Production of Monoclonal Antibodies 1
Wayne M. Yokoyama, Michelle Christensen, Gary Dos Santos, Diane Miller,2 Jason Ho,3 Tao Wu,3 Michael Dziegelewski,3 and Francisca A. Neethling3 1
Washington University School of Medicine, St. Louis, Missouri StemCell Technologies, Inc., Vancouver, British Columbia, Canada 3 Boehringer Ingelheim Pharmaceuticals, Inc., Ridgefield, Connecticut 2
ABSTRACT This unit describes the production of monoclonal antibodies beginning with immunization, cell fusion, and selection. Support protocols are provided for screening primary hybridoma supernatants for antibodies of desired specificity, establishment of stable hybridoma lines, cloning of these B cell lines by limiting dilution to obtain monoclonal lines, and preparation of cloning/expansion medium. An alternate protocol describes cell fusion and one-step selection and cloning of hybridomas utilizing a semi-solid methylcellulosebased medium (ClonaCell-HY from StemCell Technologies). Curr. Protoc. Immunol. C 2013 by John Wiley & Sons, Inc. 102:2.5.1-2.5.29. Keywords: monoclonal antibodies r B cell hybridomas r limiting dilution r semi-solid methylcellulose-based medium
INTRODUCTION Highly specific antibodies can be obtained by fusing immune B cells from the spleen with tumor cells to produce hybridomas, each of which will then secrete a single antibody. The desired antibody-producing hybridoma can be identified by a screening process. If this hybridoma is subjected to a cloning process in which clones are selected, such that all progeny are derived from a single cloned parental cell, a monoclonal antibody is obtained. Monoclonal antibodies have high specificity and can be produced in large quantities. Thus, these biological reagents have been used extensively as probes in a wide range of applications including the characterization of novel cell-surface and soluble proteins and carbohydrates, as enzyme catalysts, and for targeting in immunotherapy (see Commentary). This unit describes the production of monoclonal antibodies beginning with protocols for immunization (Basic Protocol 1) and cell fusion and selection (Basic Protocol 2). Alternate Protocol 1 describes an alternative immunization procedure eliminating complete Freund’s adjuvant, which is objectionable because of its extremely inflammatory characteristics. Another alternate protocol (Alternate Protocol 2) describes an alternative method of cell fusion and one-step selection and cloning of hybridomas utilizing a semisolid methylcellulose-based medium (ClonaCell-HY, a registered trademark of StemCell Technologies, Inc.; http://www.stemcell.com); yet another method of cell fusion and selection (electrofusion) is described in Alternate Protocol 3. Methods are provided for screening primary hybridoma supernatants for antibodies of desired specificity (Support Protocol 1), establishment of stable hybridoma lines (Support Protocol 2), cloning of these B cell lines by limiting dilution to obtain monoclonal lines (Support Protocol 3), recloning of hybridoma cells in semisolid medium (Support Protocol 4), and preparation of cloning/expansion medium (thymocyte-conditioned medium; Support Protocol 5). Figure 2.5.1 summarizes these stages and notes the protocols in this and subsequent units in which they are detailed. Selection and cloning of hybridomas
Current Protocols in Immunology 2.5.1-2.5.29, August 2013 Published online October 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/0471142735.im0205s102 C 2013 John Wiley & Sons, Inc. Copyright
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Basic Protocol 1 or Alternate Protocol 1 prepare antigen
immunize appropriate host prepare myeloma cell fusion partner by expansion harvest and prepare spleen cells
prepare myeloma cell fusion partner by expansion
do a booster immunization
harvest and prepare spleen cells
mix fuse and plate
Basic Protocol 2
plate in cloning/ selection medium
feed wean off HAT
Alternate Protocol 2 or 3
harvest clones in 96-well plate
Support Protocol 1
expand and freeze desired hybridomas
Support Protocol 2
expand and freeze desired hybridomas
clone by limiting dilution screen clones
reclone, if necessary
Support Protocol 4
expand and freeze positive clones Support Protocol 3 reclone by limiting dilution screen clones prepare purified MAb expand and freeze positive recloned lines obtain a stable MAb-producing hybridoma
UNITS 2.7 & 2.8 produce supernatant or ascites fluid UNIT 2.6
Figure 2.5.1 Stages of monoclonal antibody production, with references to the basic, alternate, and support protocols in this unit (as well as subsequent units) that describe the steps. Production of Monoclonal Antibodies
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using the techniques described in Basic Protocol 2 require a major commitment of time and labor. However, Alternate Protocol 2 or 3 reduces the overall time necessary to produce monoclonal antibodies by 18 to 20 days. When successful, the monoclonal antibody may be an extremely valuable reagent that will be available in large quantities. Submission of monoclonal antibodies to the American Type Culture Collection (ATCC) for distribution to the scientific community is encouraged. Moreover, the ATCC serves as a repository for cell lines should the line be lost in the investigator’s laboratory due to unforeseen circumstances. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations for the care and use of laboratory animals. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.
