bluuebunny/crossref_metadata_2025 huggingface dataset:
Anesthesia
Authors: Donovan John , Brown Patricia
Published: June 2004

Anesthetic agents are used in laboratory animals to prevent pain or distress due to an experimental procedure or for restraint to facilitate a technically difficult procedure. This unit provides three basic protocols: injectable anesthesia for mouse, rat, and hamster; inhalant anesthesia using methoxyflurane for mouse, rat, and hamster; and injectable anesthesia using ketamine/xylazine for rabbit. An describes sedation using butorphanol/acetylpromazine in the rabbit. The further describes and compares these methods of anesthesia for various applications.
Production of Polyclonal Antisera
Authors: Cooper Helen M. , Paterson Yvonne
Published: June 2004

Although the advances offered by the development of monoclonal antibody techniques have revolutionized the specificity, uniformity, and quantity of antibodies, there remain many circumstances in which polyclonal antibodies are more desirable than monoclonal antibodies. Production of polyclonal antisera takes less time and effort than production of monoclonal antibodies, requires relatively simple and readily available equipment, and produces reagents that can be used for immunoprecipitation, immunoblotting, and enzyme‐linked immunosorbent assays (ELISAs). Theandin this unit describe the production of polyclonal antisera specific for protein antigens in rabbits, rats, mice, and hamsters. Thepresents a method for preparing serum from blood.
Fragmentation of Immunoglobulin G
Authors: Andrew Sarah M. , Titus Julie A.
Published: June 2004

This unit describes procedures for fragmentation of IgG to the monovalent Fab fragment using papain digestion, and to the bivalent F(ab′)2 fragment by pepsin digestion. Alternative methods of fragmentation to F(ab′)2 include use of papain that is preactivated with cysteine and use of the enzyme ficin. These alternate methods are particularly useful for mouse IgG1 antibodies.
Purification of Immunoglobulin M and Immunoglobulin D
Authors: Andrew Sarah M. , Titus Julie A. , Coico Richard , Amin Ashok
Published: June 2004

This unit describes two classical protocols for the purification of IgM ‐ dialysis of ascites fluid, tissue culture medium, or bioreactor supernatants against distilled water to precipitate pure IgM, and ammonium sulfate precipitation. Both protocols can be followed by size‐exclusion chromatography to obtain a highly purified product. Recently an affinity method for purification of IgM has been developed using mannan binding protein, and is described here. The third approach presented is a one‐step IgD purification method, designed specifically for murine derived samples, that uses Sepharose coupled to lectin derived from the seeds of Griffonia simplicifolia‐1. This represents a simple, rapid, and gentle approach to isolating this highly labile immunoglobulin from IgD‐containing ascites or hybridoma sources.
Production of Bispecific Antibodies
Authors: Segal David M. , Bast Bert J.E.G.
Published: June 2004

Bispecific antibodies (bsAbs) contain two different binding specificities within a single molecule and can specifically bind two different molecules together. BsAbs can be produced by chemically cross‐linking purified antibodies or Fab fragments with reducible disulfide bonds or nonreducible thioether bonds, both of which are described in this unit. Protocols are also presented for producing BsAbs by fusing two antibody‐producing hybridomas that can be selected for based on drug resistance, or by double labeling with fluorochromes and FACS. Support protocols describe screening and purification of bsAbs.
Nucleic Acid Immunizations
Authors: Robinson Harriet L. , Pertmer Tamera M.
Published: June 2004

This unit provides protocols for two broadly used methods of DNA immunization: saline injections and gene gun deliveries of DNA. Saline injections deliver DNA into extracellular spaces; gene gun deliveries bombard DNA directly into cells. Support protocols present methods for preparation of DNA‐coated gold beads, creation of cartridges containing these beads, and optimization of gene gun parameters. Issues relating to plasmid vectors, DNA preparation and doses, and immunization regimens are also discussed. Expression library immunizations, genetic and conventional adjuvants, alternative boosts, neonatal immunizations, and approaches to mucosal delivery of DNA are covered in the commentary.
Isolation of Mouse Mononuclear Cells
Authors: Kruisbeek Ada M.
Published: June 2004

This unit covers techniques for preparing lymphoid cell suspensions and removing red blood cells and dead cells. These basic manipulations are required for most of the subsequent protocols in this chapter, i.e., measuring functional responses of mouse lymphocytes.
T Cell Enrichment By Nonadherence to Nylon
Authors: Hathcock Karen S.
Published: June 2004

Differential adherence properties of T cells, B cells, and accessory cells such as macrophages or other APC (antigen‐presenting cells) can be employed to enrich for T cells. While the basis for this adherence is not understood, either nylon wool or Sephadex G‐10 can be used as column matrices to which B cells and accessory cells will more readily adhere than T cells. Such procedures offer a convenient, although imprecise, means of enriching T cells through removal of accessory and B cells; use of nylon wool is preferred if both of the latter subsets are to be removed, while Sephadex is used when the goal is primarily to remove accessory cells.
Fractionation of T and B Cells Using Magnetic Beads
Authors: Thornton Angela M.
Published: June 2004

This unit describes the purification of mouse T cells, B cells, and T cell subsets using magnetic bead separation. Isolation of cell subsets using magnetic beads is quick, simple, and reliable and can result in high yields of very pure cells.
Isolation of Dendritic Cells
Authors: Inaba Kayo , Swiggard William J. , Steinman Ralph M. , Romani Nikolaus , Schuler Gerold
Published: June 2004

