Determining the Class and Subclass of Monoclonal Antibodies Using Antibody Capture on Anti-Immunoglobulin Antibody-Coated Plates

By definition, a monoclonal antibody should only be of a single class or subclass. Each class of antibody is associated with specific functions, and it can be useful to know the class/subclass of the monoclonal antibody produced by a specific hybridoma. In this protocol, class/subclass-specific antibodies are used to capture the monoclonal antibody from hybridoma supernatant. If the antibody is clonal, it will only be bound by one anti-heavy-chain capture antibody and one anti-light-chain antibody. No antigen is required.

Generating Monoclonal Antibodies

Antibodies that are produced by hybridomas are known as monoclonal antibodies. Here we introduce methods for generating and screening monoclonal antibodies, including developing the screening procedure and producing hybridomas.

Immunizing Animals

The traditional method for generating polyclonal and monoclonal antibodies requires the immunization of an animal. Selecting the best species of animal and getting that animal's immune system to respond to a target antigen with an antibody response are essential to obtaining good-quality antibodies and hybridomas. There are only a limited number of opportunities for a researcher to intervene to manipulate and tailor the response to a particular antigen. Here we present advice and methods for designing the way in which the antigen is presented to the immune system (i.e., the immunization protocol), including the choice of animal, the antigen dose, the use of adjuvants, the route and number of injections, and the period between injections.

Hybridoma Screening by Antibody Capture: Flow Cytometry/FACS with Permeabilized Cells to Detect Intracellular Binding

Flow cytometry or fluorescence-activated cell sorting (FACS) can be used to identify hybridomas secreting monoclonal antibodies to internal cellular proteins, but the cells must be permeabilized before the hybridoma supernatants are applied. In using this technique, useful controls are positive and negative cell lines with primary and secondary antibodies as well as positive and negative cell lines with secondary antibody alone.

Hybridoma Screening by Antibody Capture: Flow Cytometry/FACS with Whole Cells to Detect Cell-Surface Binding

If the antigen of interest is a cell-surface protein, flow cytometry or fluorescence-activated cell sorting (FACS) can be used to identify hybridomas secreting monoclonal antibodies to these proteins. Two alternative protocols are presented here-staining in individual tubes and staining in 96-well plates.

Determining the Class and Subclass of a Monoclonal Antibody by Ouchterlony Double-Diffusion Assays

Originally, the Ouchterlony double-diffusion assays were the most common method for determining the class and subclass of a monoclonal antibody, and they still are useful, particularly when only a few assays will be performed. A sample of hybridoma tissue culture supernatant is placed in a well in a bed of agar, and class- and subclass-specific antisera are placed in other wells in a ring surrounding the test antibody. As the antibodies diffuse into the agar, they meet and multimeric immune complexes precipitate to form a visible "precipitin line."

Hybridoma Screening by Antibody Capture: Enzyme-Linked Detection (Indirect ELISA) in Polyvinyl Chloride Wells

In this antibody capture assay for hybridoma screening, the antigen is immobilized on a solid substrate (the surface of the wells in a polyvinyl chloride [PVC] microtiter plate), and antibodies in the hybridoma tissue culture supernatant are incubated with the antigen. Unbound antibodies are removed by washing, and antibody-antigen complexes are detected by secondary antibody conjugated to alkaline phosphatase (AP), which catalyzes the conversion of a chromogenic substrate to a blue/green product. Alternative secondary antibodies are necessary for experiments in which immunoglobulin-fusion proteins have been used as immunogens or screening proteins.

Hybridoma Screening by Antibody Capture: Immunohistochemistry

To determine the subcellular location of an antigen, hybridoma tissue culture supernatants can be screened using immunohistochemistry. For antibodies to have access to antigens in fixed and embedded tissue sections, the paraffin must be removed and the tissue must be rehydrated and digested before immunohistochemical staining.

Hybridoma Screening by Antibody Capture: Detection of Cell-Surface Binding by Immunofluorescence

If the antigen of interest is a cell-surface protein, immunofluorescence can be used to identify hybridomas secreting monoclonal antibodies to these proteins. They can be stained on microscope chamber slides, flat-bottomed cell culture plates (96, 48, or 24 well), or imaging plates (96 well). This protocol uses 96-well imaging plates.