IMMUNIZATION USING COMPLETE FREUND’S ADJUVANT TO PRODUCE MONOCLONAL ANTIBODIES
BASIC PROTOCOL 1
A wide variety of antigen preparations have been used successfully to produce monoclonal antibodies (see Critical Parameters for discussion of antigen preparation). The following protocol provides an immunization schedule for the production of most antibodies, although several different schedules can be used. In this protocol, emulsified antigen is injected intraperitoneally into the species of choice. A booster injection is administered 10 to 14 days after the primary immunization. Three days after the booster injection, the animals’ spleens are ready for cell fusion (Basic Protocol 2).
Materials Antigen Complete Freund’s adjuvant (CFA; Sigma) Animal: pathogen-free mouse, hamster, or rat (Armenian hamsters from Cytogen Research are recommended; see Critical Parameters for discussion of animal choice and UNIT 1.1) Incomplete Freund’s adjuvant (IFA; Sigma), optional 1 to 2-ml glass syringes with Luer-Lok tips, sterile 3-way stopcock 20- and 22-G needles, sterile Additional reagents and equipment for handling and restraint of animals (UNIT 1.3) and injection of rodents (UNIT 1.6) CAUTION: CFA is an extremely potent inflammatory agent, particularly if introduced intradermally or into the eyes. Profound sloughing of skin or loss of sight may occur. Self-injection can cause a positive TB skin test and lead to a granulomatous reaction. Use gloves and protective eyewear when handling CFA. 1. Prepare antigen using 2 × 106 to 5 × 107 cells or 1 to 50 μg protein or peptide per animal to be immunized in normal saline. Calculate antigen requirement at approximately 350 μg/mouse for the full series of immunizations.
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The antigen may be in several different forms depending on the desired property of the MAb and the method of screening (see Critical Parameters for discussion of antigen preparation and screening assays). If cells are the immunogen, wash three times in serumfree medium before immunization. Plan the immunization of several animals (enough for several fusions) so that primed and boosted animals will be ready 3 days before fusion (see Basic Protocol 2). To minimize the risk of introducing a pathogen into the rodent colony, screen cells for pathogens by antibody-production assay (UNIT 1.1).
2. Draw up antigen into a sterile 1- to 2-ml glass syringe with a Luer-Lok tip. Connect syringe to a 3-way stopcock. 3. Completely resuspend CFA to disperse the Mycobacterium tuberculosis bacilli, which settle to the bottom of the container with time. Draw up a volume of CFA equal to the antigen volume in a syringe and connect to the antigen-containing syringe. 4. Emulsify antigen and CFA by discharging antigen into CFA, then discharging back and forth until a thickened mixture results. Test whether the emulsion is stable—a stable emulsion will not disperse when a drop of it is placed in water. See UNIT 2.4 for further discussion of immunization. Figure 2.4.1 illustrates the doublesyringe device.
5. Transfer all of the CFA/antigen emulsion to one syringe and remove the other syringe and stopcock. Attach a sterile 20-G needle to the syringe containing the emulsion. 6. Inject emulsion intraperitoneally (UNIT 1.6) into the animal using 1 mg/ml, ideally 2 mg/ml Sigma Adjuvant System (Sigma S6322), reconstituted per manufacturer’s instructions Animal(s): Mice of preferred strain (e.g., Balb/C, C57Bl/6, CD1), three to five mice per cohort 1- or 2-ml syringes with 26-G needles Sterile microcentrifuge tube of adequate size to mix antigen and adjuvant 1:1 Additional reagents and equipment for ELISA (UNIT 2.1) 1. Working in a biosafety cabinet using sterile technique, thoroughly but gently mix 50 μg/mouse of antigen 1:1 (v/v) with adjuvant from the Sigma Adjuvant System. Prepare each set of immunizations fresh just before use.