This unit presents two methods for preparing dendritic cells (DCs), a highly specialized type of antigen‐presenting cell (APC). The first method involves the isolation of DCs from mouse spleen, resulting in a cell population that is highly enriched in accessory cell and APC function. A support protocol for collagenase digestion of splenocyte suspensions is described to increase the yield of dendritic cells. The second method involves generating large numbers of DCs from mouse bone marrow progenitor cells. In that technique, bone marrow cells are cultured in the presence of granulocyte/macrophage colony‐stimulating factor (GM‐CSF) to yield 5‐10 106 cells, 60% of which express DC surface markers (e.g., B‐7‐2/CD86). Additional techniques for isolating DCs from mouse spleens or other mouse tissues, as well as from human tissues, are also discussed.
Early Events in B Lymphocyte Activation
Authors: Bondada Subbarao , Troyer Amy , Chelvarajan Ralph L.
Published: June 2004

B cell activation can be quantitated indirectly by assaying antibody production or directly by measuring cellular changes that occur immediately after exposure to an activation signal. This unit provides methods for the latter (direct) approach‐‐namely, methods for quantifying early parameters of B cell activation such as increases in intracellular ionized calcium concentration [Ca2+]I, cell size, and MHC class II‐antigen expression. These assays are not so much B cell specific as activation specific; as such they are a useful means of providing information about cellular events that are independent of antibody secretion. While the protocols included in this unit have been specifically designed for use with murine B cells, they can also be used with B cells obtained from other species. In addition, the method described here also applies to T cells.
Quantitation of Functional T Cells by Limiting Dilution
Authors: Miller Richard A.
Published: June 2004

Culturing by limiting dilution (LD) can be used to estimate the proportion of T cells in a cell preparation which can respond to an activating stimulus to produce interleukin 2 or to generate cytotoxic T lymphocyte (CTL) activity. In the first basic protocol, IL‐2‐producing murine T cells are measured following stimulation by the mitogen Con A. This protocol can be modified as described in the first alternate protocol to quantitate responses to alloantigens, anti‐CD3 antibody, protein antigens such as keyhole limpet hemocyanin (KLH), and viruses. The second alternate protocol provides a modification for using human responder cells. The
second basic protocol is used for estimating the proportion of cells that can generate a clone of cytotoxic effector cells when stimulated by Con A with the addition of IL‐2.
Granule Enzyme Exocytosis Assay for Cytotoxic T Lymphocyte Activation
Authors: Taffs Rolf , Sitkovsky Michail , Takayama Hajime
Published: June 2004

Upon activation by specific target cells, cytotoxic T lymphocytes (CTL) release into the culture medium the content of cytoplasmic granules that contain serine esterases. The amount of enzyme released during CTL activation can be easily quantitated by spectrophotometric measurement of the colored product of the enzymatic degradation of a synthetic substrate. In the primary method presented here, CTL are activated with monoclonal antibodies prepared against the T cell receptor (TCR) complex, then activation is quantitated according to the amount of serine esterase released in the supernatant. Alternate protocols describe the activation of CTL by a combination of protein kinase C and calcium ionophores (a TCR‐independent approach) and by the more conventional approach of target‐cell mediation. In a third approach, β‐glucuronidase rather than esterase activity is measured, as this enzyme is also present in granules released upon CTL activation. This unit therefore includes a colorimetric assay for CTL‐induced β‐glucuronidase activity employing the substrate phenolphthalein glucuronic acid as well as a corresponding automated fluorimetric assay employing the substrate 4‐methylumbelliferyl‐D‐glucuronide. Finally, the quantitation of granule exocytosis resulting from cell damage or death induced by the activating agent, rather than CTL activation, is described.
Isolation of Mouse Small Intestinal Intraepithelial Lymphocytes, Peyer's Patch, and Lamina Propria Cells
Authors: Lefrançois Leo , Lycke Nils
Published: June 2004

The intestinal mucosal immune system is composed of three major lymphoid areas: the lamina propria (LP), which lies just underneath the basement membrane in the intestinal villi; (2) the intraepithelial compartment, which contains the intraepithelial lymphocytes (IEL) and is located just above the basement membrane, between the columnar epithelial cells; and (3) Peyer's patches (PP), lymphoid nodules (akin to lymph nodes) embedded in the gut wall, separated from the LP and IEL. The LP, PP, and IEL lymphoid populations form a complex, interconnected network that responds to immunological insults in the intestine. Therefore, these lymphocyte populations should be analyzed when studying the immunological status of the intestine, for example in oral immunization or in intestinal disease (including infectious disease and tumors). This unit details techniques for isolation of IEL, PP cells, and LP cells from the small intestine of the mouse.
In Vivo Depletion of CD4‐ and CD8‐Specific T Cells
Authors: Kruisbeek Ada M.
Published: June 2004

In vivo depletion of CD4‐ and CD8‐specific T cells is a means of studying the role of these subpopulations in the initiation and effector phases of particular in vivo immune responses. In this unit, a protocol is provided for harvesting anti‐CD4 or anti‐CD8 monoclonal antibody‐ producing ascites fluid or tissue culture supernatant from rat or mouse T cell hybridomas. The antibody (preferably IgG) is then purified and injected intraperitoneally into adult mice. Depletion of the appropriate subset of T cells is verified by flow cytometry analysis of lymph node and spleen cell suspensions in pilot experiments. Once conditions have been established, depleted mice can be used to study the impact of T cell subsets on in vivo immune responses. The depleted condition is maintained by repeated injections of the monoclonal antibody.
Contact Hypersensitivity
Authors: Gaspari Anthony A. , Katz Stephen I.
Published: June 2004