Hybridoma Screening by Antibody Capture: Detection of Intracellular Binding by Immunofluorescence

Hybridoma screening by immunofluorescence stainings of whole cells can be adapted to screen for antibodies to internal proteins by permeabilizing the cells before applying the hybridoma supernatants. In this protocol, cells are attached to a solid support (glass slides), which makes them easy to manipulate and transfer between different reagent solutions.

Hybridoma Screening by Antigen Capture: Immunoprecipitation

Immunoprecipitation is rarely used for screening hybridoma fusions because the assays are tedious and time-consuming. However, it can be useful when working with complex antigens because the precipitated antigen is normally detected after sodium dodecyl sulfate (SDS)-polyacrylamide electrophoresis and thus it is simple to discriminate between true and false positives. Furthermore, the assay provides information regarding the molecular weight of the antigen.

Hybridoma Screening by Antigen Capture: Reverse Dot Blot

A dot blot is widely used to determine the productivity of a given hybridoma. This assay can also be used to screen a fusion or subclone plate for productive hybridoma clones. First, a nitrocellulose membrane is coated with an affinity-purified goat or rabbit anti-mouse immunoglobulin and then incubated with hybridoma tissue culture supernatant. Monoclonal antibodies in the supernatant are then "captured" on the coated nitrocellulose membrane surface and detected by screening with horseradish peroxidase (HRP).

Hybridoma Screening by Antigen Capture: Capture or Sandwich ELISA in 96-Well Plates

In an antigen capture assay for hybridoma screening, the detection method identifies the presence of the antigen. Often this is achieved by labeling the antigen directly. In this assay, the polyvinyl chloride (PVC) wells of a high-binding-capacity ELISA plate are first coated with an affinity-purified rabbit anti-mouse immunoglobulin and then incubated with hybridoma tissue culture supernatant. Monoclonal antibodies in the supernatant are "captured" on the coated PVC surface and detected by screening with biotin- or histidine (His)-tagged antigen. The antigen can be labeled to a high specific activity and thus very little antigen is required for this procedure.

Adoptive Transfer Immunization of Mice

This procedure is designed to enrich and expand antibody-forming cells for use in generating monoclonal antibodies. Gamma-irradiation is used to wipe out the immune system in a recipient animal, after which spleen cells that have reverted to memory cells are obtained from syngeneic donor animals and transferred to the irradiated animal, allowing the implanted immune cells to take over. This method can produce an 80-fold enrichment of antibody-producing cells over that obtained in the original immunized animal.

Selecting the Antigen

The classical method for generating polyclonal or monoclonal antibodies relies on the in vivo humoral response of animals. Here we describe the factors that antigens can have that might influence the strength and quality of an antibody response. This introduction is divided into three sections: (1) an overview of immunogenicity, (2) choosing the best form for the immunogen, and (3) methods for modifying antigens to make them more immunogenic.

Hybridoma Screening by Antibody Capture: Dot Blot

A dot blot is an appropriate hybridoma screening procedure when the antigen is a protein that is available in purified form. The antigen is bound directly to a nitrocellulose sheet and incubated with hybridoma tissue culture supernatant. A dot blot is widely used to determine the productivity of a given hybridoma, and this is described here. This assay can also be used to screen a fusion or subclone plate for productive hybridoma clones.

Hybridoma Screening by Antibody Capture: A High-Throughput Western Blot Assay

Antibody capture assays are often the easiest and most convenient of the hybridoma screening methods. In this procedure, proteins in solution or in a cell lysate are separated according to size by gel electrophoresis and then transferred by blotting to a nitrocellulose sheet. Antigen bound to the solid substrate is incubated with the primary antibody, and the resultant antibody-antigen complexes are detected by a horseradish peroxidase (HRP)-conjugated secondary antibody and a chemiluminescent substrate for HRP.

Monitoring PD-1 Phosphorylation to Evaluate PD-1 Signaling during Antitumor Immune Responses

PD-1 expression marks activated T cells susceptible to PD-1-mediated inhibition but not whether a PD-1-mediated signal is being delivered. Molecular predictors of response to PD-1 immune checkpoint blockade (ICB) are needed. We describe a monoclonal antibody (mAb) that detects PD-1 signaling through the detection of phosphorylation of the immunotyrosine switch motif (ITSM) in the intracellular tail of mouse and human PD-1 (phospho-PD-1). We showed PD-1⁺ tumor-infiltrating lymphocytes (TILs) in MC38 murine tumors had high phosphorylated PD-1, particularly in PD-1⁺TIM-3⁺ TILs. Upon PD-1 blockade, PD-1 phosphorylation was decreased in CD8⁺ TILs. Phospho-PD-1 increased in T cells from healthy human donors after PD-1 engagement and decreased in patients with Hodgkin lymphoma following ICB. These data demonstrate that phosphorylation of the ITSM motif of PD-1 marks dysfunctional T cells that may be rescued with PD-1 blockade. Detection of phospho-PD-1 in TILs is a potential biomarker for PD-1 immunotherapy responses.

