescent leukocyte or nucleated cell cluster (R3). Panel D demonstrates the HbF expression of the red cell population gated as shown in panel B with the fetal red cell region of analysis indicated by the bracketed region (R4).
6.17.8 Supplement 28
Current Protocols in Cytometry
Red fluorescence (HbF-PE-Cy5)
Red fluorescence (HbF-PE-Cy5)
104 103
102
101
100 0
10
1
2
3
10 10 10 Green fluorescence
4
10
104 103
102
101
100 100
101 102 103 Thiazole orange
104
Figure 6.17.4 F-reticulocyte analysis using dual staining for HbF and RNA content. Blood samples stained with anti-HbF (PE-Cy5) to identify F cells and non-F cells based upon level of red fluorescence are seen in both panels. The reticulocyte component of both red cell types can be calculated by comparing the green fluorescence of autofluorescence in samples without thiazole orange staining (left panel) to that seen with co-staining with thiazole orange (right panel).
Stain specimens 1. Stain two aliquots of each blood sample as described (see Basic Protocol 1, steps 1 to 17) using PE-Cy5 (Tri-color)-conjugated anti-HbF monoclonal antibody. Before beginning fixation, perform a standard thiazole orange reticulocyte analysis (see Support Protocol 3). 2. Resuspend the cell pellet in one tube by vortexing in 500 µl of 1% formaldehyde fixative. Store at 4°C shielded from light until ready to run on the flow cytometer. This tube is now ready for data acquisition.
3. Resuspend the cell pellet in the remaining tube by vortexing in 0.5 ml Retic-COUNT reagent (or an alternative thiazol orange solution) from the Retic-COUNT kit. Incubate 30 min in the dark at room temperature. Acquire data 4. Prepare a calibrated and standardized flow cytometer that has been properly quality controlled according to laboratory procedure. 5. Run the tube not exposed to thiazole orange first, while the second tube is incubating with the Retic-COUNT reagent (or an alternative thiazole orange solution). Be sure to run a blank tube containing sheath fluid after every patient specimen. 6. Run the thiazole orange/1Hb-F dual-stained tubes after the formaldehyde-fixed tubes and within 1 hr of the initial incubation with thiazole orange. Be sure to run a blank tube containing sheath fluid after every patient specimen to avoid specimen carryover. Use the formaldehyde-fixed sample as a guide to adjust cursor or quadstat positions to define regions for F reticulocytes (thiazole orange and HbF dual-positive) and reticulocytes of the non-F cell population (thiazole orange positive and HbF negative).
Phenotypic Analysis
6.17.9 Current Protocols in Cytometry
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7. Collect log FS, log SS, log green fluorescence (thiazole orange), and log red fluorescence (HbF-PE-Cy5) in listmode. Set the threshold on the FS signal and collect 50,000 events at high flow rate for each specimen. The same voltage settings for the detectors are used consistently for standardizing results. A change in the laser, the FS detector, the SS PMT, the green fluorescence PMT, or the red fluorescence PMT will require adjustment of these settings. When selecting a new voltage setting, place the negative population for both green fluorescence and red fluorescence fully within the first decade. It will be necessary to adjust the green from red compensation to achieve histogram displays as shown in Figure 6.17.4. SUPPORT PROTOCOL 3
RETICULOCYTE ASSAY USING THIAZOLE ORANGE A standard thiazole orange reticulocyte analysis is performed for each patient specimen just prior to beginning the fixation process for the HbF staining procedure. This provides assurance that the result of the F-reticulocyte assay correlates with standard reticulocyte analysis results. See Basic Protocols 1 and 3 for materials. Stain samples 1. Label two 12 × 75–mm polystyrene tubes for each patient specimen. One tube is for the stained sample and one for the unstained (control) sample.
2. Add 1 ml Retic-COUNT reagent (or alternative thiazol orange solution) to the stained sample tube and 1 ml PBS/0.1% BSA to the unstained (control) sample tube. 3. Add 5 µl well-mixed whole blood from the same specimen to each tube. Cap and vortex gently. 4. Incubate both tubes 30 min in the dark at room temperature. Samples are now ready to run on the flow cytometer.
Acquire data 5. Perform daily quality control for the flow cytometer according to laboratory procedure, ensuring the instrument is in acceptable working order. 6. Set up a protocol that collects log FS, log SS, and log green fluorescence. 7. Adjust FS and SS such that the red blood cells are in the upper right of the histogram and fully on scale. 8. Using a control tube, adjust the green fluorescence signal such that the peak is in the first decade of the single-parameter histogram (Fig. 6.17.5). If necessary, increase the FS threshold to exclude noise and debris. 9. Acquire 50,000 events from the unstained sample tube using a low flow rate. Save the listmode data. Run all unstained tubes before stained tubes. Do not change any settings. 10. Acquire 50,000 events from the stained sample tube using a low flow rate. Save the listmode data. Analyze data 11. Gate on the red cell population based on light-scatter properties. Define the reticulocyte population using the unstained sample by setting the cursor such that 0.1% of the cells lie to the right of the cursor on the thiazole orange fluorescence signal. Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
12. Use this cursor position to analyze the thiazole orange–stained sample.
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500
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Figure 6.17.5 Thiazole orange staining of blood samples for reticulocytes. Thiazole orange can label both the platelets (cluster in middle) and red cells (upper right cluster) shown by light scatter parameters in the left panel. The red-cell cluster is gated to analyze the green fluorescence distribution or RNA content of the cells, with reticulocytes being defined by autofluorescence of red cells as shown in the right panel.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Alsever’s solution Dissolve 0.50 g citric acid (C6H8O7), 20.50 g dextrose (C6H12O6), 4.20 g sodium chloride (NaCl), and 8.00 g trisodium citrate (C6H5Na3O7⋅2H2O) in ∼600 ml water. Add 0.33 g chloramphenicol, 2.00 g inosine, and 0.50 g neomycin sulfate; mix well. Adjust to 1 liter with water and sterilize using a 0.2-µm vacuum filter. Adjust pH to 7.2, using aliquots to determine pH and aseptic technique to avoid contaminating the filtered solution. Store ≤3 months at 4°C; discard if any visible growth is detected. Anti-HbF antibody dilution FITC-conjugated anti-HbF antibody (Caltag Laboratories or Maine Biotechnology Services): use 5 or 10 µl undiluted antibody per test as determined by titration. Anti-HbF optimal concentration is usually ∼2.5 µg/ml, but it is determined by examining a series of dilutions (two fold dilutions between 0.1 and 10 µg/ml) with samples with a mixture of fetal red cells and adult red cells containing at least 1% fetal red cells. The antibody concentration should be sufficient to provide the optimal fluorescence separation between the negative adult RBCs and the positive fetal cells. Additionally, the concentration should be sufficient to have no change in fetal red cell fluorescence intensity for samples up to 10% fetal cells. Glutaraldehyde, 0.05% 50 µl 25% glutaraldehyde (Sigma; store at −20°C) 25 ml PBS, pH 7.4, without BSA (CellGro or Sigma) Mix well and keep at 4°C Make fresh working solution each day just prior to use and use cold Stability past 2 hr is uncertain
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Formaldehyde fixative, 1% Combine 15 ml of 10% formaldehyde (methanol free; Polysciences) and 135 ml PBS/0.1% BSA (see recipe). Sterilize using a 0.2-µm vacuum filter. Adjust pH to 7.4 when solution reaches room temperature, using aliquots to measure pH and aseptic technique to avoid contaminating the filtered solution. Store ≤1 week at 4°C. PBS/BSA, 0.1% 1 packet phosphate-buffered saline powder (Sigma; available through Diamedix) 1000 ml H2O 1.0 g BSA (Sigma) Mix well and sterilize using a 0.2-µm vacuum filter Adjust pH to 7.4 when solution reaches room temperature Store ≤1 week at 2° to 8°C Triton X-100, 0.1% 50 µl Triton X-100 (Sigma) 49.95 ml PBS/0.1% BSA (see recipe) Mix well and store ≤1 month at 4°C Discard at detection of visible growth Use cold COMMENTARY Background Information
Enumeration of Fetal Red Blood Cells, F Cells, and F Reticulocytes in Human Blood
Fetomaternal hemorrhage and fetal RBC detection The most important clinical use of fetal RBC detection is the diagnosis and quantitation of fetomaternal hemorrhage (FMH; Sebring and Polesky, 1990; Davis, 2001a,b ; Davis et al., 2001). FMH occurs normally throughout pregnancy in minute amounts, with increasing volumes during the later stages of gestation (Giacoia, 1997). Any significant difference in the RBC antigenicity between fetus and mother can result in allosensitization of the maternal immune system either before or after parturition. The maternal antibodies to the fetal RBC antigens may be clinically silent or cause lifethreatening autoimmune sequelae for current or subsequent pregnancies (e.g., erythroblastosis fetalis, early abortion). Such sensitization can occur with any RBC antigen mismatch, but the highest frequency and profound clinical consequences occur with Rh or D-antigen mismatches. Detection and enumeration of fetal RBCs is an essential part of the management of those patients with FMH treated with Rh immune globulin (RhIG) preparations, such as RhoGam (Ortho Pharmaceutical; Polesky and Sebring, 1981; Sebring and Polesky, 1990). The use of Rh immune globulin prophylaxis is a universal practice, but dosing amounts and schedules have regional variations (Hartwell, 1998; Lee et al., 1999). Hence, the sensitivity
and specificity of detection assays for FMH is a critical factor in therapeutic efficacy and subsequent clinical outcome. Heretofore, the most widely used assay for FMH detection has been the visual microscopic counting Kleihauer-Betke (KB) method, which is based upon the differences in solubility properties in acid conditions between fetal hemoglobin (HbF) and adult hemoglobin (Kleihauer et al., 1957). The KB technique is easily performed by most clinical laboratories, but lacks sensitivity and exhibits poor reproducibility and precision (CVs of 50% to 100%). The problematic nature of the KB assay was demonstrated in the recent proficiency surveys (HBF) conducted by the College of American Pathologists, in which two surrogate blood samples were selected at values either side of the 0.6% fetal RBC level, indicative of a FMH of ∼30 ml, typically used in the U.S. as the trigger level for additional Rh immune globulin therapy. The performance by laboratories using the KB method in this survey was less than optimal for a laboratory test used for therapeutic monitoring, as ∼12% of labs reported results of ≤0.6% for the sample with >0.8% fetal RBC and >45% of labs reported results of ≥0.6% for the sample with ≤0.4% fetal RBC. Such performance in this laboratory survey indicates that clinical practice using the KB assay is likely associated with a significant amount of both under- and over-administration of Rh immune globulin to patients with signifi-
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cant FMH. Similar reported experiences with the KB assay have also brought into question the clinical validity of this manual method (Emery et al., 1995; Ducket and Constantine, 1997). Flow cytometric methods have recently been developed that improve both sensitivity and precision by using monoclonal antibodies to HbF, i antigen, and D antigen (Nance et al., 1989; Thorpe et al., 1994; Lloyd-Evans et al., 1996; Davis et al., 1998; Navenot et al., 1998; Nelson et al., 1998; Campbell et al., 1999; Chen et al., 2002). These methods reliably detect low levels of fetal RBCs in maternal circulation (560
547-563
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a Green fluorescence of AO-DNA has a long tail toward higher wavelengths. b Value depends on base composition.
Nucleic Acid Analysis Contributed by Zbigniew Darzynkiewicz, Gloria Juan, and Edward F. Srour Current Protocols in Cytometry (2004) 7.3.1-7.3.16 C 2004 by John Wiley & Sons, Inc. Copyright
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BASIC PROTOCOL 1
DIFFERENTIAL STAINING OF DNA AND RNA OF UNFIXED CELLS WITH ACRIDINE ORANGE This protocol describes the application of AO to differential staining of DNA and RNA. The cells to be stained with AO can be either prefixed in ethanol (see Alternate Protocol 1) or permeabilized with the nonionic detergent Triton X-100 as described here. The permeabilization is done at low pH in the presence of serum proteins. At low pH, most histones dissociate from DNA, making DNA more accessible to AO and thereby improving stoichiometry and accuracy in DNA detection (Darzynkiewicz et al., 1984b). Following treatment with the permeabilizing solution, the cells are stained with AO dissolved in phosphate–citric acid buffer. The high molarity and excess of the buffer neutralize the acid maintaining the low pH in the first step. In the presence of EDTA, AO at the proper concentration interacts with cellular RNA to form condensed complexes that luminesce red, with maximum emission above 630 nm. At the same time, interactions of AO with DNA result in green fluorescence. Thus this metachromatic fluorochrome differentially stains RNA and DNA (see Support Protocol).
Materials Cells to be stained (APPENDIX 3B): ≤106 cells/ml suspended in tissue culture medium containing 10% (v/v) serum or 1% (w/v) BSA Cell permeabilizing solution (see recipe), ice cold Acridine orange (AO) staining solution (see recipe), ice cold Flow cytometer equipped either with a 488- or 457-nm argon-ion laser (or both lasers) or with a mercury arc lamp NOTE: In performing the following steps, carefully avoid pipetting, vortexing, or any mechanical agitation of cells, to prevent cell lysis. 1. Set up the flow cytometer with excitation at 488 nm, using emission filters and a dichroic mirror that discriminate green fluorescence (measured at 530 ± 15 nm) and red luminescence (measured preferably above 640 or 650 nm). Maximum absorption by AO occurs at ∼455 to 490 nm. The 488-nm line of the argon ion laser is the most commonly used excitation wavelength. In instruments having a mercury or xenon lamp, blue excitation filters can be used (for example, a BG 12 short-pass filter transmitting below 470 nm or band-pass combination fitters transmitting between 460 and 500 may be used). Optimal excitation can be achieved using two lasers, one tuned to 488 nm (DNA detection, green fluorescence) and another to 457 nm (RNA detection, red luminescence).
2. Transfer a 0.2-ml aliquot of the original cell suspension to a small glass or plastic tube (e.g., 2- or 5-ml volume). Chill on ice. The 0.2-ml aliquot should have ≤2 × 105 cells suspended in tissue culture medium containing 10% (v/v) serum or 1% (w/v) BSA. The serum or BSA protects cells from lysis by detergent in step 3.
3. Gently add 0.4 ml ice-cold cell permeabilizing solution. Wait 15 sec, keeping cells on ice. 4. Gently add 1.2 ml ice-cold AO staining solution. Keep cells on ice in the dark. 5. Measure and record cell fluorescence in the flow cytometer during the 2 to 10 min after addition of AO staining solution. Differential Staining of DNA and RNA
The sample should be kept on ice prior to and during the measurement. Vortexing or syringing cells in the permeabilizing solution, especially in the absence of any serum or proteins in the original cell suspension, results in disintegration of plasma membranes and isolation of cell nuclei. The RNA content of isolated nuclei, therefore, can be measured
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after plasma membrane disruption in this way. Visual inspection of the nuclei under phasecontrast or UV light microscopy is essential to estimate the efficiency of the isolation, which can be controlled by selecting either an optimal time and speed of vortexing or an optimal number of syringings. DNA frequency histograms (green fluorescence) can be deconvoluted to obtain the proportion of cells in G1 versus S versus G2 /M (see UNIT 7.5).
DIFFERENTIAL STAINING OF FIXED CELLS WITH ACRIDINE ORANGE In the instances when cells have to be fixed (e.g., for storage or transportation), staining with AO is done on fixed cells according to the following protocol.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Cells to be stained PBS (APPENDIX 2A), ice cold 70% ethanol, ice cold Centrifuge, 4◦ C Additional reagents and equipment for trypsinizing adherent cells (UNIT 5.2 or APPENDIX 3B) or dissociating cells from tissues (UNIT 5.2) 1a. For cells in suspension culture or hematologic samples: Rinse cells once with ice-cold PBS and suspend in ice-cold PBS at ∼106 cells/ml. 1b. For cells attached to tissue culture plates: Collect cells from flasks or petri plates by trypsinization, pool the trypsinized cells with cells floating in the medium (mostly detached mitotic and dead cells), and rinse once with medium containing serum to inactivate the trypsin (see UNIT 5.2 or APPENDIX 3B for details of this procedure). Suspend cells in ice-cold PBS at ∼106 cells/ml. Other means of trypsin inactivation such as addition of protease inhibitors may also be used.
1c. For cells isolated from solid tumors: Rinse cells free of any enzyme used for cell dissociation and suspend in ice-cold PBS at ∼106 cells/ml. The final cell suspension should be well dispersed (no aggregates), with a density ≤5 × 106 cells/ml. The cells should not be stored on ice for longer than 30 min before fixation.
2. With a Pasteur pipet transfer 1 ml cell suspension to a 15-ml conical glass tube containing 10 ml ice-cold 70% ethanol. Fix cells ≥2 hr on ice. To minimize cell clumping, rapidly injecting the cell suspension into the fixative, rather than layering onto the surface and then mixing, is preferred. The reverse order (i.e., addition of ethanol to cell suspensions) results in more extensive cell loss due to cell adherence to the glass surface and aggregation. The time of fixation in ethanol may vary, but at least 2 hr should be allowed for cells to be fixed. Cells may also be stored in ethanol at 4◦ C for several months.
3. Centrifuge tubes 5 min at 300 × g, 4◦ C. Remove all ethanol, rinse cells once with ice-cold PBS, and suspend in ice-cold PBS at a density of 20 nsec (Darzynkiewicz and Kapuscinski, 1990). These spectral properties of AO in complexes with ss nucleic acids are suggestive of intersystem crossing (phosphorescence; T1 to S0 transition) rather than fluorescence. The solidstate nature of the complexes, similar to freezing AO in solution, may facilitate the intersystem crossing by partially eliminating collision quenching, which otherwise occurs due to the long lifetime of the T1 excited state in the presence of oxygen and solvents. In the cell, large sections of rRNA and tRNA have ds conformation. Therefore, to obtain differential staining of DNA versus RNA with AO, these sections have to be selectively denatured, under conditions in which
DNA still remains double stranded. This is accomplished by treatment of cells with AO in the presence of the chelating agent EDTA (Darzynkiewicz et al., 1976). By breaking RNA-protein interactions in ribosomes that stabilize ds RNA, EDTA promotes denaturation of ds RNA, which occurs as a result of interaction with AO. The RNA-selective denaturing properties of AO result from the fact that this ligand has higher affinity to ss RNA than ss DNA (Kapuscinski and Darzynkiewicz, 1989). The denaturation itself results from the fact that, at increased dye/phosphate (D/P) ratio, binding of AO to ss nucleic acids becomes thermodynamically preferable; the weaker but more numerous 1:1 (D/P) interactions of AO with ss sections thermodynamically dominate the stronger but fewer (1:4) sites of AO intercalation to ds regions (Kapuscinski and Darzynkiewicz, 1989). Thus, increasing D/P promotes denaturation of ds RNA sections. Cell staining with AO is generally done in salt solutions of relatively high ionic strength (0.1 to 0.2 M NaCl). Because of the significant electrostatic component in binding of AO to nucleic acids, competitive interactions between the AO+ and Na+ result, and it is the concentration of free AO in solution (rather than absolute D/P calculated on the basis of molar ratios of the dye and nucleic acid in the sample) that is of importance for selective RNA denaturation (Darzynkiewicz and Kapuscinski, 1990). To recapitulate, AO has two distinct functions in the mechanism of metachromatic staining of RNA, namely, it (1) denatures ds RNA and (2) differentially stains RNA (after its conversion to ss form) versus DNA. At a given ionic strength, selective RNA denaturation can be achieved only within a narrow concentration range of free dye. Too low an AO concentration produces incomplete RNA denaturation (both DNA and portions of RNA then stain green), whereas too high a concentration also leads to denaturation of DNA (both RNA and DNA stain red). Thus acridine orange, in contrast to most other dyes used in cytometry, requires very stringent conditions, especially with regard to concentration of AO and ionic strength of the staining solutions. The protocol of cell staining described in Basic Protocol 1 and Alternate Protocol 1 has been established after testing a variety of ionic conditions, pH, and dye concentrations (Darzynkiewicz et al., 1976; Traganos et al., 1977).
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Differential staining with PY–Hoechst 33342 Historically, PY has been widely used in absorption microscopy as a dye that in combination with methyl green specifically stains cellular RNA (Scott, 1967). More recently, it also found an application in flow cytometry as a fluorochrome of RNA (Tanke et al., 1980; Shapiro, 1981). The interactions of PY with nucleic acids, which are responsible for its specificity to RNA, as in the case of AO (see above) also are complex (Kapuscinski and Darzynkiewicz, 1987). The following binding and spectral characteristics of PY are of importance for its role as an RNA fluorochrome: 1. Intercalary mode of binding. PY binds by intercalation to ds nucleic acids. Its binding affinity to ds RNA is severalfold higher than to ds DNA. In the intercalated form, whether bound to RNA or DNA, the dye has maximal absorption between 547 and 563 nm and fluoresces with maximum emission between 565 and 574 nm; the variation is due to differences in base composition of the nucleic acids (Kapuscinski and Darzynkiewicz, 1987). Its quantum yield also varies widely with changes in base composition. In contrast to AO, which stains total cellular RNA, PY used as fluorochrome can detect only ds sections of RNA and is sensitive to the AU/GC base ratio. 2. Condensation and precipitation of nucleic acids. Like AO binding, binding of PY to ss nucleic acids results in condensation (precipitation) of the product. The mechanism of condensation and the structure of the complexes are similar to those generated by AO. In contrast to AO, however, PY fluorescence is nearly totally quenched in these complexes (Kapuscinski and Darzynkiewicz, 1987). In absorption microscopy these products are characterized by lavender color. 3. Stoichiometry of RNA detection. The stoichiometry and thermodynamics of binding of PY to ds and ss nucleic acids are similar to those of AO. Thus, PY can denature the ds sections of nucleic acids, rendering them single-stranded and causing their condensation and agglomeration (Kapuscinski and Darzynkiewicz, 1987). At increasing PY concentration, therefore, the fluorescence of PY bound to ds RNA is suppressed because of the progressive denaturation of the ds sections (“self-extinguishing” effect of PY). Similarly, as in the case of AO, selective stainability of RNA with PY can be obtained at only a relatively narrow range of dye concentrations. 4. DNA stainability with PY. PY, having affinity to ds DNA, can also stain DNA.
Its binding to DNA, however, can be suppressed by DNA-specific ligands such as methyl green and Hoechst 33342 (Tanke et al., 1980; Shapiro, 1981). In the presence of these dyes, therefore, PY can be used as a specific RNA fluorochrome. Dual cell staining with Hoechst 33342 and PY as described in Basic Protocol 2 provides the basis for simultaneous detection of DNA and RNA in flow cytometry (Shapiro, 1981; Darzynkiewicz et al., 1987). Comparison of AO and PY–Hoechst 33342 methods Each method is characterized by different advantages and limitations that should be considered. Quantitative aspect of the methods. The stoichiometry of RNA measurement is better assured by the AO methodology (Bauer and Dethlefsen, 1981), because total RNA content is stained by AO, and because there is less variation in quantum yield resulting from differences in base composition or conformation of RNAs than in the case of PY. Differences in sensitivity of RNA detection. Because there is significant spectral overlap between AO bound to DNA and AO bound to RNA, the ability to detect small amounts of RNA under standard measuring conditions is better provided by the PY–Hoechst 33342 technique. Excitation of AO-stained cells at two different wavelengths, however, raises the sensitivity of the AO method to or above that of PY. Quantum yields and fluorescence intensities of cells stained with AO are higher than with PY. Specificity in DNA content measurement. Hoechst 33342 is a more specific DNA stain than AO; the latter stains glycosaminoglycans (Darzynkiewicz and Kapuscinski, 1990) as well as DNA and RNA. Resolution of DNA measurements by AO in cells containing excessive amounts of glycosaminoglycans (primary fibroblasts, mast cells, and keratinocytes) is therefore low. Analysis of RNA conformation. Both AO and PY can be used to reveal changes in the conformation of RNA. The PY–Hoechst 33342 technique detects changes associated with assemblage of polyribosomes (Traganos et al., 1988), whereas the AO method can be applied to measure the degree of double strandedness of RNA and its sensitivity to heat denaturation (melting profile). Instrument requirements. The AO methodology (Basic Protocol 1, Alternate Protocol 1) has the advantage of requiring simpler and less costly instruments. The protocol can be
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performed with excitation at a single wavelength or with a mercury lamp instead of a laser. RNA content standards RNA and DNA content measured in any cells can be expressed quantitatively by comparison with the content of standard, calibrated cells. Nonstimulated peripheral blood lymphocytes appear to offer the best standard. RNasetreated and untreated lymphocytes should be measured to establish the extent of RNasespecific red luminescence. The cells to be compared have to be measured under conditions identical to those used for the lymphocytes. Their RNA index should be expressed as a multiplicity of the lymphocyte RNA content. The lymphocytes may also serve as a calibrator of DNA content, to estimate the DNA index from the mean (modal) intensity of the green fluorescence of the G0 /G1 population.
Differential Staining of DNA and RNA
Applications of RNA analysis In the majority of cell types, ∼80% of total cellular RNA is rRNA. Most of the remaining RNA is tRNA, and only a minor fraction of total RNA is mRNA. The content of total cellular RNA, therefore, is primarily an indicator of the number of ribosomes per cell and reflects cell translational potential. Nuclear RNA, being predominantly pre-rRNA, also is associated with cell capacity to synthesize proteins, and its increase often precedes the buildup of ribosomal machinery in the cytoplasm. RNA content measurement, either of the whole cell or of the isolated nucleus, is a marker of the overall translational capacity of the cell. Differences in RNA content between individual cells have two different origins. One is associated with the tissue type– or cell differentiation–related constitutive level of the translational activity. Thus, cells known to secrete, or produce for internal use, large quantities of tissue-specific protein (e.g., plasma cells or neurons) are generally characterized by high RNA content. In these cases the cellular RNA content is a reflection of a phenotype of the differentiated cell. Its measurement, therefore, can discriminate between cells of different tissues in the sample or be a marker of cell differentiation. The most common application of RNA measurement, in this context, is to identify reticulocytes, the immature red blood cells which retain RNA; in contrast, mature red blood cells have no measurable RNA (Tanke et al., 1980). The second cause of variability in RNA content is related to cell reproduction. Divid-
ing cells double their constituents, including the number of ribosomes, during the cell cycle. Progression through the cell cycle is thus associated with an increase in cellular RNA content, which occurs throughout interphase at a relatively constant rate proportional to the rate of cell proliferation (Darzynkiewicz et al., 1984a). Cellular RNA content, therefore, is a reflection of cell maturity in the cycle, and as such allows the discrimination of earlyG1 from late-G1 cells (Darzynkiewicz et al., 1980). Because growth in cell size (number of ribosomes) and the rate of proliferation are generally coupled, the RNA parameter, therefore, is indirectly a marker of cell proliferation. Noncycling cells withdrawn from the cell cycle (quiescent cells, G0 , G1Q ) have on average 5- to 10-fold fewer ribosomes than their cycling counterparts (Johnson et al., 1974). Thus, the difference in RNA content allows one to identify the noncycling cells and can be used as a marker of their mitogenic stimulation (Darzynkiewicz et al., 1976). Tumorous transformed cells, on the other hand, even noncycling ones, are characterized by an RNA content significantly higher than that of normal quiescent cells (Stanners et al., 1979). There is a growing body of evidence that the RNA content of tumor cells has prognostic value in many malignancies (reviewed by Darzynkiewicz, 1988).
Critical Parameters and Troubleshooting Differential staining with AO 1. Preservation of intact cells during permeabilization with Triton X-100. Disintegration (lysis) of unfixed cells during permeabilization by Triton X-100 is prevented by the presence of serum or serum albumin (Darzynkiewicz et al., 1976). It is recommended, therefore, to have the cells suspended in PBS or culture medium that contains 10% to 20% (v/v) serum or 1% (w/v) bovine serum albumin prior to addition of the permeabilizing solution. Furthermore, vigorous shaking, pipeting, or vortexing cell suspensions after addition of the detergent should be avoided (Darzynkiewicz et al., 1976). 2. Critical concentration of AO. Differential staining of RNA versus DNA requires a proper concentration (∼20 mM) of free (unbound) AO in the final staining solution as well as at the time of fluorescence measurement, i.e., at the moment of cell intersection with the laser beam in the flow cytometer. The
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Figure 7.3.1 Differential staining of RNA and DNA. Lymphocytes were analyzed (A) 0 hr, (B) 16 hr, (C) 24 hr, and (D) 48 hr after stimulation with phytohemagglutinin (PHA). The cells were stained with AO according to Basic Protocol 1. Insets represent DNA content frequency histograms (green fluorescence) of the respective cell populations.
following problems associated with this requirement may occur: (a). When the cell number (density) in the original suspension exceeds 2 × 106 cells/ml (or even less when cells are highly hyperdiploid and/or have excessive RNA content), the amount of bound AO is high and therefore the free dye concentration may be significantly reduced (the “mass action” law). RNA denaturation is then incomplete, and some RNA can stain green. In such cases, dilute the cell suspension to obtain a lower concentration. (b). With some instruments (e.g., most cell sorters) in which cell measurements take place in air outside the nozzle, a significant diffusion
of dye from the sample to the sheath fluid takes place before the cell reaches the laser beam (see Fig. 1.1.6). This breaks the equilibrium and lowers the actual AO concentration in the sample at the time of cell measurement. Dye diffusion is also a problem in some instruments that have narrow sample streams and long flow channels (e.g., Cytofluorograf 50 made by Ortho Diagnostics). The solution is to increase the AO concentration in the AO solution (up to 20 µg/ml) and to increase the sample flow rate to compensate for the diffusion. Wherever possible, use channels with favorable geometry (wider sample stream and/or shorter distance between the nozzle and intersection with
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Figure 7.3.2 Protocol 2.
Differential Staining of DNA and RNA
Differential staining of RNA and DNA with PY–Hoechst 33342 as described in Basic
the laser beam). The optimal dye concentration for a particular instrument can be established by preparing a series of staining solutions with increasing AO concentrations (e.g., from 5 to 20 µg/ml) and determining the concentration at which cells in G0 /G1 cell cluster have the same green fluorescence (the lowest coefficient of variation [CV] of the green fluorescence mean value, corresponding to a lack of correlation between green and red luminescence). On the bivariate DNA/RNA cytograms (see Figs. 7.3.1 and 7.3.2), the G0 /G1 cell cluster ought to be horizontal (or vertical if axes are reversed) but never skewed (diagonal). 3. Overlap of red and green emission spectra of AO. One of the limitations of the AO technique is the relatively low sensitivity of RNA detection. This is primarily due to overlap of the emission spectra: the green fluorescence of AO intercalated to DNA has a long tail toward higher wavelengths. Therefore, RNA measurements in cells (or cell nuclei) characterized by a high DNA/RNA ratio lack sensitivity, being obscured by the high component of AO bound to DNA. There are two ways to improve the sensitivity of RNA measurements: (1) Use long-pass filters transmitting above 640 or 650 nm, rather than 610 or 620 nm, to measure red luminescence. This significantly reduces the DNA-associated spectral component. (2) When possible use two lasers for excitation, one tuned to 457 nm to excite the red luminescence and another tuned to 488 nm to excite the green fluorescence of AO. 4. Contamination of sample tubing by AO. Like other cationic, strongly fluorescing fluorochromes (e.g., rhodamine 123), AO is adsorbed to surfaces of the sample flow tubing. Its subsequent release from the tubing interferes with later sample measurement, especially with cells of low fluorescence intensity.
To avoid this problem, after measuring samples stained with AO and prior to measuring samples stained with other fluorochromes, rinse the sample flow line with bleach (e.g., 10% Clorox), then 50% ethanol, then PBS, each for 10 min. Alternatively, if possible, have some sample flow tubing dedicated exclusively to AO use. Differential staining with PY–Hoechst 33342 In many respects the critical points of cell staining are similar for PY and AO. The most crucial aspect of the PY–Hoechst 33342 methodology is a stringent requirement for the appropriate PY concentration. Too low a concentration of the dye (or too dense a cell suspension) cannot ensure stoichiometry of staining because of the paucity of PY in the solution (i.e., dye/binding site ratio 600 nm) filter is recommended for detecting PI emission.
10. Measure cell fluorescence in the flow cytometer. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. Analyze the data (Fig. 7.5.1) using DNA content frequency histogram deconvolution software. DNA CONTENT ANALYSIS OF FIXED CELLS WITH DAPI This protocol is similar to Basic Protocol 1, except the cells are stained with DAPI rather than PI. Thus, there is no need for cell treatment with RNase. Excitation of DAPI, however, requires a UV light source, which is not universally available. Emission of DAPI is measured at blue wavelengths.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Cells to be stained DAPI/Triton X-100 staining solution (see recipe), freshly made Flow cytometer with UV light illumination source (e.g., mercury arc lamp, laser tuned to 340 to 380 nm) Fix cells with ethanol 1. Fix cells in 70% ethanol (see Basic Protocol 1, steps 1 to 5). Stain cells with DAPI 2. Centrifuge the ethanol-suspended cells 5 min at 200 × g. Decant ethanol thoroughly. 3. Suspend the cell pellet in 5 ml PBS, wait 60 sec, and centrifuge again, 5 min at 200 × g. 4. Suspend cell pellet in 1 ml DAPI/Triton X-100 staining solution. Keep 30 min in the dark. Perform flow cytometry 5. Set up and adjust flow cytometer for UV excitation at 340 to 380 nm and detection of DAPI emission at blue wavelengths. For excitation, an UG 1 optical filter (short-pass, 390 nm) may be used when the light source is a mercury arc or xenon lamp. For detecting DAPI, a band-pass filter at 470 ± 20 nm is recommended.
6. Measure cell fluorescence in the flow cytometer. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. Analyze the data (Fig. 7.5.1) using software that deconvolutes DNA content frequency histograms.