Contact hypersensitivity is a simple in vivo assay of cell‐mediated immune function in which exposure of epidermal cells to exogenous haptens results in a delayed‐type hypersensitive reaction that can be measured and quantified. The Langerhans cell is the critical antigen‐presenting cell in this reaction which initiates sensitization to haptens by presenting antigens to CD4‐bearing T lymphocytes which, in turn, secrete lymphokines and recruit other cells to the site of the reaction. In the protocol described here, mice are shaved and the skin of their abdomens is exposed to a hapten. After 6 days (the afferent phase), the baseline ear thickness is measured prior to initiation of the efferent phase. Finally, the ear is treated epicutaneously with the hapten solution and ear thickness is measured in ˜24 hr. The change in ear thickness after allergen treatment can be used to calculate the percent suppression of contact hypersensitivity.
Skin Allograft Rejection
Authors: Rosenberg Amy S.
Published: June 2004

Skin allograft rejection is a test of the competence of T lymphocytes to mediate in vivo tissue destruction, which in turn reflects their role in critical functions such as anti‐viral and tumor immunity. The tail‐skin graft procedure described here is useful predominantly because of the ease of preparation and resistance to ischemic (nonspecific) necrosis. Additionally, it is not necessary to sacrifice the donor mouse. However, rejection of tail‐skin grafts should not be used to test for genetic homogeneity in breeding experiments or to detect minor histocompatibility (minor‐H) antigens because tail skin is less sensitive than trunk skin in detecting such differences. The trunk‐skin graft procedure detailed here more difficult to perform than the tail‐skin graft. It requires extensive tissue preparation and is more susceptible to ischemic necrosis. Trunk‐skin grafts are especially useful for detecting minimal histocompatibility differences.
Measurement of T and B Cell Turnover with Bromodeoxyuridine
Authors: Tough David F. , Sprent Jonathan
Published: June 2004

The rate of turnover of T and B cells can be measured in terms of the time between two mitoses or between mitosis and death. In vivo, this is accomplished by administering DNA precursors and measuring the rate at which cells become labeled with these precursors.
5‐bromo‐2'‐deoxyuridine (BrdU) is a thymidine analog that is incorporated into the DNA of cells during the S phase of the cell cycle and can be subsequently detected with a specific monoclonal antibody. In this method, BrdU is given to experimental animals either continuously or as a pulse, depending on the goal of the experiment. Subsequently, cells are stained in vitro with an anti‐BrdU antibody and labeled cells detected by flow cytometry. By combining cell surface staining for multiple lymphocyte markers with anti‐BrdU staining, it is possible to determine the lifespan of phenotypically defined subsets.
Flow Cytometry Analysis Using the Becton Dickinson FACS Calibur
Authors: Holmes Kevin L. , Otten Gillis , Yokoyama Wayne M.
Published: June 2004

This unit details the operation of a FACS Calibur flow cytometer for cell analysis. The operation of the Becton Dickinson FACS Calibur and CELLQuest software version 3.0 is described, but the unit is general enough to be helpful for users of all flow cytometers. The FACS Calibur replaces both the FACSCan and FACSort; the information presented here is also applicable to older BD instruments. In this unit, particular emphasis is placed on data acquisition rather than data analysis. Single‐color analysis using fluorescein isothiocyanate (FITC)‐conjugated antibodies is described along with procedures to check instrument performance and sensitivity, single‐color (FITC) analysis with simultaneous live/dead discrimination using propidium iodide (PI), simultaneous two‐color analysis using FITC‐ and phycoerythrin (PE)‐conjugated antibodies, two‐color FITC/PE analysis with simultaneous live/dead discrimination using PI, simultaneous three‐color immunofluorescence with FITC, PE, and the red fluorescent dyes, and finally, simultaneous four‐color immunofluorescence with FITC, PE, red fluorescence dyes, and APC.
Analysis of Cellular DNA Content by Flow Cytometry
Authors: Darzynkiewicz Zbigniew , Huang Xuan
Published: June 2004

Cellular DNA content can be measured by flow or laser‐scanning cytometry with the aim of (1) revealing cell distribution within the major phases of the cell cycle, (2) estimating the frequency of apoptotic cells with fractional DNA content, and/or (3) disclosing the DNA‐ploidy of the measured cell population. In this unit, simple and universally applicable methods for staining fixed cells are presented, as are methods that utilize detergents and proteolytic treatment to permeabilize cells. Additionally, supravital cell staining with Hoechst 33342, which is primarily used for sorting live cells for subsequent culturing based on DNA‐content differences, is also described. Also presented are methods for staining of cell nuclei isolated from paraffin‐embedded tissues, and deconvolution of DNA‐content‐frequency histograms to estimate the percentage of cells in major phases of the cell cycle and frequency of apoptotic cells with fractional DNA content.
Multiparameter FACS Analysis
Authors: Roederer Mario
Published: June 2004