Induction of Ascites in Mice

Smaller animals such as rats, mice, hamsters, and guinea pigs are usually poor choices for polyclonal antibody production because only small volumes of serum can be obtained. This problem can be reduced by inducing the formation of ascites in mice, which can provide up to 10 mL of ascites fluid from a single animal. Antibody titers in ascites fluids are almost as high as serum titers.

Inducing, Collecting, and Storing Ascites

Ascitic fluid (also called ascites) is an intraperitoneal fluid extracted from mice that have developed a peritoneal tumor. For antibody production, the tumor is induced by injecting hybridoma cells into the peritoneum, which serves as a growth chamber for the cells. The hybridoma cells grow to high densities and continue to secrete the antibody of interest, thus creating a high-titered solution of antibodies for collection. A single mouse may yield as much as 10 mL of ascitic fluid or as little as 1 mL per batch. Antibody concentrations will typically be between 1 and 10 mg/mL. The most common problem encountered in storing ascites is contamination of these solutions with bacteria or fungi. This can be prevented by the addition of sodium azide.

Growing Hybridomas

This introduction discusses the techniques used to grow and maintain myeloma and hybridoma cell lines, the production and collection of monoclonal antibodies, and methods for drug selection used in hybridoma work.

Making Weak Antigens Strong: Preparing Immune Complexes for Injection

If antibodies against a particular antigen are available, that antigen can be purified and used for further immunizations, and antigens thus purified can show enhanced immunogenicity. Purified immune complexes can be injected directly, or while coupled to beads; the presence of antibodies and/or beads stimulates phagocytosis and usually will not influence the response. This method provides a useful means of antigen enrichment for a variety of applications, such as using antibodies raised against a denatured antigen to harvest a native protein for further immunizations, or when using a monoclonal antibody as an intermediate to the preparation of polyclonal antisera. Injecting antibody-coated antigens has also been used to mask a particularly immunodominant epitope on an antigen, and thereby develop a response against other epitopes. The amount of antigen needed to elicit a strong response using immune complexes will vary from one compound to another. Doses as low as 50 ng of antigen have been used successfully when delivered this way.

Decoy Immunization for Mice, Rats, and Hamsters

Decoy immunization relies on misdirecting the T and B cells of the immune system away from immunogenic regions against which one does not want to generate antibodies, allowing the desired region to be profiled. Essentially, it involves immunizing animals with two forms of the targeted protein on opposite sides of the body. One form, with the region of interest removed (i.e., a modified protein), is injected into the left side of animal, whereas the intact protein of interest is injected into the right side. In theory, undesired B cells are drawn to the left side of the animal, leaving the desired B cells to be drawn to the right (specifically, in the procedure presented here, to the right popliteal lymph node).

Selecting for and Checking Cells with HGPRT Deficiency for Hybridoma Production

For drug-selective media to work for hybridoma selection, myeloma cells expressing a mutation abrogating the function of their HGPRT gene (and subsequently unable to produce purines for DNA biosynthesis) are used. HGPRT will recognize 8-AG as a substrate and convert it to the monophosphate nucleotide. The 8-AG-containing nucleotide is then processed further and incorporated into DNA and RNA, where it is toxic. Therefore, cells with a functional HGPRT enzyme grown in the presence of 8-AG will die. Cells that are deficient in HGPRTase cannot incorporate 8-AG in vivo and thus continue to grow. Cells that have been selected for resistance to 8-AG should be checked periodically to ensure that they maintain sensitivity to drugs that block the de novo synthesis of DNA. In addition, all myeloma cell lines should be checked periodically for reversion of their drug selection markers. Any line that is not killed completely by drug selection should either be reselected or replaced with a new line.

Making Weak Antigens Strong: Coupling Antigens to Red Blood Cells

Coupling antigens to red blood cells (RBCs) can increase the immunogenicity of weak antigens. Their size slows dispersal; that, and their particulate nature, also make them good targets for phagocytosis. If the source of the cells is different from the animal to be injected, they can also provide good targets for MHC class II-T-cell receptor binding. The choice of coupling method will depend on the antigen, but because of the complexity of proteins found on the surface of the RBCs, almost all chemical groups are available for coupling. Commonly used coupling methods include tannic acid, chromic chloride, and glutaraldehyde.