Nucleic Acid Analysis
7.5.3 Current Protocols in Cytometry
BASIC PROTOCOL 2
DNA CONTENT ANALYSIS OF SAMPLES UTILIZING DETERGENTS AND TRYPSIN This protocol uses detergent-based lysis and staining solutions, which improve DNA content analysis by flow cytometry compared to staining of intact cells as in Basic Protocol 1 and Alternate Protocol 1. This protocol is suitable for tissue samples, whereas Alternate Protocol 2 provides a simplified detergent-based method designed for fixed cells or cells from suspension cultures. Cells are collected via aspiration from tissue samples into a sucrose-citrate buffer that contains DMSO, which allows for long-term storage of samples if needed. After samples are supplemented with an internal DNA standard (a mixture of chicken and trout erythrocytes), cells are lysed, digested with trypsin, and stained with PI. The stained nuclei are then subjected to flow cytometry. Materials Tissue sample with cells to be stained (e.g., resected tumor) Citrate/DMSO buffer (see recipe) Internal DNA content standard (e.g., chicken or trout erythrocytes; see recipe) Cell lysis solution with trypsin (see recipe) Trypsin-inactivating solution (see recipe), ice cold Propidium iodide (PI)/spermine staining solution (see recipe), ice cold 0.5 × 25–mm needles (25 G × 1 in.) 10-ml disposable syringes 38 × 12.5–mm polypropylene tubes with caps 24- to 34-µm nylon mesh Flow cytometer with 488-nm argon ion laser fluorescence excitation source Collect cells from tissue samples 1. Insert the needle into the tissue sample and apply suction by syringe. Move the needle several times forward and back in different directions. Do not draw the aspirate into the syringe: keep the material within the needle. The 1-in. needle should be adequate for collecting cells in vitro from resected tumors. Longer needles may be needed for aspirating cells in vivo. Alternatively, specimens may be frozen in dry ice, stored at −60° to −80°C, thawed to room temperature, and then aspirated.
2. Flush the aspirate plug with 200 µl citrate/DMSO buffer into a 38 × 12.5–mm polypropylene tube. The tubes may be capped and mechanically shaken to further disperse the cells.
3. Count the cells in a hemacytometer. Repeat aspirations (steps 1 and 2) until 106 cells per sample are obtained. The samples may be analyzed the same day (steps 5 to 11) or stored at −40° to −80°C for up to several years. The DMSO in the citrate cell collection buffer protects cells during prolonged storage.
Lyse cells and stain with PI 4. Thaw stored samples in a 37°C water bath and transfer to room temperature. DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis
5. Add the internal DNA content standard. The mixture of chicken and trout erythrocytes used as the internal DNA content standard (chicken-to-trout cell ratio 3:5), which may be thawed to room temperature in a 37°C water
7.5.4 Current Protocols in Cytometry
bath after retrieval from −40° to −80°C storage, if necessary, is added to the 200 ìl sample of tumor cells in a proportion of ∼1:5 with respect to the number of studied cells.
6. Add 1.8 ml cell lysis solution with trypsin to 200 µl of tumor aspirate cells in citrate/DMSO buffer. Cap the tubes and incubate 10 min at room temperature, mixing gently by inverting the tubes five to ten times. 7. Add 1.5 ml ice-cold trypsin-inactivating solution. Mix gently 10 min at room temperature. 8. Add 1.5 ml ice-cold PI/spermine staining solution. Wrap tubes in aluminum foil or keep them in the dark, on ice. Mix gently and filter the sample through 24 to 34-µm nylon mesh to remove debris and cell aggregates. 9. Store cells on ice. Measure cell fluorescence between 20 min and 4 hr after adding the staining solution. Perform flow cytometry 10. Set up and adjust flow cytometer to provide excitation with blue light and detection of PI emission at red wavelengths. For excitation, the 488-nm argon ion laser line can be used. Use a BG 12 optical filter (short-pass, 470 nm) when the source of illumination is mercury arc or xenon lamp. For detecting emission of PI, a long-pass (>600 nm) filter is recommended.
11. Measure cell fluorescence in a flow cytometer. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. Analyze the data (Fig. 7.5.1) using software that deconvolutes DNA content frequency histograms. DNA CONTENT ANALYSIS OF SAMPLES UTILIZING DETERGENTS This protocol uses detergent only, rather than detergent with trypsin as in Basic Protocol 2, to lyse cells and aid staining of DNA for flow cytometric analysis. In this simplified method, cells in suspension are mixed with staining solution that contains DAPI, buffers, and Triton X-100; the DNA content of the stained nuclei is then measured by flow cytometry, using UV excitation. This method also allows simultaneous analysis of DNA and protein if the protein-specific dye sulforhodamine 101 is included in the staining solution.
ALTERNATE PROTOCOL 2
Additional Materials (see also Basic Protocol 2) Cells to be stained DAPI/PIPES staining solution (see recipe) Flow cytometer with UV light illumination source (e.g., mercury arc lamp, laser tuned to 340 to 380 nm) Additional reagents and equipment for preparing cell suspensions from tissue cultures (UNIT 7.4) Stain cells with DAPI 1. Mix 0.2 ml cell suspension (105 to 106 cells) with 2 ml DAPI/PIPES staining solution. Keep the sample ≥10 min on ice. Cells used in this protocol may be collected directly from tissue culture flasks or plates (UNITS 7.3 & 7.4) and suspended in PBS to ∼5 × 106 cells/ml. In addition, cells may be fixed ≥2 hr in 70% ethanol (see Basic Protocol 1), then rinsed and resuspended in 0.2 ml PBS. The advantage of ethanol fixation is that it offers the possibility of sample storage or transport prior to analysis.
Nucleic Acid Analysis
7.5.5 Current Protocols in Cytometry
The DAPI/detergent staining solution may be supplemented with sulforhodamine 101 (0.02 mg/ml final concentration in the staining solution) to allow simultaneous measurement of DNA and protein. Fixed cells should be used for analysis of protein content. Regardless of the dye(s) used, cell fluorescence should be measured within 10 to 60 min after staining.
Perform flow cytometry 2. Set up the flow cytometer for UV excitation at 340 to 380 nm and detection of DAPI fluorescence at blue wavelengths. For UV excitation, use an UG 1 optical filter when the source of excitation is mercury arc or xenon lamp. For detecting DAPI emission, a band-pass filter at 470 ± 20 nm is recommended. Fluorescence of sulforhodamine (which like DAPI also is excited with UV light) is at red wavelengths > 600 nm.
3. Measure cell fluorescence in a flow cytometer within 60 min of staining. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. Analyze the data (Fig 7.5.1) using software which deconvolutes DNA content frequency histograms (see Chapter 10). BASIC PROTOCOL 3
SUPRAVITAL STAINING OF DNA Supravital staining of DNA offers the possibility of sorting of live cells on the basis of differences in their DNA content. This protocol uses Hoechst 33342 to measure DNA by flow cytometry in live cells. The actual procedure of cell staining is simple. Cells suspended in culture medium are incubated in the presence of 2.0 to 5.0 µg/ml Hoechst 33342 for 20 to 90 min. Cell fluorescence is then measured directly, without any additional treatments, centrifugations, etc. Materials Cells to be stained, 106 cells/ml suspended in tissue culture medium 1 mg/ml Hoechst 33342 staining solution (see recipe) Flow cytometer with UV light illumination source Stain cells with Hoechst 33342 1. Add Hoechst 33342 staining solution to cells suspended in tissue culture medium (106 cells/ml) to obtain a final dye concentration of 2 µg/ml. Incubate 20 min at 37°C. Perform flow cytometry 2. Set up and adjust flow cytometer for UV excitation at 340 to 380 nm and detection of Hoechst 33342 at blue wavelengths. An UG 1 optical filter may be used when the source of excitation is a mercury arc or xenon lamp. For detecting the blue fluorescence of Hoechst 33342, a band-pass filter at 470 ± 20 nm is recommended.
3. Measure cell fluorescence in the flow cytometer. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. When intensity of cell fluorescence or resolution of cells in the cell cycle phases is inadequate, prolong the staining time (up to 90 min) and/or increase the dye concentration in the medium (up to 5 ìg/ml). The same sample may be reanalyzed after prolonged incubation and/or addition of more staining solution. DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis
This protocol is predominantly used for sorting live cells. However, because sensitivity of cells to Hoechst 33342 varies depending on the cell type, it is possible that viability and cell cycle sorted progression of cells may be affected by the staining procedure.
7.5.6 Current Protocols in Cytometry
DNA CONTENT ANALYSIS OF PARAFFIN-EMBEDDED SAMPLES This protocol describes DNA content analysis of archival samples embedded in paraffin blocks. The technique is based on preparation of thick microtome sections of the paraffin-embedded material, solubilization and extraction of paraffin from the sections, tissue rehydration in graded ethanols, and isolation of nuclei by proteolytic digestion of the tissue. Samples are then stained with DAPI and subjected to flow cytometry.
BASIC PROTOCOL 4
Materials Paraffin-embedded tissue blocks Xylene or xylene substitute (e.g., Histo-Clear, National Diagnostics) 100%, 95%, 80%, and 50% ethanol Protease solution (see recipe), freshly made 0.15 M NaCl, for diluting nuclei if needed DAPI/phosphate staining solution (see recipe), freshly made Microtome 57-µm nylon mesh bags, 1 × 1 cm 1- to 2-mm-diameter glass beads Phase-contrast or differential interference–contrast (Nomarski optics) microscope Flow cytometer with UV light illumination source Prepare paraffin sections 1. Cut a standard thin section (5 to 10 µm from the paraffin-embedded tissue block), adjacent to the subsequent section that will be subjected to nuclear isolation. Process by routine hematoxylin and eosin (HE) staining. 2. Examine the thin section by light microscopy and select the area (e.g., tumor site) to be processed by flow cytometry. On the basis of examination of the thin section, with a scalpel trim the block from the undesired tissue. 3. Mount the paraffin block on a microtome. Cut sections 50 to 100 µm thick. The sections may curl up as they come from the microtome knife. Depending on the size (area) of the section and the cell density in the tissue, one to four thick sections are generally adequate for DNA analysis.
Isolate cell nuclei from paraffin sections 4. With forceps transfer the tissue sections into 1 × 1–cm fine mesh (57 µm) nylon bags. Add one or two 1- to 2-mm glass beads to prevent floating of bags on the surface of solutions used in subsequent steps. 5. Immerse bags in 20 ml xylene or xylene substitute and mix 60 min on a slowly rotating shaker at room temperature. CAUTION: Xylene is toxic. Wear gloves and keep lids on jars. When possible, xylene should be substituted by less toxic reagents such as Histo-Clear. Keep xylene and ethanol solutions in aliquots of 20 ml in closed glass or plastic, xylene-resistant containers (e.g., Coplin jars or Erlenmeyer flasks).
6. Drain xylene and transfer the bags with sections to 20 ml of 100% ethanol. Keep 10 min at room temperature. Successively transfer the bags to 95%, 80%, and 50% ethanol, keeping the bags 20 min in each solution. 7. Transfer each bag to a separate 15-ml tube containing distilled water. Keep 30 min at room temperature. Repeat rinse with water. Separation of bags from one another at this stage is necessary because with rehydration the tissue becomes soft and breaks up, which may cause cross-contamination of samples. Cross-contamination may be avoided by using bags made of fine nylon mesh (600 nm) filter is recommended.
7. Measure cell fluorescence in a flow cytometer. Use the pulse width–pulse area signal to discriminate between G2 cells and cell doublets and gate out the latter. Analyze the data as shown in Fig. 7.5.4.
Nucleic Acid Analysis
7.5.9 Current Protocols in Cytometry
Supplement 11
SUPPORT PROTOCOL
AGAROSE GEL ELECTROPHORESIS OF DNA EXTRACTED FROM APOPTOTIC CELLS In this protocol, low-molecular-weight DNA, extracted from the same cells that are subjected to flow cytometry (see Basic Protocol 5), is subsequently analyzed by agarose gel electrophoresis (Gong et al., 1994). Cell extraction is described in Basic Protocol 5 (i.e., cells are prefixed in 70% ethanol and, after removal of ethanol, DNA is extracted with a small volume of 0.2 M phosphate-citrate buffer, pH 7.8). In this protocol, the extract is sequentially treated with RNase A and proteinase K and then directly subjected to electrophoresis. Additional Materials (also see Basic Protocol 5) Cells to be studied 2 mg/ml DNase-free RNase A stock solution (APPENDIX 2A) 1 mg/ml proteinase K (Sigma) 6× gel loading buffer (APPENDIX 2A) 0.8% agarose gel (see recipe) DNA molecular weight standards, 100 to 1000 bp Electrophoresis buffer: 10× TBE buffer (APPENDIX 2A) Ethidium bromide staining solution (APPENDIX 2A) Extract low-molecular-weight DNA from cells 1. Fix cells in ethanol and extract low-molecular-weight DNA in DNA extraction buffer (see Basic Protocol 5, steps 1 to 3). 2. Centrifuge cells 10 min at 1500 × g. Withdraw 40 µl of supernatant and transfer to a 0.5-ml microcentrifuge tube. Remove proteins and RNA from cell extract 3. Add 5 µl of 2 mg/ml DNase-free RNase A stock solution and incubate 30 min at 37°C. Cap tube to prevent evaporation. 4. Add 5 µl of 1 mg/ml proteinase K and incubate 30 min at 37°C. Perform electrophoresis 5. Add 5 µl of 6× gel loading buffer and transfer the entire tube contents to one well of a 0.8% agarose horizontal gel. 6. Prepare and load a sample of DNA molecular weight standards in a total of 55 µl in 1× gel loading buffer. 7. Assemble gel electrophoresis apparatus, using electrophoresis buffer to fill the reservoir. Run electrophoresis 16 to 20 hr at 2 V/cm. Turn off the power when the bromphenol blue from the loading buffer migrates a distance sufficient for separation of DNA fragments. 8. To visualize the bands, stain the gel 20 to 30 min with ethidium bromide staining solution. CAUTION: Ethidium bromide is a potential carcinogen. Wear gloves when handling.
9. Transfer the gel onto a UV transilluminator. Observe after illumination. DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis
CAUTION: Ultraviolet light is dangerous to eyes and exposed skin. Wear protective eyewear and facewear.
10. Photograph the gel using a red or orange (e.g., Kodak Wratten no. 23A) emission filter and a clear UV light blocking filter (e.g., Kodak Wratten no. 2B).
7.5.10 Supplement 11
Current Protocols in Cytometry
REAGENTS AND SOLUTIONS Use distilled, deionized water for the preparation of all buffers. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Agarose gel, 0.8% Dissolve 1.6 g agarose in 200 ml hot (boiling) electrophoresis buffer (10× TBE). Cool to 55°C and pour solution onto a 15 × 15–cm sealed gel-casting platform. Insert the gel comb. Cool to room temperature. After the gel has hardened, remove the seal from the gel-casting platform and remove the gel comb. Place into an electrophoresis tank containing sufficient electrophoresis buffer to cover gel to a depth of ∼1 mm. Cell lysis solution with trypsin Dissolve 3 mg trypsin (Sigma type IX from porcine pancreas) in 100 ml detergent stock solution (see recipe) and adjust to pH 7.6. Store ≤1 year at −40° to −80°C in aliquots of 5 to 10 ml in tubes. Before use, bring to room temperature in a 37°C water bath. Avoid repeated thawings. Citrate/DMSO buffer 85.50 g sucrose (0.25 M final) 11.76 g trisodium citrate dihydrate (40 mM final) 5 ml DMSO (Sigma; 0.5% final) H2O to 1000 ml Adjust to pH 7.6 Store 1 month at 4°C To prepare the buffer, dissolve the dry ingredients and DMSO in ~800 ml water and then dilute to 1000 ml.
DAPI/phosphate staining solution, for nuclei isolated from paraffin tissue blocks Add 4 µl of 1 mg/ml DAPI to 80 ml water. Add 11.4 mg anhydrous Na2HPO4 (0.8 M final) and 0.82 g citric acid monohydrate (40 mM final), and mix to dissolve. The solution should be pH 7.4. Add water to 100 ml. Prepare freshly. The final DAPI concentration in this staining solution is 2 ìg/ml. A DAPI stock solution, made by dissolving 1 mg DAPI in 1 ml water, may be stored several months in dark or foil-wrapped bottles at or below −20°C.
DAPI/PIPES staining solution, for detergent-lysed cells Add 100 µl of 1 mg/ml DAPI to 100 ml PIPES/Triton X-100 buffer (see recipe). Store several weeks in dark or foil-wrapped bottles at 0° to 4°C. A DAPI stock solution, made by dissolving 1 mg DAPI in 1 ml water, may be stored several weeks in dark or foil-wrapped bottles at or below −20°C. For simultaneous staining of DNA and protein, add 100 ìl of 1 mg/ml DAPI to 100 ml PIPES/Triton X-100 buffer (see recipe) and then add 2 mg sulforhodamine 101 (Molecular Probes); stir to dissolve. Store several weeks in dark or foil-wrapped bottles at 0° to 4°C.
DAPI/Triton X-100 staining solution, for ethanol-fixed cells To 10 ml of 0.1% Triton X-100 in PBS add 10 µl of 1 mg/ml DAPI (Molecular Probes). Prepare freshly. A DAPI stock solution, made by dissolving 1 mg DAPI in 1 ml water, can be stored several months in dark or foil-wrapped bottles at −20°C.
Nucleic Acid Analysis
7.5.11 Current Protocols in Cytometry
Detergent stock solution 1 g trisodium citrate dihydrate (3.4 mM final) 1 ml Nonidet P-40 (Sigma; 0.1% final) 522 mg spermine tetrahydrochloride (1.5 mM final) 61 mg Tris (Sigma 7-9; 0.5 mM final) H2O to 1000 ml Store ≤1 year at 0° to 4°C Dissolve the dry ingredients and NP-40 in ~800 ml water and then dilute to 1000 ml.
DNA extraction buffer (0.2 M phosphate citrate buffer, pH 7.8) 192 ml 0.2 M Na2HPO4 8 ml 0.1 M citric acid Store several months at 4°C Hoechst 33342 staining solution, 1 mg/ml Dissolve 1 mg Hoechst 33342 (Molecular Probes) in 1 ml water. Store in dark or foil-wrapped bottles several months at 0° to 4°C. Internal DNA Content Standard Chicken and trout erythrocytes are convenient internal DNA content standards. Chicken blood is acquired by heart puncture and collected with heparin (50 U/ml blood). The blood is diluted with citrate/DMSO buffer (see recipe) to obtain 1.5 × 106 cells/ml, as counted with a hemacytometer. Rainbow trout blood is obtained by caudal vein puncture of an anesthetized fish, immediately mixed with citrate/DMSO buffer, and adjusted to 2.5 × 106 cells/ml. These solutions are then mixed 1:1 (v/v) to obtain 2 × 106 cells/ml. The proportion of chicken to trout erythrocytes in such a mixture provides approximately similar height peaks on DNA frequency histograms. The standard cells can be kept frozen, in small aliquots, at −40° to −80°C. PIPES/Triton X-100 buffer 3.02 g PIPES (Calbiochem; 10 mM final) 5.84 g NaCl (0.1 M final) 406 mg MgCl2⋅6H2O (2 mM final) 1 ml Triton X-100 (Sigma; 0.1% final) H2O to 1000 ml Adjust to pH 6.8 Store ≤1 year at 0° to 4°C Dissolve the dry ingredients and Triton X-100 in ~800 ml water, adjust pH with NaOH or HCl, and then dilute to 1000 ml.
Propidium iodide (PI)/spermine staining solution, for detergent-lysed cells Dissolve 20 mg PI (Molecular Probes) and 116 mg spermine tetrahydrochloride (Sigma) in 100 ml detergent stock solution (see recipe) and adjust to pH 7.6. Store ≤1 year at −40° to −80°C in small aliquots in 5- to 10-ml foil-wrapped tubes. Before use, thaw in a 37°C water bath and then keep on ice, protected from light. Propidium iodide (PI)/Triton X-100 staining solution with RNase A, for ethanolfixed cells To 10 ml of 0.1% (v/v) Triton X-100 (Sigma) in PBS add 2 mg DNase-free RNase A (Sigma) and 200 µl of 1 mg/ml PI (e.g., Molecular Probes). Prepare freshly. DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis
A stock solution of PI, made by dissolving 1 mg PI in 1 ml water, can be stored several months at 0° to 4°C. If the RNase is not DNase-free, boil a solution of 2 mg RNase A in 1 ml water for 5 min.
7.5.12 Current Protocols in Cytometry
Protease solution Dissolve 100 mg Sigma XXIV bacterial protease in 80 ml water. Add 1.58 g Tris⋅Cl (Sigma; 0.1 M final) and 0.41 g NaCl (0.7 M final) and dissolve. Adjust to pH 7.2 and add water to 100 ml. Prepare freshly. The protease solution makes use of the “Carlsberg subtilisin.”
Trypsin-inactivating solution Dissolve 50 mg chicken egg white trypsin inhibitor (Sigma) and 10 mg RNase A (Sigma) in 100 ml detergent stock solution (see recipe) and adjust to pH 7.6. Store ≤1 year at −40° to −80°C in small aliquots in 5- to 10-ml tubes. Before use, bring to 0° to 4°C in a 37°C. COMMENTARY Background Information Choosing a particular protocol among those presented in this unit depends primarily on the sample type (unfixed or fixed cells, paraffinembedded tissue blocks) and the necessity for sample storage (or transport) between cell collection and analysis. The discussion below describes characteristics of each of the methods and its applicability to different material. Analysis of fixed samples In Basic Protocol 1 DNA content is measured in prefixed cell samples. The preference for analysis of fixed cells often is dictated by the need to store or transport samples (e.g., clinical samples of solid or hematologic tumors). Extended storage of unfixed cells, unless done at low temperatures following cell suspension in cryopreservative media, leads to cell deterioration and DNA degradation. Fixed cells, on the other hand, often can be stored for months if not years without much deterioration. The fixative essentially has two functions: (1) it preserves the cells by preventing their lysis and autolytic degradation, and (2) it makes the cells permeable and their DNA accessible to the fluorochrome. Precipitating fixatives (ethanol, methanol, acetone) are preferred over cross-linking agents (formaldehyde, glutaraldehyde). This is because cross-linking of chromatin has deleterious effects on the stoichiometry of DNA staining. Precipitating fixatives, though inferior in terms of stabilization and preservation of the low-molecular-weight constituents within the cell, adequately stabilize undamaged DNA. It should be stressed, however, that damaged DNA, especially DNA having large numbers of double-strand breaks (e.g., as present in apoptotic cells, see Basic Protocol 5), leaks from the ethanol-fixed cells during their hydration and subsequent staining.
Although absolute alcohols or acetone, or a mixture of absolute ethanol and acetone (1:1), can be used and may be preferred for some applications (e.g., to obtain better stabilization and retention of particular proteins for their immunocytochemical detection), they induce more extensive cell aggregation, and the aggregates cannot be easily dissociated after hydration of the fixed cells. Fixation of cells in 70% to 80% ethanol (at 0° to 4°C), on the other hand, results in less cell clumping and is generally preferred in situations when the analysis is limited to DNA content alone. Sample storage at −20° to −40°C, especially when prolonged (months), appears to be more advantageous compared to storage at room temperature. A variety of DNA fluorochromes (UNIT 4.2) can be used to stain DNA in the fixed cells. Staining with dyes that react with both DNA and RNA, such as PI used in Basic Protocol 1, requires preincubation of cells with RNase. For cytometry, PI requires blue light as the fluorescence excitation source, which is conveniently provided by the 488-nm line of the argon ion laser available on most flow cytometers. Alternate Protocol 1 employs DAPI instead of PI for staining DNA in fixed cells. The advantage of DAPI is its greater specificity toward DNA, which often is reflected by lower coefficient of variation (CV) values of the mean DNA content of G1 cell populations. A disadvantage of DAPI is the requirement for UV excitation, which may not be possible in all flow cytometers. Analysis of detergent-lysed samples The major advantage of detergent-based methods is better accuracy in DNA content estimates. Exposure of live cells to detergents results in rupture of the plasma membrane and leakage of cytoplasmic constituents. Thus, iso-
Nucleic Acid Analysis
7.5.13 Current Protocols in Cytometry
DNA Content Measurement for DNA Ploidy and Cell Cycle Analysis
lated nuclei, rather than whole cells, are stained. Because several cytoplasmic constituents either are autofluorescent or nonspecifically interact with DNA fluorochromes, the specificity of DNA staining by methods based on cell permeabilization by detergents is superior compared to that of methods based on cell fixation. This is reflected by the high accuracy of DNA content estimates, which is represented by low values of the CV of the mean DNA content of cells having uniform DNA content, such as the G1 cell population. It has to be taken into account, however, that lysis of mitotic cells, which lack a nuclear envelope, leads to dispersion of individual chromosomes. Thus, mitotic cells may not be detected by methods utilizing detergents or hypotonic solutions. Furthermore, the presence of isolated chromosomes or chromosome aggregates may contribute to an increased frequency of detection of objects with low fluorescence values, generally classified as debris or apoptotic cells. Likewise, the lysis of apoptotic cells, which have fragmented nuclei, releases numerous nuclear fragments from a single cell. This generally precludes application of detergent-based methods for analysis of the frequency of apoptotic cells on the basis of fractional DNA content, and an alternative method must be sought (see Basic Protocol 5). Further improvement in the accuracy of DNA content analysis is seen after mild and controlled proteolysis of detergent-lysed cells. It is likely that the proteolytic step removes nuclear proteins known to restrict the accessibility of DNA to many fluorochromes, resulting in improved stoichiometry of DNA staining (Darzynkiewicz et al., 1984). This approach was perfected by Vindeløv (1983a,b,c,d; Vindeløv and Christiansen, 1994), who developed a highly accurate method of cellular DNA content analysis. These authors also pioneered in introducing internal DNA content standards as an intrinsic part of the staining protocol. Their methodology, presented in Basic Protocol 2, is now widely used, especially in clinical settings for DNA content analysis of tumor samples. Alternate Protocol 2 is a simplified detergent method that is more applicable to uniform cell populations such as tissue culture cells. Vindeløv’s procedure was designed for fine needle aspiration of normal tissue and tumor biopsies. The aspiration has two functions: collection of cells from the tissue and cell disaggregation. The aspiration can be done either in vivo (needles longer than 1 in. may be needed) or in vitro, from the resected tumor. In addition,
the specimen may initially be frozen at dry ice temperature, stored at −60° to −80°C, thawed to room temperature, and then aspirated. The presence of the cryopreservative dimethylsulfoxide (DMSO) in the citrate buffer used for collecting cells in Basic Protocol 2 protects cells from damage if the samples are stored at low temperatures prior to staining. Alternate Protocol 2 employs only detergent and is simpler than the detergent/proteolytic enzyme procedure of Basic Protocol 2. It also offers excellent resolution for uniform cell populations such as tissue culture cells (the CV of the mean fluorescence of G1 populations is typically 590 nm. For MI-stained nuclei, the corresponding settings are: excitation 400 to 450 nm, beam splitting at 490 nm, and emission detection at >510 nm. The released nuclei should appear intact and homogeneously stained.
7.6.2 Supplement 2
Current Protocols in Cytometry
1b. For protoplasts: Resuspend protoplasts in homogenization buffer at a concentration of 106 protoplasts/ml. Include reference protoplasts of known DNA content as needed. Harkins et al. (1990) provide information about protoplast preparation. The protoplasts break as a consequence of the detergent present in the homogenization buffer. Protoplast viability is an important consideration. Protoplasts should not be employed without gradient purification and typically should be 90% to 100% viable as determined via staining with fluorescein diacetate (Harkins et al., 1990).
2. Filter nuclear suspension through 15-µm nylon mesh. This step is critical for removing large debris, which otherwise might block the flow cell. For nuclei of plants having genome sizes much larger than tobacco (i.e., >4.8 × 109 base pairs/haploid genome or >10 pg DNA/2C nucleus), larger mesh sizes (40 to 60-ìm) are appropriate. The nylon mesh can be conveniently placed over the tip of a standard 5-ml disposable syringe and held in place using the plastic cover provided with disposable syringes, after cutting the tip off this cover to produce a ring. Alternatively, filter units can be made using ordinary 1-ml pipet tips. Cut off the tapering tip end and gently press the cut edge on a preheated hot plate until the plastic softens (since this method produces plastic fumes, work in a fume hood). Immediately press the hot edge onto the middle of a mesh square cut a bit larger than the pipet tip. Check that the seal is fully round and complete.
3a. To stain with propidium iodide: Add 1 mg/ml PI stock to a final concentration of 50 to 200 µg/ml and 1 mg/ml RNase stock to a final concentration of 10 µg/ml. Let sample stand 5 min prior to flow cytometric analysis. If unfixed CRBCs (see Support Protocol 1) are to be used as internal standards, they should be mixed with the cells prior to the addition of PI (or MI, see below). RNase must be added to avoid binding of PI to RNA. Since mithramycin binds specifically to DNA, RNase is not used in step 3b.
3b. To stain with mithramycin: Add 0.1 mg/ml MI stock to a final concentration of 10 µg/ml. Let the sample stand for 5 min prior to flow cytometric analysis. The precise concentrations of PI or MI to be employed, relative to the mass of tissue homogenized, should be determined through saturation analysis (see Critical Parameters). The time of incubation prior to flow analysis should be sufficient to permit the fluorescence signal to become invariant with respect to time.
Set up and align the flow cytometer 4. Power up the computer, cytometer, and laser. Adjust laser-emission wavelength to 488 nm (for PI) or 457 nm (for MI). For air-cooled lasers, 20-mW power output at 488 nm is sufficient. For water-cooled lasers, 100-mW output at 457 nm is employed.
5. Prepare an adequate amount of sheath fluid that closely matches the sample fluid in ionic strength and filter through an 0.22-µm Millipore GSWP 047 filter. Empty the waste tank and fill the sheath tank with the filtered sheath fluid. The sheath fluid should have the same ionic composition as the sample fluid (in this case the homogenization buffer) but detergents should be omitted.
6. Design and load a protocol for alignment using fluorescent microspheres. A Coulter Elite is normally equipped with four photomultiplier tubes (PMTs). In the standard filter configuration, 90° light scatter is assigned to PMT1, green (505- to 545-nm) fluorescence to PMT2, orange (555- to 595-nm) fluorescence to PMT3, and red (670- to 680-nm) fluorescence to PMT4. The flow cell has an orifice of 100-ìm diameter. For PI,
Nucleic Acid Analysis
7.6.3 Current Protocols in Cytometry
Supplement 2
the protocol should include acquisition of single-parameter histograms of forward-angle light scatter (FS), a single fluorescence channel (PMT3; peak and integral signal, 555- to 595-nm), and biparametric histograms of PMT3 fluorescence versus time. For MI, the protocol should substitute PMT2 (integral signal, 505- to 545-nm) for PMT3. Further blocking filters (BG38) can be employed to eliminate chlorophyll autofluorescence.
7. Align the cytometer using fluorescent microspheres diluted 1⁄10 with deionized water. Collect uniparametric histograms of fluorescence emission (peak and integral signal) and FS at a sample flow rate of 70 particles/sec, with the FS discriminator set to 100 and all other discriminators switched off. Adjust the optics until population coefficients of variation (CVs) for pulse integral and FS are minimized (typically 5% to 7%). Various supplements to the homogenization buffer can be helpful in suppressing these interferences, including (final concentrations in buffer): 15 mM 2-mercaptoethanol (Dolezel et al., 1994), 5% (v/v) polyvinylpyrrolidone (MW 40,000; PVP 40), 10 mM ascorbic acid, or 10 mM dithiothreitol (Bharathan et al., 1994). These supplements may be added individually or in combination; the ideal combination is determined empirically.
Analyze flow cytometric data 11. Determine the channel number of the sample G1 nuclear DNA peak and that of the internal standard. The peaks are usually symmetrical, so either the mode position or calculated mean can be employed.
12. Calculate the somatic (2C) DNA content of the sample according to the equation: sample 2C DNA content (pg DNA) = Analysis of Nuclear DNA Content and Ploidy in Higher Plants
sample G1 peak mean × standard 2C DNA content standard G peak mean 1
7.6.4 Supplement 2
Current Protocols in Cytometry
If necessary, DNA content values can be expressed in base pairs, using the following formula: 1 pg DNA = 0.965 × 109 bp (see Internet Resources). Whereas genome sizes in base pairs are conventionally reported in terms of the haploid size (1C) of the genome, mass values for genome size are typically reported per 2C value or even per 4C value. Care should therefore be taken in converting these values. Because of base-pair specificity effects, the fluorochrome employed for the DNA content measurement should always be identified.
13. Carry out ploidy analysis if desired. Ploidy analysis represents a special case in which the DNA contents of the unknown plant nuclei are compared to those from the same species having known ploidy.This comparison can be made either between two analyses performed under identical conditions, or, in some cases, through mixing the two (or more) samples for simultaneous measurement.
14. Carry out cell-cycle analysis, if desired, using cell-cycle analysis software. Cell-cycle analysis can be conveniently done on uniparametric flow histograms of plant nuclear DNA contents using commercially available software. This assumes an absence of endoreduplication. Alternatively, the proportions of cells within the various phases of the cell cycle can be determined by using a scissor to cut out from the appropriate plots the various areas corresponding to G1, S, and G2 cells, and weighing the paper. This approach can also be used for analysis of endoreduplicated cells, in which case the graphing of logarithmic uniparametric plots is required.