The purpose of this unit is three‐fold: (1) toprovide a brief historical perspective to illustrate the demands for this technology and to lay the foundation for its application; (2) to explain the hurdles that were surmounted to achieve the current level of multiparametric analysis which serves to alert researchers to potential problems they may encounter when they either bring this technology to their own laboratories, or when they use extant instrumentation in another laboratory; and (3) to illustrate some of the complexities that arise even when all of the major technical hurdles have been overcome. The majority of the unit is focused on “high‐end” multiparameter analysis (termed “polychromatic flow cytometry,” or PFC); i.e., experiments utilizing more than four fluorescence detectors (in addition to scattered light parameters).
Measurement of Mouse and Human Interleukin 5
Authors: Harriman Gregory R.
Published: June 2004

Two fundamentally different types of assays are available for quantitating interleukin 5 in biological samples. One type, a bioassay, is based on the ability of IL‐5 to enhance B cell proliferation and immunoglobulin secretion or eosinophil proliferation and differentiation. The other type of assay, an ELISA, uses antibodies against IL‐5 to capture and quantitate IL‐5 in samples. Advantages and disadvantages of each assay are discussed. The bioassay described in this unit, utilizing BCL1 cells as the indicator line, has been designed primarily to assay murine IL‐5; however, it can also be used to measure human IL‐5. The IL‐5 ELISA, while sensitive and specific, provides no information about biological activity.
Measurement of Interleukin 6
Authors: Nordan Richard P. , Richards C.D. , Gauldie Jack
Published: June 2004

Interleukin 6 (IL‐6) is a pluripotent cytokine with multiple effects on many different cell types. It is produced by a variety of cells in response to immunological and other stimuli. This unit describes a simple and sensitive assay for human, rat, rabbit, pig, and mouse IL‐6, based on IL‐6‐dependent proliferation of a murine B cell hybridoma cell line, B9. Support protocols discuss maintenance of B9 cells, preparation of IL‐6 standards, and production of IL‐6‐containing supernatant. In addition, IL‐6 ELISA kits for the measurement of human or mouse IL‐6 are available from a number of companies. These products are reliable, highly sensitive, and specific, and thus should be considered as an excellent (although more expensive) alternative, keeping in mind that bioactivity is not assessed with this approach
Biological Responses to Chemokine Superfamily Members
Authors: Taub Dennis D. , Schaffer Eric
Published: June 2004

A detailed summary of all the chemokine subfamily members including alternative names, receptor‐ligand pairs, receptor cellular distribution, and leukocyte responsiveness is presented here along with assays to measure the biological activities of chemokines on each of the major leukocyte subsets (i.e., microchemotaxis chambers, fluorescence‐based assays, use of nitrocellulose filters and membranes coated with extracellular matrix proteins, cellular adhesion assays, mobilization of intracellular free calcium, actin polymerization, degranulation assays, and the use of chemokine receptor‐specific antibodies in flow cytometric analysis and radiolabeled chemokine binding assays).
Measurement of Mouse and Human Interleukin 9
Authors: Renauld Jean‐Christophe , Van Snick Jacques
Published: June 2004

This unit describes two proliferation assays to detect or quantitate human and murine interleukin 9 (IL‐9). The first is based on the ability of IL‐9 to stimulate the proliferation of the TS1h9RA3 cell line, a murine IL‐9‐dependent cell line transfected with the human IL‐9 receptor. An alternate protocol is based on the ability of IL‐9 to stimulate the proliferation of the human megakaryoblastic leukemia cell line, M‐O7e. M‐O7e cells depend on either human IL‐3 or granulocyte/macrophage colony‐stimulating factor (GM‐CSF) for growth, although other cytokines including IL‐2, ‐4, ‐6, and ‐9 and steel factor are also weakly mitogenic (relative to IL‐3 and GM‐CSF). Thus, although M‐O7e cells can readily be used to quantitate levels of IL‐9 in the absence of other cytokines, analysis in the presence of complex mixtures of cytokines (e.g., natural sources) requires the use of specific antibodies against IL‐9 and the other cytokines, as described in this unit.
Measurement of Human and Mouse Interleukin‐12
Authors: Gately Maurice K. , Chizzonite Richard , Presky David H.
Published: June 2004

This unit describes functional assays for measurement of bioactive IL‐12 and ELISAs for measurement of IL‐12 protein. The functional assays are based on the ability of IL‐12 to stimulate proliferation of PHA‐activated T lymphoblasts (“PHA blasts”). The ELISAs are technically simpler to perform than the functional assays, but cannot distinguish bioactive from inactive cytokine.
ELISPOT Assay to Detect Cytokine‐Secreting Murine and Human Cells
Authors: Klinman Dennis M. , Nutman Thomas B.
Published: June 2004

The filter immunoplaque assay, otherwise called the enzyme‐linked immunospot assay (ELISPOT), was initially developed to detect and quantitate individual antibody‐secreting B cells. Recent modifications have improved the sensitivity of the ELISPOT assay such that cells producing as few as 100 molecules of specific protein per second can be detected. The ELISPOT assay utilizes two high‐affinity cytokine‐specific antibodies directed against different epitopes on the same cytokine molecule: either two monoclonal antibodies or a combination of one monoclonal antibody and one polyvalent antiserum. ELISPOT generates spots based on a colorimetric reaction that detects the cytokine secreted by a single cell. The spot represents a “footprint” of the original cytokine‐producing cell. Spots are permanent and can be quantitated visually, microscopically, or electronically.
Detection of Cytokine Receptors by Flow Cytometry
Authors: Zola Heddy
Published: June 2004