2. Draw the mixture up into syringe, ensuring that all air is dispelled before administering to the mouse. 3. Administer the first immunization subcutaneously (UNIT neck.
on the back of the
4. At 14-day intervals, boost intraperitoneally (as described in Basic Protocol 1) with two further 50-μg doses of antigen diluted 1:1 with adjuvant as described above. 5. At a time point 3 to 7 days after the third immunization, assess serum antibody titers by a suitable assay such as ELISA (UNIT 2.1). 6. If the titer is sufficient, perform a final boost intraperitoneally at same dose mixed in adjuvant, 14 days after the third immunization. Recover the spleen 3 to 5 days after the final boost and proceed to cell fusion (Basic Protocol 2 or Alternate Protocol 2 or 3). If the titer is not high enough, perform 3 additional IP immunizations at 14-day intervals, then perform a serum titer determination again. If the titer is now sufficient, administer IP final boost 14 days after the last immunization. Recover the spleen 3 to 5 days later and proceed to fusion. Serum immune response to the antigen may vary in a cohort of mice, inbred and outbred. Conduct as thorough an evaluation of the serum as possible and consider performing additional tests if available. If, after 2 series of immunizations, there is still insufficient antibody titer, the immunization approach may have to be reevaluated.
CELL FUSION AND SELECTION OF HYBRIDOMAS While animals should be immunized as soon as the decision has been made to produce a monoclonal antibody and the antigen prepared, do not perform cell fusion until the screening assay (Support Protocol 1) has been perfected. Artifactual results that may arise from conditioned medium must be identified before cell fusion, because after a fusion there is only a finite amount of time available to assay for the desired monoclonal antibody. In many cases, this screening assay can also be used to determine specific polyclonal antibody titer, indicating which animal should be used for spleen harvest and fusion. Prior to cell fusion, the partner (myeloma) cell line is expanded and a booster injection of antigen is administered to the primed animals. On the day of fusion, the spleens are harvested. Spleen cells and partner cells are washed, harvested, and mixed. Cell fusion
BASIC PROTOCOL 2
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is performed at 37◦ C aided by the presence of polyethylene glycol (PEG) or by transient pulsing in an electric field (a process known as electrofusion; see Alternate Protocol 3). The resulting pellet is harvested and plated into tissue culture plates. After incubation with hypoxanthine, aminopterin, and thymidine (HAT) medium and feeding over ∼2 weeks, the hybridomas are ready for screening (Support Protocol 1).
Materials SP2/0-Ag14 myeloma cell line (drug-marked, nonsecretory; ATCC #CRL 1581) Complete DMEM-10 and -20 media (APPENDIX 2A) with 10 mM HEPES and 1 mM sodium pyruvate Primed animal; mouse, hamster, or rat (10 to 14 days after primary immunization; (see Basic Protocol 1) Complete DMEM medium (APPENDIX 2A), serum-free 50% polyethylene glycol (PEG), sterile Ammonium chloride solution (see recipe) Complete DMEM-20/HEPES/pyruvate/HAT (or HT) medium (see recipe) 175-cm2 flasks Fine-mesh metal screen or 70 μm synthetic screen 50-ml conical polypropylene centrifuge tubes Beckman TH-4 rotor or equivalent 96-well flat-bottom microtiter plates Additional reagents and equipment for animal euthanasia (UNIT 1.8), spleen removal (UNIT 1.10), and counting cells and assessing cell viability by trypan blue exclusion (APPENDIX 3B) Prepare myeloma cells (1 week before fusion) 1. One week before fusion, begin expansion of SP2/0-Ag14 myeloma cell line (the fusion partner cell line) in complete MEM-10/HEPES/pyruvate (see Critical Parameters). By the day cell fusion is to be performed, the following total number of myeloma cells must be available (in multiple 175-cm2 flasks containing 100 ml each), depending upon the source of the primed animal: mouse spleen, 1 × 108 cells in two or three flasks; hamster spleen, 2 × 108 cells in three or four flasks; and rat spleen, 5-10 × 108 cells in ten flasks. Two mouse or hamster spleens, or one rat spleen, will provide enough cells for the fusion (see step 7).
Boost primed animal (3 days before fusion) 2. Three days before fusion, boost primed animal(s) according to step 7 of Basic Protocol 1. Prepare reagents and split myeloma cells (1 day before fusion) 3. One day before fusion, prepare all reagents and media, particularly 50% PEG. 4. One day before fusion, split SP2/0-Ag14 myeloma cells (from step 1) into fresh complete DMEM-10/HEPES/pyruvate medium. Vigorous growth of the SP2/0-Ag14 cells is generally required for good fusion.