ANALYSIS OF SOMATIC DNA CONTENT IN PLANT TISSUES USING AN ARC LAMP–BASED FLOW CYTOMETER The following alternate protocols are all for use with arc lamp–based flow cytometers and involve staining with 4′,6-diamidino-2-phenylindole (DAPI) or propidium iodide (PI). Alternate Protocol 1 is for isolation of nuclei using lysis buffer LB01 (Dolezel et al., 1989), which works well with most plant species and tissues. For a very few species, resolution of DNA histograms is not satisfactory using LB01. In those cases, Alternate Protocol 2, which is a modification of a procedure originally developed by Otto (1990), can provide improved resolution (also see Dolezel and Göhde, 1995). The disadvantages of Alternate Protocol 2 are the it is quite laborious and that it is not universally applicable (i.e., some species give histograms with increased levels of background and high CVs). If none of these procedures works, the authors recommend the use of Partec buffer (de Laat et al., 1987). This buffer gives very good resolution with Arabidopsis thaliana tissues. The procedure for analysis using Partec buffer is identical to that for LB01 buffer (see Alternate Protocol 1). The flow cytometer should be set up according to the manufacturer’s instructions for analysis of PI- or DAPI-derived fluorescence. The steps described below for preparing the flow cytometer are for Partec flow cytometers. Nuclear DNA Content Analysis Using LB01 Buffer or Partec Buffer and Arc Lamp–Based Flow Cytometer
ALTERNATE PROTOCOL 1
The following instructions are for preparation of cells with either LB01 (Dolezel et al., 1989) or Partec buffer (de Laat et al., 1987). Additional Materials (also see Basic Protocol) Lysis buffer LB01 or Partec buffer (see recipes) with fluorochrome, ice-cold 1 mg/ml propidium iodide (PI) stock solution (see recipe) or 0.1 mg/ml 4′,6-diamidino-2-phenylindole (DAPI) stock solution (see recipe) and 1 mg/ml ribonuclease (RNase) stock solution (see recipe)
Nucleic Acid Analysis
7.6.5 Current Protocols in Cytometry
Supplement 2
Particles for instrument alignment: e.g., microspheres or fixed chicken red blood cells (CRBCs; see Support Protocol 2) stained with the fluorochrome used (also see UNIT 1.3) Glass petri dish 42-µm pore-size nylon mesh Flow cytometer equipped with high-pressure mercury-arc lamp Filters and dichroic mirror appropriate for fluorochrome used Prepare suspensions of nuclei 1a. For intact plant tissue, cell cultures, or callus: Weigh a small amount of plant material (typically 20 mg) and chop with a new razor blade or a sharp scalpel in 1 ml of ice-cold fluorochrome-containing LB01 lysis buffer or Partec buffer in a glass petri dish (see Basic Protocol, step 1a, for chopping technique). It is preferable to include the DNA fluorochrome (DAPI or PI) in the buffer. Alternatively, the stain may be added immediately after filtration (step 2). Note that commercially available Partec buffer contains 4 ìg/ml DAPI. The actual quantity of plant material to be used for nuclei isolation depends both on the type of tissue and on the species, and must be determined experimentally (larger quantities are usually needed for callus or cultured cells).
1b. For protoplasts: Resuspend protoplasts in ice-cold fluorochrome-containing LB01 lysis buffer or Partec buffer to a concentration of 1 × 105 to 1 × 106 protoplasts/ml. For protoplasts the concentration of detergent (Triton X-100) in LB01 buffer should be increased to 0.5% (v/v); this improves the release of the nuclei from the protoplasts. Nuclei cannot be released from “collapsed” protoplasts; hence protoplast viability is an important consideration. Typically the protoplasts should be 90% to 100% viable as determined using FDA (Harkins et al., 1990).
2. Filter the suspension through a 42-µm nylon mesh. If the DAPI or PI was not initially added to the LB01 or Partec buffer, add 0.1 mg/ml 4′,6-diamidino-2-phenylindole (DAPI) stock solution (see recipe) to a final concentration of 2 ìg/ml or 1 mg/ml propidium iodide (PI) stock solution (see recipe) to a final concentration of 50 ìg/ml along with 1 mg/ml RNase stock solution (see recipe) to a final concentration of 50 ìg/ml.
3. Store on ice prior to analysis (a few minutes to 1 hr). Set up and align the flow cytometer 4. Empty the waste container of the flow cytometer and fill the sheath-fluid container with distilled water. Older Partec cytometers (e.g., PAS II) ran under vacuum and sheath fluid had to be deaerated. This is not necessary with the newer models.
5. Choose the optical filter set that corresponds to the fluorochrome used to stain nuclear DNA. For DAPI-stained samples, use UG1 as the excitation filter, TK420 as the dichroic mirror, and GG435 as a barrier filter (a dry 20× objective is sufficient for most applications). For PI-stained samples, use BP520 as an excitation filter, TK575 as the dichroic mirror, and RG590 as a barrier filter; the use of a 40×/1.25 glycerol-immersion objective is recommended. Analysis of Nuclear DNA Content and Ploidy in Higher Plants
6. Set the amplification to linear mode. 7. Prepare particles suitable for instrument alignment (e.g., microspheres or fixed CRBCs stained with appropriate fluorochrome; see Support Protocol 2 and UNIT 1.3).
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Current Protocols in Cytometry
8. Using the particles prepared in step 7 and a low data rate (20 to 50 events/sec), adjust the gain of the instrument so that the peak corresponding to the G1 nuclei appears approximately at one-fifth of the distance across the x axis (e.g., channel 50 on a 256 scale). It is important to run the analyses at a low data rate to achieve the highest resolution.
9. Check the resolution (the population coefficients of variation of the fluorescent peaks) and the linearity of the instrument. For CRBC nuclei, the coefficient of variation should be 1 year, defined as infertile), respectively (Table 7.13.3). A couple requesting counselling on their chances of pregnancy may be advised that their chances of pregnancy are greatly reduced if the percent sperm with abnormal chromatin is above 30%. Figure 7.13.4 consists of cartoon illustrations of normal fluorescence populations and common variations when sperm are measured by the SCSA. Panel A is a classical SCSA cytogram with the following numbered areas: (1) The main population of cells in a semen sample that, under the conditions imposed, remain as a coherent population. Although the DNA content in these cells is the same haploid amount, the cytogram cluster is elliptical in shape due to the optical artifact discussed in the text. This artifact is of little to no consequence to the outcome and interpretation of SCSA data. (2) This area contains the cells outside the main population, or COMPαt. These cells typically move out and downward from the main population at a ~45° angle, showing the increase of red fluorescence at the expense of green fluorescence. Some samples show a continuous
cluster ranging from just outside the main population to ones with very high red and very low green values. Alternatively, there are several discrete clusters in the COMPαt population with two shown in this illustration. (3) This area represents seminal debris consisting of broken cellular components and other particulate matter stained with AO. For proper SCSA analysis, the debris signals must be resolved from the sperm signals. Since the COMPαt population forms a ~45° slope, downward and to the right, an effective means to delineate between these cell clusters is to draw a 45° computer gate between them at the bottom edge of the main population. Note that this is also an active gate during acquisition so that 5000 or more sperm signals are accumulated and any debris signals excluded. This is most important in samples with a high ratio of debris to sperm so that the same number of sperm cells are statistically analyzed per sample. (4) The sperm signals appearing in areas 4 and 5 have high AO stainability and are termed sperm with high green stainability (% HIGRN). Chromatin in these sperm is probably not fully condensed, thus allowing a greater accessibility by intercalating DNA dyes. The percentage of HIGRN sperm is a ratio of the number of cells in areas 4 and 5 divided by the total number of cells contained in the gating region (includes sperm populations 1, 2, and 4) plus population 5 × 100. (Note that for most SCSA parameters the cytogram gating region contains the total number of sperm cells in areas 1, 2, and 4). The % HIGRN region starts at ~70%-75% of the green fluorescence scale. (5) This area contains sperm that also have higher DNA stainability, aggregates of sperm, early sperm forms, and possible somatic cells. The authors have not sorted this population for light microscopic identification; thus, a conclusion on how much of the region to include in this calculation has not been finalized. Panel B of Figure 7.13.5 is the same as panel A except that area (6) represents a semen sample with excessive bacterial contamination. Due to random size clumping of the bacteria, a straight line of signal is seen to the left of the sperm population. When present, this population is gated out during acquisition in order to accumulate 5000 sperm signals. Note that this tends to be confounded with cells in the upper high green stainability area also. Panel C of Figure 7.13.5 is also the same as A except that area (7) is an approximation of a sperm sample that has been compromised by excessive freeze/thaw cycles or left in a thawed state for an extended time period. This popula-
7.13.24 Supplement 13
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tion has an increased DNA stainability and shifts a bit to the right, as discussed in the text, but this compromised state is also revealed by a telltale hook usually present at the top of the population cluster. Samples showing this artifact are removed from the database. When all the samples in a box have this telltale artifact, inquiry usually confirms that a freezer failure or similar event occurred. Panel D of Figure 7.13.5 is a composite of possible sperm populations.
Time Considerations For fresh sperm samples, following collection of the sample, and a 30-min period for semen liquefaction, an aliquot of the sample can be measured by the SCSA, or be placed into a cryotube and plunged directly into liquid nitrogen or an ultracold freezer for storage. Since cryoprotectants are not needed, this is accomplished within a couple of minutes. Longitudinal studies of changes in sperm parameters are possible with the SCSA, as the sperm can be stored frozen until all samples are collected. The time of course depends on the length of the experimental study. The SCSA requires paying very strict attention to time. Once a sample that has been frozen is warmed for thawing, the process is committed. As mentioned in the text, a sample may be refrozen once or more for a nonexacting measurement of SCSA parameters. Thawing the sample takes ~30 to 60 sec and preparing the sample for flow cytometry measurement and equilibration in the flow cytometer takes 3 min. Data on 5000 cells can be collected in less than 1 min. When a flow operator becomes proficient, 5 to 6 samples measured in duplicate can be run per hour. Data analysis on each sample takes ∼10 min.
Literature Cited Aravindan, G.R., Bjordahl, J., Jost, L.K., and Evenson, D.P. 1997. Susceptibility of human sperm to in situ DNA denaturation is strongly correlated with DNA strand breaks identified by single-cell electrophoresis. Exp. Cell Res. 236:231-237. Ballachey, B.E., Hohenboken, W.D., and Evenson, D.P. 1987. Heterogeneity of sperm nuclear chromatin structure and its relationship to fertility of bulls. Biol. Reprod. 36:915-925 Ballachey, B.E., Saacke, R.G., and Evenson, D.P. 1988. The sperm chromatin structure assay: Relationship with alternate tests of sperm quality and heterospermic performance of bulls. J. Androl. 9:109-115.
Darzynkiewicz, Z., Traganos, F., Sharpless, T., and Melamed, M.R. 1975. Thermal denaturation of DNA in situ as studied by Acridine Orange staining and automated cytofluorometry. Cell Res. 90:411-428. Engh, E., Clausen, O.P.F., Scholberg, A., Tollefsrud, A., and Purvis, K. 1992. Relationship between sperm quality and chromatin condensation measured by sperm DNA fluorescence using flow cytometry. Int. J. Androl. 15:407-415. Estop, A.M., Munne, S., Jost, L.K., and Evenson, D.P. 1993. Alterations in sperm chromatin structure correlates with cytogenetic damage of mouse sperm following in vitro incubation. J. Androl. 14:282-288. Evenson, D.P. 1997. Sperm nuclear DNA strand breaks and altered chromatin structure: Are there concerns for natural fertility and assisted fertility in the andrology lab? Moving Beyond Boundaries: Clinical Andrology in the 21st Century. Andrology Laboratory Workshop, Postgraduate Course, Baltimore, Md. Evenson, D.P. 1999a. Alterations and damage of sperm chromatin structure and early embryonic failure. In Towards Reproductive Certainty: Fertility and Genetics Beyond 1999. Proceedings of the 11th World Congress on In Vitro Fertilization and Human Reproductive Genetics (R. Jannsen and D. Mortimer, eds.) pp. 313-329. Parthenon Publishing Group, New York. Evenson, D.P. 1999b. Loss of livestock breeding efficiency due to uncompensable sperm nuclear defects. Reprod. Fertility Dev. 11:1-15. Evenson, D.P. and Darzynkiewicz, Z. 1990. Acridine orange induced precipitation of mouse testicular sperm cell DNA reveals new patterns of chromatin structure. Exp. Cell Res. 187:328-334. Evenson, D.P. and Jost, L.K. 1993. Hydroxyurea exposure alters mouse testicular kinetics and sperm chromatin structure. Cell Prolif. 26:147159. Evenson, D.P. and Jost, L.K. 1994. Sperm chromatin structure assay: DNA denaturability. In Methods in Cell Biology, Vol. 42: Flow Cytometry (Z. Darzynkiewicz, J.P. Robinson, and H.A. Crissman, eds.) pp. 159-176. Academic Press, Orlando, Fla. Evenson, D.P. and Melamed, M.R. 1983. Rapid analysis of normal and abnormal cell types in human semen and testis biopsies by flow cytometry. J. Histochem. Cytochem. 31:248-253. Evenson, D.P., Darzynkiewicz, Z., and Melamed, M.R. 1980. Relation of mammalian sperm chromatin heterogeneity to fertility. Science 240:1131-1133. Evenson, D.P., Klein, F.A., Whitmore, W.F., and Melamed, M.R. 1984. Flow cytometric evaluation of sperm from patients with testicular carcinoma. J. Urol. 132:1220-1225.
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Evenson, D.P., Higgins, P.H., Grueneberg, D., and Ballachey, B. 1985. Flow cytometric analysis of mouse spermatogenic function following exposure to ethylnitrosourea. Cytometry 6:238-253. Evenson, D.P., Baer, R.K., Jost, L.K., and Gesch, R.W. 1986a. Toxicity of thiotepa on mouse spermatogenesis as determined by dual parameter flow cytometry. Toxicol. Appl. Pharmacol. 82:151-163. Evenson, D.P., Darzynkiewicz, Z., Jost, L., Janca, F., and Ballachey, B. 1986b. Changes in accessibility of DNA to various fluorochromes during spermatogenesis. Cytometry 7:45-53. Evenson, D.P., Baer, R.K., and Jost, L.K. 1989a. Flow cytometric analysis of rodent epididymal spermatozoal chromatin condensation and loss of free sulfhydryl groups. Mol. Reprod. Dev. 1:283-288. Evenson, D.P., Baer, R.K., and Jost, L.K. 1989b. Long term effects of triethylenemelamine exposure on mouse testis cells and sperm chromatin structure assayed by flow cytometry. Environ. Mol. Mutagen. 14:79-89.
Evenson, D., Jost, L., and Sailer, B. 1995b. Flow cytometry of sperm chromatin structure as related to toxicology and fertility. Proceedings of the Seventh International Spermatology Symposium, Cairns, Australia. Evenson, D.P., Jost, L.K., Zinaman, M.J., Clegg, E., Purvis, K., de Angelis, P., and Clausen, O.P. 1999. Utility of the sperm chromatin structure assay (SCSA) as a diagnostic and prognostic tool in the human fertility clinic. Hum. Reprod. 14:1039-1049. Evenson, D.P., Jost, L.K., Corzett, M., and Balhorn, R. 2000a. Effect of elevated body temperature on human sperm chromatin structure. J. Andrology. Submitted for publication. Evenson, D.P., Jost, L.K., and Varner, D.D. 2000b. Stallion sperm nuclear protamine-SH status and susceptibility to DNA denaturation are not strongly correlated. J. Fertility Reprod. Suppl. In press.
Evenson, D.P., Janca, F.C., Jost, L.K., Baer, R.K., and Karabinus, D.S. 1989c. Flow cytometric analysis of effects of l,3-dinitrobenzene on rat spermatogenesis. J. Toxicol. Environ. Health 28:81-98.
Fossa, S.D., De Angelis, P., Kraggerud, S.M., Evenson, D., Theodorsen, L., and Claussen, O.P. 1997. Prediction of post-treatment spermatogenesis in patients with testicular cancer by flow cytometric sperm chromatin structure assay. Commun. Clin. Cytometry 30:192-196.
Evenson, D.P., Janca, F.C., Baer, R.K., Jost, L.K., and Karabinus, D.S. 1989d. Effect of l,3-dinitrobenzene on prepubertal, pubertal and adult mouse spermatogenesis. J. Toxicol. Environ. Health 28:67-80.
Gledhill, B.L., Lake, S., Steinmetz, L.L., Gray, J.W., Crawford, J.R., Dean, P.N., and VanDilla, M.A. 1976. Flow microfluorometric analysis of sperm DNA content: Effect of cell shape on the fluorescence distribution. J. Cell Physiol. 87:367-376.
Evenson, D.P., Jost, L., Baer, R., Turner, T., and Schrader, S. 1991. Individuality of DNA denaturation patterns in human sperm as measured by the sperm chromatin structure assay. Reprod. Toxicol. 5:115-125.
Gorzcyca, W., Gong, J., and Darzynkiewicz, Z. 1993. Detection of DNA strand breaks in individual apoptotic cells by the in situ terminal deoxynucleotidyl transferase and nick translation assays. Cancer Res. 53:945-951.
Evenson, D.P., Emerick, R.J., Jost, L.K., KayongoMale, H., and Stewart, S.R. 1993a. Zinc-silicon interactions influencing sperm chromatin integrity and testicular cell development in the rat as measured by flow cytometry. J. Anim. Sci. 71:955-962.
Grajewski, B., Cox, C., Schrader, S.M., Murray, W.E., Edwards, R.M., Turner, T., Smith, J.M., Shekar, S., Evenson, D., Simon, S.W., and Conover, D.L. 2000. Semen quality and hormone levels among radiofrequency heat sealer operators. J. Occupational Environ. Med. In press.
Evenson, D.P., Jost, L.K., and Baer, R.K. 1993b. Effects of methyl methanesulfonate on mouse sperm chromatin structure and testicular cell kinetics. Environ. Mol. Mutagen. 21:144-153. Evenson, D.P., Jost, L.K., and Gandy, J.G. 1993c. Glutathione depletion potentiates ethyl methanesulfonate-induced susceptibility of rat sperm DNA denaturation in situ. Reprod. Toxicol. 7:297-304. Evenson, D.P., Thompson, L., and Jost, L. 1994. Flow cytometric evaluation of boar semen by the sperm chromatin structure assay as related to cryopreservation and fertility. Theriogenology 41:637-651. Sperm Chromatin Structure Assay for Fertility Assessment
nal axes flow cytometers. Cytometry 19:295303.
Evenson, D., Jost, L., Gandour, D., Rhodes, L., Stanton, B. Clausen, O.P., De Angelis, P., Coico, R., Daley, A., Becker, K., and Yopp, T. 1995a. Comparative sperm chromatin structure assay measurements on epiillumination and orthogo-
Larson, K.L., Brannian, J.D., Timm, B.K., Jost, L.K., and Evenson, D.P. 1999. Density gradient centrifugation and glass wool filtration of semen remove sperm with damaged chromatin. Hum. Reprod. 14:2015-2019. Larson, K., DeJonge, C., Barnes, A., Jost, L., and Evenson, D. 2000. Relationship between assisted reproductive techniques (ART) outcome and status of chromatin integrity as measured by the sperm chromatin structure assay (SCSA). Hum. Reprod. In press. Lemasters, G.K., Olsen, D.M., Yiin, J.H., Lockey, J.E., Shukla, R., Selevan, S.G., Schrader, S.M., Toth, G.P., Evenson, D.P., and Huszar, G.B. 1999. Male reproductive effects of solvent and fuel exposure during aircraft maintenance. Reprod. Toxicol. 13:155-166.
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Martin, D. and Lenardo, M. 2000. Morphological, biochemical, and flow cytometric assays of apoptosis. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 14.13.1-14.13.21. John Wiley & Sons, New York. Potts, R.J., Newbury, C.J., Smith, G., Notarianni, L.J., and Jefferies, T.M. 1999. Sperm chromatin damage associated with male smoking. Mutat. Res. 423:103-111. Rubes, J., Lowe, X., Moore, D., Perreault, S., Slott, V., Evenson, D., Selevan, S., and Wyrobek, A.J. 1998. Cigarette-smoking lifestyle is associated with increased sperm disomy in teenage men. Fertility Sterility 70:715-723. Sailer, B.L., Jost, L.K., Erickson, K.R., Tajiran, M.A,. and Evenson, D.P. 1995a. Effects of X-ray irradiation on mouse testicular cells and sperm chromatin structure. Environ. Mol. Mutagen. 25:23-30.
Sassa, and I.G. Sipes, eds.) pp. 3.5.1-3.5.6. John Wiley & Sons, New York. Tejada, R.I., Mitchell, J.C., Norman, A., Marik, J.J., and Friedman, S. 1984. A test for the practical evaluation of male fertility by acridine orange (AO) fluorescence. Fertil. Steril. 42:87-91. Vine, M.F., Hulka, B.S., Everson, R.B., and Evenson, D. 2000. An assessment of DNA damage in the sperm of smokers and nonsmokers using the sperm chromatin structure assay. Cancer Epidemiol. Biomarkers Prevention. In press. Wyrobek, A.J., Schrader, S.M., Perrault, S.D., Fenster, L., Huszar, G., Katz, D.F., Osorio, A.M., Sublet, V., and Evenson, D. 1997. Assessment of reproductive disorders and birth defects in communities near hazardous chemical sites. III. Guidelines for field studies of male reproductive disorders. Reprod. Toxicol. 11:243-259.
Key References Evenson et al., 1980. See above.
Sailer, B.L., Jost, L.K., and Evenson, D.P. 1995b. Mammalian sperm DNA susceptibility to in situ denaturation associated with the presence of DNA strand breaks as measured by the terminal deoxynucleotidyl transferase assay. J. Androl. 16:80-87.
The first article correlating SCSA data and fertility.
Selevan, S.G., Borkovec, L., Slott, V.L., Zudova, Z., Rubes, J., Evenson, D.P., and Perreault, S.D. 2000. Semen quality and reproductive health of young Czech men exposed to seasonal air pollution. Submitted for publication.
Evenson et al., 1991. See above.
Evenson et al., 1999. See above. The most definitive paper on the SCSA and its application to human fertility clinics.
First longitudinal study utilizing the SCSA and showing the within-donor repeatability of samples over time and the repeatability of the SCSA when measuring the same sample several times.
Shapiro, H.M. 1985. Practical Flow Cytometry. Alan R. Liss, New York, p. 36.
Evenson et al., 1995a. See above.
Shapiro, H.M. 1988. Practical Flow Cytometry, 2nd ed. Alan R. Liss, New York, pp. 67-68, 205-207.
For more details on instrumentation already used with the SCSA.
Spanò, M., Kolstad, H., Larsen, S.B., Cordelli, E., Leter, G., Giwercman, A., and Bonde, J.P. 1998. ASCLEPIOS Study Group: The applicability of the flow cytometric sperm chromatin structure assay in epidemiological studies. Hum. Reprod. 13:2495-2505.
Contributed by Donald Evenson and Lorna Jost South Dakota State University Brookings, South Dakota
Sutherland, J.E and Costa, M. 1999. Assays for DNA damage. In Current Protocols in Toxicology (M.D. Maines, L.G. Costa, D.J. Reed, S.
This research is based upon work supported by Environmental Protection Agency Grant No. R827019, United States Department of Agriculture Sabbatical Grant No. 9803911, National Science Foundation EPSCoR Grant OSR 9452894, and South Dakota Futures Funds. This is South Dakota Agricultural Experiment Station Publication No. 3168 of the journal series.
Nucleic Acid Analysis
7.13.27 Current Protocols in Cytometry
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Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling and Multivariate Flow Cytometry
UNIT 7.14
A key parameter of cell culture is the quantitation of cell proliferation and cell survival. This can be determined by allowing proliferating cells to incorporate halogenated DNA precursors, such as 5-bromodeoxyuridine (BrdU) and 5-iododeoxyuridine, into DNA during passage through the S phase of the cell cycle. In the methods described in this unit, the halogenated DNA precursors can be detected by virtue of their property of quenching Hoechst dye fluorescence (Latt, 1973). After staining with Hoechst 33258 or 33342, cells can be analyzed with a flow cytometer (Kubbies and Rabinovitch, 1983; Rabinovitch, 1983). This way, cells that have undergone one, two, or three cell cycles during the period of labeling with halogenated DNA precursors (i.e., incorporation of halogenated DNA precursors during one, two, or three S phases) can be resolved. Counterstaining cells with a dye that exhibits fluorescence proportional to DNA content and in a different range of the emission spectrum than that of the Hoechst dyes (i.e., green, yellow, or red fluorescence emission), allows resolution of cells in the G1, S, and G2 phases of the cell cycle. Combining the information from the two DNA dyes allows one to distinguish cells in the G1, S, and G2 phases of three consecutive cell cycles (Rabinovitch et al., 1988). Recently, protocols have been developed that allow this type of information to be obtained from cells distinguished simultaneously on the basis of differential expression of up to two surface antigens or green fluorescent protein (GFP). In addition, calibration for the sample volume analyzed allows determination of the absolute number of proliferating and nonproliferating cells in a sample. With this technique, the proliferative survival of cells (i.e., the number of surviving and proliferating cells) can be determined. Presented in this unit are four protocols for analysis of cell proliferation and cell survival by flow cytometry: (1) a generic protocol to analyze the proliferative history of cells after continuous labeling with BrdU based on cell permeabilization and staining with Hoechst 33258 and ethidium bromide (see Basic Protocol); (2) a protocol to analyze the proliferative history of GFP-expressing versus nonexpressing cells in a cell culture (see Alternate Protocol 1); (3) a protocol to analyze the proliferative history of cell surface antigen–expressing versus non-expressing cells in a cell culture (see Alternate Protocol 2); and (4) a protocol to determine the absolute number of proliferating and nonproliferating cells in a sample by calibration for the sample volume analyzed (see Alternate Protocol 3). NOTE: BrdU is a photosensitizing drug. To avoid photochemical damage to the stained cells, all incubations are performed in subdued light. NOTE: All protocols described have been performed on cultured animal cells; limited data exist regarding the use of these methods for plant cells and in yeast. Expertise is assumed for basic techniques in flow cytometry as well as in cell culture and harvesting (of both suspension cultures and adherent cells; APPENDIX 3B), immunocytochemistry (Watkins, 1989), and fluorescence microscopy (UNIT 2.4). CAUTION: DMSO and dye solutions are potentially toxic to humans. Use nitrile gloves and wear eye protection at all stages of handling. Seek medical advice if dye or dye solutions are ingested or inhaled. The dyes mentioned are for in vitro use only; do not administer either externally or internally. All staining solutions should be poured through a funnel with a filter containing activated charcoal in a fume hood. The nonfluorescent filtrate can be poured down the sink. When the passing solution becomes fluorescent, the Contributed by Martin Poot, M. Rosato, and Peter S. Rabinovitch Current Protocols in Cytometry (2001) 7.14.1-7.14.9 Copyright © 2001 by John Wiley & Sons, Inc.
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filter should be incinerated or disposed of according to applicable rules for environmental hygiene and a fresh filter should be installed. BASIC PROTOCOL
ANALYSIS OF CELL PROLIFERATION BY CONTINUOUS 5-BrdU INCORPORATION FOLLOWED BY HOECHST 33258 AND ETHIDIUM BROMIDE FLOW CYTOMETRY Incorporation of BrdU into DNA leads to quenching of the Hoechst dyes. After excitation with UV light (∼360 nm), the Hoechst dye emits blue fluorescence (∼450 nm) and ethidium bromide emits red fluorescence (590 to 630 nm). Ethidium bromide can also be excited at 488 nm with an argon laser. This protocol allows determining the distribution of cells among the G1, S, and G2 compartments of three successive cell cycles. Materials 10 mM 5-bromodeoxyuridine (BrdU) in distilled water Cells in appropriate culture medium containing 10% FBS (APPENDIX 2A) Generic Hoechst staining buffer (see recipe) 10% (v/v) Nonidet P-40 (or IGEPAL, Sigma) 1 mg/ml ethidium bromide (Sigma) in distilled water 15-ml screw-capped centrifuge tubes Flow cytometer with either a mercury arc lamp or an argon laser (tuned to 360 nm) as excitation source; alternatively, two time-resolved argon lasers (tuned to 360and 488-nm excitation wavelengths, respectively) can be used 12 × 75–mm polypropylene flow cytometer sample tubes Computer and appropriate software for data collection and processing Label cells with BrdU 1. Add 10 mM BrdU to a final concentration of 100 µM to the cell culture medium of each culture of interest, wrap the plates or flasks with aluminum foil, and continue cell culture for the desired time period. It is important that the BrdU not be depleted during cell growth, and for this reason the starting cell concentration should not exceed 100,000 cells/ml or 2,500 cells/cm2. Since BrdU is a photosensitizer, all staining steps have to be performed in subdued light. The duration of BrdU labeling depends on the cell cycle time of the cells under study and on the kind of information required (see Commentary). For instance, if detailed information regarding cell cycle compartment transition times is required, several samples spanning at least two cell cycle periods have to be taken. In some cell types it may be necessary to include an equimolar amount of deoxycytidine (see Commentary).
Prepare cells 2. Harvest BrdU-labeled cells by standard procedures in 15-ml screw-capped centrifuge tubes and centrifuge 5 min at 200 × g, room temperature. Proceed to cell staining. Pellets may be resuspended in cell culture medium supplemented with 10% FBS and 10% DMSO and stored at −20°C for up to 6 months without detectable degradation.
3. Resuspend cell pellets at 0.5-1.0 × 106 cells/ml in generic Hoechst staining buffer, add an aliquot of 10% NP-40 solution to a final concentration of 0.1%, and incubate ≥15 min at room temperature in the dark. Since BrdU is a photosensitizer all staining steps have to be performed in subdued light. Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling
4. After 15 min staining in the dark, add 1 mg/ml ethidium bromide to a final concentration of 2.0 µg/ml and continue staining for another 15 min, room temperature, in the dark.
7.14.2 Supplement 15
Current Protocols in Cytometry
Set up flow cytometer 5. Set up and optimize the flow cytometer. Collect ultraviolet (360 nm)-excited blue fluorescence (Hoechst fluorescence) and 488 nm-excited fluorescence between 590 and 630 nm (ethidium bromide). Carefully resuspend cell sample and disrupt cell clumps by gently pipetting up and down a few times in 12 × 75–mm polypropylene flow cytometer sample tubes immediately before analysis. To avoid “bleeding” of Hoechst fluorescence into the ethidium bromide channel, time-resolved sample excitation (with spatially separated UV and 488-nm laser beams) can be used. Ethidium bromide can be excited by both 360- and 488-nm light. In addition, ethidium bromide will be excited by blue Hoechst 33342 fluorescence via fluorescence energy transfer. Thus, excitation can be made with UV light only, but the visual representation is clearer with dual-wavelength excitation. During harvesting, cells tend to clump; to obtain meaningful data on a per-cell basis it is essential to resuspend cells immediately before analysis. If this procedure is not sufficient to disrupt cell clumps, it is advised that they be broken up by aspirating the samples through a 0.5-mm internal diameter 21-G needle.
6. Collect ≥10,000 signals for each sample on a computer and appropriate software. If detailed information regarding cell cycle distributions among two or three successive cell cycles is required, one should acquire 2 or 3 times 10,000 signals.
ANALYSIS OF CELL PROLIFERATION OF GFP-EXPRESSING VERSUS NONEXPRESSING CELLS IN A SINGLE-CELL CULTURE Upon excitation with 488-nm laser light, GFP emits green fluorescence (∼510 nm). This fluorescence can be detected in samples that are stained with Hoechst 33258 and 7-AAD dyes. With this protocol, the distribution of GFP-expressing and nonexpressing cells among the G1, S, and G2 compartment of three successive cell cycles can be determined.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol) GFP-containing vector/virus cells 2% (w/v) paraformaldehyde solution (see recipe) Phosphate buffered saline (PBS; APPENDIX 2A) 1% (w/v) saponin (Sigma) in PBS 1 mg/ml 7-aminoactinomycin D (7-AAD; Calbiochem) in DMSO Label cells with BrdU 1. After transfection/transduction with a GFP-containing vector/virus, label cells with 100 µM BrdU as described in the Basic Protocol, step 1. Prepare and fix cells 2. Harvest BrdU-labeled cells by standard procedures in 15-ml screw-capped centrifuge tubes and centrifuge 5 min at 200 × g, room temperature. 3. Pipet 7 ml of 2% paraformaldehyde into a second 15-ml screw-capped cell culture tube. 4. Resuspend the cell pellet in 7 ml PBS and add cell suspension dropwise to the tube with 1 ml of 2% paraformaldehyde while vortexing at maximum speed. The final paraformaldehyde concentration is 0.25%. Let the tube sit 15 min at room temperature to fix cells. This procedure is intended to minimize the formation of cell clumps during the fixation step.
5. Centrifuge samples 5 min at 200 × g, room temperature, decant supernatant, and resuspend cell pellets in PBS.
Nucleic Acid Analysis
7.14.3 Current Protocols in Cytometry
Supplement 15
At this stage samples can be stored overnight or several days at 4°C.
6. Centrifuge cells, resuspend in generic Hoechst buffer, and add an aliquot of 1% saponin solution to a final concentration of 0.05%. Saponin is a mild detergent that dissolves cholesterol molecules in the cell membranes; this makes the nuclear membranes permeable to 7-AAD.
7. Incubate ≥1 hr at room temperature in the dark, add an aliquot of 1 mg/ml 7-AAD stock solution to a final concentration of 20 µg/ml, and continue staining 1 hr more. Set up flow cytometer 8. Set up and optimize the flow cytometer. Collect ultraviolet (360 nm)-excited blue fluorescence (Hoechst fluorescence), 488 nm-excited green GFP fluorescence (∼510 nm), yellow autofluorescence (∼575 nm), and 488 nm-excited red 7-AAD fluorescence (>630 nm). To avoid “bleeding” of Hoechst fluorescence into the GFP and the 7-AAD channels, time-resolved sample excitation (with spatially separated UV and 488-nm laser beams) must be used. The fluorescence emission maximum of GFP is 508 nm. Cellular autofluorescence covers the region between 500 and 600 nm, but is usually stronger >550 nm. Thus, the yellow autofluorescence can be used to compensate for the influence of green autofluorescence on GFP detection. This allows improved detection of weak GFP fluorescence (see Commentary). During harvesting, cells tend to clump; to obtain meaningful data on a per-cell basis it is essential to resuspend cells immediately before analysis. If this procedure is not sufficient to disrupt cell clumps, it is advised that the samples be passed at least ten times through a 0.5-mm internal diameter 21-G needle. ALTERNATE PROTOCOL 2
ANALYSIS OF CELL PROLIFERATION OF ANTIGEN-EXPRESSING VERSUS NONEXPRESSING CELLS This protocol enables analysis of the proliferative history of cell surface antigen-expressing versus nonexpressing cells. The method involves cell surface antigen labeling with antibodies, followed by cell fixation, detergent treatment, and staining with Hoechst 33258 and 7-AAD. Additional Materials (also see Basic Protocol) FBS-PBS: 2% fetal bovine serum and 0.1 % NaN2 in PBS Primary and secondary antibodies for desired antigens 2% (w/v) paraformaldehyde (see recipe) Phosphate buffered saline (PBS; APPENDIX 2A) 1% (w/v) saponin (Sigma) in PBS 1 mg/ml 7-aminoactinomycin D (7-AAD; Calbiochem) in DMSO Label cells with BrdU 1. After isolation, label cells in culture with 100 µM BrdU as described in the Basic Protocol, step 1. Prepare and fix cells 2. Harvest BrdU-labeled cells by standard procedures in 15-ml screw-capped centrifuge tubes and centrifuge 5 min at 200 × g, room temperature.
Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling
3. Resuspend cell pellets in FBS-PBS, centrifuge again, and resuspend again in CFMPBS at a concentration of 4 × 106 cells/ml. 4. Stain cells with monoclonal antibodies for desired cell surface antigens, such as CD3, CD4, CD8, CD14, CD19, and CCR5, either directly conjugated with fluorescein
7.14.4 Supplement 15
Current Protocols in Cytometry
isothiocyanate (FITC) and phycoerythrin (PE) or with fluorescently labeled secondary antibodies (see UNIT 6.2). 5. Pipet 1 ml 2% paraformaldehyde into a new 15-ml screw-capped cell culture tube. 6. After immunostaining, resuspend cells in 7 ml PBS and add cell suspension dropwise to the tube with 1 ml 2% paraformaldehyde while vortexing at maximum speed (final paraformaldehyde concentration, 0.25%). Let the tubes sit 15 min at room temperature to fix cells. This procedure is intended to minimize the formation of cell clumps during the fixation step.
7. Centrifuge samples 5 min at 200 × g, room temperature, decant supernatant, and resuspend cell pellets in PBS. Wash cells two times with ice-cold FBS-PBS. At this stage samples can be stored overnight or several days at 4°C.
8. Centrifuge cells, resuspend in generic Hoechst buffer, and add an aliquot of 1% saponin solution to a final concentration of 0.05%. Saponin is a mild detergent that dissolves cholesterol molecules in the cell membranes; this makes the nuclear membranes permeable to 7-AAD.
9. Incubate ≥1 hr at room temperature in the dark, add an aliquot of 1 mg/ml 7-AAD stock solution to a final concentration of 20 µg/ml, and continue staining 1 hr more. Set up flow cytometer 10. Set up and optimize the flow cytometer. Collect ultraviolet (360 nm)-excited blue fluorescence (Hoechst fluorescence), 488 nm-excited green (∼525 nm) and yellow fluorescence (∼575 nm) from the FITC- and PE-labeled antibodies, respectively, and 488 nm-excited red fluorescence (>630 nm) from the 7-AAD. To avoid “bleeding” of Hoechst fluorescence into the green (FITC), yellow (PE), and red (7-AAD) channels, time-resolved sample excitation (with the UV laser beam set as the second excitation source) must be used. If only FITC labeling is used (no PE), then 488 nm-excited yellow autofluorescence can be collected and compensation used to improve the detection of low-level FITC staining, as described for GPF detection in Alternate Protocol 1. During harvesting, cells tend to clump; to obtain meaningful data on a per-cell basis, it is essential to resuspend cells immediately before analysis. If this procedure is not sufficient to disrupt cell clumps, it is advised that they be broken up by aspirating the samples using a 0.5-mm internal diameter 21-G needle.
ANALYSIS OF PROLIFERATIVE SURVIVAL In this protocol, the absolute number of proliferating and nonproliferating cells in a sample is determined by calibration for the sample volume analyzed. This calibration is achieved by adding a known number of standard particles (e.g., chicken erythrocyte nuclei; CEN) to the buffer in which all cell pellets are resuspended. The procedure can be used in combination with the generic protocol (see Basic Protocol) or with the protocols for the analysis of GFP-expressing or surface-marker-expressing samples (see Alternate Protocols 1 and 2).
ALTERNATE PROTOCOL 3
Nucleic Acid Analysis
7.14.5 Current Protocols in Cytometry
Supplement 15
Additional Materials (also see Basic Protocol) Chicken erythrocyte nuclei (CEN; BioSure Controls) Label cells with BrdU 1. Add 100 µM BrdU to the cell culture medium of each culture of interest, wrap the plates or flasks with aluminum foil, and continue cell culture for the desired time period. Since BrdU is a photosensitizer, all staining steps have to be performed in subdued light. The duration of BrdU labeling depends on the cell cycle time of the cells under study and on the kind of information required (see Commentary). For instance, if detailed information regarding cell cycle compartment transition times is required, several samples spanning at least two cell cycle periods have to be taken.
Prepare cells 2. Harvest BrdU-labeled cells by standard procedures in 15-ml screw-capped centrifuge tubes and centrifuge 5 min at 200 × g, room temperature. Proceed to cell staining. Pellets may be resuspended in cell culture medium supplemented with 10% FBS and 10% DMSO and stored at −20°C for up to 6 months without detectable degradation.
3. Resuspend CEN in provided storage buffer by vigorously vortexing the dropper bottle with CEN. Add 1 µl CEN per ml of Hoechst buffer. 4. Resuspend cell pellets at 0.5-1.0 × 106 cells per ml in CEN-supplemented generic Hoechst staining buffer and incubate ≥15 min, room temperature, in the dark. Since BrdU is a photosensitizer, all staining steps have to be performed in subdued light.
5. After 15 min of staining in the dark, add an aliquot of 1 mg/ml ethidium bromide stock solution to a final concentration of 2.0 µg/ml, and continue staining 15 min longer in the dark. Set up flow cytometer 6. Set up and optimize the flow cytometer. Collect ultraviolet (360 nm)-excited blue fluorescence (Hoechst fluorescence) and 488 nm-excited fluorescence between 590 and 630 nm (ethidium bromide). Carefully resuspend the cell sample and disrupt cell clumps by gently pipetting up and down a few times immediately before analysis. To avoid “bleeding” of Hoechst fluorescence into the ethidium bromide channel, time-resolved sample excitation (with the UV laser beam set as the second excitation source) can be used, as described in the Basic Protocol. During harvesting, cells tend to clump; to obtain meaningful data on a per-cell basis, it is essential to resuspend cells immediately before analysis. If this procedure is not sufficient to disrupt cell clumps, it is advised that they be broken up by aspirating the samples at least ten times through a 0.5-mm internal diameter 21-G needle.
7. Collect at least 10,000 cell signals for each sample on a computer with the appropriate software. If detailed information regarding cell cycle distributions among two or three successive cell cycles is required, one should acquire 2 or 3 times 10,000 signals. The number of CEN signals is usually between 5% and 10% of total signals, such that between 500 and 3,000 CEN signals will be acquired. If fewer than 500 CEN signals are acquired, a larger total number of signals should be collected to allow for accurate enumeration of CEN. Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Generic Hoechst buffer 100 mM Tris⋅Cl, pH 7.4 154 mM NaCl 1 mM CaCl2 0.5 mM MgCl2 0.2% (w/v) bovine serum albumin (BSA) 1.2 µg/ml Hoechst 33258 (Sigma) Store up to 1 month at 4°C in the dark Paraformaldehyde solution, 2% Weigh out 10 g of paraformaldehyde (J.T. Baker). Measure 500 ml of PBS (APPENDIX 2A) and adjust pH to 12.0 with NaOH or KOH. Heat PBS, pH 12, to 60°C in fume hood, slowly add the paraformaldehyde powder, and stir for 5 to 10 min. Allow to cool to room temperature in the fume hood. At room temperature adjust pH to 7.4. Store up to 1 month at 4°C in the dark. COMMENTARY Background Information Halogenated analogs of deoxyuridine (such as BrdU) are incorporated into DNA in place of thymidine. This incorporation into DNA leads to a specific distortion of the geometry of the DNA helix (Loontiens et al., 1990) such that the fluorescence of Hoechst dyes is quenched (Latt, 1973). This quenching of Hoechst fluorescence allows cells that have incorporated BrdU to be distinguished from those that have not. Analysis of Hoechst 33258-stained cells by flow cytometry has become a simple way to reliably enumerate cells that have incorporated BrdU (Kubbies and Rabinovitch, 1983; Rabinovitch et al., 1988). Counterstaining cells with a dye that exhibits fluorescence proportional to DNA content and fluoresces in a different range of the emission spectrum than that of the Hoechst dyes (i.e., green, yellow, or red fluorescence emission) allows resolution of cells in the G1, S, and G2 phases of the cell cycle. This methodology resolves cells that have undergone one, two, or three cell cycles during the period of labeling with BrdU (i.e., have incorporated BrdU during one, two, or three S phases) and simultaneously distinguishes cells in the G1, S, and G2 phases of these three consecutive cell cycles (Rabinovitch et al., 1988).
Critical Parameters and Troubleshooting Incorporation of BrdU into DNA may lead to an enhanced sensitivity of cells towards cer-
tain drugs (Poot et al., 1991) and to UV light. In this way, BrdU may elicit a cytotoxic response that may lead to a high level of G2 cells. It should be noted that G2 arrest will not destroy the utility of this assay for detecting the proportion of noncycling (G0/G1) cells, but it will prevent the assessment of the progression of cells from one cycle to the next. If the latter is important in the experimental design, it is recommended that each new cell type and drug to be investigated be first tested with a range of BrdU concentrations for BrdU-dependent G2 arrest before the BrdU method is used on a routine basis. If a high level of G2-phase cells is observed, the BrdU concentration in the cell culture medium should be reduced. It has been suggested that BrdU may perturb cellular nucleotide pools in some cells, such as human fibroblasts, which then may affect cell cycling (Poot et al., 1994). To avoid this, addition of 65 µM deoxycytidine to the cell labeling medium has been recommended (Poot et al., 1994). A too-low concentration of ethidium bromide (see Basic Protocol) or 7-AAD (see Alternate Protocols 1 and 2) may lead to a poor coefficient of variation (CV) in this fluorescence parameter. This can easily be remedied by adding more ethidium bromide or 7-AAD and reanalyzing the sample. In addition, increasing the staining time (in particular with paraformaldehyde-fixed samples) may improve CVs for both Hoechst and for ethidium bromide (see Basic Protocol) or 7-AAD (see Alternate Protocols 1 and 2) fluorescence.
Nucleic Acid Analysis
7.14.7 Current Protocols in Cytometry
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Poor quenching of Hoechst fluorescence may be due to poor BrdU incorporation or to insufficient Hoechst staining (Kubbies and Rabinovitch, 1983). The first may result from unusually high levels of thymidine in the serum used to culture the cells. In this case a higher BrdU concentration has to be used. Insufficient Hoechst staining may result from too high a cell density. Diluting the sample with Hoechst staining buffer and adding a compensating amount of ethidium bromide or 7-AAD may improve the cytogram. When samples show excessive clumping it is recommended that samples be passed several times through a syringe with a 0.5-mm internal diameter 21-G needle. If difficulties in distinguishing GFP-expressing and nonexpressing cells are experienced, the experimenter should consult the unit on reporters of gene expression (UNIT 9.12).
Anticipated Results With optimal BrdU labeling and after appropriate staining, a cytogram similar to Figure
7.14.1 should be obtained. It should be noted that the signal clusters representing the G0/G1 and the G1 cells of the second and third cell cycle are on a horizontal line, since they all have identical EB-fluorescence intensity. This result allows for easier analysis of the distribution of signals among the G1, S, and G2 compartments of each cell cycle (see section on data processing below) and is obtained only when two spatially separated laser beams (i.e., UV and 488 nm) are used. If single (UV) excitation is used, the G0/G1 and the G1 clusters appear on a line that passes through the origin of the Hoechst/ethidium bromide cytogram. If the cytograms show distorted images, the experimenter should consult the troubleshooting guide. Processing of data The cytogram shown in Figure 7.14.1 represents the result of an experiment in which cells have progressed through three consecutive cycles. After harvesting, the sample was processed according to Alternate Protocol 3.
128 1st cycle
112 6
Ethidium bromide fluorescence
7 G2/M
96
8
G2/M
3rd cycle S
80 S 64
48
5
G1 2nd cycle
G0/G1
32
16 CEN 0 0
Analysis of Cell Proliferation and Cell Survival by Continuous BrdU Labeling
16
32
80 48 64 Hoechst fluorescence
96
112
128
Figure 7.14.1 Bivariate cytogram of cultured human cells stained with Hoechst 33258 dye (x-axis) and ethidium bromide (y-axis) with chicken erythrocyte standard added (see Alternate Protocol 3). The signal cluster corresponding to resting cells is labeled G0/G1, the cluster of first-cycle cells (S + G2/M) is labeled as such, as are the G1, S, and G2/M cells in the 2nd and 3rd cell cycles, and the cluster of chicken erythrocyte nuclei is labeled CEN.
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Analysis software was used to obtain the numbers of cells in the following cell cycle compartments: (1) cells that have not incorporated BrdU (G0/G1); (2) cells that have incorporated BrdU, but not divided (1st cycle); (3) cells with incorporated BrdU that have divided once (2nd cycle); (4) cells that have divided twice (3rd cycle); and (5) chicken erythrocyte nuclei (CEN). From these data the following parameters can be derived. The percentage of proliferating cells. This number is most meaningful if expressed as the percentage of cells originally in the culture that proliferated. This can be calculated by adding together the number of cells that have incorporated BrdU, but have not divided (1st cycle S plus G2/M), plus the number of cells that have divided once (2nd cycle cells) divided by two (since two cells in the second cycle have resulted from one cell in the first cycle), plus the number of cells that have divided twice (3rd cycle cells) divided by four (since four cells in the third cycle have resulted from one cell in the first cycle), divided by the total number of cells originally plated (the above number plus the G0/G1 cells) multiplied by 100%. Proliferative survival. This is the number of proliferating cells (see above) normalized to the number of chicken erythrocyte nuclei (CEN). In this way the number of proliferating (and surviving) cells can be determined. If the actual number of CENs added to the culture is known, the absolute number of proliferating and surviving cells can be calculated; however it is often easiest to express this number relative to the proliferative survival of untreated control cells. This parameter is particularly informative in systems in which cell death by apoptosis or necrosis takes place. The impact of cell death on the cell total number in a culture is thus assessed. The CEN population is shown “circled” in Figure 7.14.1. Identification of CEN against a background of debris from the larger cells is facilitated by simultaneous analysis of forward and right-angle scatter signals; gating on these two scatter signals plus the two fluorescence parameters usually allows unambiguous identification of CEN (not shown).
Time Considerations For staining and analyzing 12 samples, 1 hr for sample staining and 1 hr for flow cytometric analysis should be allocated. Staining of paraformaldehyde-fixed samples should be ≥2 hr before analysis by flow cytometry.
Literature Cited Kubbies, M. and Rabinovitch, P.S. 1983. Flow cytometric analysis of factors which influence the BrdUrd-Hoechst quenching effect in cultivated human fibroblasts and lymphocytes. Cytometry 3:276-281. Latt, S.A. 1973. Microfluorometric detection of deoxyribonucleic acid replication in human metaphase chromosomes. Proc. Natl. Acad. Sci. U.S.A. 70:3395-3399. Loontiens, F.G., Regenfuss, P., Zechel, A., Dumortier, L., and Clegg, R.M. 1990. Binding characteristics of Hoechst 33258 with calf thymus DNA, poly[d(A-T)], and d(CCGGAATTCCGG): Multiple stoichiometries and determination of tight binding with a wide spectrum of site affinities. Biochemistry 29:9029-9039. Poot, M., Schuster, A., and Hoehn, H. 1991. Cytostatic synergism between bromodeoxyuridine, bleomycin, cisplatin and chlorambucil demonstrated by a sensitive cell kinetic assay. Biochem. Pharmacol. 41:1903-1909. Poot, M., Hoehn, H., Kubbies, M., Grossmann, A., Chen, Y., and Rabinovitch, P.S. 1994. Cell-cycle analysis using continuous bromodeoxyuridine labeling and Hoechst 33358-ethidium bromide bivariate flow cytometry. Methods Cell Biol. 41:327-340. Rabinovitch, P.S. 1983. Regulation of human fibroblast growth rate by both noncycling cell fraction transition probability is shown by growth in 5-bromodeoxyuridine followed by Hoechst 33258 flow cytometry. Proc. Natl. Acad. Sci. U.S.A. 80:2951-2955. Rabinovitch, P.S., Kubbies, M., Chen, Y.C., Schindler, D., and Hoehn, H. 1988. BrdU-Hoechst flow cytometry: A unique tool for quantitative cell cycle analysis. Exp. Cell Res. 174:309-318. Watkins, S. 1989. Immunohistochemistry. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 14.6.1-14.6.13. John Wiley & Sons, New York.
Contributed by Martin Poot, M. Rosato, and Peter S. Rabinovitch University of Washington Seattle, Washington
Nucleic Acid Analysis
7.14.9 Current Protocols in Cytometry
Supplement 15
Ultraviolet-Induced Detection of Halogenated Pyrimidines (UVID)
UNIT 7.15
Halogenated pyrimidines such as bromodeoxyuridine (BrdU) are used to identify DNAsynthesizing cells. Immunocytochemical detection of these DNA precursors when combined with counterstaining of DNA allows one to distinguish S-phase cells from cells in G0/G1 and G2/M. Their detection with monoclonal antibodies, however, requires the denaturation of DNA to make them accessible to the antibody (UNIT 7.7). The harsh conditions needed to denature DNA (acid or heat treatment) can lead to cell loss and abrogation of other cellular markers and antigens. Partial photolysis of BrdU with ultraviolet-B light (UV-B) induces extensive damage to nuclei which incorporated the halogenated pyrimidine. Under hypotonic conditions certain monoclonal antibodies can detect BrdU in the unfolding chromatin. In contrast to other methods, such as treatment with DNase or SBIP (Li et al., 1994a), this approach requires no DNA denaturation step nor any form of enzymatic treatment and allows for the detection of other cellular markers. The Basic Protocol can be used with either coagulative fixation (ethanol) or fixation with cross-linking agents (paraformaldehyde), and allows the simultaneous detection of a wide range of antigens. In the authors’ experience, when using the UVID method, paraformaldehyde offers some advantages over ethanol for certain surface markers and often improves the retention of intracellular antigens. ULTRAVIOLET-INDUCED DETECTION OF BrdU IN ETHANOL- OR PARAFORMALDEHYDE-FIXED CELLS
BASIC PROTOCOL
This protocol describes the ultraviolet-induced detection (UVID) of BrdU in ethanol- or paraformaldehyde-fixed cells. Exponentially growing cells are incubated with BrdU, harvested, and irradiated with UV-B light to partially photolyze the DNA sections that contain incorporated BrdU and to induce nuclear damage. Cells are then fixed in cold ethanol. Hypotonic conditions subsequently allow the detection of BrdU with monoclonal antibodies. The simultaneous detection of other cellular markers is optional. For surface antigens, cells should be stained after UV irradiation and prior to fixation. Intracellular antigens can be detected by adding the antibody to the hypotonic buffer. The counterstaining of DNA with 7-aminoactinomycin D (7-AAD) is optional and is performed simultaneously with the hypotonic treatment. Materials 10 mM BrdU stock solution (see recipe) Exponentially growing cells Phosphate buffered saline (PBS; APPENDIX 2A) with and without 0.1% BSA 70% (v/v) high purity ethanol, −20°C 0.2% (w/v) paraformaldehyde in PBS, 20°C (see recipe) Sodium tetraborate solution (see recipe) Anti-BrdU FITC antibody 7-AAD solution (optional; see recipe) 37°C, 5% CO2 incubator Tabletop centrifuge and swinging bucket rotor Hand-held 8-W UV-B lamp with an intensity of ∼22 W/m2 measured 0.5 cm above the filter surface 14-ml clear polypropylene tubes (e.g., Sarstedt) Contributed by Hans-Joerg Hammers and Peter Schlenke Current Protocols in Cytometry (2001) 7.15.1-7.15.6 Copyright © 2001 by John Wiley & Sons, Inc.
Nucleic Acid Analysis
7.15.1 Supplement 16
Clear tape or rubber band Microcentrifuge Vortex mixer Tubes suitable for use with flow cytometer Distilled water Flow cytometer equipped with a 488-nm argon laser and filters for detection of FITC (525 nm), phycoerythrin (PE; 575 nm), and 7-AAD (650-nm long-pass or 675-nm band-pass) NOTE: Not every clone can be used with the UVID approach. The authors recommend the Bu20a clone (Dako), but the 3D4 clone (PharMingen) and the B44 clone (Becton Dickinson) are acceptable as well. Label DNA-synthesizing cells with BrdU 1. Add 30 µl 10 mM BrdU stock solution per 10 ml exponentially growing cells to obtain 30 µM BrdU final concentration in the culture. Mix well. Optimal incorporation of BrdU requires culture conditions that facilitate exponential growth; these conditions are cell-type dependent. As a rule of thumb, density in suspension cultures should not exceed 1 × 106 cells/ml and cell confluence in adherent cell cultures should be avoided. Up to 4 × 106 cells per sample can be processed; however, for optimal results, 2 × 106 cells per sample are recommended. Avoid any subsequent light exposure of the cell culture.
2. Incubate another 60 min in a 37°C, 5% CO2 incubator. 3. Harvest cells and centrifuge in a tabletop centrifuge with a swinging bucket rotor at 20°C, selecting time and g force appropriate for the cell line used (e.g., 40 min at 300 × g). Partially photolyze BrdU with UV-B CAUTION: Take precautions when working with UV-B. Do not look into light source and avoid irradiation of skin. Work under a hood and use maximal protection from UV light if possible. 4. Turn on hand-held 8-W UV-B lamp and warm up for 5 min. 5. Remove supernatant and resuspend cells in 12 ml PBS/0.1% BSA per sample. 6. Transfer cells into 14-ml clear polypropylene tubes. 7. Place tubes directly on the filter surface of the upright standing UV-B lamp. Fix tube with clear tape or rubber band. Take care that any labeling or frosting on the tube is facing away from the lamp surface. Irradiate 5 min (approximate dose of 6.6 kJ/m2). Under these conditions, with the use of a hand-held 8-W UV-B lamp, cells are irradiated with an average intensity of ∼22 W/m2 (∼2.2 mW/cm2). Cells are irradiated in an upright standing position to avoid partial shielding by the tube cap.
8. Centrifuge cells 10 min at 300 × g, room temperature. After UV irradiation and before fixation, the cells are more sensitive and require gentle handling and pipetting. Optimally, centrifugation conditions should be the same as step 3.
Ultraviolet-Induced Detection of Halogenated Pyrimidines (UVID)
Fix cells 9. Remove supernatant. Carefully resuspend cells in 1000 µl PBS and transfer into microcentrifuge tubes. 10. Microcentrifuge 3 min at 3000 rpm. Remove supernatant and resuspend in 100 µl PBS.
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For ethanol fixation 11a. Place tube containing the cell pellet on vortex mixer and add dropwise 700 µl 70% (v/v) high purity ethanol, −20°C. 12a. Place tube into a −20°C freezer and fix 60 min. For paraformaldehyde fixation 11b. To the tube containing the cell pellet, add 1000 µl 0.2% paraformaldehyde in PBS. 12b. Fix cells 15 min at 20°C. The temperature and time play a crucial role in avoiding overfixation of the cells, especially in laboratories without air conditioning. Alternatively, higher concentrations at lower temperatures can be used (e.g., 0.5% paraformaldehyde on ice for 10 min). Overfixation may result in decreasing sensitivities or abrogation of the BrdU detection.
13. Microcentrifuge 3 min at 3000 rpm. Remove supernatant. 14. Wash with 1000 µl PBS. 15. Microcentrifuge 3 min at 3000 rpm. Remove supernatant and resuspend in 200 µl PBS. NOTE: After this step, cells can be stored overnight at 4°C.
16. Transfer cells to a flow cytometer tube. Stain with anti-BrdU under hypotonic conditions 17. Add 1000 µl distilled water and 50 µl sodium tetraborate solution. 18. Add appropriate amount of anti-BrdU FITC antibody (10 µl for the Bu20a clone and 20 µl for the 3D4 and B44 clones). 19. (Optional) Add 20 µl 7-AAD stock solution. 20. Incubate at room temperature at least 30 min for ethanol-fixed cells or at least 60 min for paraformaldehyde-fixed cells. Cells fixed with paraformaldehyde are stable for at least 2 to 3 hr, while ethanol-fixed cells should be stable overnight.
21. Resuspend by vortexing. Analyze cells on the flow cytometer 22. Set up the flow cytometer with excitation at 488 nm, using dichroic, band-pass, and/or long-pass filters to measure green fluorescence (525 nm) and deep-red fluorescence (>650 nm or 675-nm band-pass). If additional cellular markers are detected with conjugated antibodies, discriminate those signals in the appropriate range (∼575 nm for PE). Compensate for spectral overlap with FITC and 7-AAD (UNIT 1.14). 23. Set the discriminator on 7-AAD fluorescence. 24. If possible use doublet discrimination features (e.g., integral versus peak 7-AAD fluorescence). 25. Measure at low sample flow pressure and acquire data for more than 10,000 cells if possible.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
7-AAD stock solution, 0.5 mg/ml Allow 1 mg 7-AAD (Sigma-Aldrich) to dissolve completely overnight in 2 ml PBS (APPENDIX 2A). Store at 4°C and keep protected from light. BrdU stock solution, 10 mM Dissolve 31 mg BrdU (Sigma-Aldrich) in 10 ml PBS (APPENDIX 2A). Store up to 3 months at 4°C. Keep protected from light by either using a brown bottle or wrapping in aluminum foil. Paraformaldehyde solution, 0.2% Dissolve 100 mg paraformaldehyde (Serva Biochemicals) in 50 ml PBS (APPENDIX 2A). Heat to 50°C until dissolved. Prepare fresh and store at 20°C until use. Sodium tetraborate solution Dissolve 1.9 g sodium tetraborate decahydrate (Na2B4O7⋅10H2O; Sigma-Aldrich) in 50 ml distilled water. Adjust pH with HCl to 8.0. COMMENTARY Background Information
Ultraviolet-Induced Detection of Halogenated Pyrimidines (UVID)
The use of halogenated pyrimidines such as BrdU is a valuable tool to discriminate S-phase cells from other phases of the cell cycle and to provide information about the cell cycle kinetics (Dolbeare, 1995a,b, 1996). Its detection, however, requires denaturation of DNA with harsh physical measures, such as heat or acid treatment, that often abrogate the simultaneous detection of other cellular markers. Alternative strategies employ enzymatic treatment, but suffer from specific disadvantages. Digestion of DNA with DNase I can lead to over- and underdigestion (Carayon and Bord, 1992; Penit and Vasseur, 1993; Takagi et al., 1993) and strand-break labeling of photolyzed BrdU (SBIP) can be confounded with apoptotic events (Li et al., 1994a,b, 1996). The ultraviolet-induced detection (UVID) of BrdU is the first nonenzymatic approach that is able to preserve cellular markers and to avoid enzymespecific artifacts (Hammers et al., 2000). The principle used by this approach is the selective induction of severe oxidative damage to the nucleus and chromatin, which in turn allows the binding of some anti-BrdU clones to their epitope under hypotonic conditions (with these protocols 68 mM NaCl). The mechanism is not fully understood at this time, but data suggest that the unfolding of the chromatin fibers in the hypotonic environment permits the effect. The detection is reversible as the further addition of sodium chloride or other salts will
decrease the signal. The composition of the hypotonic buffer and the pH play an important role as well and were chosen to maximize the signal-to-noise (S/N) ratio. After UV irradiation, cells are fixed with either a coagulative or a cross-linking fixative. Ethanol is the classic coagulative fixative and provides the highest S/N ratio since coagulative fixation does not interfere with chromatin decondensation. It also allows the extraction of low-molecular-weight (LMW) DNA from apoptotic nuclei, identifying them as a subG0/G1 peak. In the authors’ experience, ethanol-fixed cells seem to be more stable under the hypotonic conditions as well and might be advantageous for some antigens when longer incubation periods are necessary. Paraformaldehyde, on the other hand, is currently the most commonly used cross-linking fixative in flow cytometry. It usually provides excellent preservation of cellular markers, but higher concentrations or prolonged incubation times lead to rigid cross-linking of chromatin and inhibit the necessary structural changes that allow the detection of BrdU. For this reason the S/N ratio of paraformaldehyde-fixed cells tends to be lower than with ethanol fixation and therefore strict adherence to the protocol is essential; however, good preservation of surface antigens, low frequency of doublet formation, and a potential to retain intracellular antigens make this approach preferred for many applications.
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104
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BrdU FITC
103 102 101
G2M
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S
0 0
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G1 S G2M 0
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Figure 7.15.1 Left: Exclusion of doublets in the DNA peak versus DNA integral histogram. Middle: Typical bivariate cell-cycle analysis with anti-BrdU FITC and DNA counterstaining. The signal intensity of mid-S phase cells and of the G1 cells can used to calculate a signal-to-noise (S/N) ratio. Right: The corresponding DNA histogram.
Critical Parameters and Troubleshooting BrdU incorporation into DNA-synthesizing cells: The most important single parameter is the condition of the cells to be labeled. Optimal cell culture conditions and appropriate cell densities are imperative to ensure exponential growth and the optimal incorporation of BrdU analog. If cell lines are used, make sure that optimal conditions are applied at least 24 hr before the addition of the precursor. Partial photolysis of BrdU with UV-B: In the authors’ experience the photolysis of BrdU with UV is surprisingly robust as long as frequencies in the UV-B range (280 to 320 nm) are used. UV-C (254 nm) induces the detection as well, but tends to be more destructive and the S/N ratios are lower. UV-A alone cannot replace UV-B; only in combination with Hoechst dyes does UV-A sufficiently photolyze BrdU. The authors found that irradiation for 5 min with a simple 8-W UV-B hand lamp produces reliable and reproducible results. If other UV-B sources are employed, experimentally determine the irradiation time that produces the highest S/N. When the irradiation is prolonged, a decrease of the S/N ratio and damage of the cells is observed. Irradiated cells are generally more sensitive to shear stress and should be treated gently until fixation. Fixation of UV-treated cells: Fixation with cross-linking agents such as paraformaldehyde
requires well-defined conditions. The optimal margin for fixation (time and paraformaldehyde concentration) is narrow, and too extensive fixation results in a decrease in sensitivity of BrdU detection. Hypotonic treatment: The hypotonic treatment described here was successful for a wide variety of different cell types. It should be stressed, however, that cells once exposed to the hypotonic buffer are more sensitive to shear stress than are the unexposed cells. Their resuspension after centrifugation should be carried out with a vortex mixer rather than a pipetting device. In most cases, cells remain stable in the hypotonic buffer for ∼24 hr, but analysis after one hour is preferred. Ethanol fixation seems to be more stabilizing than the mild paraformaldehyde fixation. Keep in mind that the required incubation with BrdU antibody for paraformaldehyde-fixed cells is twice as long as for ethanol-fixed cells. DNA counterstaining: 7-AAD, a GC-specific intercalator, can be used to counterstain the DNA and does not interfere with the BrdU detection. Surprisingly and in contrast to aciddenatured DNA, the nonspecific intercalating dye propidium iodide (PI) strongly suppresses the anti-BrdU FITC signal and cannot be used. The use of 7-AAD has certain advantages, since this dye has a very large Stokes’ shift which allows the simultaneous measurement of other
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cellular markers with PE-conjugated antibodies. Anti-BrdU FITC clones: It should be noted that only a handful of anti-BrdU clones can be used with the UVID procedure. This indicates that the BrdU epitope detected by UVID is different from the one that becomes accessible after DNA denaturation. The authors recommend the Bu20a clone from Dako. Other clones such as 3D4 (PharMingen) and B44 (Becton Dickinson) can be used as an alternative.
Anticipated Results Pulse-labeling of cells (incubation with BrdU for 60 min) leads to the detection of S-phase cells with a characteristic DNA content that places them on the DNA content histograms between G1 and G1/M cells. Successful experiments then depict the typical horseshoelike pattern of the BrdU-FITC versus DNA content scatterplots or contour maps (Fig. 7.15.1).
Time Considerations BrdU incorporation requires 60 min, but 40 min can be sufficient. The UV treatment should take no longer than 5 min. Fixation with ethanol calls for a 40 to 60 min fixation period with 30 min hypotonic treatment. Fixation with paraformaldehyde takes only 15 min, but the hypotonic treatment must be increased to 60 min. Overall time, including the BrdU labeling period should be around 2.5 hr, with the paraformaldehyde protocol being slightly faster (∼15 min) than the ethanol protocol. Hands-on time is considerably shorter, because most of the time required is for incubation purposes.
Literature Cited Carayon, P. and Bord, A. 1992. Identification of DNA replicating lymphocyte subsets using a new method to label the bromo-deoxyuridine incorporated into the DNA. J. Immunol. Methods 147:225-230. Dolbeare, F. 1995a. Bromodeoxyuridine: A diagnostic tool in biology and medicine. Part I: Historical perspectives, histochemical methods and cell kinetics. Histochem. J. 27:339-369.