This unit describes three protocols for detecting cytokine receptors on human peripheral blood mononuclear cells or murine splenic lymphocytes via immunofluorescence labeling. The three protocols have in common the use of phycoerythrin as a highly effective fluorescent dye, multiple‐layer staining to increase sensitivity, and selection of reagents for maximum sensitivity. The first procedure uses monoclonal antibody (MAb) against receptor protein, detected with biotinylated anti‐Ig (directed against the species in which the MAb is made), which in turn is stained with phycoerythrin‐streptavidin (PE‐SA). The second procedure uses biotinylated cytokines in the first step, followed by PE‐SA. Finally, a third procedure describes multiparameter analysis involving second and third fluorophores introduced as direct antibody conjugates, which is used to study subset expression of receptors. Instructions are also provided for titrating MAbs and cytokines, as well as how to evaluate detection reagents such as anti‐Ig conjugates and fluorophores.
Measurement of Interleukin 15
Authors: Paxton Raymond J.
Published: June 2004

Interleukin 15 (IL‐15) is a small helical cytokine that was first characterized by its ability to stimulate proliferation of the murine T cell line CTLL‐2, but is now known to have activity on a variety of cell types. This unit describes two protocols that can be used to measure IL‐15 in culture supernatants and serum samples. The first method uses an enzyme‐linked immunosorbent assay (ELISA) that is specific for human IL‐15, although it can also detect simian IL‐15. The second method relies on the ability of IL‐15 to support proliferation of CTLL‐2 cells. The CTLL‐2 assay is slightly more sensitive than the ELISA, but it is not specific for IL‐15, because CTLL‐2 cells also respond to IL‐2 and IL‐4. The CTLL‐2 assay can be used to measure human, simian, or murine IL‐15.
Measurement of Interleukin 17
Authors: Patera Andriani C. , Cohen Jeffrey I.
Published: June 2004

This unit describes two procedures for measurement of interleukin‐17 (IL‐17). IL‐17 is a pleiotropic proinflammatory cytokine that binds to its ubiquitously expressed receptor, IL‐17R, resulting in the induction of several other cytokines including IL‐6 and IL‐8. The unit guides the reader on the use of individual reagents and commercial kits available for detection of murine and human IL‐17 by ELISA or Western blotting and presents step‐by‐step protocols for each of these approaches. In addition to detection of IL‐17 protein, a functional assay is described which measures the biological activity of IL‐17 by induction of IL‐6 from fibroblasts.
Detection and Isolation of Cytokine Secreting Cells Using the Cytometric Cytokine Secretion Assay
Authors: Assenmacher Mario , Löhning Max , Radbruch Andreas
Published: June 2004

The cell‐surface affinity matrix technology, otherwise called “secretion assay” or “capture assay,” represents a new, innovative method for the analysis and enrichment of viable cells according to secreted molecules such as antibodies or cytokineof T cells to induce cytokine secretion is included. Alternate protocols provide very rapid and simple procedures for the direct detection of antigen‐specific T cells from small numbers of PBMCs or whole blood without magnetic enrichment.
The In Vivo Cytokine Capture Assay for Measurement of Cytokine Production in the Mouse
Authors: Finkelman Fred , Morris Suzanne , Orekhova Tatyana , Sehy David
Published: June 2004

Because most cytokines are utilized, catabolized, or excreted shortly after they are produced, it has been difficult to directly measure in vivo cytokine production. Consequently, it has been necessary to infer in vivo cytokine secretion levels from the results of ex vivo assays of cytokine secretion, assays that measure tissue levels of cytokine mRNA, or assays that stain tissues for cytokine protein levels. Results of these assays provide important and useful information, but do not necessarily reflect in vivo cytokine secretion. To better determine in vivo cytokine production, the in vivo cytokine capture assay (IVCCA) was developed. IVCCA facilitates measurement of cytokines in serum by increasing their in vivo half‐lives. This increases the sensitivity of measurement of in vivo cytokine production 30‐ to 1,000‐fold. The first protocol described in this unit is for luminescence‐based ELISA, while the second is for an absorbance‐based method.
Isolation of T Cells Using Rosetting Procedures
Authors: Kanof Marjorie E.
Published: June 2004

This unit describes a procedure for separating T cells from other mononuclear cells by exploiting the unique ability of cells to bind to and form rosettes with sheep red blood cells (SRBC). This isolation method also allows recovery of the nonrosetting cell population (B lymphocytes, monocytes, and macrophages). Neuraminidase‐ and 2 ‐aminoethylisothiouronium bromide (AET)‐treated SRBC are used for rosetting because of enhanced binding to T cells.
Isolation of B Cell Populations
Authors: Splawski Judy B. , Lipsky Peter E. , Eisenstein Eli M. , Chua Kevin S.
Published: June 2004

This unit describes procedures for the isolation of pure human B cell populations. Two broad approaches to the isolation of pure human B cell populations are available, one based on techniques of negative selection and the other using positive selection. The negative selection approach has the advantage that the purified B cells obtained are less likely to be functionally altered by the isolation procedure. In addition, this approach does not require the use of expensive antibodies, which are usually the key element of a positive selection procedure. On the other hand, negative selection has the disadvantage that cell types not specifically targeted for removal (such as eosinophils) may be present in the final cell population. The positive selection method is an immunomagnetic positive selection using magnetic beads conjugated directly to an antibody specific for the B cell membrane determinant CD19. Also presented is a Support Protocol detailing the preparation of neuraminidase‐treated sheep red blood cells for use in rosetting.
Measurement of Proliferative Responses of Cultured Lymphocytes
Authors: James Stephen P.
Published: June 2004