Preparing mFc- and hFc-Fusion Proteins from Mammalian Cells

Fc-fusion proteins are composed of an immunoglobulin Fc domain that is directly linked to the antigen of interest. Typically, these vectors will contain an amino-terminal signal sequence that permits trafficking to the cell surface and secretion into the media and a carboxy-terminal Fc receptor that enables purification on Protein A-Sepharose. Fc-fusion proteins have several applications in protein microarrays, oncological therapies, and vaccine and antibody development. Presence of the Fc domain significantly increases the plasma life of the fusion partner, which prolongs therapeutic activity. Furthermore, the Fc domain enhances the solubility and stability of the partner molecule. Because Fc-fusion proteins are secreted into the culture medium, purification by affinity chromatography is relatively easy and cost-effective. Immunizing a murine host with mFc-fusion protein generates an antigen-specific immune response because the Fc domain is recognized as "self" by the host.

Preparing Live Cells for Immunization

Generating monoclonal antibodies against cell-surface (i.e., membrane) proteins can be challenging because membrane and membrane-associated proteins often lose their native conformation during the purification process. This also makes fusion screening very difficult. One widely used technique to overcome this issue is to overexpress the target protein in HEK 293T cells and then immunize the host with these cells. The advantage of immunizing with native cells is that the target protein is expressed and presented to the immune system in a correctly folded form with all of its secondary posttranslational structure in place. This is essential for conformational or discontinuous epitopes, and for transmembrane proteins that weave in and out of the cell membrane multiple times. Transient or stable transfectants can be used for immunization and for screening using fluorescence-activated cell sorting, western blot, or immunoprecipitation. Although transfectants often have higher expression levels than do native cells, care should be taken to ensure that the transfectant expresses a functionally active version of the target protein, as otherwise minor folding issues or modifications in structure can result in antibodies that recognize the transfected, but not the native, protein. Care must also be taken when using cells as immunogens because numerous antigenic proteins coimmunize with the target protein. Screening hybridomas using the same cells and counterscreening them on untransfected cells will enable the selection of specific hybridomas.

Testing Hybridoma Cells for Mycoplasma Contamination

Because mycoplasmas are a diverse group of organisms and are difficult to culture, several different strategies for detecting mycoplasma contamination have been developed. To date, no one test is suitable for detecting all of the possible mycoplasmas that may contaminate hybridoma or myeloma cultures. Therefore, it is sensible to consider using several methods. The most commonly used techniques are described here. Mycoplasma screening by growth on microbial medium can be performed on agar plates or in broth culture. Cultures are grown under both aerobic and anaerobic conditions because some common strains of Mycoplasma prefer the lack of oxygen (Mycoplasma buccale, Mycoplasma faucium, M. orale, and M. salivarium). Mycoplasma colonies form a characteristic "fried egg" appearance on agar plates, and this is the diagnostic feature used to confirm mycoplasma contamination. The colonies are small and are most easily seen with an inverted microscope. A quicker method for testing for mycoplasma takes advantage of the DNA-intercalating dye Hoechst 33258. Fixed cells are stained with the dye, and contaminated cultures are detected by the bright, punctate cytoplasmic staining of the Mycoplasma DNA. Finally, commercial kits for the detection of mycoplasmas using colorimetric assays or reporter cells are also described.

Ridding Hybridoma Cells of Mycoplasma Contamination

Resistance to the most common antibiotics is what makes elimination of mycoplasma contamination so difficult, but not impossible. Different species of Mycoplasma have varied sensitivities to each of the major classes of antibiotics. One method presented here entails selection in antibiotic-containing medium combined with single-cell cloning over activated macrophage feeders. Its success rate is as high as 70%. As a method of last resort, growing hybridoma cells as ascites tumors is one of the most effective methods of removing mycoplasma contamination. The mycoplasmas are removed from the hybridoma cell surface by the immune system of the mouse. Mice must be of the same genetic background as the hybridomas. This technique is the same as the method for ridding cells of bacterial or fungal infection, and the success rate is perhaps 80%.