Dolbeare, F. 1995b. Bromodeoxyuridine: A diagnostic tool in biology and medicine. Part II: Oncology, chemotherapy and carcinogenesis. Histochem. J. 27:923-964. Dolbeare, F. 1996. Bromodeoxyuridine: A diagnostic tool in biology and medicine. Part III: Proliferation in normal, injured and diseased tissue, growth factors, differentiation, DNA replication sites and in situ hybridization. Histochem. J. 28:531-575. Hammers, H.J., Kirchner, H., and Schlenke, P. 2000. Ultraviolet-induced detection of halogenated pyrimidines: Simultaneous analysis of DNA replication and cellular markers. Cytometry 40:327335. Li, X., Traganos, F., Melamed, M.R., and Darzynkiewicz, Z. 1994a. Detection of 5-bromo-2deoxyuridine incorporated into DNA by labeling strand breaks induced by photolysis (SBIP). Int. J. Oncol. 4:1157-1161. Li, X., Traganos, F., and Darzynkiewicz, Z. 1994b. Simultaneous analysis of DNA replication and apoptosis during treatment of HL-60 cells with camptothecin and hyperthermia and mitogen stimulation of human lymphocytes. Cancer Res. 54:4289-4293. Li, X., Melamed, M.R., and Darzynkiewicz, Z. 1996. Detection of apoptosis and DNA replication by differential labeling of DNA strand breaks with fluorochromes of different color. Exp. Cell Res. 222:28-37. Penit, C. and Vasseur, F. 1993. Phenotype analysis of cycling and postcycling thymocytes: Evaluation of detection methods for BrdUrd and surface proteins. Cytometry 14:757-763. Takagi, S., McFadden, M.L., Humphreys, R.E., Woda, B.A., and Sairenji, T. 1993. Detection of 5-bromo-2-deoxyuridine (BrdUrd) incorporation with monoclonal anti-BrdUrd antibody after d eox yribo nu clease treatment. Cytometry 14:640-648.
Key Reference Hammers et al., 2000. See above. This article describes the original work and gives examples of how to combine the method with the simultaneous detection of surface markers.
Contributed by Hans-Joerg Hammers and Peter Schlenke University of Luebeck Luebeck, Germany
Ultraviolet-Induced Detection of Halogenated Pyrimidines (UVID)
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Analysis of DNA Content and Green Fluorescent Protein Expression
UNIT 7.16
Analysis of DNA content by flow cytometry is a powerful and frequently used method to investigate cell-cycle progression. Cell-cycle effects of a specific gene can be studied by measuring cellular DNA content in cells transfected with a vector that drives expression of that gene. To distinguish between transfected and nontransfected cells on the flow cytometer, the gene of interest is generally co-transfected with a reporter gene such as green fluorescent protein (GFP), an intracellular reporter molecule widely utilized for assessment of gene transfer and expression. While wild-type GFP is not well suited for the 488-nm argon-ion laser excitation of routine flow cytometry, “red-shifted” variants of GFP (e.g., EGFP from Clontech Laboratories and Green Lantern from Life Technologies—see also UNIT 9.12) have excitation maxima around 490 nm and emit fluorescence in the green range of the spectrum. Thus, cells carrying the gene of interest that co-express red-shifted GFP can be readily identified on standard flow cytometers equipped with fluorescein filter sets and can consequently be selected by fluorescence-activated cell sorting for further analysis. As the fluorescence intensity of red-shifted GFP expression correlates with gene expression, low-expressing cells can be discriminated on the fluorescent scale from cells with high gene expression and both these subpopulations can be analyzed and sorted differentially. Simultaneous measurement of red-shifted GFP and propidium iodide (PI) fluorescence proves difficult, because PI, the most commonly used dye for cell-cycle analysis by flow cytometry, requires that cells be permeabilized for DNA staining; however, red-shifted GFP accumulates in the cytoplasm and then readily leaks out of permeabilized cells. In contrast, fixation of cells with formaldehyde in high concentration for red-shifted GFP retention leads to DNA histograms with unacceptably high coefficients of variations (CV) for the G0/1 peaks. Thus, an appropriate cell preparation technique for detection of red-shifted GFP expression and cell-cycle progression requires a delicate balance between retaining GFP fluorescence and obtaining adequate DNA histogram resolution. Alternate use of a vital DNA dye (e.g., Hoechst 33342) that enters live cells can prevent problems associated with cell membrane permeabilization. The first protocol (see Basic Protocol) describes a method for cell fixation/permeabilization for combined measurement of red-shifted GFP expression and PI DNA content. Cells are first fixed with formaldehyde followed by treatment with ethanol for cell membrane solubilization, and then stained with PI for DNA content. The second protocol (see Alternate Protocol) describes staining of unpermeabilized cells with the vital dye Hoechst 33342 for combined red-shifted GFP fluorescence and cell-cycle analysis. It is important that this unit be read in conjunction with UNIT 7.5, UNIT 9.5, and UNIT 9.12, which provide further background on nucleic acid analysis and on reporter genes.
Nucleic Acid Analysis Contributed by Ingrid Schmid and Kathleen M. Sakamoto Current Protocols in Cytometry (2001) 7.16.1-7.16.10 Copyright © 2001 by John Wiley & Sons, Inc.
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BASIC PROTOCOL
DNA CONTENT ANALYSIS IN COMBINATION WITH ASSESSMENT OF RED-SHIFTED GREEN FLUORESCENT PROTEIN EXPRESSION IN FIXED/PERMEABILIZED CELLS In this protocol, cells are fixed with 1% formaldehyde, permeabilized with 70% ethanol, and then stained with PI in the presence of ribonuclease A for DNA content. Materials Cells Phosphate buffered saline (PBS; APPENDIX 2A), ∼4°C Fixation solution, ∼4°C (see recipe) 70% ethanol, −20°C Propidium iodide (PI) working solution (see recipe) Centrifuge, 2° to 8°C 37°C water bath 70-µm nylon mesh (e.g., Fisher Scientific’s Spectra/Mesh N, Becton Dickinson’s Falcon Nylon Cell Strainers) Flow cytometer with 488-nm excitation and 530/30 and 585/42 nm band-pass filters or equivalent Additional reagents and equipment for counting cells (APPENDIX 2A) and retroviral transduction of red-shifted GFP into cells (UNIT 9.12). Fix cells with formaldehyde 1. Count cells (APPENDIX 3A). See UNIT 9.12 for retroviral transduction of red-shifted GFP into cells and see Kain (2000) for strategies for cell transfection.
2. Place ∼106 cells into a 12 × 75-mm test tube and wash once with phosphate buffered saline (PBS) by centrifuging 5 min at 300 × g, 2° to 8°C. Preferably, the PBS should be cold (4°C); however, this is not absolutely necessary as the cells will stop growing when exposed to UV light.
3. Remove supernatant by aspiration or rapid decanting and add 500 µl cold (∼4°C) PBS to the cell pellet. Mix gently. Add 500 µl cold (∼4°C) fixation solution and mix again. Incubate 1 hr at 2° to 8°C. Different concentrations of formaldehyde fixative may be needed for optimal retention of red-shifted GFP in various cell types, and for obtaining acceptable coefficients of variations on DNA histograms as discussed below (see Commentary).
Permeabilize cells with ethanol 4. Centrifuge cells 5 min at 300 × g, 2° to 8°C. Remove supernatant by aspiration or rapid decanting and wash once with cold PBS. Add 1 ml 70% ethanol at −20°C dropwise to the cell pellet with the tube sitting on a vortex mixer. Incubate cell suspension overnight at 2° to 8°C. A cell pellet may not be visible after the fixation step with formaldehyde, because fixed cells aggregate less well and therefore tend to spread out at the bottom of the tube. For this reason, careful aspiration is preferable to decanting. Vortexing cells gently during the addition of ethanol can reduce the formation of cell clumps; however, avoid extensive vortexing, because it can lead to cell disruption. Incubation times with 70% ethanol can be shorter than overnight, but should not be less than 2 hr. Analysis of DNA Content and Green Fluorescent Protein Expression
5. Centrifuge cells 5 min at 300 × g, 2° to 8°C. Remove supernatant by aspiration or rapid decanting and add 1 ml propidium iodide (PI) working solution. Incubate cell suspension 30 min in a 37°C water bath in the dark. If needed, filter samples through a 70-µm nylon mesh to remove clumps.
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Acquire fluorescence data on the flow cytometer 6. Set up flow cytometer with 488-nm excitation. Use forward versus side scatter to look at cell size versus cell granularity. Collect red-shifted GFP fluorescence in log amplification with a 530/30 nm band-pass or similar filter, and PI fluorescence in linear amplification with a 585/42 nm band-pass or a similar filter that collects fluorescence above 590 nm. Cells that have been fixed and permeabilized will have altered light scattering properties. Usually, they have less forward-scatter and more side-scatter signal on the flow cytometer compared to untreated cells. Use orange (PI) fluorescence area and the pulse width in addition to height for discrimination of doublets. Alternatively, current DNA analysis software can provide aggregate modeling algorithms to compensate for cell clumps during DNA histogram deconvolution. Always use nonGFP-expressing cells as controls for background fluorescence. To set up photomultiplier tube voltages and compensation of spectral emission overlap between red-shifted GFP and PI, use GFP-expressing cells (i.e., GFP+) that were not stained with PI, and cells that do not express red-shifted GFP that were stained with PI alone. Accurate setup of fluorescence compensation is critical for correct correlation of GFP expression with cell-cycle status (also see UNIT 9.12). Acquire samples on the cytometer at a low flow rate for improved coefficients of variation on DNA histograms. For samples that contain few GFP+ cells, set acquisition counters to collect a minimum of 10,000 GFP+ events. In most cases, at least 10,000 events are needed to obtain accurate estimation of cell frequencies within different cell cycle phases after mathematical modeling for DNA histogram deconvolution (Ott, 1993; Shankey et al., 1993). For best results, analyze the cells on the flow cytometer as soon as possible. Samples can be held longer (e.g., for up to one week at 2° to 8°C); however, the time period will depend on the cell type and the retention of red-shifted GFP.
DNA CONTENT ANALYSIS IN COMBINATION WITH ASSESSMENT OF RED-SHIFTED GREEN FLUORESCENT PROTEIN EXPRESSION IN UNFIXED CELLS
ALTERNATE PROTOCOL
In this protocol cells are stained at 37°C with the cell-permeant DNA dye Hoechst 33342 for combined GFP and DNA content analysis. A cytometer with UV excitation is required. Additional Materials (see also Basic Protocol) Culture medium, 37°C 1 mg/ml Hoechst 33342 stock solution (see recipe) Flow cytometer with 488-nm and 325-nm excitation and 530/30 and 424/44 nm band-pass filters or equivalent Additional reagents and equipment for concurrent dead cell discrimination (UNIT 9.2; optional). Stain cells for DNA content 1. Count cells (APPENDIX 3A). See UNIT 9.12 for retroviral transduction of red-shifted GFP into cells and see Kain (2000) for strategies for cell transfection.
2. Place ∼106 cells into a 12 × 75-mm test tube and centrifuge 5 min at 300 × g, room temperature. 3. Remove supernatant by aspiration or rapid decanting and add to the cell pellet 500 µl of the same medium that was used for growing the cells, prewarmed to 37°C. Mix gently. Add 5 µl 1 mg/ml Hoechst 33342 stock solution and mix again. Incubate 45 min at 37°C. The optimal Hoechst 33342 dye concentration and staining time for different cell types vary, as dye uptake depends on cellular metabolic rates; therefore, both have to be
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determined empirically. In general, dye concentrations between 1 and 10 ìg/ml, and incubation times between 20 and 90 min, will produce DNA histograms with acceptable coefficients of variations (see Commentary and UNIT 7.5 for further details). Because Hoechst DNA staining is performed on unfixed cells, it is possible to use other nonvital DNA dyes (e.g., PI or 7-aminoactinomycin D), for concurrent dead cell discrimination (UNIT 9.2). Both PI and 7-aminoactinomycin D (7-AAD) are excited by the 488-nm line of an argon-ion laser and have emission spectra that can be effectively separated from the emissions of red-shifted GFP and Hoechst 33342, respectively.
Acquire fluorescence data on the flow cytometer 4. Set up flow cytometer with 488-nm excitation and with 325-nm ultraviolet excitation. Use forward versus side scatter to look at cell size versus cell granularity. Collect red-shifted GFP fluorescence in log amplification with a 530/30 nm band-pass or similar filter and Hoechst fluorescence in linear amplification with a 424/44 nm band-pass or similar filter that collects fluorescence between 400 nm and 500 nm. Always use nonGFP-expressing cells as controls for background fluorescence. Use blue (Hoechst) fluorescence width versus area in addition to height for discrimination of doublets. Alternatively, current DNA analysis software can provide aggregate modeling algorithms to compensate for cell clumps during DNA histogram deconvolution. Fluorescence emissions of red-shifted GFP and Hoechst 33342 do not overlap; therefore, compensation between their fluorescence channels is not needed. It is advisable to use cells that were not stained with Hoechst to control for effects that Hoechst dyes can have on cellular background fluorescence. Acquire samples on the cytometer at a low flow rate for improved coefficients of variation on DNA histograms. For samples that contain few GFP+ cells, set acquisition counters to collect a minimum of 10,000 GFP+ events. In most cases, at least 10,000 events are needed to obtain accurate estimation of cell frequencies within different cell cycle phases after mathematical modeling for DNA histogram deconvolution (Ott, 1993; Shankey et al., 1993).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Fixation solution Add 2 g high-purity paraformaldehyde (Polysciences; 2% w/v final) to 100 ml PBS (APPENDIX 2A). Heat to 70°C in a fume hood until the paraformaldehyde completely dissolves (∼1 hr). Cool to room temperature and adjust pH to 7.2 with 0.1 M NaOH or 0.1 M HCl. Store at 2° to 8°C protected from light. The solution is stable for at least 1 month. Check pH periodically. Do not heat the solution above 70°C. For best results, use only very pure preparations of paraformaldehyde (i.e., electron microscopy grade from Polysciences). See Commentary for further details.
Hoechst 33342 stock solution, 1 mg/ml Dissolve 1 mg Hoechst 33342 powder (Molecular Probes) in 1 ml distilled water. Store at 2° to 8°C protected from light for up to 1 month.
Analysis of DNA Content and Green Fluorescent Protein Expression
Propidium iodide (PI) stock and working solutions Stock solution: Dissolve 1 mg PI in 1 ml PBS (APPENDIX 2A). Add 2.5 mg DNase-free ribonuclease A (Sigma-Aldrich) to the solution. Store at 2° to 8°C protected from light for up to 1 month. Working solution: Dilute PI stock solution 1:25 with PBS (APPENDIX 2A). Final concentration of PI is 40 µg/ml. Final concentration of ribonuclease A is 100 µg/ml. Make fresh before staining. Protect from light.
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COMMENTARY Background Information Verification on the flow cytometer of cellular uptake and expression of a candidate gene after transfection requires co-transfection with a marker gene (see Kain, 2000, and UNIT 9.12). One method commonly used for assessment of the frequency of transfected cells is co-transfection with the E. coli lacZ reporter gene that encodes β-galactosidase (Fiering et al., 1991; UNIT 9.5); however, this technique requires the addition of a fluorogenic substrate and can lead to considerable cell death in the cell preparation (Klein et al., 1997). A second method involves co-transfection with genes for the expression of cell surface antigens such as CD19 (Adams et al., 1997) or CD20 (van den Heuvel and Harlow, 1993). Nevertheless, for antigen detection, the cells need to be stained with fluorescently tagged monoclonal antibodies, and since CD20 is a calcium channel (Bubien et al., 1993), its expression can lead to undesirable side effects such as alterations of cell-cycle progression (Kanzaki et al., 1995). More recently, green fluorescent protein (GFP), which shows bright, light-stimulated fluorescence independent from other co-factors and substrates, has been isolated from the jellyfish Aequorea victoria (Chalfie et al., 1994); however, wildtype GFP is excited maximally at 396 nm and has only a minor excitation peak around 475 nm. Thus, it is not well suited for routine 488nm flow cytometry excitation (Ropp et al., 1995). Generation of variants of GFP that have red-shifted excitation maxima around 490 nm and show enhanced fluorescence intensity as compared to wild-type GFP has overcome these limitations (Heim et al., 1995; Ropp et al.,1995; Cormack et al., 1996; Stauber et al., 1998). Cloning of red-shifted variants of GFP into various expression vectors suited for many different cell types has led to the use of this novel genetic reporter system with species-independent expression in a multitude of applications (UNIT 9.12). The development of variants of wild-type GFP that either have altered excitation maxima (e.g., optimized ultraviolet excitation), or show different emission characteristics (e.g., blue emission or yellow-green emission that can be separated effectively from the emission of red-shifted GFP), provides the possibility of simultaneous flow cytometric analysis of the expression of multiple genes in single cells (Ropp et al., 1996; Lybarger et al., 1998).
In order to study the effect of a gene of interest or its expression level on cell-cycle progression, it is desirable to measure redshifted GFP expression in combination with DNA content; however, the unique nature of GFP makes this approach difficult. GFP is expressed in live cells, and due to its small size of 26.9 kD, will leak out of the cytosol when cells are permeabilized with ethanol (Kalejta et al., 1997; Chu et al., 1999). In contrast, most fluorescent DNA dyes suited for high-resolution cell-cycle analysis by flow cytometry cannot pass through intact cell membranes and require permeabilization for cellular entry. To address these problems, researchers have developed membrane-localized green fluorescent protein variants that remain detectable after cell permeabilization with ethanol for combined GFP and cell-cycle analysis (Kalejta et al., 1997, 1999; Jiang and Hunter, 1998). Similarly, it has been shown that red-shifted GFP targeted to the endoplasmic reticulum can be measured in combination with DNA content in unfixed cells that are permeabilized by digitonin (Pestov et al., 1999). In contrast, Chu et al. (1999) demonstrated that it is possible to retain cytoplasmic red-shifted GFP with low-concentration formaldehyde fixation followed by alcohol treatment to achieve adequate cell-cycle profiles. Furthermore, the use of vital DNA dyes that enter unpermeabilized cells, such as the Hoechst compounds (Arndt-Jovin and Jovin, 1977) and the newly developed, deep-red fluorescing anthraquinone number 5 (DRAQ5; Smith et al., 2000), can avoid compromising GFP analysis.
Critical Parameters and Troubleshooting Two critical parameters define any protocol that can be used successfully for simultaneous measurement of cytoplasmic GFP expression and DNA content by staining with nonvital dyes. The first parameter is the need for cell fixation to retain GFP. The second is the requirement for sufficient cell membrane permeabilization to allow nonvital DNA dyes like PI access to the DNA for adequate nucleic acid staining. Conversely, intrinsic cellular factors of dye uptake and efflux are the main determinants for a successful outcome of experiments of combined assessment of GFP expression and cell cycle using vital DNA staining. Nucleic Acid Analysis
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Analysis of DNA Content and Green Fluorescent Protein Expression
Reagents for cell fixation and permeabilization The standard fixative for flow cytometric applications is formaldehyde. It cross-links proteins and when used without detergents or alcohols for membrane solubilization does not permit sufficient dye access to DNA. The preferred reagent for formaldehyde cell fixation is a buffered solution made from paraformaldehyde powder, because some commercial formalin preparations may contain formic acid generated either during manufacturing or by exposure to excessive heat or light. Unbuffered solutions of formalin can contain methanol as a stabilizing agent. Both substances can have detrimental effects on the detection of redshifted GFP fluorescence, which, although it is stable up to pH 11.5, decreases markedly below pH 7.0 (see Clontech Laboratories manual), and which may be altered by methanol denaturation. For laboratories without access to a fume hood for the preparation of formaldehyde fixative from paraformaldehyde, high-quality electron microscopy formalin sealed into ampules is available from Polysciences. GFP is not affected by formaldehyde treatment (Chalfie et al., 1994). The main protocol of this unit (see Basic Protocol), first described by Chu et al. (1999), uses a 1-hr fixation with buffered 1% formaldehyde solution. One percent formaldehyde fixative maintains redshifted GFP fluorescence after ethanol cellmembrane solubilization. Nevertheless, its concentration is low enough to avoid excessive cross-linking of chromosomal proteins and DNA, which has been shown to adversely affect the resolution of DNA histograms (Schmid et al., 1991). For this reason, some commercial fixation and permeabilization reagent systems (e.g., Permeafix from Ortho Diagnostics Systems and Fix and Perm from Caltag Laboratories) are unsuitable for combined analysis of red-shifted GFP expression and cell cycle because their solutions contain cross-linking fixatives at high concentrations. These systems may be able to retain red-shifted GFP fluorescence, but are not optimized for DNA staining and consequently will lead to unacceptably broad peaks on DNA histograms (Kalejta et al., 1997; Schmid, 2000). The procedure outlined in the Basic Protocol uses cell membrane solubilization with 70% ethanol instead of detergent treatment. Because alcohols coagulate proteins and solubilize lipids, they simultaneously fix and permeabilize cellular membranes, as opposed to detergents,
which cannot provide the added fixation that may aid in GFP retention. Vital DNA staining In contrast to nonvital nucleic acid stains such as PI, the benzimidazole compounds Hoechst 33258 and Hoechst 33342 are DNAspecific dyes that are able to cross intact cell membranes (Arndt-Jovin and Jovin, 1977; UNIT 7.5). Of these two dyes, Hoechst 33342 has been used most commonly for determination of DNA content in living cells by flow cytometry (Loken, 1980; Shapiro, 1981; Crissman et al., 1990). Hoechst 33342 is more hydrophobic than 33258, and therefore crosses cell membranes more easily. Moreover, it tends to exert less toxicity than Hoechst 33358, although its general toxicity is highly cell-type dependent (Fried et al., 1982). Hoechst dyes require ultraviolet (UV) excitation, which until recently was available only from large argon-ion lasers supplied on cell sorters. The commercial availability of benchtop flow cytometers equipped with air-cooled helium-cadmium UV-emitting lasers has made their use more practical. Stoichiometric vital Hoechst 33342 DNA staining generally requires dye concentrations between 5 and 10 µM, with the optimal concentration dependent on differences in the balance between cellular dye uptake and/or efflux. As optimal Hoechst staining concentrations are cytotoxic for most cell types (Loken, 1980; Fried et al., 1982), selection and recovery of cells with intact reproductive capacity after cell sorting can be difficult (Loken, 1980; Shapiro, 1995). Because staining with Hoechst 33342 can be performed without cell fixation and permeabilization, this dye readily permits concurrent staining of dead cells which have lost membrane integrity with nonvital DNA dyes such as PI or 7-AAD (for further details see UNIT 9.2) and makes the Alternate Protocol particularly suited for use with GFP-containing cells that show loss of fluorescence of the marker gene after cell membrane permeabilization; however, the precision of DNA histograms from intact cells is not as good as the high-resolution cell-cycle profiles that can be achieved with cell preparations that have been permeabilized. In addition, some cell types have been shown to be refractive to vital DNA staining with Hoechst dyes (Crissman et al., 1990). Moreover, it has been reported that live staining with Hoechst dyes can result in redistribution of DNA-binding proteins, possibly due to alterations in chromatin structure caused by their
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interactions with DNA (Baumann and Reyes, 1999). Thus, experiments that use Hoechst compounds to study the expression of proteins related to cell cycle by combined GFP and DNA content analysis must be carefully monitored by fluorescence microscopy; otherwise, incorrect protein localization may occur.
cellular GFP fluorescence. Thus, it is advisable to confirm the location of GFP fluorescence with fluorescence microscopy. This is an important confirmation step for proteins with known localization, as flow cytometry cannot offer this information.
Anticipated Results Troubleshooting The Basic Protocol has successfully been used on murine leukemic cells, but may not be appropriate for all cell types and for all GFP applications. To determine the effect of cell fixation and permeabilization on red-shifted GFP expression levels, use as controls untreated cells transfected with a red-shifted GFPcontaining plasmid. If red-shifted GFP fluorescence is lost or diminished extensively after cell membrane permeabilization, try to increase the concentration of formaldehyde in the fixation solution gradually. If this is unsuccessful or leads to inadequate PI DNA histograms, try to apply the protocol as outlined in UNIT 9.12 to the cell preparation. To avoid problems that can occur with fixation/permeabilization of GFP-containing cells, use vital DNA dyes such as the Hoechst dyes (Arndt-Jovin and Jovin, 1977; Loken, 1980; Shapiro, 1981) for simultaneous detection of red-shifted GFP and DNA content as described in the Alternate Protocol; however, Hoechst dyes require UV excitation, and as dye uptake and retention vary among cell types, staining conditions have to be optimized individually. Even with individual optimization, certain cell types will yield unsatisfactory DNA histograms, or vital Hoechst DNA staining may not be attainable (Crissman et al., 1990), although on occasion, improvement in the resolution of Hoechst 33342 DNA histograms through addition of membrane potential–modifying compounds such as DiOC5(3) has been accomplished (Crissman et al., 1990; UNIT 7.5). In these difficult cases, the new vital DNA dye DRAQ5 (Smith et al., 2000) may offer a valuable alternative, particularly because DRAQ5 can be excited with 488-nm argon-ion lasers available on standard flow cytometers. Finally, it may be necessary to modify the transfection/transduction procedure and utilize GFP variants that are targeted to cellular membranes (Kalejta et al., 1997, 1999; Jiang and Hunter, 1998; Pestov et al., 1999). Furthermore, any fixation/permeabilization procedure as well as overexpression of the GFP fusion protein (Baumann and Reyes, 1999) may lead to a redistribution of the localization of intra-
Figure 7.16.1 shows the application of the Alternate Protocol to the combined analysis of EGFP and DNA content in 293 T cells. Figure 7.16.1A shows the background fluorescence of nontransduced cells and Figure 7.16.1B shows EGFP expression in cells that were retrovirally transduced with an EGFP-containing vector. Dead cells which have lost membrane integrity are stained with PI. It is critical to set up the flow cytometer with appropriate compensation between green (FL-1) EGFP fluorescence and orange (FL-2) PI fluorescence; for instance, in the example shown here, compensation of FL-2 out of FL-1 is set to 0.4% and compensation of FL-1 out of FL-2 to 18.8%, respectively. Note that regions instead of quadrant markers are used to accurately determine the frequency of live EGFP+ cells. Markers as shown in Figure 7.16.1A and B, set according to the autofluorescence levels in the control sample, lead to underestimation of EGFP+ cells because they do not include EGFPdim+ cells (i.e., with lowlevel EGFP expression) in the positive quadrant. As a result, in the sample shown in panel B, the frequency of 39% EGFP+ cells falling within R1 would be reduced to 29% of cells contained in the lower right quadrant. After the addition of Hoechst 33342 and incubation at 37°C for 45 min, there are slightly more dead cells in the nonEGFP-containing and EGFP-containing sample and both show a small increase in background fluorescence (Fig. 7.16.1C and D) compared to cells that were not stained with Hoechst 33342 (Fig. 7.16.1A and B); however, the frequency and the relative mean fluorescence intensity (RFL) of EGFP+ cells as shown in panel D (%EGFP+ = 42, RFL = 207) compare well to cells displayed in panel B (%EGFP+ = 39, RFL = 201). By setting a gate on EGFP-negative cells versus EGFP+ cells (Fig. 7.16.1E) their DNA content can then be compared (Fig. 7.16.1F: %G1 = 52, %S = 36, %G2+M = 12; Fig. 7.16.1G: %G1 = 49, %S = 39, %G2+M = 12). Results obtained from other experiments may differ markedly from results shown here, as the frequency of GFP+ cells and their expression level vary considerably depending on cell type, experimental conditions, GFP variant, transfection plasmid,
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Figure 7.16.1 Combined analysis of EGFP expression and DNA content in 293 T cells (kindly provided by Dr. K. Morizono) using vital DNA staining with Hoechst 33342 as described (see Alternate Protocol) with 2 µg/ml PI added for dead cell discrimination. EGFP fluorescence versus PI fluorescence of nontransduced cells, not stained with Hoechst 33342 (A) or stained with Hoechst 33342 (C). EGFP fluorescence versus PI fluorescence of cells transduced with an EGFP-containing vector, not stained with Hoechst 33342 (B) or stained with Hoechst 33342 (D). Dot plot (E) of EGFP fluorescence versus Hoechst 33342 fluorescence area displays the same data file as shown in (D) and is gated on PI-negative live cells and on Hoechst fluorescence width versus area for doublet discrimination. Histogram of Hoechst fluorescence of EGFP-negative live cells contained within gate R2 (F) as shown in (E). Histogram of Hoechst fluorescence of EGFP-positive live cells contained within gate R3 (G) as shown in (E).
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promoter sequences, and flow cytometry instrumentation.
Heim, R., Cubitt, A.B., and Tsien, R.Y. 1995. Improved green fluorescence. Nature 373:663-664.
Time Considerations
Jiang, W. and Hunter, T. 1998. Analysis of cell-cycle profiles in transfected cells using a membrane targeted GFP. BioTechniques 24:348-354.
The Basic Protocol will take ∼2 hr on the first day, and 1 hr the following day for sample preparation. If the incubation with 70% ethanol is shortened to 2 hr instead of overnight, the procedure can be completed in ∼5 hr on the same day. Preparation of samples for the Alternate Protocol will take ∼20 min up to 90 min, based on the optimal incubation time for Hoechst 33342 staining. Flow cytometer setup time will differ depending on the instrumentation used; furthermore, the acquisition time per sample can vary widely, because it is contingent on the frequency of GFP+ cells in the cell preparation.
Literature Cited Adams, P.D., Lopez, P., Sellers, W.R., and Kaelin, W.G. Jr. 1997. Fluorescent-activated cell sorting of transfected cells. Methods Enzymol. 283:5972. Arndt-Jovin, D.J. and Jovin, T.M. 1977. Analysis and sorting of living cells according to deoxyribonucleic acid content. J. Histochem. Cytochem. 25:585-589. Baumann, C.T. and Reyes, J.C. 1999. Tracking components of the transcription apparatus in living cells. Methods 19:353-361. Bubien, J.K., Zhou, L.-J., Bell, P.D., Frizzell, R.A., and Tedder, T.F. 1993. Transfection of the CD20 cell surface molecule into ectopic cell types generates a Ca2+ conductance found constitutively in B lymphocytes. J. Cell Biol. 121:1121-1132. Chalfie, M., Tu, Y., Euskirchen, G., Ward, W.W., and Prasher, D.C. 1994. Green fluorescent protein as a marker for gene expression. Science 263:802805. Chu, Y.-W., Wang, R., Schmid, I., and Sakamoto, K.M. 1999. Analysis with flow cytometry of green fluorescent protein expression in leukemic cells. Cytometry 36:333-339. Cormack, B.P., Valdivia, R.H., and Falkow, S. 1996. FACS-optimized mutants of the green fluorescent protein (GFP). Gene 173:33-38. Crissman, H.A., Hofland, M.H., Stevenson, A.P., Wilder, M.E., and Tobey, R.A. 1990. Supravital cell staining with Hoechst 33342 and DiOC5(3). Methods Cell Biol. 33:89-95.
Kain, S.R. 2000. Flow cytometric analysis of GFP expression in mammalian cells. In In Living Color: Protocols in Flow Cytometry and Cell Sorting (R.A. Diamond and S. DeMaggio, eds.) pp. 199-226. Springer-Verlag, Berlin. Kalejta, R.F., Shenk, T., and Beavis, A.J. 1997. Use of a membrane-localized green fluorescent protein allows simultaneous identification of transfected cells and cell cycle analysis by flow cytometry. Cytometry 29:286-291. Kalejta, R.F., Brideau, A.D., Banfield, B.W., and Beavis, A.J. 1999. An integral membrane green fluorescent protein marker, Us9-GFP, is quantitatively retained in cells during propidium iodide-based cell cycle analysis by flow cytometry. Exp. Cell Res. 248:322-328. Kanzaki, M., Shibata, H., Mogami, H., and Kojima, I. 1995. Expression of calcium-permeable cation channel CD20 accelerates progression through the G1 phase in Balb/c 3T3 cells. J. Biol. Chem. 270:13099-13104. Klein, D., Indraccolo, S., von Rombs, K., Amadori, A., Salmons, B., and Günzburg, W.H. 1997. Rapid indentification of viable retrovirustransduced cells using the green fluorescent protein as a marker. Gene Ther. 4:1256-1260. Loken, M.R. 1980. Simultaneous quantitation of Hoechst 33342 and immunofluorescence on viable cells using a fluorescence activated cell sorter. Cytometry 1:136-142. Lybarger, L., Dempsey, D., Patterson, G.H., Piston, D.W., Kain, S.R., and Chervenak, R. 1998. Dualcolor flow cytometric detection of fluorescent proteins using single-laser (488-nm) excitation. Cytometry 31:147-152. Ott, L. 1993. An Introduction to Statistical Methods and Data Analysis, 4th Edition. Duxbury Press, Belmont, Ca. Pestov, D.G., Polonskaia, M., and Lau, L.F. 1999. Flow cytometric analysis of the cell cycle in transfected cells without cell fixation. BioTechniques 26:102-106. Ropp, J.D., Donahue, C.J., Wolfgang-Kimball, D., Hooley, J.J., Chin, J.Y.W., Hoffman, R.A., Cuthbertson, R.A., and Bauer, K.D. 1995. Aequorea green fluorescent protein analysis by flow cytometry. Cytometry 21:309-317.
Fiering, S.N., Roederer, M., Nolan, G.P., Micklem, D.R., Parks, D.R., and Herzenberg, L.A. 1991. Improved FACS-Gal: Flow cytometric analysis and sorting of viable eukaryotic cells expressing reporter gene constructs. Cytometry 12:291-301.
Ropp, J.D., Donahue, C.J., Wolfgang-Kimball, D., Hooley, J.J., Chin, J.Y.W., Cuthbertson, R.A., and Bauer, K.D. 1996. Aequorea green fluorescent protein: Simultaneous analysis of wild-type and blue-fluorescing mutant by flow cytometry. Cytometry 24:284-288.
Fried, J., Doblin, J., Takamoto, S., Perez, A., Hansen, H., and Clarkson, B. 1982. Effects of Hoechst 33342 on survival and growth of two tumor cell lines and on hematopoietically normal bone marrow cells. Cytometry 3:42-47.
Schmid, I. 2000. Intracellular antigen detection by flow cytometry. In In Living Color: Protocols in Flow Cytometry and Cell Sorting (R.A. Diamond and S. DeMaggio, eds.) pp. 524-531. Springer-Verlag, Berlin.