Measurement of proliferative responses of human lymphocytes is a fundamental technique for the assessment of their biological responses to various stimuli. Most simply, this involves measurement of the number of cells present in a culture before and after the addition of a stimulating agent. This unit contains several different prototype protocols to measure the proliferation response of lymphocytes following exposure to mitogens, antigens, allogeneic or autologous cells, or soluble factors. Each of these protocols can be used in conjunction with an accompanying support protocol which contains methods for pulsing cultures with [3H]thymidine and determining incorporation of [3H]thymidine into DNA. The protocols described here provide an estimate of DNA synthesis and cell proliferation in an entire cell population, but do not provide information on the proliferation of individual cells.
Induction of Polyclonal Immunoglobulin Synthesis
Authors: James Stephen P.
Published: June 2004

This unit is designed to examine the effects of T cells or lymphokines on B cell differentiation in situations where the antigen specificity of the B cells is not of interest. In these cases, antibody production induced by polyclonal stimuli (e.g., mitogens, antibodies, Epstein‐Barr virus (EBV), or lymphokines) can be measured instead of antigen‐specific immunoglobulin production. Because B cells in the circulation and tissues are pleomorphic (containing subpopulations of cells that may be resting, cells that may have had prior antigenic exposure, and cells that may have undergone prior isotype commitment), the antibody responses of these subpopulations to various growth and differentiation factors differ. Therefore, the choice of which lymphocyte subpopulation to culture and which activation signal to use is determined by the particular experimental question.
Measurement of Polyclonal Immunoglobulin Synthesis Using ELISA
Authors: Nutman Thomas B.
Published: June 2004

The enzyme‐linked immunosorbent assay (ELISA) is a simple, specific, highly reproducible, and nonradioactive method for measuring immunoglobulin levels in cell culture supernatants and other biologic fluids. It can be used to measure the common immunoglobulin classes (IgG, IgM, and IgA) as described in the basic protocol, as well as classes and subclasses found in low levels in these fluids (IgG4 and IgE), as detailed in the alternate protocols.
Measurement of Polyclonal Immunoglobulin Synthesis Using the Reverse Plaque Technique
Authors: Tosato Giovanna , Pike Sandra E. , Blaese Michael
Published: June 2004

This unit describes the reverse hemolytic plaque assay, an effective method for measuring the number of immunoglobulin (Ig)‐secreting cells present in a cell population at any particular time. Cell populations that can be assayed using the technique include peripheral blood mononuclear cells or cells from tissues such as the tonsils. The basic protocol is divided into three stages. First, protein A‐sensitized sheep red blood cells (SRBC), guinea pig complement, and anti‐Ig antibody are prepared. Test samples are then combined and incubated with the SRBC, complement, and antibody in appropriate chambers. Finally, the resulting plaques are scored. A support protocol describes the preparation of plaquing chambers.
Isolation and Functional Analysis of Neutrophils
Authors: Clark Robert A. , Nauseef William M.
Published: June 2004

This unit describes the isolation of high‐purity neutrophils (or polymorphonuclear leukocytes; PMN) and the assays that can be performed to assess their function. An assay for the in vitro study of phagocytic capacity is presented, and microbicidal assays using lysostaphin and differential centrifugation are also described. During phagocytosis or on exposure to soluble agonists, neutrophils exhibit a marked increase in oxidative metabolism which is mediated by activation of NADPH oxidase and results in the formation of superoxide anion as well as hydrogen peroxide and other reactive species. The nitroblue tetrazolium reduction slide test is presented, and is a nonquantitative screening test for the presence of reactive oxygen products. The quantitation of superoxide by the kinetic and static assays are detailed, and a “broken cell” assay is also outlined.Finally, measurement of hydrogen peroxide formation is described.
Isolation of Human Basophils
Authors: Metcalfe Dean D.
Published: June 2004

Human basophils are normally the rarest of all circulating leukocytes and thus are among the most difficult of the blood elements to purify. In this unit, basophils are isolated from leukocyte populations, resulting in preparations of cells of which 5% to 50% are basophils and the rest are lymphocytes along with neutrophils and monocytes. This technique is advantageous because of its speed and because it minimizes basophil stimulation and histamine release.
51Cr Release Assay of Antibody‐Dependent Cell‐Mediated Cytotoxicity (ADCC)
Authors: Nelson David L. , Kurman Carole C. , Serbousek Deborah E.
Published: June 2004

Antibody‐dependent cell‐mediated cytotoxicity (ADCC) is an immunologic cytotoxic effector mechanism that is dependent on the cooperative interaction of humoral and cellular effector elements. This unit describes an assay of ADCC activity that can be used as a test for immunocompetence in effector cells or to test the activity of a monoclonal antibody to mediate ADCC. In this form of cytotoxicity, effector cells with receptors for the Fc portion of immunoglobulin produce target cell lysis by attachment to the Fc portion of antibodies that are bound to target cells via their antigen‐combining sites. Therefore, an ADCC assay involves three essential components: labeled target cells, antibodies with specificity for target‐cell surface antigens, and effector‐cell populations. The basic protocol describes a method of measuring ADCC effector activity in lymphoid cells (peripheral blood mononuclear cells, or PBMC) that employs
51Cr‐labeled target cells. The three components are mixed in microtiter‐plate wells and lysis of the target cells is detected by measuring the release of radioactivity into the cell supernatant. Support protocols describe procedures for preparing anti‐target cell antiserum and 51Cr‐labeled target cells.
Cell Adhesion Under Hydrodynamic Flow Conditions
Authors: Gopalan Priya K. , Jones David A. , McIntire Larry V. , Smith C. Wayne
Published: June 2004