Ridding Hybridoma Cell Lines of Contaminating Microorganisms

The long tissue culture manipulations involved in hybridoma production and maintenance sometimes can lead to contamination. If uncontaminated frozen stocks are not available, then it may be possible to rescue these cultures. Three potential methods for rescuing the lines and their success rates are provided here. If the contamination has been detected early, then the use of drugs (antibiotics) to halt the growth of the contaminating organism may succeed. For 96-well plates that have mild to moderate infections, macrophages may help clear infection. If the contamination is gross, passing the cell line through a mouse and recovering the hybridoma cells through subcloning may be the only method that may succeed.

NEDD9 Is a Novel and Modifiable Mediator of Platelet-Endothelial Adhesion in the Pulmonary Circulation

Rationale: Data on the molecular mechanisms that regulate platelet-pulmonary endothelial adhesion under conditions of hypoxia are lacking, but may have important therapeutic implications. Objectives: To identify a hypoxia-sensitive, modifiable mediator of platelet-pulmonary artery endothelial cell adhesion and thrombotic remodeling. Methods: Network medicine was used to profile protein-protein interactions in hypoxia-treated human pulmonary artery endothelial cells. Data from liquid chromatography-mass spectrometry and microscale thermophoresis informed the development of a novel antibody (Ab) to inhibit platelet-endothelial adhesion, which was tested in cells from patients with chronic thromboembolic pulmonary hypertension (CTEPH) and three animal models in vivo. Measurements and Main Results: The protein NEDD9 was identified in the hypoxia thrombosome network in silico. Compared with normoxia, hypoxia (0.2% O2) for 24 hours increased HIF-1α (hypoxia-inducible factor-1α)-dependent NEDD9 upregulation in vitro. Increased NEDD9 was localized to the plasma-membrane surface of cells from control donors and patients with CTEPH. In endarterectomy specimens, NEDD9 colocalized with the platelet surface adhesion molecule P-selectin. Our custom-made anti-NEDD9 Ab targeted the NEDD9-P-selectin interaction and inhibited the adhesion of activated platelets to pulmonary artery endothelial cells from control donors in vitro and from patients with CTEPH ex vivo. Compared with control mice, platelet-pulmonary endothelial aggregates and pulmonary hypertension induced by ADP were decreased in NEDD9-/- mice or wild-type mice treated with the anti-NEDD9 Ab, which also decreased chronic pulmonary thromboembolic remodeling in vivo. Conclusions: The NEDD9-P-selectin protein-protein interaction is a modifiable target with which to inhibit platelet-pulmonary endothelial adhesion and thromboembolic vascular remodeling, with potential therapeutic implications for patients with disorders of increased hypoxia signaling pathways, including CTEPH.

cDNA Immunization of Mice, Rats, and Hamsters

Genetic immunization has been useful in vaccine technology and can also be used to generate immune responses to novel proteins. It is particularly useful when a properly folded protein is difficult to isolate or make in recombinant form and the required antibody must recognize a conformational epitope. Injecting cDNA allows the protein to be expressed in native form by the animal's own cells in vivo and then presented to the immune system. Genetic immunization is most effective for the generation of antibodies if the cDNA encodes for secreted or cell-surface proteins to make them accessible to the immune system. It also has been successful in generating antibody responses to difficult protein targets such as G-coupled protein receptors, ion channel proteins, and other multiple membrane-spanning proteins. High-affinity antibodies tend to be favored because the proteins are expressed at low levels and are constantly present for presentation to the immune system. In addition, several months can be saved obtaining sufficient amounts of properly folded, soluble recombinant protein or high-expressing transfected cells.

Liquid Nitrogen Storage of Hybridoma Cells

Hybridoma and myeloma cell lines can be stored by slowly freezing cells in an appropriate solution of nutrients and a cryoprotectant such as glycerol or dimethyl sulfoxide (DMSO). In this protocol, cells are centrifuged at 4°C, resuspended in cold freezing solution (10% DMSO in FBS), and then transferred to an appropriate freezing vial. The vials are slowly frozen to -70°C in Styrofoam racks and then stored in liquid nitrogen (LN₂). Cells stored in LN₂ will remain viable for years. Once a frozen vial has been removed from LN₂ storage, it should be thawed as described, grown out into log phase, and refrozen.