Check myeloma cells and prewarm reagents (day of fusion) 5. Use an inverted microscope to check the SP2/0-Ag14 myeloma cells to make sure they are growing vigorously (refractile and not pyknotic), they are not contaminated (no obvious bacteria or fungi), and there are enough cells for the fusion. Production of Monoclonal Antibodies
It is better to postpone the fusion than to perform an ill-advised fusion, since the entire selection and screening effort will take ∼3 weeks.
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6. Prewarm the following in a 37◦ C water bath:
Three 400- and three 600-ml beakers, each containing ∼100 ml H2 O 20 ml sterile complete serum-free DMEM 5 ml sterile 50% PEG solution. Harvest spleen and prepare cells 7. Sacrifice boosted animal(s) (UNIT 1.8) and aseptically harvest spleen(s) (UNIT 1.10). Do not use anesthetics for sacrifice. Instead, use cervical dislocation for mouse, or CO2 asphyxiation for mouse, hamster, or rat to avoid introducing an anesthetic into the bloodstream and therefore into the cultures.
8. Transfer spleen to a sterile 100-mm-diameter petri dish filled with 10 ml sterile complete serum-free DMEM. Perform all subsequent steps in a laminar flow hood.
9. Tease spleen into a single-cell suspension by squeezing with angled forceps or by chopping with fine-tipped dissecting scissors. Remove debris and disperse cells further by passage through a fine-mesh metal screen. 10. Transfer spleen cell suspension to a sterile 50-ml conical centrifuge tube and fill with sterile complete serum-free DMEM. Do not use protein- or HEPES-containing medium because the PEG will precipitate proteins and HEPES can be toxic to cells during fusion.
11. Centrifuge 5 min in TH-4 rotor at 1500 rpm (500 × g), room temperature, and discard supernatant. 12. Lyse red blood cells (RBC) by resuspending pellet in 5 ml ammonium chloride solution. Let stand 5 min at room temperature. 13. Add 45 ml sterile complete serum-free DMEM, and centrifuge as in step 11. 14. Resuspend pellet in 50 ml sterile complete serum-free DMEM. Centrifuge as in step 11. Repeat DMEM addition and centrifuging once (each repeat is a wash). 15. While spleen cells are being washed, separately harvest the SP2/0-Ag14 myeloma cells (from step 5) by transferring the cells to 50-ml conical centrifuge tubes. Centrifuge as in step 11. Resuspend myeloma cells in DMEM and pool all cells into one 50-ml conical centrifuge tube. Wash myeloma cells three times as in step 14. 16. Separately resuspend the spleen and myeloma cells in 10 ml complete serum-free DMEM. Count cells and assess viability in each cell suspension using a hemacytometer and trypan blue exclusion (APPENDIX 3B); there should be nearly 100% viability in both suspensions. 17. On basis of cell counts (from step 16), calculate the amount of complete DMEM20/HEPES/pyruvate needed to plate cells at ∼2.5 × 106 total cells/ml. Prewarm this amount of complete DMEM-20/HEPES/pyruvate in 37◦ C water bath. Prepare 96well flat-bottom plates by labeling them sequentially: one plate is required for each 10 ml of final cell suspension.
Perform cell fusion 18. Mix SP2/0-Ag14 myeloma and spleen cells at a 1:1 ratio in a 50-ml conical centrifuge tube. Fill the tube with complete serum-free DMEM. Other cell ratios work. Successful fusions have been performed with a ratio of myeloma/spleen cells as low as 1:20.
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19. Centrifuge cell mixture 5 min at 500 × g, room temperature. 20. While cells are in the centrifuge, prepare three 37◦ C double-beaker water baths in the laminar flow hood by placing a 400-ml beaker (from step 6) containing 100 ml of 37◦ C water into 600-ml beaker containing 75 to 100 ml of 37◦ C water. Place the tubes of prewarmed 50% PEG solution and complete serum-free DMEM (from step 6) into two of the 37◦ C water baths in the hood. 21. Aspirate and discard supernatant from the mixed-cell pellet (from step 19). 22. Perform the cell fusion at 37◦ C by placing the tube containing the mixed-cell pellet in one of the double-beaker water baths in the laminar flow hood. 23. Using a 1-ml pipet, add 1 ml prewarmed 50% PEG to the mixed-cell pellet dropby-drop over 1 min, stirring the cells with the pipet tip after each drop. Stir for an additional minute. 24. Using a clean pipet, add 1 ml prewarmed complete serum-free DMEM to the cell mixture drop-by-drop over 1 min, stirring after each drop. Repeat once with an additional 1 ml of prewarmed complete serum-free DMEM. 25. With a 10-ml pipet, add 7 ml prewarmed complete serum-free DMEM drop-by-drop over 2 to 3 min. Macroscopic clumps of cells should be obvious at this point.