This unit describes a hydrodynamic assay to study the relative importance of various receptor/ligand interactions in cell‐cell and cell‐substrate adhesion and to quantitate the strength of their binding. The basic protocol describes how to assemble the single‐chamber flow system with the substrate, add the cells in suspension, and record the experiment on videotape. Alternate protocols present assays to determine how monoclonal antibodies and stimulating and inhibiting agents affect the substrate and the perfusing cells in suspension. Another alternate protocol details the use of the double‐chamber flow system. Support protocols describe how to construct the single‐ and double‐chamber flow systems and how to analyze the data from an experiment. Recording and analyzing the flow experiment requires the use of video equipment and, optionally, a computer and imaging software.
Isolation of Human Intestinal Mucosal Mononuclear Cells
Authors: Fiocchi Claudio , Youngman Kenneth R.
Published: June 2004

The aim of this procedure is to obtain large numbers of isolated, viable, and functional mononuclear cells that are representative of the lymphoid population present in the mucosa of the human gastrointestinal tract under physiological and pathological conditions. The basic protocol is based on the use of surgically resected small and large bowel specimens, and consists of two basic stages: (1) a combination of chemical, enzymatic, and mechanical treatments to dissociate intestinal tissue and free the mononuclear cells from the surrounding interstitial framework; and (2) separation, isolation, and purification of viable mucosal lamina propria mononuclear cells from other cellular and amorphous components. The proportion of viable cells obtained can be increased by including extra separation steps using nylon wool columns or Percoll gradients, as described.
Isolation and Generation of Human Dendritic Cells
Authors: Tedder Thomas F. , Jansen Paul J.
Published: June 2004

Dendritic cells are highly specialized antigen‐presenting cells (APC), which may be isolated or generated from human blood mononuclear cells. Although mature blood dendritic cells normally represent ˜0.2% of human blood mononuclear cells, their frequency can be greatly increased using the cell enrichment methods described in this unit. More highly purified dendritic cell preparations can be obtained from these populations by sorting of fluorescence‐labeled cells. Alternatively, dendritic cells can be generated from monocytes by culture with the appropriate cytokines, as described here. In addition, a negative selection approach is provided that may be employed to generate cell preparations that have been depleted of dendritic cells to be used for comparison in functional studies.
Measurement of Prostaglandins and Other Eicosanoids
Authors: Stenson William F.
Published: June 2004

Eicosanoids are arachidonic acid metabolites formed through the cyclooxygenase or one of the lipoxygenase metabolic pathways. They include the prostaglandins (PG), leukotrienes (LT), thromboxanes (Tx), peptidoleukotrienes, and 5‐hydroxyeicosatetraenoic acid (5‐HETE). This unit describes procedures for measuring individual eicosanoids using several commercial kits. These include kits for an enzyme immunoasssay (ELISA) and a radioimmunoassay (RIA). A method is also described that can be used to analyze the full range of eicosanoids produced in a biological system, and a support protocol presents an acetone/chloroform extraction method for purifying eicosanoids prior to assay. Another support protocol describes a different extraction protocol in which samples are run over an octadecylsilyl silica column.
Solubilization of Lymphocytes
Authors: Weissman Allan M.
Published: June 2004

Purification and study of transmembrane proteins require isolation of these structures from their lipid environment. This isolation is carried out through the use of detergents. In this unit, several approaches to solubilization of membrane proteins are presented. Solubilization of whole lymphocytes, using conditions aimed at minimizing the disruption of protein‐protein interactions, is described with an optional step that may be useful when the disruption of protein interactions is desired as part of a purification protocol. In some situations, it may be desirable to purify membranes prior to their solubilization or to determine the physical relationship between proteins, which can be accomplished by a cross‐linking.
Purification of Organelles from Mammalian Cells
Authors: Castle J. David
Published: June 2004

Organelle purification procedures capitalize on the differences in size, density, and (occasionally) surface charge density of individual types of organelles. Most fractionation procedures that are based on centrifugation involve some combination of procedures that distinguish both size and density. Initially, a homogenate is prepared in isoosmotic (or slightly hyperosmotic) sucrose or some other predominantly nonelectrolyte medium. A wide range of procedures have been used to fractionate tissue homogenates. The protocols in this unit emphasize different fractionation techniques that have been used for rat liver, an abundant tissue that has been a favorite of many investigators and has served as the source of many organelle preparations of excellent purity. For selected procedures, examples have been given using other tissue sources (e.g., glandular tissues that maintain protein storage granules for regulated secretion) or, where particularly favorable, cultured cells.
Diagonal Gel Electrophoresis
Authors: Samelson L.E.
Published: June 2004