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Schmid, I., Uittenbogaart, C.H., and Giorgi, J.V. 1991. A gentle fixation and permeabilization method for combined cell surface and intracellular staining with improved precision in DNA quantification. Cytometry 12:279-285.
Stauber, R.H., Horie, K., Carney, P., Hudson, E.A., Tarasova, N.I., Gaitanaris, G.A., and Pavlakis, G.N. 1998. Development and applications of enhanced green fluorescent protein mutants. BioTechniques 24:462-471.
Shankey, V.T., Rabinovitch, P.S., Bagwell, B., Bauer, K.D., Duque, R.E., Hedley, D.W., Mayall, B.H., and Wheeless, L. 1993. Guidelines for implementation of clinical DNA cytometry. Cytometry 14:472-477.
van den Heuvel, S. and Harlow, E. 1993. Distinct roles for cyclin-dependent kinases in cell cycle control. Science 262:2050-2054.
Shapiro, H.M. 1981. Flow cytometric estimation of DNA and RNA content in intact cells stained with Hoechst 33342 and Pyronin Y. Cytometry 2:143-150.
Chu et al., 1999. See above.
Shapiro, H.M. 1995. Parameters and probes. In Practical Flow Cytometry, pp. 254-255. John Wiley & Sons, New York. Smith, P.J., Blunt, N., Wiltshire, M., Hoy, T., Teesdale-Spittle, P., Craven, M.R., Watson, J.V., Amos, W.B., Errington, R.J., and Patterson, L.H. 2000. Characteristics of a novel deep red/infrared fluorescent cell-permeant DNA probe, DRAQ5, in intact human cells analyzed by flow cytometry, confocal and multiphoton microscopy. Cytometry 40:280-291.
Key Reference Describes the procedure presented in the Basic Protocol.
Contributed by Ingrid Schmid and Kathleen M. Sakamoto UCLA School of Medicine Los Angeles, California
Analysis of DNA Content and Green Fluorescent Protein Expression
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Analysis of Viral Infection and Viral and Cellular DNA and Proteins by Flow Cytometry Viruses are obligate intracellular parasites that require the host cell replication, transcription, and translation machinery for the production of new viral progeny. They come in many sizes and shapes, and have a wide range of hosts, which include bacteria, plants, and animals. Each viral group provides a unique series of viral-cellular interactions, leading to unique cellular pathologies and/or replication of the virus. As knowledge of viruses has increased so has understanding of the diversity and types of virus-cell interactions that exist. Studies have provided insight not only into virus replication and control of host functions but also into cellular regulatory events, since the virus must utilize the host metabolic systems. A considerable amount of information has been gathered regarding cellular functions such as eukaryotic replication, transcription, translation, and regulation of these events. The ability to assay viral events in each cell in a population provides information that is useful in developing a knowledge base to understand viral replication and pathologies. Flow and static cytometry coupled with other technologies provides the opportunity to monitor the infection process on a single-cell basis.
UNIT 7.17
BASIC PROTOCOL
Flow and static cytometry monitor the infection and quantitate the viral-cellular events, on intact, single, fixed cells, for DNA, RNA, and proteins. This protocol was developed for the study of mammalian viruses, utilizing SV40 as the prototype (Jacobberger et al., 1986; Lehman et al., 1988; Laffin and Lehman, 1994), and to correlate viral proteins (T antigen and the VPs), cellular and viral DNA, and cell cycle proteins, such as the cyclins, PCNA, Cdks, Cdk inhibitors, pRb and p53, as infection proceeded (Friedrich et al., 1993; Whalen et al., 1999; Lehman et al., 2000). Cells are cultured, harvested using trypsinEDTA to obtain a single-cell suspension, and fixed. The cells are stained with a suitable monoclonal or polyclonal primary antibody, then labeled with a FITC-conjugated secondary antibody to the protein of interest and with propidium iodide (PI) for DNA. Positive and negative controls should always be included. A wide range of cells have been analyzed (see below). CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. Materials Cells of interest, monolayer-grown or suspension-grown PBS (see recipe) 10× trypsin-EDTA (Life Technologies), 37°C Wash solution (see recipe) Methanol, ice cold Primary antibody (see recipe) Secondary antibody (see recipe) RNase A solution (see recipe) Propidium iodide (PI; see recipe) Inverted microscope to view monolayer cells and the trypsin-EDTA dispersion 37°C, 5% CO2 incubator 1.0- and 1.5-ml microcentrifuge tubes Variable microcentrifuge (e.g., Fisher Model 95A) Contributed by John M. Lehman, Thomas D. Friedrich, and Judith Laffin Current Protocols in Cytometry (2001) 7.17.1-7.17.9 Copyright © 2001 by John Wiley & Sons, Inc.
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∼50-µm pore size nylon mesh (Nytex) Flow cytometer with 488-nm excitation and filters for collection of green (535-nm band-pass filter) and red (640-nm long-pass filter) fluorescence Harvest cells 1. Remove growth medium from cells and wash cells with PBS. A wide range of cells have been analyzed, including monkey kidney cells (CV-1), human diploid fibroblasts (IMR-90, WI38, etc.), mouse fibroblasts, Syrian hamster fibroblasts, Chinese hamster fibroblasts, and numerous human and animal tumor cell lines. These cells have mostly been monolayer-grown cells; however, suspension-grown cells have also been utilized.
2. Add 1 to 5 ml prewarmed 1× trypsin-EDTA in PBS to cells. Incubate 30 to 60 sec and decant solution. To prevent cell loss and clumping, do not leave cells in trypsin-EDTA any longer than necessary.
3. Place cells in the 37°C, 5% CO2 incubator until they detach; observe with an inverted microscope. Add 1.0 ml wash solution and shake gently to resuspend cells. 4. Transfer cells to a 1.5-ml microcentrifuge tube. Centrifuge in a variable microcentrifuge 15 sec at 5000 rpm, room temperature. Remove supernatant and resuspend the pellet in 1 ml PBS. Repeat centrifugation. Fix cells 5. Resuspend pellet in 20 to 100 µl PBS and add 900 µl ice-cold (−20°C) methanol. Different fixatives can be utilized. Methanol was chosen for the SV40 large T antigen because it maintains antigenicity, results in minimal clumping of the cells during the staining protocol, and provides a strong fluorescence signal from the stained cells.
6. Count cells and adjust concentration to 1 × 106 cells/ml with PBS. Store at −20°C until ready to stain. The length of storage at −20°C may be variable depending on the viral antigen studied. The SV40 large T antigen maintains antigenicity for up to 1 year, and the fluorescence signals generated are comparable over this period.
Stain with primary antibody 7. Microcentrifuge the stored cells in 1.5-ml tubes 15 sec at 5000 rpm, room temperature, to remove the fixative and wash once with 1 ml PBS. Repeat centrifugation. All centrifugation steps are similar for this protocol.
8. Add 0.5 ml primary antibody and suspend cells by gently vortexing. Vortexing is an important step and may be variable depending upon cell type. Gentle vortexing provides a single-cell suspension with minimal clumping and loss of cells.
9. Incubate ≥30 min at 37°C. The time can vary from as short as 15 min to as long as overnight at 4°C. Each antigen and antibody may have different requirements, so staining time and antibody concentration must be experimentally determined. With the SV40 T antigen, the antibody was incubated 30 min at room temperature in most experiments.
Analysis of Viral Infection by Flow Cytometry
10. Wash in PBS to remove unbound antibody. Microcentrifuge 15 sec at 5000 rpm, room temperature, and resuspend pellet in wash solution with gentle vortexing. Repeat centrifugation and resuspension step at least two times.
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Stain with secondary antibody 11. Add 0.5 ml secondary antibody to each tube and vortex gently. 12. Incubate 30 to 60 min at 37°C, or until a detectable signal is obtained. Depending on the amount of antigen per cell and concentration of antibody, the time required to obtain a detectable fluorescence signal may vary.
13. Wash to remove excess antibody. Repeat centrifugation, wash, and vortex as described in step 10. Numerous secondary antibodies from a number of companies have been utilized with good to excellent results. Selection of secondary antibody is dependent upon sensitivity, specificity, stability, and signal intensity detected.
Stain DNA 14. Add 0.5 ml RNase A solution and 0.5 ml PI solution. Vortex gently. Keep cells at 4°C in the dark and analyze within 4 hr. Just before running on the flow cytometer, filter cells through 50-µm nylon mesh to minimize clumps and provide a single-cell population. Cells are stained with PI to detect the quantity of DNA per cell for characterization of cell-cycle changes occurring with infection. This also provides an opportunity to measure the increase in viral DNA as infection proceeds, since both cellular and viral DNA will intercalate the dye between the bases of DNA.
Analyze cells 15. Collect listmode data for green fluorescence (FITC) with a 535-nm band-pass filter and for red fluorescence (PI) with a 640-nm long-pass filter. Gate on light scatter versus red (DNA) fluorescence to eliminate noncellular debris. Then analyze these cells using red area versus red peak to select single cells. Display these gated cells in a third dual histogram of red area versus green area (T antigen; see Fig. 7.17.1). Collect 10,000 to 50,000 cells for each data point. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Phosphate buffered saline without Ca2+ and Mg2+ (PBS) 8.0 g NaCl 0.2 g KCl 1.2 g Na2HPO4 0.2 g KH2PO4 Dissolve in 1 liter distilled water, adjust pH to 7.4, filter sterilize through a 0.22-µm filter, and store ≤1 year at room temperature. Primary antibody (1° Ab) Many of the primary antibodies whether monoclonal or polyclonal may be obtained from the American Type Culture Collection, Oncogene Research Products, and Santa Cruz Biotechnology, Inc. A large number of antibodies have been prepared to many viruses and may be obtained from the above listed companies or directly from the investigators that have published on the antibody. The antibodies used were stable 4 to 5 years stored in small aliquots at −20°C. Nucleic Acid Analysis
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Propidium iodide (PI) solution 10.0 mg propidium iodide (Sigma) 100 ml PBS (see recipe) 20 µl Triton X-100 0.1 g sodium azide This solution has been stored 2 to 3 years at 4°C. Ribonuclease A (RNase A) solution 0.1 g RNase A (Sigma) 100 ml PBS (see recipe) 20 µl Triton X-100 0.1 g sodium azide Boil 1 hr and store ≤1 year at 4°C. Secondary antibody (2° Ab) These antibodies may be purchased from a number of companies and were generally fluorescein isothiocyanate (FITC) labeled. Recently, Alexa Fluor 488 (Molecular Probes) conjugated to secondary antibodies has been used with excellent results. Wash solution 100 ml normal goat serum 900 ml PBS (see recipe) 200 µl Triton X-100 1.0 g sodium azide Heat-inactivate goat serum 60 min at 56°C, filter sterilize through a 0.22-µm filter, and store 6 months at 4°C. COMMENTARY Background Information
Analysis of Viral Infection by Flow Cytometry
The papovaviruses, which include the papillomaviruses and polyomaviruses (Cole, 1996), are small DNA viruses. The polyomavirus virion, SV40, is surrounded by a protein coat composed of the viral proteins VP1, VP2, and VP3, which cover the double-stranded (ds) closed circular DNA (5243 bp) associated with the cellular histones H2A, H2B, H3, and H4 in a chromatin complex. The SV40 genome codes for at least four other proteins. Two major proteins are the T antigens, small t antigen and large T antigen, which are important in virus replication and transformation. Recently, another T antigen, a tiny t, has been described. A fourth late protein, agnoprotein, is involved in the spread of the virus between cells. These viruses have two major options when they infect a cell: (1) cell lysis with production of progeny viruses (permissive infection) and (2) transformation (nonpermissive infection). Because the majority of viral events occur in the nucleus, they are classified as nuclear viruses. The replication cycle (permissive or lytic infection) involves the following steps: (1) adsorption of the virus to the cell; (2) virion entry
and uncoating (nucleus); (3) transcription and translation of early viral mRNA (T antigens); (4) replication of viral DNA; (5) transcription and translation of late mRNA (coat proteins); and (6) assembly of the viral DNA and late proteins into the virion. Transformation (nonpermissive) includes: (1) steps 1 to 3 of the replication cycle; (2) absence of viral DNA replication; (3) integration of all or a portion of the viral genome into the host DNA; (4) failure to transcribe and translate the late mRNA and proteins; and (5) expression of the transformed phenotype by the infected cells. The transformed cell phenotype displays the following characteristics: increased saturation density, growth in soft agar, increased cloning efficiency, loss of contact inhibition, reduced growth factor requirements, cell membrane changes, chromosomal changes, immortality, and tumorigenicity. A considerable amount of the information regarding the transformed cell was first identified with this viral-transforming model and has provided the basis for defining and providing assay systems to study various models of neoplasia.
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The viral components and products may be studied directly after disruption of the virion and/or infected cell to allow the assay and characterization of the viral DNA, RNA, and proteins. A number of procedures have been utilized to study viruses and their effects on the host cell. These include Southern, northern, and western gel analysis, immunoprecipitation, and analytical ultracentrifugation, to name just a few. However, these procedures require disruption of the cell, precluding the analysis of viral events at the single-cell level as the infection proceeds. An area of particular interest in the viral lytic cycle is the production of specific viral proteins and DNA content changes as infection proceeds (Lehman et al., 1988, 1993; Friedrich et al., 1994). The availability of monoclonal and/or polyclonal antibodies directed to both viral and cellular proteins offers the ability to assay a specific molecule in the intact (fixed) cell and the opportunity to correlate viral events with cellular events such as progression through the cell cycle. These studies have provided insight into how the infectious agent utilizes its genetic information and products (protein and RNA) in conjunction with hostcell proteins to initiate and complete viral replication or initiate and maintain the transformed cell. Interest centered on the viral proteins synthesized during the infection and the DNA content changes (both cellular and viral) that occurred as infection proceeds. Cells for these studies were derived from monolayer cell cultures of infected, mock-infected, and transformed populations. The protocol has been used for many other viral agents, such as polyoma virus, cytomegalovirus (CMV), and human immunodeficiency virus (HIV), as well as any viral protein to which a monoclonal or polyclonal antibody is available (Elmendorf et al., 1988; McSharry et al., 1990). Both the lytic and the transforming cell models of SV40 infection have been studied with this technique. Knowledge of the cell model system and how to obtain a single cell suspension is critical. Another important aspect of the procedure is the availability of specific antibody to the viral proteins. SV40 virus has available numerous polyclonal and monoclonal antibodies made to the epitopes of the viral proteins.
Critical Parameters and Troubleshooting This procedure has been utilized in experimental cell-virus model systems, both mono-
layer and suspension culture. Clinical samples—including blood, and bladder and lung washes—have been assayed for specific viral proteins utilizing this technique. Archival material (paraffin-embedded) may be utilized if the antigen (viral) is stable after fixation, paraffin embedding, and removal of paraffin. Solid tissue may also be utilized if a single-cell suspension can be prepared. Clumping is a major concern in all flow cytometry assays and considerable effort may be necessary to obtain a viable single-cell preparation for a particular virus-cell system. The major events leading to clumping are associated with the initial trypsin-EDTA treatment to detach the cells from the substrate. Therefore, careful attention should be addressed to the length of trypsin treatment, temperature, and concentration of the trypsinEDTA. Addition of serum to inactivate the trypsin-EDTA and immediate fixation of the cells will minimize clumping. Once fixed, cells may be retained as a single-cell suspension by paying particular attention to resuspension of the pellet and the centrifugation step. The investigator who has the most experience with the particular virus-cell system under study best determines these steps. Each cell type may present a new set of problems that may need to be addressed with an experiment to determine the optimal conditions. To define the fixation and antibody dilutions required, the assay is performed on cells grown on coverslips, which allows monitoring of the staining by immunofluorescence microscopy. This also provides an opportunity to determine the specificity, location of antigen, and optimal dilution of antibody, both primary and secondary, for the antigen of interest. The use of coverslips and fluorescent microscopy provides an initial dilution, which is used for the flow cytometry assay. Generally, for flow cytometry, the antibodies can be diluted further (10- to 100-fold), suggesting that the assay is more sensitive. This protocol requires that the antibodies be specific and saturating. Therefore, the cell number (antigen) must be constant so that if most cells contain the antigen, antibody is available to react with all epitopes within or on the cells. This may not be an issue if only a small number of cells in the population are antigen positive. The authors have always kept the cell number constant; usually the ideal concentration is 1-1.5 × 106 cells/ml. This concentration allows for cell loss during the staining protocol and for collection of 104 to 105 cells for flow cytometry analysis. The primary
Nucleic Acid Analysis
7.17.5 Current Protocols in Cytometry
Supplement 17
12 hr
36 hr
16 hr
52 hr
T antigen
1000
1 0
127
0
127
0
127
0
127
DNA content Figure 7.17.1 DNA content (linear scale) and T antigen (log scale) of CV-1 cells infected with SV40. Cells were harvested 12, 16, 36, and 52 hr postinfection. Cells above the line drawn across the histograms are infected cells expressing T antigen; they were stained with monoclonal antibody to T antigen (PAb101) followed by Alexa Fluor 488 conjugated to secondary antibody. Cells below the line are uninfected cells. These data demonstrate the progress of infection with (1) the increase in the number of T antigen–positive cells; (2) the increase in quantity of T antigen per cell; and (3) the increase in DNA content per cell. At 12 hr postinfection (pi), the majority of the population is G1 and uninfected. By 16 hr, the cells are in G1, but the majority are T-antigen positive. As the infection proceeds, the DNA reaches a total of 10 to 12C with accumulation of replicating viral DNA and increase in cellular DNA. This protocol allows the detection of infected and uninfected cells, and the determination of quantity of T antigen/cell, DNA content/cell, and progress of the infection on a single-cell basis. Multiple parameters would allow characterization of the replication, transcription, and translation of virus per cell.
Analysis of Viral Infection by Flow Cytometry
antibodies are generally monoclonal and usually prepared from either cell-culture fluid of hybridomas or mouse ascites fluid. Following a 50% cut with ammonium sulfate, the antibodies are desalted and/or purified on a protein A Sepharose column and stored at −20°C in the presence of 0.1% sodium azide. This preparation should not be contaminated with proteases and/or DNases, which will lower the antibody titer and decrease or destroy the DNA staining. A number of secondary antibodies from different companies have been utilized, requiring some experimentation to determine the optimal antibody and concentration. These studies have primarily utilized indirect immunofluorescence, but the authors have also in some studies used a direct immunofluorescence assay (primary antibody conjugated to a fluorochrome). The utilization of the direct procedure will (1) shorten the procedure without loss of specificity; (2) minimize the number of steps in the protocol; and (3) reduce cell loss since the procedure requires fewer steps. The majority of the 2° antibodies have been labeled with fluorescein isothiocyanate (FITC). However, recently, Molecular Probes developed secondary antibody conjugated to Alexa Fluor 488 dye, which provides an increased and stable signal compared to FITC and therefore requires lower concentration of antibody. Furthermore, it is also important to include a positive and negative cell population (control)
in the assay to allow monitoring of the staining protocol of each experiment. This provides a control for the specificity, saturation of the antibody, and reproducibility of average fluorescence value of the control cells. Inclusion of these controls (positive and negative cells) provides a standard to which many experiments, over time, may be compared. If the values of these cells vary, this allows the determination of which step in the protocol is responsible. The authors have always included positive and negative cells as a control for each experiment run on the flow cytometer. The size of the cells should be considered since the size may change during the progress of the infection. Size can be discerned from the scatter parameter, which is routinely included with the saved listmode data. This is important information to know so that size and background fluorescence may be taken into consideration when calculating the results of the experiment. CV-1 cells infected with SV40 increase in DNA, protein, size, etc., as the infection proceeds. It is important to determine the amount of autofluorescence and nonspecific binding of the antibodies by staining both positive and negative cells. The following is performed on positive/negative cells: (1) one series stained with 1° antibody and PI, but no secondary antibody; (2) one series with secondary antibody and PI, but no 1°; (3) one series with PI
7.17.6 Supplement 17
Current Protocols in Cytometry
A
1000
1000
T antigen
mock
54 hr
100
100
10
10
1
1 0
B
32
96
127
0
450
32
64
96
127
450 mock
405
Cell number
64
54 hr
405
360
360
315
315
270
270
225
225
180
180
135
135
90
90
45
45
0
0 0
256
512
768 1023
0
256
512
768 1023
DNA
Figure 7.17.2 (A) T antigen and DNA content of mock- and SV40-infected CV-1 cells. (B) DNA content profiles. By 54 hr pi, the majority of the cells are infected and expressing T antigen. The lower histograms demonstrate DNA content increases obtained in an SV40-permissive infection.
only; and (4) one series with an unrelated antibody of the same isotype as the 1°, with the secondary and PI. These data then allow selection of proper controls for each antibody and cell-virus system once the specificity and background have been determined. It is important to know the level of background fluorescence in order to calculate specific fluorescence/cell versus background/nonspecific fluorescence per cell. Once this measurement is performed, one can calculate the fluorescence attributable to the antigen of interest. The T antigen (and VPs) was an excellent choice as the initial viral protein assayed by this procedure, since it provided a log signal ∼40 to 60 channels above the control (uninfected cell) signal. If the signal for the antigen is in the linear range, careful attention must be focused on the background/nonspecific fluorescence. Therefore, the above experiment to determine the background/nonspecific staining is critical and may require
background subtraction for each channel (data point) collected. Proper safety precautions should be taken, since a virus is an infectious agent. The laboratory and investigators should maintain the proper conditions in handling an infectious agent. The following standards and guidelines are recommended by NIH for use in developing and implementing health and safety operating procedures and practices for both personnel and facilities, and they serve to supplement prevailing federal, state, and local laws and regulations: Biosafety in Microbiological and Biomedical Laboratories, U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, and the National Institutes of Health. HHS Publication No. (CDC) 93-8395. Investigators should follow these procedures and be knowledgeable about handling the agent of interest. For this protocol the cells and virus are fixed with methanol (or
Nucleic Acid Analysis
7.17.7 Current Protocols in Cytometry
Supplement 17
another fixative), thus eliminating the infectious nature of the agent and its products. However, investigators must maintain proper conditions prior to fixation. If the material is not fixed, the investigator must adhere to the biosafety standards for that agent and be concerned with aerosol generation during flow cytometry analysis.
Anticipated Results The results shown in Figures 7.17.1 through 7.17.3 are examples of the types of data that may be obtained. Figure 7.17.1 provides twocolor data of DNA content and T antigen as the infection proceeds. These histograms allow assessment of the efficiency of the infection by gating the infected versus the uninfected population, which provides the number of infected cells and quantity of viral antigen (T antigen) made per cell and population. Figure 7.17.2 shows the results for DNA versus T antigen of control and infected cells. The changes in DNA content between these two populations are
shown in the bottom panel. These results provide data on the progress of the infection and DNA content changes while monitoring the T antigen-positive/negative cells. Figure 7.17.3 shows the DNA content shifts of the infected population compared to a control CV-1 population. The ability to perform this assay enabled the conclusion that multiple S phases occurred during the SV40 permissive infection of CV-1 cells.
Time Considerations
Fixation will require ∼15 to 30 min depending on how much time the trypsin-EDTA needs to disperse the monolayer into a single suspension. The primary and secondary antibody stainings are usually 30 min each with 15 to 30 min for washes to remove the antibodies. The PI staining requires 30 min prior to filtration and running on the flow cytometer. Therefore the procedures for one sample may be completed within 2.5 to 3 hr. However, staining multiple samples, up to 30 to 60 per experiment,
Cell number
24 hr
48 hr
72 hr
DNA
Analysis of Viral Infection by Flow Cytometry
Figure 7.17.3 DNA profiles of uninfected cells (gray) and SV40-infected cells (black) at 24, 48, and 72 hr pi. The infected population has a total increase in DNA content of 10-12C: ∼2C from replication of SV40 DNA (1 × 106 copies) and an additional 6 to 8C of cellular DNA (2C from the first S phase and 4 to 6C from a second S phase resulting from a block to mitosis by the virus replication process).
7.17.8 Supplement 17
Current Protocols in Cytometry
takes considerably longer, and some experiments can extend over 2 days. Usually, the primary antibody is left to stain at 4°C overnight with the rest of the procedure performed the next day at room temperature, except for the 2° antibody, which is incubated at 37°C.
Literature Cited Cole, C. 1996. Polyomavirinae: The viruses and their replication. In Fundamental Virology, 3rd ed. (B.W. Fields, D.M. Knipe, and P.M. Howley, eds.) pp. 917-945. Lippencott-Raven Press, Philadelphia.
McSharry, J.J., Costantino, R., Robbiano, E., Echols, R.M., Stevens, R., and Lehman, J.M. 1990. Detection and quantitation of human immunodeficiency virus-infected peripheral blood mononuclear cells by flow cytometry. J. Clin. Microbiol. 28:724-733. Whalen, B., Laffin, J., Friedrich, T.D., and Lehman, J.M. 1999. SV40 small t antigen enhances rereplication and alters expression of G1 regulatory proteins in permissive CV1 cells. Exp. Cell Res. 251:121-127.
Key References Jacobberger et al., 1986. See above.
Elmendorf, S., McSharry, J., Laffin, J., Fogleman, D., and Lehman, J.M. 1988. Detection of an early cytomegalovirus antigen with two-color quantitative flow cytometry. Cytometry 9:254-260.
Initial description of the protocol for viral protein and DNA detection.
Friedrich, T.D., Laffin, J., and Lehman, J.M. 1993. Hypophosphorylated retinoblastoma gene product accumulates in SV40 infected CV-1 cells acquiring a tetraploid DNA content. Oncogene 8:1673-1677.
A review of the protocol with specific comments addressing the steps and potential problems.
Friedrich, T.D., Laffin, J., and Lehman, J.M. 1994. Induction of tetraploid DNA content by simian virus 40 is dependent on a T antigen function in G2 phase of the cell cycle. J. Virol. 68:40284030. Jacobberger, J.W., Fogleman, D., and Lehman, J.M. 1986. Analysis of intracellular antigens by flow cytometry. Cytometry 7:356-364. Laffin, J. and Lehman, J.M. 1994. Detection of intracellular virus and viral product. Methods Cell Biol. 41:543-557. Lehman, J.M., Laffin, J., Jacobberger, J., and Fogleman, D. 1988. Analysis of simian virus 40 infection of permissive CV-1 cells by quantitative two-color fluorescence with flow cytometry. Cytometry 9:52-59. Lehman, J.M., Friedrich, T.D., and Laffin, J. 1993. Quantitation of simian virus 40 T antigen correlated with the cell cycle of permissive and nonpermissive cells. Cytometry 14:401-410. Lehman, J.M., Laffin, J., and Friedrich T.D. 2000. Simian virus 40 induces multiple S phases with the majority of viral DNA replication in the G2 and second S phase in CV-1 cells. Exp. Cell Res. 258:215-222.
Laffin and Lehman, 1994. See above.
Internet Resources http://library.thinkquest.org/23054/basics/ index.html Information on general virology. http://www.asmusa.org/ Web page for The American Society for Microbiology (Virology). http://www.isac-net.org/ Web site for the International Society for Analytical Cytology. http://www.atcc.org/ Web site for the American Type Culture Collection: cells, viruses, and hybridomas. http://www.orcbs.msu.edu/biological/BMBL/ BMBL-1.htm Site providing Biosafety in Microbiological and Biomedical Laboratories publication.
Contributed by John M. Lehman, Thomas D. Friedrich, and Judith Laffin Albany Medical College Albany, New York
Nucleic Acid Analysis
7.17.9 Current Protocols in Cytometry
Supplement 17
Apoptosis Signaling Pathways In the past few years much has been learned about the molecular signals governing apoptosis, or programmed cell death, in lymphocytes and other cells of the immune system (for reviews see van Parijs and Abbas, 1996; Lenardo et al., 1998). Two major pathways, active and passive apoptosis, have been identified. Active apoptosis, also termed propriocidal cell death or antigen-induced cell death, occurs when cells are stimulated through a family of TNF-related receptors termed death receptors. These receptors are up-regulated in activated lymphocytes and trigger apoptosis chiefly in effector cells that have been recently stimulated through the antigen receptor. Passive or lymphokine withdrawal apoptosis occurs when activated lymphocytes are deprived of essential growth cytokines and does not require death receptors. Cell death initiated by either pathway is carried out by a unique family of intracellular cysteine proteases, the caspases. Defects in each of these pathways produce distinct pathologies and can synergize to allow accumulation of excess and autoimmune lymphocytes (Reap et al., 1995).
ACTIVE APOPTOSIS Death receptor–induced apoptosis serves to eliminate potentially harmful lymphocytes during responses against chronically expressed antigens. To allow the initial expansion of cells against these antigens, death receptor–induced apoptosis is regulated at a number of points during lymphocyte activation. Expression of Fas (CD95) and tumor necrosis factor (TNF) receptors is up-regulated during initial lymphocyte activation, but sensitivity to apoptosis lags behind this step, indicating that other mechanisms must be operative. The state of T cells in the cell cycle may be important, as progression through G1 and into S phase has been shown to be necessary for T cell receptor (TCR)–induced apoptosis (Boehme and Lenardo, 1993; Lissy et al., 1998). In activated T cells, Fas-ligand (FasL) and TNF expression are up-regulated after reengagement of antigen receptors, which allows cell “suicide” through Fas/FasL interactions, even on a single cell (Brunner et al., 1995; Dhein et al., 1995; Ju et al., 1995). However, mixing experiments have shown that only those T cells that are restimulated actually undergo apoptosis, indicating that a “competency-todie” signal that is separate from FasL up-regu-
lation emanates from the TCR (Hornung et al., 1997). B cells are also sensitized to undergo apoptosis when stimulated through surface Ig, but do not express FasL, and are thus dependent on other cell types to mediate Fas-induced cell death (Rathmell et al., 1995; Foote et al., 1996). These mechanisms cooperate to allow elimination of chronically stimulated B and T cells during viral infections and also participate in the elimination of self-reactive activated peripheral lymphocytes, while bystander cells activated by other stimuli are relatively preserved. In this way, death receptor signaling restrains the proliferation of antigen-specific effector cells during an immune response. The importance of death receptor signaling is illustrated by the phenotype of lpr/lpr or gld/gld mice, which are deficient in Fas and FasL, respectively (Watanabe-Fukunaga et al., 1992; Lynch et al., 1994). These mice have massive accumulation of abnormal T cells and autoantibody-mediated disease. A similar syndrome has been described in humans resulting from dominant negative mutations in Fas (Fisher et al., 1995). There are now five described death receptors in humans (Table 7.18.1), and their relative roles in apoptosis in different cell types is just beginning to be studied. Death receptors, like other TNF family members, recruit intracellular signaling molecules after oligomerization at the plasma membrane in response to soluble or membranebound ligands. In the case of Fas, as illustrated in Figure 7.18.1, this involves recruitment of the adapter molecule FADD and caspase-8 through protein-protein interaction domains. Fas recruits FADD via the death domain, and FADD recruits the pro-caspase-8 or FLICE molecule via death effector domains. Recent structural studies have shown that the death domain and death effector domains share the same basic structure containing six α-helices, but heterotypic interaction between proteins with these domains has not been found. Viral and cellular proteins containing death effector domains have recently been identified (FLICE inhibitory proteins, or FLIPs). These proteins can interfere with the FADD/caspase-8 interaction and block apoptosis induced by a number of death receptors (Bertin et al., 1997; Han et al., 1997; Hu et al., 1997; Irmler et al., 1997; Shu et al., 1997; Srinivasula et al., 1997; Thome
Contributed by Richard M. Siegel and Michael J. Lenardo Current Protocols in Cytometry (2002) 7.18.1-7.18.10 Copyright © 2002 by John Wiley & Sons, Inc.
UNIT 7.18
Nucleic Acid Analysis
7.18.1 Supplement 21
Supplement 21
TRAMP, WSL, APO-2, LARD
TRAIL-R1
TRAIL-R2
TRAIL-R3, DcR1
TNFR2
DR3
DR4
DR5
TRID
aND, not defined.
TRAIL- DcR2 R4
CD120b
TNFR1
42 (predicted)
27 (predicted)
45 (predicted)
50 (predicted)
45 (predicted)
75-80
55-60
45-50
Fas
DR1, APO-1, CD95 DR2, CD120a
Mol. wt. (kDa)
TRAIL
TRAIL
TRAIL
TRAIL
TWEAK (APO-3)
TNF
TNF
FasL
Ligand
Can heterodimerize with DR4, induced by DNA damage GPI-linked extracellular receptor; blocks TRAIL-induced apoptosis
NDa
NDa
Incomplete death domain; blocks TRAIL-induced apoptosis
May bind FADD, but no requirement for FADD in DR4-induced death found in FADD knockout mice
NDa
In mature CD8 T cells, appears to mediate most TCR-induced apoptosis Expression restricted to lymphocytes; induced after DNA damage via p53-dependent mechanism
No intracellular domain
References
Pan et al. (1997); Walczak et al. (1997); Wu et al. (1997) Degli-Esposti et al. (1997b); Pan et al. (1997); Sheridan et al. (1997) Degli-Esposti et al. (1997a); Marsters et al. (1997)
Schneider et al. (1997a,b); Yeh et al. (1998)
Chinnaiyan et al. (1996); Kitson et al. (1996); Screaton et al. (1997)
Zheng et al. (1995)
Trauth et al. (1989); Itoh et al. (1991); Oehm et al. (1992) Cytotoxicity can be modulated by NF-κB Loetscher et al. (1990); Schall et al. (1990)
Most consistently pro-apoptotic
Comments
TRADD, FADD (indirect), TNFR1, caspase-10
TRADD, RIP (indirect), FADD (indirect), TRAF2 (indirect) TRAF-1/2
FADD, caspase-8
Interacting molecules
Death Receptors and Their Signaling Pathways
Molecule Other names
Table 7.18.1
Apoptosis Signaling Pathways
7.18.2
Current Protocols in Cytometry
death ligand + receptor
ALPS mutants/decoy receptors
death receptor
death receptors (e.g., Fas/CD95)
pro-caspase-8
FADD active caspase-8 (activator caspases)
FLIP lymphokine withdrawal
14-3-3 BAD
BAD-P
Akt*
BID
signal integration
Bcl-2 Bcl-x
mitochondrial permeability transition
pro-caspase-9 (APAF-3) active caspase-9
p35, IAPs caspase-3,-6,-7 (effector caspases)
dATP
cytochrome c (APAF-2)
Bcl-2 APAF-1 Bcl-x
cleavage of cellular substrates (e.g., ICAD, PARP, DNAPK, actin)
cytoskeletal reorganization/cellular shrinkage
execution
plasma membrane phosphaditylserine exposure DNA cleavage
Figure 7.18.1 Induction of apoptosis by death receptors and lymphokine withdrawal. Four major steps in apoptosis signaling are depicted. The Fas/CD95 receptor is shown as an example of deathreceptor signaling. Ligand binding triggers trimerization and recruitment of FADD through the death domain (grey rectangle). Pro-caspase-8 is then recruited through death effector domain interactions (hexagons). Oligomerization of caspase-8 triggers its proteolytic cleavage into the active p17 and p11 subunits. Apoptosis can then proceed via a direct pathway (right arrow), involving cleavage of effector caspases, or an indirect pathway, requiring release of cytochrome c and activation of caspase-3 via APAF-1. The indirect pathway of death-receptor signaling, possibly mediated by BID, is shown by the left arrow. One possible pathway by which lymphokine withdrawal might trigger apoptosis is also depicted. Other mechanisms, such as a change in the Bcl-2/Bax ratio, have also been described (Broome et al., 1995). Once effector caspases are activated, the execution phase of apoptosis inevitably proceeds. Abbreviations: Akt*, activated Akt; ALPS, autoimmune lymphoproliferative syndrome; DNA-PK, DNA protein kinase; ICAD, inhibitor of caspase-activated DNase; PARP, poly(ADP-ribose) polymerase.