26. Centrifuge 5 min at 500 × g, room temperature. 27. While the cells are in the centrifuge, rewarm the beaker water baths to 37◦ C and place in the hood. Place prewarmed complete DMEM-20/HEPES/pyruvate (from step 17) in the beaker water bath. 28. Discard the supernatant (from step 26). Place tube in the beaker water bath. 29. With a clean 10-ml pipet, forcefully discharge 10 ml prewarmed complete DMEM20/HEPES/pyruvate to the cell pellet. 30. Repeat step 29 until the total volume of prewarmed complete DMEM-20/HEPES (calculated in step 17) is added. If necessary, allow clumps to settle and disrupt with the pipet tip. Further warming of cell suspension is no longer required. If the total volume exceeds 50 ml, gently aspirate and transfer to another sterile container such as a tissue culture flask.
31. Gently aspirate 10 ml of cell suspension with a 10-ml pipet. Add 2 drops (100 to 125 μl) of suspension to each well of a 96-well flat-bottom plate (continue until entire suspension is plated). Incubate overnight in a humidified 37◦ C, 5% CO2 incubator. Vigorous pipetting of the cell suspension should be avoided at this point, as the newly formed hybrids are unstable. Moreover, the vigorous addition of cells to the wells with repeating micropipettor is not advised. Use a pipet aid and hold the 10-ml pipet at a 45◦ angle with the tip 1 to 2 cm above the well, bracing the pipet with a finger from the opposite hand. To avoid introducing contaminants, do not hold hands above the plate. A steady, even flow of drops from the pipet will allow the most efficient delivery of cell suspension or medium to the wells. Use a fresh pipet to withdraw additional cell suspension. As an optional step to minimize fibroblast overgrowth, permit the fibroblasts in bulk-fused cell suspension to adhere overnight to tissue culture flasks before seeding the 96-well plates. Production of Monoclonal Antibodies
Many investigators select their hybridomas under bulk conditions—i.e., they incubate large numbers of cells per well in larger plates or flasks. This makes feeding easier, but allows fast-growing hybridomas to overgrow the others. Since nonproducing hybridomas
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tend to grow faster, especially in hamster-mouse fusions, hybridomas are isolated initially in multiple small wells in this protocol. The primary hybridomas tend to be monoclonal. This is especially important when screening procedures are used that require differential reactivities, e.g., to different cell lines by flow cytometry analysis or to different antigen preparations. In those cases, multiple hybridomas per well will obscure the reactivity of the MAb of interest.
Monitor and feed cells 32. After one day of incubation, check wells under an inverted microscope. If seeded with the appropriate number of cells, there should be a nearly confluent monolayer of highly viable cells on the bottom and obvious clumps of cells.
33. Add 2 drops complete DMEM-20/HEPES/pyruvate/HAT to each well with a 10-ml pipet (see step 31). Place in humidified 37◦ C, 5% CO2 incubator. Use a separate pipet for each microtiter plate and keep the same covers with each plate to ensure that each plate remains a separate unit and to avoid spreading contamination. It cannot be overemphasized that it takes practice and meticulous attention to possible sources of contaminants to keep these plates sterile during the subsequent 2-to 3-week feeding and monitoring schedule. If plates become contaminated, discarding them is advised. Alternatively, contamination in one or two wells may be treated by aspirating the contents of the contaminated well with a sterile Pasteur pipet attached to a vacuum flask, rinsing the well with 70% ethanol, and wiping with a sterile cotton swab. Wash twice with ethanol. Finally, blot the well dry with the sterile cotton swab and blot the appropriate area of the cover with a sterile cotton swab soaked in 70% ethanol. Do not open contaminated plates while other plates are in the hood.
34. On days 2, 3, 4, 5, 7, 9, and 11, aspirate half the volume of each well using a sterile, short Pasteur pipet attached to a vacuum flask, holding pipet at a 45◦ angle and touching tip to surface of supernatant at the point where the liquid meets the opposite wall of the well. Feed the cells by adding 2 drops complete DMEM20/HEPES/pyruvate/HAT from a 10-ml pipet (see steps 31 and 33) to each well, and return to humidified 37◦ C, 5% CO2 incubator. Use a separate Pasteur pipet for each plate to minimize spreading contamination. Since the frequency of successful viable hybridoma formation is ≤10−5 , when HAT is added, profound cell death should be apparent at days 2 and 3 and the remaining viable cells should not be readily apparent until they have expanded. By day 7 to 9 for mouse-mouse fusions, day 11 for rat-mouse fusions, and day 14 for hamster-mouse fusions, clusters of hybridoma cells should become visible under the inverted microscope. If profound cell death is not apparent on days 2 and 3, check the medium containing HAT and the parental cell line by incubating an aliquot of the parental myeloma line with the medium containing HAT. The feeding schedule is not rigid except for the first 4 days, when it is necessary to remove the toxic products of cell death. Thereafter, feedings will depend on the actual number of cells deposited in the wells, efficiency of fusion, and appearance and growth of hybridomas. Do not allow wells to become yellow (acidic) for more than a day. Examine plates daily, even if cells are not scheduled to be fed, and feed plates if acidic wells are noted.