Diagonal gel electrophoresis is a form of two‐dimensional analysis useful for investigating the subunit composition of multisubunit proteins containing interchain disulfide bonds. Proteins are electrophoresed in the first dimension in a slab or tube gel under nonreducing conditions. The proteins are then reduced in the gel and this piece of gel is layered onto a second gel and electrophoresed. In the second gel, the proteins migrate at right angles to their original, first‐dimension migration. The majority of cellular proteins are not disulfide‐linked and will fall on the “diagonal” in this system; that is, they migrate approximately equal distances in both directions during electrophoresis and lie approximately on the diagonal line connecting opposite corners of the gel. Upon reduction, component subunits of proteins connected by interchain disulfide bonds will resolve below the diagonal because the individual subunits migrate faster than the disulfide‐linked complex during the second electrophoresis.
Immunoblotting and Immunodetection
Authors: Gallagher Sean , Winston Scott E. , Fuller Steven A. , Hurrell John G.R.
Published: June 2004

Immunoblotting (often referred to as western blotting) is used to identify specific antigens recognized by polyclonal or monoclonal antibodies. This unit presents procedures for electrophoretically transferring antigens from a denaturing polyacrylamide gel in a tank or a semidry transfer apparatus to a nitrocellulose, PVDF, or nylon membrane. The process can be monitored by reversible staining or by Ponceau S staining, both of which are described here. A protocol for blotting previously stained gels is also described. The transferred proteins are bound to the surface of the membrane, providing access for reaction with immunodetection reagents. All remaining binding sites are blocked by immersing the membrane in a solution containing either a protein or detergent blocking agent. After probing with the primary antibody, the membrane is washed and the antibody‐antigen complexes are identified with horseradish peroxidase (HRPO) or alkaline phosphatase enzymes coupled to the secondary anti‐IgG antibody (e.g., goat anti‐rabbit IgG). The enzymes are attached directly or via an avidin‐biotin bridge to the secondary antibody, and protocols are provided for both methods. Chromogenic or luminescent substrates are then used as described to visualize the activity. Finally, a method for stripping and reprobing membranes is presented.
Release of Saccharides from Glycoconjugates
Authors: Powell Leland D. , Varki Ajit P. , Freeze Hudson H.
Published: June 2004

Glycosidases are specific enzymes that can partially or completely remove sugar chains from cell‐surface glycoconjugates. The enzymes can be exoglycosidases (which remove a terminal saccharide unit from an oligosaccharide chain), endoglycosidases (which cleave within an oligosaccharide chain, releasing an oligosaccharide fragment), or glycoamidases (which cleave between an oligosaccharide unit and its N‐linkage to a protein). Commonly used examples of each enzyme are presented in this unit. Sialidase digestion of purified proteins is described and an Alternate Protocol details the application of the technique to intact cell suspensions. A support protocol describes testing sialidase activity in a variety of buffers. Basic protocols are detailed for the digestion of intact glycoproteins and glycopeptides by the most common endoglycosidases and glycoamidases: Endoglycosidase H (Endo H), Endoglycosidase F2 (Endo F2), and Peptide:N‐glycosidase F (PNGase F). The applications described in the second and third support protocols employ these enzymes alone or as part of a sequential digestion. The second support protocol describes how to use partial digestions with one enzyme or sequential digestions with different enzymes to estimate the number or types of N‐linked carbohydrate chains on a protein. The third support protocols describes sample preparation and digestion followed by gel‐filtration chromatography to recover the released radiolabeled oligosaccharide chains for subsequent analysis.
Biotinylation and Analysis of Membrane‐Bound and Soluble Proteins
Authors: Lisanti M.P. , Sargiacomo M.
Published: June 2004

Coupling of a biotin moiety to proteins allows sensitive detection of those proteins with a variety of commercially available avidin or streptavidin conjugates. This unit presents protocols for biotinylation of cell‐surface proteins and proteins in solution. A major advantage of these techniques is that they do not require use of radioisotopes (although use of radioiodinated streptavidin is optional for biotin detection).
Synthetic Peptides for Production of Antibodies that Recognize Intact Proteins
Authors: Grant Gregory A.
Published: June 2004

Antibodies that recognize intact proteins can be produced through the use of synthetic peptides based on short stretches of the protein sequence, without first having to isolate the protein. The steps to produce an effective antibody include: (1) designing the peptide sequence based on the sequence of the protein; (2) synthesizing the peptide; (3) preparing the immunogen either by coupling the synthetic peptide to a carrier protein or through the use of a multiple antigenic peptide (MAP); (4) immunizing the host animal; (5) assaying antibody titer in the host animal's serum; and (6) obtaining the antiserum and/or isolating the antibody. This unit covers steps 1 and 3. Once the coupling procedure has been performed, it is possible to determine the approximate degree of coupling by amino acid analysis. Also presented are methods for assaying free sulfhydryl content and for reducing disulfide bonds in synthetic peptides.
B Cell Epitope Mapping Using Synthetic Peptides
Authors: Carter John Mark , Loomis‐Price Larry
Published: June 2004

Synthetic peptides are used for identification of functional B cell epitopes in antibody preparations. ELISA‐type assays are used to identify sequences of proteins comprising antibody‐binding regions. This unit describes three peptide display formats. Pepscan (pins) and SPOTs (cellulose membranes) may be used as solid‐phase support media for peptide synthesis, followed by ELISA directly on the resulting peptide array. Alternatively, peptides may be cleaved from the array and tested in a standard microplate‐based antibody capture ELISA format. The discussion includes choosing the peptide sequences by length and overlap, as well as determination of the minimum essential sequence for antibody binding. This method is highly effective for continuous epitopes and is often also useful for discontinuous epitopes.