Collecting and Storing Hybridoma Tissue Culture Supernatants

For most immunochemical methods, tissue culture supernatants will be the most useful source of monoclonal antibodies. The supernatants are not contaminated with high levels of other antibodies, and the concentration is high enough for most assays if used undiluted. This protocol describes the procedure of collecting tissue culture supernatants. When collecting supernatants for antibodies, allow the individual cultures to grow until the hybridomas die. This will allow collection of higher-titer supernatants. In general, antibodies are resistant to the proteases that are released from dying cells, so allowing the cells to die should not affect the quality of the antibodies. If extraneous IgG molecules will alter any of the assays for which the supernatants are being prepared, use medium with fetal bovine serum or use serum-free medium. The yield of this method is ∼20-50 µg of antibody/mL of supernatant. The most common problem encountered in storage of tissue culture supernatants after collection is contamination with bacteria or fungi. This can be prevented by the addition of sodium azide as described.

Sarkosyl Preparation of Antigens from Bacterial Inclusion Bodies

In many cases, solubility and proper folding of fusion proteins expressed in bacteria pose a major challenge in protein purification and crystallization. This is especially true when the fusion proteins are of eukaryotic origin. They form aggregates or become packaged into inclusion bodies, which makes protein purification extremely difficult. Sarkosyl is widely used to extract misfolded proteins from inclusion bodies in soluble form.

Preparing GST-, His-, or MBP-Fusion Proteins from Bacteria

An excellent source of antigens is to overexpress cloned genes in bacteria. A wide variety of coding regions can be expressed in bacteria either on their own or as fusion proteins. Indeed, it is often convenient to use vectors that express the antigen fused to an affinity tag, such as glutathione-S-transferase (GST), maltose-binding protein (MBP), or poly-histidine (e.g., 6×His). Each of these tags reliably binds to a specific affinity column enabling the antigen to be conveniently eluted under appropriate conditions. It should be noted that GST adds ∼25 kDa and MBP adds 40 kDa to the expressed protein. Because bacteria will reliably express proteins that are <80 kDa, the mass of the affinity tag should be included when designing your recombinant protein construct.

Standard Immunization of Rabbits

Rabbits can be immunized by administering biweekly injections of a purified antigen, cultured cells, or cDNA. The minimum amount of antigen capable of inducing a response will depend on the nature of the antigen and on the host, but for rabbits, the minimum dose will be in the range of 10 μg per injection, although 100 μg per injection will be used more commonly. If a pure, soluble protein antigen is being used and is abundant, then a dose of 0.5-1 mg in adjuvant at each immunization is a sensible general recommendation. The injection sites on the rabbit are shaved and disinfected before immunization. Adjuvants are mixed with the immunizing antigen for the first two immunizations only, and Complete Freund's adjuvant is only used with the first immunization; subsequent immunizations are performed in phosphate-buffered saline (PBS) or normal saline, with or without Incomplete Freund's adjuvant. Once a good titer has developed against the antigen of interest, regular boosts and bleeds are performed to collect the maximum amount of serum. For rabbits, boosts should be spaced every 6 wk, and serum samples of 20-40 mL should be collected ∼10-12 d after each boost; typically, a single sample bleed from a rabbit will yield 25 mL of serum.

Repetitive Immunization at Multiple Sites (RIMMS) of Mice, Rats, and Hamsters

The repetitive immunization at multiple sites (RIMMS) protocol capitalizes on the animal's innate immune system, which is genetically preprogrammed to recognize many antigens. By repetitively immunizing the animal, B cells that recognize the antigen are kept continuously expanding until the lymph nodes are harvested for hybridoma generation. This is a good method for making a more diverse repertoire of antibodies or antibodies directed against conformational epitopes.

Standard Immunization of Mice, Rats, and Hamsters

Mice, rats, or hamsters are immunized by giving biweekly injections of a purified antigen, cultured cells, or cDNA. For mice, if a pure, soluble protein antigen is being used and is abundant, a dose of 50-100 µg in adjuvant at each immunization is a sensible general recommendation; for rats and hamsters, a dose of 100-200 µg is sufficient. Lower doses can be used for antigens with higher immunogenicity. Adjuvants (Freund's, Ribi, Hunter's TiterMax, ImmunEasy, or Alum) should be mixed with the immunizing antigen for the first two immunizations only; Complete Freund's adjuvant is only used with the first immunization. Subsequent immunizations are performed in phosphate-buffered saline (PBS) or normal saline, with or without Incomplete Freund's adjuvant. The choice of adjuvant is dependent on the subclass of immunoglobulin required. Over the course of the 6-wk immunization schedule, each animal usually receives a total of six injections (three subcutaneous and three intraperitoneal). Once a good titer has developed against the antigen of interest, regular boosts and bleeds are performed to collect the maximum amount of serum. For rats and hamsters, boosts should be spaced every 2-3 wk, and serum samples of 400-500 µL should be collected 10-12 d after each boost. For mice, boosts should be spaced every 2-3 wk, and serum samples of 200-300 µL should be collected 10-12 d after each boost.