Nucleic Acid Analysis
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Apoptosis Signaling Pathways
et al., 1997; Rasper et al., 1998). Clustering of caspase-8 in the Fas signaling complex triggers proteolytic autoactivation of the protease encoded by this protein, which initiates a cascade of caspase activation to produce apoptosis. Other death receptors can activate caspases but produce distinct responses, probably because they associate with different adapter molecules (Table 7.18.2). The exact connections between specific death receptors and the intracellular signaling machinery is currently under study. The execution phase of apoptosis is associated with activation of the caspase family of cysteine proteases (for reviews see Cohen, 1997; Nicholson and Thornberry, 1997). The caspases are a distinct family of cysteine proteases that cleave substrates at characteristic sequences containing aspartate in the P1 position (Table 7.18.3). Each caspase is produced as a pro-form that encodes a prodomain of variable length attached to peptides encoding the large and small catalytic subunits of the enzyme. These segments are separated by spacer segments and caspase cleavage sites. Biochemical and genetic studies have enabled the death-associated caspases to be divided into a number of subgroups. Activator caspases (caspase-8, -9, and -10) appear to function primarily by activating other downstream caspases, as their cleavage specificities closely match the cleavage sites of other caspases. This is understood most clearly in the Fas pathway, in which experimental activation of caspase-8 through oligomerization can carry out the function of the Fas signaling complex, trigger downstream caspases, and produce apoptosis (Martin et al., 1998; Muzio et al., 1998; Yang et al., 1998). Effector caspases (caspase-3, -6, and -7) are the major active caspases present in apoptotic cells, and have specificities matching cellular substrates whose cleavage is known to be important in generating the apoptotic phenotype. Some specific features of apoptosis, such as DNA cleavage, have been linked to caspases (Enari et al., 1998; Sakahira et al., 1998). Most likely, the cleavage sites and short prodomains of effector caspases prohibit autoprocessing, making their activity dependent on upstream events (Thornberry et al., 1997). Caspase-2 has substrate preferences resembling the effector caspases but has a long prodomain that can bind the adapter molecule RAIDD through a caspase recruitment domain (CARD), a protein–protein interaction domain similar to the death effector domain (Duan and Dixit, 1997). Thus, this caspase may be a more upstream effector
caspase. Studies with caspase-deficient mice created through gene targeting studies suggest that there is significant redundancy between caspases that varies between cell types. Caspase-3-deficient mice have impaired neuronal apoptosis but normal lymphocyte apoptosis (Kuida et al., 1996), whereas caspase-2-deficient animals seem to have a specific defect in germ-cell apoptosis (Bergeron et al., 1998).
PASSIVE APOPTOSIS Although also dependent on caspases, passive apoptosis in lymphocytes does not occur through death-receptor signaling. Instead, this type of cell death occurs in response to the withdrawal of trophic cytokines. During an immune response, much of the cell loss that occurs after peak expansion may be caused by lymphokine withdrawal. Artificially maintaining high interleukin 2 (IL-2) concentrations can prolong the lifespan of T cells responding to a superantigen challenge (Kuroda et al., 1996). The Bcl-2 oncoprotein and related molecules are known to suppress this form of apoptosis. Overexpression of Bcl-2 can prolong B cell survival as well as memory B cell responses, indicating the physiological importance of this death pathway (McDonnell et al., 1990; Nunez et al., 1990, 1991). Although it has been known for many years that anti-apoptotic proteins of the Bcl-2 family could block passive apoptosis, the signals by which lymphokine withdrawal initiates apoptosis are unclear. One possible mechanism was found in IL-3-dependent cell lines, in which IL-3 maintains phosphorylation of the pro-apoptotic Bcl-2 family member BAD through the phosphatidylinositol-3′-kinase (PI3K)/Akt pathway. 14-3-3 proteins sequester phosphorylated BAD, preventing interaction with other Bcl-2 family members on intracellular membranes. After IL-3 withdrawal, BAD is dephosphorylated and released from the 143-3 protein, and can then form inhibitory dimers with other Bcl-2 family members to block the anti-apoptotic function of these proteins (del Peso et al., 1997). In other cell types, levels of Bcl-2 have been shown to drop after lymphokine withdrawal, which may also predispose these cells to apoptosis (Broome et al., 1995). Whereas death receptors can activate caspases directly, Bcl-2 family proteins control a separate pathway associated with the mitochondrial release of cytochrome c and possibly other pro-apoptotic factors. Although cytochrome c is normally an essential component of mitochondrial respiration at the mitochon-
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Nucleic Acid Analysis
7.18.5
Current Protocols in Cytometry
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27
74
34
23 (predicted)
FADD
RIP
TRADD
RAIDD (CRADD) DAXX
120
Mol. wt. (kDa)
TNF-R1 via death domain RIP via death domain Fas or TNF-R1 death domains
TNF-R1 via death domain and TRADD
Fas via C-terminal death domain
Upstream molecule Death domain protects from Fas-mediated apoptosis; death effector domain induces apoptosis without Fas cross-linking Induces cell death via death domain
Caspase-8 (and caspase-10) via death effector domain
No direct downstream molecule identified
References
Hsu et al. (1995)
Stanger et al. (1995); Impaired NF-kB Ting et al. (1996); activation and hypersensitivity to death Kelliher et al. (1998) following TNF treatment
Boldin et al. (1995); Impaired Fas-mediated apoptosis; block in T cell Chinnaiyan et al. (1995) mitogenesis
Effects of blocking or deficiency
Ahmad et al. (1997); Duan and Dixit (1997) Activates JNK; synergizes with Inhibition of Fas-induced Yang et al. (1997) Fas for induction of apoptosis apoptosis
(1) TRAF-2 via intermediate domain; substrates not clear (IKK is not a direct substrate); (2) RAIDD via death domain TRAF-2, RIP, FADD Causes cell death through recruitment of FADD ICH-1 via CARD domain Induces apoptosis
Effects of overexpression
Downstream molecule
Adapter Proteins in the Death Pathway
Name
Table 7.18.2
7.18.6
Supplement 21
Current Protocols in Cytometry
DXXD, DEXD
Effector
Activator
Caspase-3 (CPP32, Yama, apopain), caspase-6 (Mch2), caspase-7 (Mch3, ICE-LAP3, CMH-1)
Caspase-8 (FLICE, MACH, Mch5), caspase-10 (Mch4, FLICE-2)
Activator
DEHD
Effector/ activator
Caspase-2 (ICH-1, NEDD-2)
Caspase-9 (ICE-LAP-6, Mch6)
(W/L)EHD
ICE-related
Caspase-1 (ICE), caspase-4 (TX, ICE-REL II, ICH-2), caspase-5 (ICE-REL III) caspase-11, caspase-12
Caspase-3, PARP
LETD Caspases 1-10 (caspase-8), IEAD (caspase-10) LEHD
Inhibitors
RAIDD?, caspase-3
YVAD-CHO Caspase-11 required for caspase-1 activation; growth-factor withdrawal and ischemia can activate ICE subfamily proteases
Activators
APAF-1/ cytochrome c
Oligomerization, aggregation in complexes with FADD
Caspase-3 and -6 may be major active species present in apoptotic cells; caspase-3 deficiency disturbs neuronal apoptosis during development
Essential in germ cell apoptosis; may inhibit neuronal apoptosis
Secreted IL-1 probably required for apoptosis induction
Comments
Forms a complex with Bcl-2 family members and APAF-1 (ced-4 homologue), which cleaves caspase-3 when active
zVAD-FMK, Crm Important for death A (caspase-8 >> receptor–activated caspase-10), FLIPs apoptosis (role in other forms of apoptosis not clear)
Upstream caspases DEVD-CHO Many: PARP, (e.g., caspase-8, -9) c-IAP family S-REBP, lamin (caspase-6 only), DNA-PK, actin, PKC, D4-GDI, ICAD, Rb, caspase-9
Pro-IL-1β, γ-interferon inducing factor
Ideal Protein targets cleavage site
Subfamily
Caspase Subfamilies
Caspase(s)
Table 7.18.3
Li et al. (1997); Zou et al. (1997); Pan et al. (1998)
Boldin et al. (1996); Muzio et al. (1996); Scaffidi et al. (1997)
Nicholson et al. (1995); Kuida et al. (1996); Takahashi et al. (1996); Faleiro et al. (1997); Rosen and Casciola-Rosen (1997); Sakahira et al. (1998)
Bergeron et al. (1998)
Friedlander et al. (1996); Wang et al. (1998)
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Nucleic Acid Analysis
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INTERNET RESOURCES http://www.apopnet.com A commercial site with links to tutorials, images, and reagent resources. http://www.access.digex.net/∼regulate/apolist.html A listing of apoptosis sites maintained by Trevigen, a biotechnology company.
Contributed by Richard M. Siegel and Michael J. Lenardo National Institute of Allergy & Infectious Diseases Bethesda, Maryland
Apoptosis Signaling Pathways
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Current Protocols in Cytometry
Flow Cytometry of Apoptosis
UNIT 7.19
This unit describes the most common methods applicable to flow cytometry that make it possible to: (1) identify and quantify dead or dying cells, (2) reveal a mode of cell death (apoptosis or necrosis), and (3) study mechanisms involved in cell death. Gross changes in cell morphology and chromatin condensation, which occur during apoptosis, can be detected by analysis with laser light beam scattering. An early event of apoptosis, dissipation of the mitochondrial transmembrane potential, can be measured using a number of fluorochromes that are sensitive to the electrochemical potential within this organelle (see Basic Protocol 1). Another early event of apoptosis, caspase activation, can be measured either directly, by immunocytochemical detection of the epitope that characterizes activated caspase (see Basic Protocol 2), or indirectly by immunocytochemical detection of the caspase-3 cleavage product, the p85 fragment of poly(ADP-ribose) polymerase (see Basic Protocol 4). Exposure of phosphatidylserine on the exterior surface of the plasma membrane can be detected by the binding of fluoresceinated annexin V (annexin V-FITC); this assay is combined with analysis of the exclusion of the plasma membrane integrity probe propidium iodide (PI; see Basic Protocol 5). Also described are methods of analysis of DNA fragmentation based either on DNA content of cells with fractional (“sub-G1”) DNA content (see Basic Protocol 6 and Alternate Protocol 1) or by DNA strand-break labeling (Terminal deoxynucleotidyltransferase–mediated dUTP Nick End Labeling, TUNEL; or In Situ End Labeling, ISEL; see Basic Protocol 7). Still another hallmark of apoptosis is the activation of tissue transglutaminase (TGase), the enzyme that crosslinks protein and thereby makes them less immunogenic. Methods for analyzing TGase activation are presented in Basic Protocol 8 and Alternate Protocol 2. STRATEGIC PLANNING The choice of a particular method often depends on the cell type, the nature of the inducer of apoptosis, the desired information (e.g., specificity of apoptosis with respect to the cell cycle phase or DNA ploidy), and technical restrictions. For example, sample transportation or prolonged storage before the measurement requires prior cell fixation, thereby eliminating the use of “supravital” methods that rely on analysis of freshly collected live cells. Positive identification of apoptotic cells is not always simple. Apoptosis was recently defined as a caspase-mediated cell death (Blagosklonny, 2000). Activation of caspases, therefore, appears to be the most specific marker of apoptosis (Shi, 2002). The detection of caspase activation, either directly (e.g., by antibody that is reactive with the activated enzyme; see Basic Protocol 2) or indirectly by the presence of poly(ADP-ribose) polymerase (PARP) cleavage product (PARP p85; see Basic Protocol 4), provides the most definitive evidence of apoptosis. Extensive DNA fragmentation is also considered as a specific marker of apoptosis. The number of DNA strand breaks in apoptotic cells is so large that intensity of their labeling in the TUNEL reaction (see Basic Protocol 7) ensures their positive identification and discriminates them from cells that have undergone primary necrosis (Gorczyca et al., 1992). However, in the instances of apoptosis when internucleosomal DNA degradation does not occur (Collins et al., 1992; Catchpoole and Stewart, 1993; Ormerod et al., 1994; Knapp et al., 1999), the number of DNA strand breaks may be inadequate to distinguish apoptotic cells by the TUNEL method. Likewise, in some instances of apoptosis, DNA fragmentation stops after the initial DNA cleavage to fragments of 50 to 300 kb (Collins et al., 1992, Oberhammer et al., 1993). The frequency of DNA strand breaks in nuclei of these cells is low, and therefore, they may not be easily detected by the TUNEL method. Contributed by Piotr Pozarowski, Jerzy Grabarek, and Zbigniew Darzynkiewicz Current Protocols in Cytometry (2003) 7.19.1-7.19.33 Copyright © 2003 by John Wiley & Sons, Inc.
Nucleic Acid Analysis
7.19.1 Supplement 25
The ability of cells to bind annexin V is still another marker considered to be specific to apoptosis. One should keep in mind, however, that use of the annexin V binding assay is hindered in some instances, e.g., when the plasma membrane is damaged during cell preparation or storage, leading to the loss of asymmetry in distribution of phosphatidylserine across the membrane. Furthermore, macrophages and other cells engulfing apoptotic bodies may also be positive in the annexin V assay (Marguet et al., 1999). Apoptosis can be recognized with greater certainty when the cells are subjected to several assays probing different apoptotic attributes (Hotz et al., 1994). For example, the assay of plasma membrane integrity (exclusion of PI) and annexin V binding combined with analysis of PARP cleavage or DNA fragmentation may provide a more definitive assessment of the mode of cell death than can be determined by each of these methods used alone. A plethora of kits designed to label DNA strand breaks and applicable to flow cytometry are available from different vendors. Most of these kits were designed by the authors (Gorczyca et al., 1992; Li and Darzynkiewicz, 1995). For example, Phoenix Flow Systems, BD PharMingen, and Alexis Biochemicals provide kits to identify apoptotic cells based on a single-step procedure utilizing either TdT and FITC-conjugated dUTP (APO-DIRECT; Li et al., 1995) or TdT and BrdUTP, as described in Basic Protocol 7 (APO-BRDU; Li and Darzynkiewicz, 1995). A description of the method, which is nearly identical to the one presented in this unit, is included with the kit. Another kit (ApopTag), based on a two-step DNA strand-break labeling with digoxygenin-16-dUTP by TdT, also designed by the authors (Gorczyca et al., 1992), was initially offered by ONCOR, later by Intergen, and most recently by Serologicals. BASIC PROTOCOL 1
MITOCHONDRIAL TRANSMEMBRANE POTENTIAL (∆ψm) MEASURED BY RHODAMINE 123 OR DiOC6(3) FLUORESCENCE The critical role of mitochondria during apoptosis is associated with the release of two intermembrane proteins, cytochrome c and apoptosis-inducing factor (AIF), that are essential for sequential activation of pro-caspase 9 and pro-caspase 3 (Liu et al., 1996; Yang et al., 1997). AIF is also involved in proteolytic activation of apoptosis-associated endonuclease (Susin et al., 1997). Still another protein, Smac/Diablo, that interacts with the inhibitors of caspases, thereby promoting apoptosis, is released from mitochondria (Deng et al., 2002). Dissipation (collapse) of mitochondrial transmembrane potential (∆ψm), also called the permeability transition (PT), likewise occurs early during apoptosis (Cossarizza et al., 1994; Kroemer, 1998; Zamzani et al., 1998). However, a growing body of evidence suggests that this event may be transient when associated with the release of cytochrome c or AIF, and mitochondrial potential may be restored for some time in the cells with activated caspases (Finucane et al., 1999; Scorrano et al., 1999; Li et al., 2000).
Flow Cytometry of Apoptosis
The membrane-permeable lipophilic cationic fluorochromes such as rhodamine 123 (R123) or 3,3′-dihexyloxacarbocyanine iodide [DiOC6(3)] can serve as probes of ∆ψm (Darzynkiewicz et al., 1981, 1982; Johnson et al., 1980). When live cells are incubated in their presence, the probes accumulate in mitochondria, and the extent of their uptake, as measured by intensity of cellular fluorescence, reflects ∆ψm. A combination of R123 and PI discriminates among live cells that stain only with R123, early apoptotic cells that have lost the ability to accumulate R123, and late apoptotic/necrotic cells whose plasma membrane integrity is compromised and that stain only with PI (Darzynkiewicz et al., 1982; Darzynkiewicz and Gong, 1994). The specificity of R123 and DiOC6(3) as ∆ψm probes is increased when they are used at low concentrations (0.5 AND THETA10,000 bacteria, acquire data for 5 min at 35 particles/sec. Low flow rates are preferred to maintain constant illumination over the small sample stream when the diameter of the focused laser beam is narrow for optimizing bacterial resolution (Robertson and Button, 1989) and to minimize artifacts caused by intercellular stain.
12. Replay listmode files to obtain a bivariate histogram of FS intensity versus DAPI fluorescence intensity (Fig. 11.9.1A). Set region boundaries for the bacterial population(s) of interest and for the cluster of 0.6-µm spheres to obtain data for each group. Gate on the cluster of 0.9-µm spheres to produce a histogram of SS for determining event frequency (Fig. 11.9.1B). Determine calibration factor 13. Prepare a radiolabeled cell sample at 106 cells/ml as described above (steps 1, 3, and 10). Detailed methodology for producing a calibration standard is beyond the scope of this protocol. The calibration standard should be similar in shape and composition to bacteria in the samples, and should be treated (preserved, diluted, and analyzed) in the same manner.
14. Measure representative bacteria by microscopy to determine the axial ratio (length/width) needed to formulate the standard curve. Calibrating for axial ratio is usually sufficient. However, if the composition of the dry matter varies a lot, or if a value for wet mass has to be determined, it is best to calibrate for each species (see discussion in Robertson et al., 1998).
15. Collect flow data as described for the sample (see steps 11 and 12). Collect radioactivity data using a liquid scintillation counter, so that the dry mass per cell can be calculated for the standard. The specific procedure for measuring radioactivity will vary depending on the sample being analyzed (see step 18). Generally, a 1-ml sample should be measured and should contain sufficient radioactivity (e.g., 5,000 to 100,000 counts/ml).
16. Run the BASIC program in Table 11.9.1 with input for axial ratio, FS collection angle, and excitation wavelength (footnote b).
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11.9.5 Current Protocols in Cytometry
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B 15
0.6
10
0.6
5
0.0 0.0
0.3
0
Dry mass (pg/cell)
Particle volume (V, µm3)
A
0.4 C. oligotrophus AR = 3 λ = 360 nm K = 1.618 x 10–5
0.2 0.0
0
10 20 30 FS intensity (I )
40
0.0
0.1 0.2 0.3 FS intensity (I )
Figure 11.9.2 Standard curve for dry cell mass from forward scatter intensity. (A) Theoretical dependency of FS intensity (I) on particle volume (V). Individual values (open circles) computed with the BASIC program in Table 11.9.1 with input for an axial ratio (AR) of 3, a collection angle of 0.5° to 20°, and an excitation wavelength of 360 nm. The fitted curve is described by Equation 11.9.1, where Xdry ≈ V, KIf = I, a = 1.62 × 10−4, b = 0.0144, c = 0.480, and d = 0.274. (B) Calibration of the curve in (A) with formaldehyde-fixed oligobacterium Cycloclasticus oligotrophus (diamond). Xsdry = 0.022 pg/cell and If = 58.7, giving K = 1.618 ×10−5 from Equation 11.9.1 (Robertson et al., 1998).
17. Import data for volume (V) and FS intensity (I) from BASIC into software with curve-fitting capability. Substitute Xdry (also from BASIC) for V, and plot and formulate the theoretical curve (Fig. 11.9.2A) according to Xdry = e
a ln( KIf )3 + b ln( KIf )2 + c ln( KIf ) + d
Equation 11.9.1
Where K is the proportionality constant or calibration factor used to convert FS intensity measured by flow cytometry (If) to intensity expected from theory (KIf = I), and a, b, c, and d are constants determined by the curve-fitting program (Robertson et al., 1998). Dry mass (Xdry) is substituted for V, because Xsdry varies with V for organisms of similar composition, because cell density can vary among species, and because light scatter intensity is a function of dry mass (Koch et al., 1996). Xsdry is used in this step as a general term for dry mass and is calculated by the algorithm.
18. Determine the dry mass per cell of calibration standard (Xsdry) from the measured radioactivity. Xsdry has been determined for the radioactivity of the marine isolate Cycloclasticus oligotrophus maximally labeled with 14C (dpm 14C/cell) after extended growth on radiolabeled acetate with high, known specific activity (dpm 14/g C) used as the sole carbon source, and from the contribution of carbon to dry weight obtained by CHN analysis (g dry wt/g C), as shown in Equation 11.9.2 (Robertson et al., 1998). s Xdry =
Bacterial Biomass from Forward Light Scatter Intensity
dpm dpm
14 14
C / cell × g dry wt /g C C/g C
Equation 11.9.2
11.9.6 Supplement 9
Current Protocols in Cytometry
19. Substitute the mean Xsdry of the radiolabeled cells (step 18) and the mean If per cell into Equation 11.9.1 to determine K. Analyze sample data and calculate biomass 20. Using sample data from steps 11 and 12, find the mean FS intensity of the bacterial population of interest (B) and of the internal standard 0.6-µm spheres (S). Normalize the data (B/S) for comparison among samples and to account for day-to-day variation. For the whole bacterial population in Figure 11.9.1A, B = 288 and S = 967.
21. Determine If for the sample for a specified value of S. Calculations for cell mass are simplified if K is computed for a specific value of intensity of the internal standard spheres. In work with small aquatic bacteria, the authors specify an intensity of 950 for the spheres. For the whole bacterial population in Figure 11.9.1A, If = (B/S) × 950 = (288/967) × 950 = 283.
22. Calculate Xsdry for bacteria in any particular sample using Equation 11.9.1, with K as determined in step 19 and If as calculated in step 21. For the whole bacterial population in Figure 11.9.1A, If × K = 283 × 1.618 × 10−5 = 4.58 × 10−3, giving a mean value of 50.0 fg/cell for Xsdry.
23. Determine the bacterial population density (N) from the ratio of the number of events recorded for the bacterial population(s) of interest in the bivariate histogram (Fig. 11.9.1A, left panel) to the number of events for the 0.9-µm standard spheres in the gated histogram of SS intensity (Fig. 11.9.1C). For the total bacterial population in Figure 11.9.1A/B, N = (8993/1560) × 1 × 105/ml = 5.76 × 105 bacteria/ml.
24. Calculate bacterial biomass from N × Xsdry. For the total bacterial population in Figure 11.9.1A, biomass = (5.76 × 105 bacteria/ml) × 50 fg/cell = 28.8 ìg/liter.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
DAPI/Triton X-100 staining solution Prepare an aqueous stock solution of 0.5 mg/ml 4′,6-diamidino-2-phenylindole (DAPI; Sigma), and store up to 6 months at −20 °C in the dark. Prepare a 5% (v/v) aqueous solution of Triton X-100 and store indefinitely at −20°C. For staining solution, mix 50 µl DAPI stock solution in 1 ml Triton X-100 stock solution and pass through a 0.2-µm filter. Prepare immediately before use. COMMENTARY Background Information Values for biomass critical to many microbiological studies have traditionally been obtained for bacteria in bulk suspension by measurements of optical density based on the relationship between dry mass and FS intensity according to Rayleigh-Gans theory (Koch, 1961; Koch and Ehrenfeld, 1968). Both the theoretical basis for the use of FS intensity measurements to determine the mass of cells
analyzed by flow cytometry (Koch et al., 1996) and an application to dilute mixed cultures have been been described (Robertson et al., 1998). A method to resolve subpopulations in a mixture of bacteria stained with DAPI (Button and Robertson, 1993) in combination with the method presented here for the determination of biomass has been used to characterize bacterial populations in Lake Zürich (Button et al., 1996) and to help characterize the marine oligobac-
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A Particle volume (V, µm3)
1.4 1.2
AR = 1
1.0 AR = 3
0.8 0.6
0.4 0.2 0.0
B Particle volume (V, µm3)
1.4 1.2
360 nm
1.0 0.8 0.6
488 nm
0.4 0.2 0.0
C Particle volume (V, µm3)
1.4 1.2 5o
1.0 0.8
10o
0.6 0.4
20o
0.2 0.0 0.0
0.2
0.4
0.6
0.8
FS intensity (I )
Figure 11.9.3 Parameters affecting the shape of the standard curve. The theoretical relationship between FS intensity and particle size as determined using the algorithm in Table 11.9.1 with various input parameters. (A) Cell shape, as indicated by the comparison of curves obtained for cells with axial ratios (AR) of 1 and 3 (cocci and rods). (B) Excitation wavelength, shown for UV (360 nm) and 488 nm. (C) FS detection angle, as computed with a lower limit of 0.5° and upper limits as shown. Intensities for the 5° and 10° upper limits are normalized to those for 20° at 0.001 µm3 for easier comparison of curve shape.
terium C. oligotrophus, isolated from Resurrection Bay, Alaska, by extinction culture (Button et al., 1998).
Critical Parameters and Troubleshooting
Bacterial Biomass from Forward Light Scatter Intensity
This method’s lower limit is set mainly by the sensitivity of the instrument, while the upper limit is dependent upon the size, shape, and refractive index of the bacteria. For the Ortho Cytofluorograf instrument configured for maximal sensitivity (Robertson and Button,
1989), the range for bacteria with an axial ratio of three and a refractive index relative to water (m) of 1.03 is 0.005 to 1.2 pg dry mass/cell (Robertson et al., 1998). The Rayleigh-Gans approximation is applicable only to microorganisms with m < 1.05 (Heller et al., 1959; Koch, 1961); it is not appropriate for standard latex particles (m = 1.19; Robertson et al., 1998). It is important that the calibration standard be as similar in cell shape and sample treatment as possible to the unknown samples, as signifi-
11.9.8 Supplement 9
Current Protocols in Cytometry
Table 11.9.2
Biomass in a Sample of M. arcticus from Figure 11.9.1
Regiona
Cell mass (fg [dry]/cell)
Population (105 cells/ml)
Biomass (µg/liter)
1 2 3
47.7 65.1 50.0
5.03 0.54 5.76
24.0 3.5 28.8
aAs indicated in Figure 11.9.1A, regions 1 and 2 are subpopulations containing cells with
one or two genome copies, respectively, and region 3 includes the entire population.
cant error can be introduced if a standard curve is formulated for rod-shaped bacteria with specified axial ratio but cocci are analyzed (Fig. 11.9.3A), if media differ in osmolarity (which would affect relative refractive index), or if there is nonuniform treatment with (or absorption of) aldehyde preservatives, which add to the cell mass (Robertson et al., 1998). The importance of using correct values for excitation wavelength and forward light scatter detection angle in the algorithm employed to produce a standard curve is shown in Figures 11.9.3B and 11.9.3C by the differences in the shapes of the theoretical curves obtained for two common excitation wavelengths and various collection angles. It appears that very narrow ranges of collection should be accurately defined in the algorithm to eliminate significant error, as shown by the large difference between the shape of the curves with an upper limit of 5° and 10°. A standard curve can be verified by independent measurements (Robertson et al., 1998). There should be agreement between dry mass determined by flow cytometry and dry mass computed from cell volume and buoyant density according to relationships previously reported (Robertson et al., 1998).
Anticipated Results Typical results are shown in Table 11.9.2, which gives the biomass of various subpopulations of M. arcticus as analyzed by flow cytometry (Fig. 11.9.1). With appropriate software (e.g., SigmaPlot), FS distributions can be easily converted to dry mass profiles (Fig. 11.9.1A, left panel; Fig. 11.9.1C) with the calculations demonstrated above. With this method, biomass as low as 5 pg in a 1-ml sample has been determined for a resolved bacterial subpopulation of 1000 cells/ml in a mixture of three species (Robertson et al., 1998).
Time Considerations Most of the effort is expended in obtaining a standard curve, which involves formulating
and fitting the applicable theoretical curve and preparing a suitable calibration standard. This may require several days. If formaldehyde-preserved cells are used, treatment should be done a day in advance of analysis by flow cytometry to ensure complete fixation (mass accumulation; Robertson et al., 1998). Staining with DAPI at 10°C takes an hour and can commence midway during the hour allowed for laser stabilization. Instrument alignment and equilibration of sample lines with DAPI take ∼20 min, and data acquisition takes 5 min for samples with ∼106 cells/ml. Sample staining at 7- to 10-min intervals can be convenient. Data analysis should be 620 nm. G1 cells (first peak) and G2 cells (second peak) are separated by a population of S-phase organisms. Sample data are shown in Figure 11.10.1. Microbiological Applications Contributed by David Lloyd Current Protocols in Cytometry (1999) 11.10.1-11.10.8 Copyright © 1999 by John Wiley & Sons, Inc.
11.10.1 Supplement 9
Number of cells
1000
500
0 0
10
20
30
40
50
60
70
80
Relative fluorescence
Figure 11.10.1 DNA histogram for Saccharomyces cerevisiae (n = 70,000) after propidium iodide staining, indicating 41,000 G1 cells (first peak) and 22,000 G2 cells (second peak). The third small peak consists of aggregated cells. The S-phase region lies between the two major peaks (Hutter and Eipel, 1978).
BASIC PROTOCOL 2
DETERMINING THE VIABILITY OF YEAST Loss of the plasma membrane electrochemical potential provides an excellent indication of cell death in a population of yeasts (Dinsdale et al., 1995, 1999; Seward et al., 1996; Willetts et al., 1997). Flow cytometry provides the most satisfactory method for enumeration of dead cells after brief exposure of the organisms to the anionic voltage-sensitive oxonol dye bis-(1,3-dibutylbarbituric acid) trimethine oxonol, often referred to as DiBAC4(3). Viable cells exclude the fluorophore so that only dead cells become fluorescent. Materials Yeast suspension Tris⋅Mg2+ buffer (see recipe) 1 µg/ml DiBAC4(3) solution (see recipe) 1. Dilute 1 ml yeast suspension in 20 mM Tris⋅Mg2+ buffer to give 105 organisms/ml. 2. Add 1 µg/ml DiBAC4(3) solution to give a final concentration of 0.1 µg/ml. 3. Incubate 5 min at room temperature. 4. Analyze using 488 nm excitation and measuring emission at 510 nm. Non-viable cells are fluorescent. Sample results are shown in Figure 11.10.2.
Flow Cytometry of Yeasts
11.10.2 Supplement 9
Current Protocols in Cytometry
A
10 20 FALS 30 40 50 60
10
fluorescence 20 30 40 50
B 60
10 20 FALS 30 40 50 60
C
10 20 FALS 30 40 50 60
10
fluorescence
20
30
40
50
fluorescence 20 30 40 50
60
D
10 20 FALS 30 40 50 60
10
fluorescence 20 30 40 50
60
E
10 20 FALS 30 40 50 60
10
10
fluorescence 20 30 40 50
60
F 60
10 20 FALS 30 40 50 60
10
fluorescence 20 30 40 50
60
Figure 11.10.2 Flow cytometry of Saccharomyces cerevisiae after DiBAC4(3)-based viability assessment of cells grown in the presence or absence of 10% ethanol. Organisms from control cultures (without added ethanol) are shown after (A) 24 hr or (B) 6 days. Results from heat-killed yeasts are shown in (C). Yeasts from cultures grown with 10% (v/v) added ethanol are shown after (D) 24 hr, (E) 3 days, and (F) 6 days. Although vitality decreases in the cultures containing ethanol, viability (i.e., ability to exclude the fluorophore) is hardly altered. FALS, forward-angle light scatter.
Microbiological Applications
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BASIC PROTOCOL 3
EVALUATING THE MITOCHONDRIAL (RESPIRATORY) FUNCTION OF YEAST The respiratory activity of yeast can be evaluated indirectly by flow cytometric measurement of rhodamine 123 uptake (Porro et al., 1994; Lloyd et al., 1996). In anaerobic fermentations (e.g., in brewing or cider-making), the capacity for uptake of this cationic dye is lost early in the process. Materials Washed yeast suspension at 1–5 × 106 organisms/ml 10 mM glucose 1 µg/ml rhodamine 123 solution (see recipe) 1. Harvest 5 ml washed yeast suspension by centrifuging 2 min at 1000 × g, room temperature. 2. Add 1 ml of 10 mM glucose and 5 ml of 1 µg/ml rhodamine 123 solution, and mix thoroughly. 3. Allow 5 min for dye uptake at room temperature. 4. Analyze using 488 or 546 nm excitation (Ar-ion or Hg-arc, respectively) and collecting fluorescence emission at 580 nm. Mitochondrial respiratory activity generates a transmembrane electrochemical potential; this drives dye accumulation in the organelles.