35. On day 14, repeat feeding as outlined in step 34 except use complete DMEM20/HEPES/pyruvate/HT to feed cells. Return to 37◦ C, 5% CO2 incubator. Cells do not require more than one change of complete DMEM-20/HEPES/pyruvate/HT. After this change, the aminopterin (from prior addition of HAT medium) is apparently diluted out enough so that the cells can survive without additional HT.
36. On day 15 and subsequently, feed wells as noted using complete DMEM20/HEPES/pyruvate without HAT or HT.
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The hybridomas are ready for screening when most of the wells containing growing cells demonstrate 10% to 25% confluence and when those with denser populations turn yellow within 2 days after feeding (see Support Protocol 1). If the screening assay requires a [3 H]thymidine incorporation assay (APPENDIX 3D), be aware that the large amount of thymidine in complete DMEM-20/HEPES/pyruvate with HAT and HT will serve as a cold-label inhibitor of [3 H]thymidine incorporation. At least three to four changes of complete DMEM-20/HEPES/pyruvate without HT are required to dilute out excess thymidine. ALTERNATE PROTOCOL 2
CELL FUSION, SELECTION, AND CLONING OF HYBRIDOMAS USING A SEMISOLID MEDIUM (ClonaCell-HY) Traditionally, monoclonal antibody development has involved selecting hybridomas in suspension cultures, identifying cultures that produce antibodies specific for the targeted antigen, and cloning the specific antibody-producing hybridoma(s) by at least one round of culture under limiting-dilution conditions (as described in Basic Protocol 2 and Support Protocols 1 to 3). This approach is laborious and time consuming, and may result in the selection of identical, duplicate clones. This section describes protocols for the use of a methylcellulose-based medium system, ClonaCell-HY, for cloning and selection of mouse hybridomas. Performing hybridoma selection and cloning simultaneously in ClonaCell-HY reduces the time and reagents necessary to obtain a monoclonal hybridoma–producing antibody against the antigen of interest. Cultures do not need feeding or maintenance during the selection process. This approach also allows all daughter cells to remain together during the selection process, decreasing the number of clones that need to be tested for antibody production. Selection of duplicate hybridomas, a common occurrence with hybridoma cloning in suspension cultures, is avoided. An additional advantage is that smaller, slow-growing clones, which can easily be lost due to overgrowth by larger, faster-growing hybridomas in traditional liquid suspension culture, remain physically separated in semisolid medium from the larger, faster-growing clones, and can thus be isolated and screened separately. NOTE: All solutions and media should be prewarmed to 37◦ C unless otherwise indicated.