Immunizing Mice and Rats with Nitrocellulose-Bound Antigen

A desired target antigen within a mixture of antigenic proteins such as a cell lysate can be separated from the mixture by gel electrophoresis. However, extracting the antigen from a section of gel and eluting it can be cumbersome and often results in significant loss of protein. Also, the animal might not tolerate the injection of the polyacrylamide gel piece containing the antigen. Alternatively, the protein can be transferred from the gel onto nitrocellulose and then injected either subcutaneously or into the peritoneal cavity. Protein antigens immobilized on nitrocellulose often make exceptionally good immunogens. This is likely because of their slow release from the paper, thus behaving somewhat like an adjuvant. Not all antigens show increased immunogenicity using this methodology, but some do. The antigen is bound to paper and can be implanted on the back of the mouse's neck, a location that makes it difficult for the mouse to disturb the surgical clip, or extracted and injected.

Detecting Protein Antigens in Sodium Dodecyl Sulfate-Polyacrylamide Gels

In some cases, a native protein can be isolated in its pure form from cell lysates or tissue preparation using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Antigens purified this way often induce good antibody responses. After electrophoresis, the band of protein of interest must be located in the gel. A variety of identification methods can be used, all of which are designed to avoid excessive fixation of the protein in the gel matrix. The choice of method depends partly on the abundance of the polypeptide. Three methods are commonly used: (1) staining side strips cut from the edge of the gel, (2) light staining of the gel itself, and (3) locating the band by radioactive labeling of the antigen. Staining strips of the gel cut from its sides avoids the need to fix the gel. When isolating abundant proteins that are well separated from other bands, staining side strips is a useful method. If the protein is not abundant or is located close to a contaminating band making a clean excision difficult, use one of the other staining methods. If the protein is reasonably abundant, then a light staining of the proteins in the gel with Coomassie Blue G will permit localization without fixing. Alternatively, the bands in the gel can be visualized by immersing the gel in sodium acetate or copper chloride. If the protein is radiolabeled with ¹²⁵I, ³²P, or ³⁵S, then use an autoradiogram as a template to excise the band of interest.

Preparing Protein Antigens from Sodium Dodecyl Sulfate-Polyacrylamide Gels for Immunization

Some native proteins can be isolated in pure form from cell lysates or tissue preparation using SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Antigens purified this way often induce good antibody responses. There are many different ways to process a gel fragment containing the protein of interest for injection. Such samples can be processed into small pieces and then injected, either by fragmenting the gel by passing it repeatedly through a syringe, or drying the entire gel slice and grinding it into a powder. Injections using the whole gel fragment should only be used with larger animals such as rabbits. For mice or other small animals, electroelution or electrophoretic transfer of the protein should be used to prepare the protein for injection. In the latter method, the protein is transferred to a suitable membrane, such as nitrocellulose or PVDF. The location of the desired protein is identified by staining, and the protein band is then cut from the surrounding membrane, minced, and dissolved in a small amount of dimethyl sulfoxide before subcutaneous injection into the animal.

Single-Cell Cloning of Hybridoma Cells by Limiting Dilution

Isolating a stable clone of hybridoma cells that all secrete the correct antibody is the most time-consuming step in the production of hybridomas. Single-cell cloning ensures that cells that produce the antibody of interest are truly monoclonal and that the secretion of this antibody can be stably maintained. The original positive well will often contain more than one clone of hybridoma cells, and many hybrid cells have an unstable assortment of chromosomes. Both of these problems may lead to the desired cells being outgrown by cells that are not producing the antibody of interest. Cloning hybridoma cells by limiting dilution is the easiest of the single-cell-cloning techniques. Two approaches are presented here, one rapid technique for generating cultures that are close to being single-cell-cloned and one for single-cell cloning directly. Even though every attempt is made to ensure that the cells are in a single-cell suspension before plating, there is no way to guarantee that the colonies do not arise from two cells that were stuck together. Therefore, limiting dilution cloning should be performed at least twice to generate a clonal population.