BASIC PROTOCOL 4
ASSAYING â-GALACTOSIDASE ACTIVITY IN VIVO A flow cytometric assay for β-galactosidase in S. cerevisiae uses the fluorogenic substrate resorufin β-D-galactopyranoside (Wittrup and Bailey, 1988). After the organisms have been permeabilized with Triton X-100, a steady state is established between product formation and leakage. Bovine serum albumin in the extracellular fluid is used to prevent reuptake by other cells. The reaction is measured at low temperature. Materials 2% (v/v) Triton X-100 Substrate solution: 2 mg/ml resorufin β-D-galactopyranoside (Molecular Probes) in dimethyl sulfoxide PBS (APPENDIX 2A), prechilled to 0°C 2% (w/v) bovine serum albumin (BSA; Sigma) in PBS (store and use at 0°C) Yeast suspension (106 cells/ml) washed twice in PBS after centrifugation from growth medium and kept at 4°C 1. Place the following (in order) into a 1.5-ml microcentrifuge tube kept on ice: 40 µl 2% Triton X-100 40 µl substrate solution 0.55 ml ice-cold PBS 0.4 ml ice-cold 2% BSA 50 µl washed yeast suspension. 2. Shake well and place in an ethanol/ice bath in the sample chamber of a flow cytometer. 3. Immediately initiate flow and measure distributions after 5 min using 568 nm excitation (Kr-ion laser) and collecting emission at >590 nm.
Flow Cytometry of Yeasts
β-Galactosidase-positive organisms show fluorescence, and fluorescence intensity is proportional to enzyme activity.
11.10.4 Supplement 9
Current Protocols in Cytometry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. Keep fluorophores as frozen solutions at −18°C in the dark. For solvents, use absolute ethanol or dimethyl sulfoxide. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
DiBAC4(3) solution, 1 ìg/ml Stock solution: Dissolve 1 mg DiBAC4(3) (Sigma) in 1 ml absolute ethanol and store up to 1 year at −18°C in the dark. Working solution: Dilute 1 µl to 1 ml in water (final 1 µg/ml) before use. Pepsin solution, 0.5% (w/v) 5.5 ml 1 N HCl 94.5 ml H2O 0.5 g pepsin Store in 1-ml aliquots for up to 1 year at −18°C Propidium iodide (PI) solution, 50 ìg/ml Stock solution: Dissolve 1 mg PI in 1 ml water and store in 50-µl aliquots for up to 1 month at −18°C in the dark. Working solution: Dilute 50 µl to 1 ml in water (final 50 µg/ml) before use. Rhodamine 123 solution, 1 ìg/ml Stock solution: Dissolve 1 mg rhodamine 123 (Sigma) in 1 ml water and store up to 1 month at −18°C in the dark. Working solution: Dilute 1 µl of to 1 ml in water (final 1 µg/ml) before use. RNase A solution, 10 ìg/ml Stock solution: Dissolve 5 mg bovine pancreas RNase A (DNase-free, Sigma) in 5 ml Tris⋅Mg2+ buffer (see recipe). Store in 0.5-ml aliquots for up to 1 week at −18°C. Working solution: Dilute 10 µl to 1 ml in Tris⋅Mg2+ buffer (final 10 µg/ml) before use. Tris⋅Mg2+ buffer 1.21 g/liter Tris base (mol. wt. 121; final 10 mM) 1 g/liter MgCl2 (5 mM) Adjust pH to 7.0 using 1 M HCl Store up to 1 month at 4°C COMMENTARY Background Information DNA-reacting fluorophores Basic Protocol 1 is suitable for use in studies of the effects of inhibitors on the DNA replication cycle of yeast (Eilam and Chernichovsky, 1988), cell cycle mutants of Schiz. pombe (Costello et al., 1986), and signaling pathway mutants (Hayashi et al., 1998). Propidium iodide (PI) is a nonspecific fluorochrome, reacting with all double-stranded nucleic acids to give complexes excitable at 488 nm and emitting above 600 nm. It has been used to measure total cell RNA, for example in Schiz. pombe (Agar and Bailey, 1982), where the RNA/DNA ratio is 100/1, and where most of the RNA is
double-stranded ribosomal RNA. After RNase treatment, PI is the most commonly employed fluorophore for cell-cycle analysis (nuclear DNA replication, see protocol above). Chromomycin A3 (Sigma) binds preferentially to GC base pairs and when excited at 457 nm emits fluorescence above 590 nm. Both Hoechst 33258 (Calbiochem) and 4′,6-diamidino-2-phenylindole (DAPI; Sigma) have a high affinity for AT base pairs. They react preferentially with mitochondrial DNA, and both fluorophores require UV excitation (329 to 425 nm; Sazer and Sherwood, 1990). DAPI has been used extensively to stain mitochondrial DNA for fluorescence microscopy (Williamson and Fennell, 1979), although in living cells
Microbiological Applications
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there may also be uptake of this dye into mitochondria by virtue of their transmembrane electrochemical potential (Pringle et al., 1989). Flow cytometry of arrested cell-division-cycle mutants of Schiz. pombe shows that care must be exercised in assigning DNA-reacting fluorophores to mitochondrial DNA (Carlson et al., 1997). Exposure of cells to Triton X-100 and hypotonic conditions after cell wall digestion was shown to remove cytoplasmic material and enable nuclear DNA content to be evaluated using either mithramycin plus ethidium bromide or Hoechst 33258. The former fluorophore mixture, used extensively for bacteria (Skarstad et al., 1985), gave the more reproducible results. Mithramycin (excited at around 440 nm using a line of the Hg arc) has a preference for GC base pairs in DNA, and effectively transfers excitation energy only to adjacent DNA-associated ethidium bromide. Similar results were also obtained with S. cerevisiae. The fluorescein diacetate/propidium iodide method also provides a technique for estimating viability of yeast (Willetts et al., 1997); other methods use ChemChrome Y (Chemunex; Brailsford and Gatley, 1993; Carter et al., 1993; Bruetschy et al., 1994; Deere et al., 1998) and the FungoLight kit (Molecular Probes; Lloyd and Hayes, 1995; Deere et al., 1998). In the last cited study, it was shown that propidium iodide exclusion and ChemChrome Y uptake gave similar results, but that for rehydrated dried yeast, DiBAC4(3) stained a higher proportion of organisms (107%) than did propidium iodide. These data compared well with those obtained by the traditional colony-counting method, but the great advantage of the flow cytometric assessment was that it took 20 min, rather than 36 hr. FungoLight, although useful for light microscopy, was not suitable for flow cytometric analyses. Cell sorting showed that viability was not affected by the dyes themselves or by the sorting procedure.
Flow Cytometry of Yeasts
Membrane potential–sensitive dyes Membrane potential–sensitive dyes, such as DiBAC4(3), can be used to assess the energy status of yeast mitochondria in vivo. Rhodamine 123, tetramethyl rhodamine ethyl ester (TMRE), 3,3′-dipropylthiacarbocyanine iodide [DiSC3(3)], and 3,3′-dipropyloxacarbocyanine iodide [DiOC3(3)] have also been used (Denksteinova et al., 1996; Plásek and Sigler, 1996). The production of fluorescent formazan from 5-cyano-2,3-ditolyl tetrazolium chloride,
a method used for bacteria (López-Amorós et al., 1995), requires prior cell permeabilization as yeasts are not permeable to this redox indicator. Other assays Intracellular flavins and NAD(P)H redox state. A dual excitation of flavins and NAD(P)H is informative about the redox state of organisms and cells. For rat liver cells, Theorell (1983) described the simultaneous flow cytometric measurement of blue and green fluorescence using excitations at 351 to 363 nm (argon UV) and at 488 nm (argon), respectively. This method can also be applied to yeast. Cellular protein. The analysis of protein distributions provides an estimate of several cell cycle parameters for steady-state and perturbed cultures (Alberghina and Porro, 1993). An excellent summary of the use of flow cytometry in investigations of the growth of yeast is given by Davey and Kell (1996). Sterol content of yeast. Fluorescein isothiocyanate–labeled nystatin A1, a macrolide reacting specifically with 3β-hydroxy sterols (80% ergosterol), has been used to measure the sterol distribution within the yeast population (Müller et al., 1992). Hydrogen peroxide production. The probe 2′,7′-dichlorofluorescin diacetate is concentrated within cells as it is hydrolyzed by esterases. Reaction with intracellular peroxides gives a product that emits intense green fluorescence when excited at 488 nm (Yurkow and McKenzie, 1993). Simultaneous viability analysis, using the propidium iodide method, indicated that aeration after a period of anoxia gives peroxide generation leading to cell damage and death. Antibiotic sensitivity testing. A flow cytometric method for detection of the sensitivity of Candida species to amphotericin has been developed (Ordónez and Wehman, 1995). The cationic membrane potential–sensitive dye 3,3′- dip en tylo xacarbocyanine iodide [DiOC5(3)] is accumulated by cells growing normally. Incubation in the presence of antibiotic for 30 min reduced dye uptake of sensitive strains in a dose-dependent manner, whereas that of resistant strains was unaffected. Propidium iodide, ChemChrome Y, and DiBAC4(3) also show promise in the flow cytometric assessment of drug sensitivity (Carter et al., 1993). Selection of novel yeast hybrids. Flow cytometry has applications in the development of improved industrial yeast strains (e.g., the rapid
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isolation of rare mating hybrids; Bell et al., 1998). Yeast as a spoilage organism. ChemChrome Y uptake has been used as a rapid and sensitive marker to detect yeasts as contaminants of soft drinks (Pettipher, 1991; Brailsford and Gatley, 1993).
Critical Parameters For live-cell assays, cells should be treated with respect (i.e., minimally perturbing conditions employed for centrifugation). All procedures should be carried out as quickly as possible and without change of temperature.
Anticipated Results The most valuable characteristic of flow cytometric data is the ability to provide information on the distribution of activities or constituent amounts in individuals of the cellular population, thereby highlighting subpopulations (e.g., of nonviable cells or of cell cycle stages where enzyme activities are very high or very low).
Time Considerations Analysis of the DNA replication cycle (Basic Protocol 1) uses fixed organisms and can be accomplished after performing other measurements, provided that storage of fixed cells (at 4°C) is for no more than 24 hr. The usefulness of the other methods using live organisms (Basic Protocols 2 to 4) lies in the speed of application: analyses require only a few minutes.
Literature Cited Agar, D.W. and Bailey, J.E. 1982. Cell cycle operation during batch growth of fission yeast populations. Cytometry 3:123-128. Alberghina, L. and Porro, D. 1993. Quantitative flow cytometry: Analysis of protein distributions in budding yeast. A mini review. Yeast 9:815823. Bell, P.J.L., Deere, D., Shen, J., Chapman, B., Bissinger, P.H., Attfield, P.V., and Veal, D.A. 1998. A flow cytometric method for rapid selection of novel yeast hybrids. Appl. Environ. Microbiol. 64:1669-1672. Brailsford, M. and Gatley, S. 1993. Rapid analysis of microorganisms using flow cytometry. In Flow Cytometry in Microbiology (D. Lloyd, ed.) pp. 171-180. Springer-Verlag, London. Bruetschy, A., Laurent, M., and Jacquet, R. 1994. Use of flow cytometry in oenology to analyse yeasts. Lett. Appl. Microbiol. 18:343-345. Carlson, C.R., Grallert, B., Bernander, R., Stokke, T., and Boye, E. 1997. Measurement of DNA content in fission yeast. Yeast 13:1329-1335. Carter, E.A., Paul, F.E., and Hunter, P.A. 1993. Cytometric evaluation of antifungal agents. In
Flow Cytometry in Microbiology (D. Lloyd, ed.) pp. 111-120. Springer-Verlag, London. Costello, G., Rodgers, L., and Beach, D. 1986. Fission yeast enters the stationary phase G0 state from either mitotic G1 or G2. Curr. Genet. 11:119-125. Davey, H.M. and Kell, D.B. 1996. Flow cytometry and cell sorting of heterogenous microbial populations: The importance of single-cell analyses. Microbiol. Rev. 60:641-696. Deere, D., Shen, J., Vesey, G., Bell, P., Bissinger, P., and Veal, D. 1998. Flow cytometry and cell sorting for yeast viability assessment and cell selection. Yeast 14:147-160. Denksteinova, B., Sigler, K., and Plásek, J. 1996. Three fluorescent probes for the flow-cytometric assessment of membrane potential in Saccharomyces cerevisiae. Folia Microbiol. 41:237242. Dinsdale, M.G., Lloyd, D., and Jarvis, B. 1995. Yeast vitality during cider fermentation: Two approaches to the measurement of membrane potential. J. Inst. Brew. 101:453-458. Dinsdale, M.G., Lloyd, D., McIntyre, P., and Jarvis, B. 1999. Yeast vitality during cider fermentation: Assessment by energy metabolism. Yeast 15:285-293. Eilam, Y. and Chernichovsky, D. 1988. Low concentrations of trifluoperazine arrest the cell division cycle of Saccharomyces cerevisiae at two specific stages. J. Gen. Microbiol. 134:1063-1069. Hayashi, M., Ohkuni, K., and Yamashita, I. 1998. Control of division arrest and entry into mitosis by extracellular alkalisation in Saccharomyces cerevisiae. Yeast 14:905-913. Hutter, K.-J. and Eipel, H.E. 1978. Flow cytometric determinations of cellular substances in algae, bacteria, molds and yeasts. Antonie van Leeuwenhoek 44:269-278. Lloyd, D. and Hayes, A.J. 1995. Vigour, vitality and viability of microorganisms. FEMS Microbiol. Lett. 133:1-7. Lloyd, D., Moran, C.A., Suller, M.T.E., Hayes, A.J., and Dinsdale, M.G. 1996. Flow cytometric monitoring of rhodamine 123 and a cyanine dye uptake by yeast during cider fermentation. J. Inst. Brew. 102:251-259. López-Amorós, R., Mason, D.J., and Lloyd, D. 1995. Use of two oxonols and a fluorescent tetrazolium dye to monitor starvation of Escherichia coli in sea water by flow cytometry. J. Microbiol. Methods 22:165-176. Müller, S., Lösche, A., and Bley, T. 1992. Flow-cytometric investigation of sterol content and proliferation activity of yeast. Acta Biotechnol. 12:365-375. Ordónez, J.V. and Wehman, N.M. 1995. Amphotericin suseptibility of Candida species assessed by rapid flow cytometric membrane potential assay. Cytometry 22:154-157. Pettipher, G.L. 1991. Preliminary evaluation of flow cytometry for the detection of yeasts in soft drinks. Lett. Appl. Microbiol. 12:109-112. Plásek, J. and Sigler, K. 1996. Slow fluorescent indicators of membrane potential: A survey of different approaches to probe response analysis. J. Photochem. Photobiol. B. 33:101-124.
Microbiological Applications
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Porro, D., Sneraldi, C., Martegani, E., Ranzi, B.M., and Alberghina, L. 1994. Flow cytometry for monitoring yeast growth. Biotechnol. Prog. 10:193-199. Pringle, J.R., Preston, R.A., Adams, A.E.M., Stearns, T., Drubin, D.G., Haarer, B.K., and Jones, E.W. 1989. Fluorescence microscopy methods for yeast. Methods Cell Biol. 31:357435. Rose, A.H. and Harrison, J.S. (eds.). 1969. The Yeasts, Vol.1: Biology of the Yeasts, Academic Press, New York. Sazer, S. and Sherwood, S.W. 1990. Mitochondrial growth and DNA synthesis occur in the absence of nuclear DNA replication in fission yeast. J. Cell Sci. 97:509-516. Seward, R., Willetts, J.C., Dinsdale, M.G., and Lloyd, D. 1996. The effects of ethanol, hexan-lol and 2 phenylethanol on cider yeast growth, viability and energy status; synergistic inhibition. J. Inst. Brew. 102:439-443. Skarstad, K., Steen, N.B., and Boye, E. 1985. Escherichia coli DNA distributions measured by flow cytometry and compared with computer simulations. J. Bacteriol. 154:656-662.
Theorell, B. 1983. Flow-cytometric monitoring of intracellular flavins simultaneously with NAD(P)H levels. Cytometry 4:61-65. Willetts, J.C., Seward, R., Dinsdale, M.G., Suller, M.T.E., Hill, B., and Lloyd, D. 1997. Vitality of cider yeast grown micro-aerobically with added ethanol, butan-l-ol or iso-butanol. J. Inst. Brew. 103:79-84. Williamson, D.H. and Fennell, D.J. 1979. Mitochondrial DNA. In Methods in Cell Biology, Vol. 12 (D.M. Prescott, ed.) pp. 335-351. Academic Press, New York. Wittrup, K.D. and Bailey, J.D. 1988. A single-cell assay of β-galactosidase activity in Saccharomyces cerevisiae. Cytometry 9:394-404. Yurkow, E.J. and McKenzie, M.A. 1993. Characterization of hypoxia-dependent peroxide production in cultures of Saccharomyces cerevisiae using flow cytometry: A model for ischaemic tissue destruction. Cytometry 14:287-293.
Contributed by David Lloyd University of Wales Cardiff, United Kingdom
Flow Cytometry of Yeasts
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Enumeration of Phytoplankton, Bacteria, and Viruses in Marine Samples
UNIT 11.11
The first applications of flow cytometry to the analysis of phytoplankton, in the mid1980s, revolutionized the study of the smallest organisms in this community—those small enough to pass through 2-µm-pore-size filters, called picophytoplankton. These tiny organisms are generally unicellular and are naturally quite concentrated in seawater, so that they can be analyzed without prior concentration or sonication. Picophytoplankton are present in all aquatic environments, although their relative contribution to the photosynthetic biomass is greatest in the central regions of oceans (90% of the total surface), which are nutrient depleted and relatively poor in chlorophyll (0.2 mg/m3). Data obtained by flow cytometry have helped confirm that picophytoplankton constitute the bulk of the photosynthetic biomass in subtropical waters. This unit presents a method for enumerating phytoplanktonic cells on the base of their natural parameters (see Basic Protocol 1). This protocol can be performed either on board ship or in the laboratory, and does not require any pretreatment of samples. If samples cannot be tested when freshly obtained, they can be preserved with formaldehyde or glutaraldehyde (see Support Protocol 1) and assayed later. Highly sensitive nucleic acid–specific stains such as TOTO-1, YOYO-1, and the SYBR Green family (all available from Molecular Probes) have also made it possible to detect and enumerate heterotrophic bacteria and, very recently, viruses in marine samples. Two further protocols detail the enumeration of bacteria (see Basic Protocol 2) and viruses (see Basic Protocol 3) in culture and in natural seawater samples. Both require fixation (see Support Protocol 3) and the use of nucleic acid–specific stains. Also included is a procedure for calibrating cytometer flow rates (see Support Protocol 2), replacing the standard approach using fluorescent microsphere standards, which is less suitable when working with seawater samples. FLOW CYTOMETRIC ENUMERATION OF PICOPHYTOPLANKTON BASED ON SCATTER AND AUTOFLUORESCENCE
BASIC PROTOCOL 1
The different populations present in a natural sample are discriminated on the basis of their scatter signals and the fluorescence of their natural phytoplanktonic pigments (see Fig. 11.11.1), which can vary throughout the water column (see Fig. 11.11.2). Flow cytometry is particularly suited to the analysis of picophytoplankton, which are difficult to study with traditional methods. Generally three major groups of these organisms, two cyanobacteria and a range of picoeukaryotes (algae), can be distinguished; see Anticipated Results for details. Marine samples may be obtained, for example, from the Provasoli-Guillard National Center for Culture of Marine Phytoplankton (CCMP), McKown Point, West Boothbay Harbor, Maine 04975, USA; http://CCMP.bigelow.org. Samples can be used fresh within 12 hr of being obtained (they should be stored at 4°C, but need not be fixed) or can be fixed and frozen (see Support Protocol 1), then thawed before being analyzed. Materials Natural marine samples or cultures, either fresh or frozen (see Support Protocol 1 for freezing procedure) 0.95-µm fluorescent microspheres (Polysciences) diluted to ∼105 beads/ml (as assessed by epifluorescence microscopy) in distilled water Seawater Contributed by Dominique Marie, Frédéric Partensky, Daniel Vaulot, and Corina Brussaard Current Protocols in Cytometry (1999) 11.11.1-11.11.15 Copyright © 1999 by John Wiley & Sons, Inc.
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0.2-µm-pore-size cartridge filter units Flow cytometer equipped with a 488-nm argon laser (e.g., FACSort, Becton Dickinson) Additional reagents and solutions for flow cytometer calibration (see Support Protocol 2) 1. If sample has been frozen, thaw at 37°C. Transfer 1 ml of sample to a suitable flow cytometer tube. If the cell suspensions are too concentrated (as may be the case with culture samples, for example), they can be diluted in seawater previously filtered through a 0.2-ìm-pore-size filter.
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Figure 11.11.1 Cytograms of scatter and fluorescence obtained for a marine sample collected in the Pacific Ocean at a depth of 65 m (OLIPAC cruise, Cast 94, 5° S to 150° W). Prochlorococcus (Proc), Synechococcus (Syn), and picoeukaryotes (Euk) are discriminated on the basis of the fluorescence of their natural pigments, chlorophyll (red) or phycoerythrin (orange). 0.95-µm beads were added as internal reference.
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Figure 11.11.2 Vertical profiles obtained for samples collected in the Pacific Ocean and analyzed fresh on board the N.O. l’Atalante during the OLIPAC cruise (Cast 94, 5° S to 150° W). Phytoplanktonic cell abundance (A), chlorophyll fluorescence (B), and side scatter (C) per cell normalized to 0.95-µm beads versus depth. Proc, Prochlorococcus; Syn, Synechococcus; Euk, picoeukaryotes.
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2. Add 10 µl of an ∼105 beads/ml suspension of 0.95-µm fluorescent microspheres (as an internal reference). 3. Filter 5 to 10 liters of seawater through 0.2-µm-pore-size cartridge filter for use as sheath fluid. Because cell scatter (especially forward scatter) is dependent on the nature of the sheath fluid, the use of filtered seawater as sheath fluid is recommended. If the fluidics system of the flow cytometer is equipped with an in-line filter, this should be removed, because it is likely to become contaminated quickly and will thereafter release particles.
4. Calibrate the flow rate of the cytometer (see Support Protocol 2). 5. Set the discriminator to red fluorescence and set all parameters on logarithmic amplification. For a surface sample from a moderately oligotrophic area, typical settings on a FACSort flow cytometer are forward scatter (FS) = E01, side scatter (SS) = 450, green fluorescence (FL1) = 650, orange fluorescence (FL2) = 650, and red fluorescence (FL3) = 650.
6. Insert the sample, allow ∼15 sec for the flow rate to stabilize, and then begin data acquisition. Data for natural samples are typically collected in listmode files for 2 to 4 min with a flow rate of 50 to 100 ìl/min.
7. Record the time of analysis to determine precisely the cell concentrations of each population. 8. Compute the absolute cell concentration for each population in a given sample as follows: Cpop = T × Npop/R × (Vtotal/Vsample) where Cpop = concentration of population in cells/µl, Npop = number of cells acquired, T = acquisition time (min), R = sample flow rate (µl/min) as determined for the sample series, Vtotal = volume (µl) of sample plus additions (fixatives, beads, etc.), and Vsample = volume of sample (µl). 9. Report parameters relative to the beads added to the samples: Xrel = Xpop/Xbeads where Xpop is the average value of a cell parameter (scatter or fluorescence) for a given population and Xbeads the same parameter for the beads. Before calculation of the ratio, Xpop and Xbeads must be expressed as linear values (not numbers of channels) after conversion from the logarithmic recording scale. BASIC PROTOCOL 2
Enumeration of Phytoplankton, Bacteria, and Viruses in Marine Samples
FLOW CYTOMETRIC ENUMERATION OF BACTERIOPLANKTON BY DNA STAINING AND FLUORESCENT DETECTION Because of its accuracy, its speed, and the lack of interference from dissolved organic matter, flow cytometry has been increasingly used to analyze heterotrophic bacteria (Shapiro, 1988; Robertson et al., 1998). In contrast to the photosynthetic prokaryotes Prochlorococcus and Synechococcus, bacteria do not contain any pigments and cannot be counted based on autofluorescence. Staining of cell DNA has been used as a means to discriminate and enumerate bacteria in natural seawater samples by epifluorescence microscopy (EFM; Hobbie et al., 1977) or flow cytometry (Button and Robertson, 1989; Monger and Landry, 1993; Li et al., 1995; Marie et al., 1997). The combination of DNA and chlorophyll fluorescence allows discrimination of autotrophic from heterotrophic
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picoplankton (Monger and Landry, 1993; Campbell et al., 1994). For details of the bacterial populations generally observed using flow cytometry, see Anticipated Results. Older UV-excited dyes, such as DAPI or Hoechst 33342, that require expensive flow cytometric equipment, are currently being superseded by a wide and continually expanding range of nucleic acid–specific dyes synthesized and manufactured by Molecular Probes. These novel dyes are excited at 488 nm, which means they are usable on small, low-cost flow cytometers equipped with air-cooled single-line argon lasers. The affinity of the cyanine dyes TOTO-1 and YOYO-1, and their monomeric equivalents YO-PRO-1 and TO-PRO-1 (all available from Molecular Probes), decreases significantly with increasing ionic strength, so they are inappropriate for direct analysis of seawater samples (Marie et al., 1996). Other dyes such as SYBR Greens I and II, SYTOX Green, and the SYTO family (all available from Molecular Probes) are less dependent on medium composition and can be used for enumerating bacteria in marine environments (Marie et al., 1997; Lebaron et al., 1998). Because SYBR Green I (SYBR-I) has a very high fluorescence yield, the authors strongly recommend the use of this dye to enumerate bacteria in marine samples. Samples must be fixed before bacterial enumeration can be performed, since fixation allows the nucleic acid–specific stain to penetrate into the cell. A 10,000-fold dilution of the commercial SYBR-I stock solution is used. NOTE: All stock solutions except the dye must be prefiltered through a 0.2-µm- (or smaller) pore-size filter to avoid contamination. Materials Natural marine samples or cultures, either fresh or frozen (see Support Protocol 1 for freezing procedure) 10% paraformaldehyde (see Support Protocol 3) and/or 25% glutaraldehyde (Sigma) DNA-specific stain such as SYBR Green I, YOYO-1, TOTO-1, or TO-PRO-1 (Molecular Probes) 0.95-µm fluorescent microspheres (Polysciences) diluted to ∼105 beads/ml (as assessed by epifluorescence microscopy) in distilled water Seawater 0.2-µm-pore-size cartridge filter units Flow cytometer equipped with a 488-nm argon laser (e.g., FACSort, Becton Dickinson) Additional reagents and solutions for flow cytometer calibration (see Support Protocol 2) 1a. If samples are live: Add 1% paraformaldehyde or 0.1% glutaraldehyde (final concentrations) and let stand 20 min. Paraformaldehyde and glutaraldehyde give equivalent results.
1b. If samples have been preserved and frozen: Thaw samples at 37°C. If the cell suspensions are too concentrated (as may be the case with culture samples, for example), they can be diluted in seawater previously filtered through a 0.2-ìm-pore-sizefilter.
2. Add SYBR-I at a final concentration of 1 part in 10,000 and incubate 15 min at room temperature in the dark. Microbiological Applications
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Figure 11.11.3 Cytograms of side scatter and fluorescence obtained for a natural sample collected in the Pacific Ocean at a depth of 65 m (OLIPAC cruise, Cast 94, 5° S to 150° W) and stained with SYBR-I. Three different bacterial populations (B-I, B-II, and B-III) can be discriminated from Prochlorococcus by the combination of the different parameters recorded. 0.95-ìm fluorescent beads were added as internal reference.
3. To 1 ml of sample, add 10 µl of an ∼105 bead/ml suspension of 0.95-µm fluorescent microspheres (as an internal reference). 4. Filter 5 to 10 liters of seawater through 0.2-µm-pore-size cartridge filter for use as sheath fluid. Distilled water can be used as sheath fluid, but for natural seawater samples, 0.2-ìm-poresize-filtered seawater is preferable, since cell scatter (especially forward scatter) is dependent on the nature of the sheath fluid. If the fluidics system of the flow cytometer is equipped with an in-line filter, this should be removed, because it is likely to become contaminated quickly and will thereafter release particles.
5. Set the discriminator to green fluorescence. 6. Calibrate the flow rate of the cytometer (see Support Protocol 2). 7. Set all parameters on logarithmic amplification. It is recommended that no more than 80,000 events be acquired in listmode, in order to avoid very large files. Typical settings on a FACSort flow cytometer are FS = E01, SS = 450, FL1 = 650, FL2 = 650, and FL3 = 650.
8. Run the sample, adjusting the flow rate and cell concentration to avoid coincidence. Enumeration of Phytoplankton, Bacteria, and Viruses in Marine Samples
Typically, the authors run samples for 1 to 2 min at a flow rate of 30 to 50 ìl/min and keep the number of events below 1000 per sec (by diluting samples that are too concentrated). Some samples, particularly those obtained in coastal areas, contain copious quantities of small particles and debris that will increase the level of background noise. This can induce
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coincidence or lead to the generation of large listmode files. In such cases, the threshold can be increased to reduce the number of events seen by the flow instrument, and/or a “bitmap” window (non-regular region) can be defined that includes the population of bacteria so that only the events belonging to this area will be recorded (see Fig. 11.11.3).
FLOW CYTOMETRIC ENUMERATION OF VIROPLANKTON BY DNA STAINING AND FLUORESCENT DETECTION
BASIC PROTOCOL 3
The existence of bacteriophages in marine environments has been known for many years (Kriss and Rukina, 1947; Spencer, 1955, 1960), but they were not really investigated until fairly recently (Bergh et al., 1989; Bratbak et al., 1990; Proctor et al., 1990). Viroplankton clearly constitute the most abundant population of biological particles in the ocean and their ecological role has only recently been investigated. These studies initially required techniques such as transmission electron microscopy (TEM) that are time consuming and allow only limited numbers of samples to be analyzed. During the past decade, investigations using epifluorescence microscopy (EFM) in conjunction with nucleic acid–specific dyes such as DAPI (Hara et al., 1991) or with cyanine dyes (Hennes and Suttle, 1995; Weinbauer and Suttle, 1997) have considerably improved knowledge of marine viruses. Very recently, flow cytometry has been successfully applied to the analysis of viruses in solution, using the nucleic acid–specific dye SYBR Green I (Marie et al., 1999). This has permitted the analysis of viruses with reduced DNA content, down to 40 Kbp (Brussaard et al., unpub. observ.). Other dyes, such as SYTOX, PicoGreen, OliGreen, SYBR Green II, SYBR Gold, or RiboGreen (all from Molecular Probes), can be used with the same efficiency as SYBR-I (Brussaard et al., unpub. observ.). For details of the viroplankton populations generally observed using flow cytometry, see Anticipated Results. NOTE: All stock solutions except the dye must be prefiltered through a 0.2-µm- (or smaller) pore-size filter to avoid contamination. Materials Natural marine samples or cultures, either fresh or frozen (see Support Protocol 1 for freezing procedure) 10% paraformaldehyde (see Support Protocol 3) or 25% glutaraldehyde (Sigma) TE buffer, pH 7.2 (APPENDIX 2A) DNA-specific stain(s) such as SYBR Green I or II, OliGreen, or RiboGreen (Molecular Probes) 0.95-µm fluorescent microspheres (Polysciences) diluted to ∼105 beads/ml (as assessed by epifluorescence microscopy) in distilled water Distilled water 0.2-µm-pore-size filtration units for plastic syringe Flow cytometer equipped with a 488-nm argon laser (e.g., FACSort, Becton Dickinson) Additional reagents and solutions for flow cytometer calibration (see Support Protocol 2) Prepare sample 1a. For fresh samples: Add 1% paraformaldehyde or 0.1% to 0.5% glutaraldehyde (final concentrations) and let stand 20 min. No significant differences have been found between results for virus enumeration performed on samples fixed with paraformaldehyde, glutaraldehyde, or a mixture of both.
1b. For fixed and frozen samples: Thaw samples at 37°C. Microbiological Applications
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2. Dilute samples in TE buffer, pH 7.2, to three different concentrations: typically 10-, 50-, and 100-fold for natural seawater samples and 100-, 1,000-, and 10,000-fold for cultured samples. Preparation of three different dilutions is necessary because the concentration of viruses is not known beforehand. Analysis must be performed with a suspension of ∼2 × 105 to 2 × 106 viruses/ml (final concentration). To avoid generating large files, samples can be run for 1 or 2 min at a rate ranging from 20 to 50 ìl/s. Different buffers have been tested for diluting virus samples. Tris-based buffers give the best result.
3. Add SYBR-I at a final concentration of 5 parts in 100,000 and incubate 15 min at room temperature in the dark. 4. To 1 ml of sample, add 10 µl of an ∼105 beads/ml suspension of 0.95-µm fluorescent microspheres (as an internal reference). For virus samples that are freshly fixed (i.e., have not been frozen), or for hard-to-stain material, it is necessary to heat the samples 10 min at 80°C in the presence of detergent (e.g., Triton X-100 at 0.1% final) to improve dye uptake.
Acquire data 5. Using distilled water as sheath fluid (even for marine samples), begin the cytometric procedure by calibrating the flow rate (see Support Protocol 2). Since samples are diluted in TE, use of seawater is not necessary.
6. Turn the discriminator to green fluorescence (FL1). Typical settings on a FACSort flow cytometer are FS = E03, SS = 600, FL1 = 600, FL2 = 650, and FL3 = 650.
7. Before starting data acquisition, wait for the sample flow rate to stabilize (this can take up to 20 sec). 8. Run the sample at a rate allowing