Materials Myeloma cell line (e.g., SP2/0, X63Ag8.653; available from ATCC) ClonaCell-HY Monoclonal Antibody Production Kit (StemCell Technologies, Inc.) containing: Medium A—ClonaCell-HY Pre-Fusion Medium and Hybridoma Expansion Medium, 500 ml Medium B—ClonaCell-HY Fusion Medium, 500 ml Medium C—ClonaCell-HY Hybridoma Recovery Medium, 100 ml Medium D—ClonaCell-HY Hybridoma Selection Medium containing HAT, 90 ml Medium E—ClonaCell-HY Hybridoma Growth Medium containing HT, 500 ml Polyethylene glycol—ClonaCell-HY PEG Solution, pretested for cell fusion, 1.5 ml Immunized mouse, 1 to 4 days after final antigen boost (Basic Protocol 1) 3% (v/v) acetic acid Liquid nitrogen (optional) Fetal bovine serum (FBS) containing 20% (v/v) DMSO Production of Monoclonal Antibodies
15- and 50-ml conical polypropylene centrifuge tubes 100-mm petri dishes Fine-mesh metal screen
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Low-speed tabletop centrifuge 3-ml and 12-ml syringes 25- and 75-cm2 tissue culture flasks 16-G blunt-ended hypodermic needles 96- and 24-well tissue culture plates Cryotubes (e.g., Nunc) Liquid nitrogen freezer (Dewar flask and canes to accommodate cryotubes; optional) Additional reagents and equipment for determining cell viability by trypan blue exclusion (APPENDIX 3B), animal euthanasia (UNIT 1.8), spleen removal (UNIT 1.10), preparing a single-cell suspension of splenocytes (UNIT 3.1), counting cells using a hemacytometer (APPENDIX 3A), assaying for antigen production from hybridoma clones by ELISA (UNIT 2.1), flow cytometry (Chapter 5), or immunoblotting (UNIT 8.10), and cryopreservation of cells (APPENDIX 3G) Prepare myeloma cells 1. Culture the parental myeloma cells in 25-cm2 tissue culture flasks with Medium A (Pre-Fusion Medium from ClonaCell-HY kit) for at least 1 week prior to fusion to ensure they are well adapted to this medium. Seed cells at a density of ∼5 × 104 cells/ml and passage every 2 days. Suggested maximum cell density is 4 × 105 cells/ml. The parental myeloma cells must not secrete any of their own immunoglobulin chains. They should be mycoplasma free and efficiently fuse to form stable hybridomas that continuously secrete specific monoclonal antibodies. Parental myeloma cells that meet these criteria (such as SP2/0 and X63Ag8.653) are widely available.
2. Calculate the cell growth rate at every passage (APPENDIX 3A). The day before the fusion, count the viable cells and split cells so that there will be at least 2 × 107 parental myeloma cells available the next day. The recommended cell density for fusion is 2 × 105 cells/ml. Only 100 ml of these cells will be needed, but 200 ml should be cultured to ensure an adequate supply.
3. Harvest the required number of parental myeloma cells in a 50-ml conical polypropylene centrifuge tube. Centrifuge 10 min at 300 × g, room temperature or 37◦ C, and remove the supernatant. Wash three times, each time by adding 30 ml of Medium B (Fusion Medium), centrifuging again as before, and removing the supernatant. Resuspend final pellet in 25 ml Medium B. This step may be performed simultaneously with or subsequent to spleen cell preparation (steps 6 to 10) to ensure that the myeloma cells do not sit for an extended period of time. It is important to remove all the serum adhering to the cells by washing with serum-free Medium B. If the serum is not removed, the PEG will not fuse the cell membranes and the fusion frequency will drop drastically.
4. Count live cells using a viability stain (APPENDIX 3B). Viability of parental myeloma cells should be >95%.
5. Calculate the volume of cell suspension that contains 2 × 107 cells, to be used in step 11. Place cells at room temperature or 37◦ C.
Harvest spleen and prepare spleen cells 6. Sacrifice immunized animal(s) (UNIT 1.8) and aseptically remove spleen(s) (UNITS 1.8 & 1.10). Place spleen in a sterile 100-mm petri dish containing 5 ml Medium A (Pre-Fusion Medium).
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IMPORTANT NOTE: The ClonaCell-HY kit has been optimized for use with mouse hybridomas. Hybridomas from other species have not been tested. Do not use anesthetics for sacrifice. Instead, use cervical dislocation or CO2 asphyxiation to avoid introducing anesthetic into the bloodstream and therefore into the cultures. It is important to collect blood from the animal to obtain serum as a source of antibodies for a positive control in subsequent screening assays. Blood can be collected from a tail bleed (UNIT 1.7) after the first intraperitoneal boost (see Basic Protocol 1, step 7) or from the heart at the time of spleen harvest. To collect blood from heart, use a sterile Pasteur pipet and place blood into a 1.5-ml microcentrifuge tube. Let blood stand at room temperature with the cap off to allow blood to clot. After 30 min, remove the blood clot with a sterile Pasteur pipet and place tube at 4◦ C. The next day, centrifuge tube for 15 min at 400 × g, 4◦ C or room temperature. Collect serum and add sodium azide to a final concentration of 0.1%. Store at −20◦ C
7. Disaggregate the spleen into a single-cell suspension (UNIT 3.1). Transfer the spleen to a fine-mesh metal screen placed on top of a 50-ml conical centrifuge tube, and use the plunger of a 3-ml syringe to gently grind the cells out of the spleen. Rinse the screen with Medium B to help cells pass through the screen. Pipet the cells up and down in the tube with a 10-ml pipet to break up lumps. Try not to cause the solution to foam. Only the spleen membrane should be left in the screen. See UNIT 3.1 for additional discussion of the above procedure. Other spleen disaggregation methods may also be used.