Single-Cell Cloning of Hybridoma Cells by Growth in Soft Agar

Single-cell cloning during hybridoma production ensures that cells that produce the antibody of interest are truly monoclonal and that the secretion of this antibody can be stably maintained. Cloning of hybridoma cells in semisolid medium is one of the most commonly used methods for producing single-cell clones. The technique is easy, but, because it is performed in two stages, it does take longer than other methods. Not all cells will grow in soft agar, and there may be a bias on the type of colony that appears. However, most of the commonly used myeloma fusion partners have relatively good cloning efficiencies in soft agar, and, consequently, so do most hybridomas. Even though every attempt is made to ensure that the cells are in a single-cell suspension before plating, there is no way to guarantee that the colonies do not arise from two cells that were stuck together. Therefore, single-cell cloning in soft agar should be repeated at least twice before the cells are considered clonal.

Preparing Feeder Cell Cultures to Support Hybridoma Growth

Feeder layer plates are prepared with cells that provide the appropriate growth factors to support the growth of hybridoma cells until they can expand in number and provide their own. Good feeder cells should have properties that allow them to be selected against during the future growth of the hybridomas. Peritoneal macrophages, myeloma cells, splenocytes, and MRC-5 cells (a human lung fibroblast line) are the most common feeder layer cells used in hybridoma fusions. Methods for their preparation are described here.

Myeloma and Hybridoma Cell Counts and Viability Checks

For most purposes, the number of hybridoma or myeloma cells can be estimated simply by observing the cells under the microscope. When an exact cell count is needed, the number can be determined by using a hemocytometer (improved Neubauer counting chambers are the most commonly used). This is a simple device in which a special coverslip rests on supports that hold it 0.1 mm above the base of the slide. The slide is engraved with a series of lines that form 1 × 1-mm squares. By counting the number of cells within the 0.1-mm³ chamber formed by the 1 × 1-mm square and the height of the coverslip, an accurate quantitation of cells per milliliter can be calculated. To determine the percentage of viable cells within a population, the cell suspension is mixed with a vital dye and observed under the microscope. Vital dyes are excluded from living cells but stain dead cells. The most common dye used for these stains is Trypan Blue.

Electro Cell Fusion for Hybridoma Production

Once a good immune response has developed in an animal and an appropriate screening procedure has been developed, the construction of hybridomas is ready to begin. The electro cell fusion (electrofusion) method uses an electrical field in the form of short, intense pulses to increase the permeability of the membrane. The resulting local perforation of the cell membrane induces the cells to fuse, forming hybridomas. Electrofusion is accomplished in three steps: Prealignment of the cells (convergence and cell contact), membrane fusion, and postalignment (rounding off the fused cells). This method has been applied successfully to hybridoma production with higher efficiency than routine polyethylene glycol fusion, allowing production of more hybrid cells.

Routes of Antigen Injection in Mice and Rats

The route of injection used to induce an antigenic response in small rodents is based on a number of practical considerations, such as the volume to be delivered, the buffers and other components co-injected with the immunogen, and the rate at which one desires the immunogen to be released into the lymphatics or circulation. For mice, large volumes are only practical with intraperitoneal injections. If adjuvants or particulate matter are included in the injection, the immunogen should not be delivered intravenously.

Administering Anesthesia to Mice, Rats, and Hamsters

Anesthesia is used to sedate animals during injections or during operations where unexpected or rapid movements would endanger the animal. Inhalant anesthetics are often the most appropriate choice to achieve anesthesia that meets the clinical and humane needs without interfering with the scientific goals of the research protocol. Isoflurane (i.e., halogenated ether)-the preferred inhalant anesthetic for small animals-is administered either by placing the animals in a closed container permeated with a mixture of isoflurane and oxygen vapor, or by using a mask (for larger animals). The depth of anesthesia is easy to assess and to control. Animals sedated by isoflurane can be handled for injections or simple operations.

Transfected Dendritic Cell Immunizations

Dendritic cells (DCs) are the most potent antigen-presenting cells (APCs). An efficient way of generating antibodies is to introduce antigens of interest into DCs and then inject them into the host. This will result in initiation of an antigen-specific immune response mediated by T-cell immunity. Apoptosis of DCs expressing transgenic proteins results in enhanced immunity through cross-presentation by endogenous DCs in vivo. The major advantage of this technology is prolonged presentation of antigens that are synthesized endogenously and their presentation by both modified DCs as well as endogenous DCs to the immune system of the host.