8. Centrifuge cell suspension 10 min at 400 × g, room temperature or 37◦ C, and remove supernatant. Wash the splenocytes three times, each time by adding 30 ml Medium B, centrifuging 10 min at 400 × g, room temperature or 37◦ C, and discarding the supernatant. Resuspend the final cell pellet in 25 ml Medium B. It is important to remove all the serum adhering to the cells by washing with serum-free Medium B. If the serum is not removed, PEG will not fuse the cell membranes and the fusion frequency will drop drastically.
9. Prepare a 1/10 dilution of cells in 3% acetic acid, e.g., by mixing 10 μl of cell suspension with 90 μl of 3% acetic acid. Count cells in this diluted sample using a hemacytometer (see APPENDIX 3A). 10. Calculate the volume of cell suspension that contains 1 × 108 cells, to be used in step 11. Place cells at room temperature or 37◦ C until fusion.
Combine myeloma cells and splenocytes for fusion 11. Add 2 × 107 parental myeloma cells and 1 × 108 viable splenocytes (as calculated in steps 5 and 10, respectively) to a 50-ml conical centrifuge tube and centrifuge 10 min at 400 × g, room temperature or 37◦ C. Aspirate supernatant. Complete removal of the supernatant is essential to avoid dilution of PEG in the next step.
Perform cell fusion by one of the two following methods
Production of Monoclonal Antibodies
Method A 12a. Break up the cell pellet obtained in step 11 gently by tapping the bottom of the tube. Add 0.5 ml of polyethylene glycol (PEG) solution from the ClonaCell-HY kit dropwise to the pellet using a 1-ml pipet. Centrifuge the mixture 3 min at 133 × g, 37◦ C or room temperature (37◦ C is preferable for this centrifugation but not essential), then aspirate off all of the PEG (during this procedure, not all cells will pellet; some will clump in the PEG; do not aspirate the clumped cells). Work quickly since cells must not be exposed to PEG for too long.
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13a. Carefully add 5 ml of Medium B dropwise to the pellet while gently swirling the tube to resuspend the cells. 14a. Slowly add 5 ml Medium C (Hybridoma Recovery Medium from the kit) to the solution. Continue to swirl the tube. 15a. Transfer the cell suspension to a 75-cm2 tissue culture flask containing 40 ml Medium C (total culture volume will be 50 ml). Incubate 16 to 24 hr at 37◦ C in 5% CO2 atmosphere. There will still be clumps of cells at this point, which will disaggregate overnight. Be gentle; fused cells are very fragile immediately after fusion.
Method B 12b. Slowly add 1 ml polyethylene glycol (PEG) solution from the ClonaCell-HY kit to the pellet obtained in step 11 dropwise over a period of 1 min, without stirring, using a 1-ml pipet. When all of the PEG has been added gently stir the cells continuously with the pipet tip for 1 min. 13b. Add 4 ml medium B to the fusion mixture dropwise over a period of 1 min, continuously stirring with the pipet tip for a total of 4 min. Slowly add 10 ml Medium B. Incubate 15 min in a water bath at 37◦ C. 14b. Slowly add 30 ml of Medium A over a period of 1 min. Centrifuge the cells 7 min at 400 × g, 37◦ C or room temperature. Discard the supernatant, add 40 ml Medium A, then centrifuge again as before and discard the supernatant to ensure all PEG is removed. 15b. Slowly resuspend the cell pellet in 10 ml Medium C. Transfer the cell suspension to a 75-cm2 tissue culture flask containing 40 ml Medium C (total culture volume, 50 ml). Incubate 16 to 24 hr at 37◦ C in 5% CO2 atmosphere. IMPORTANT NOTE: Incubating cells up to 24 hr in liquid medium allows fused cells time to go through one cell cycle and begin expressing the enzyme HPRT, which is necessary for cell survival during HAT selection. Freshly fused cells are also very fragile. Waiting a day before mixing these cells with methylcellulose (see below) will improve their survival.
Select and clone hybridoma cells 16. On the day of the fusion, place the frozen Medium D from the ClonaCell-HY kit (Hybridoma Selection Medium containing HAT) at 4◦ C and leave overnight to thaw. Before using, shake vigorously to thoroughly mix contents and leave on bench to warm up to room temperature. Do not thaw Medium D in a 37◦ C water bath, as the methylcellulose can be pulled out of solution causing the medium to become lumpy.
17. Transfer the fused cell suspension (step 15a or b) into a 50-ml conical centrifuge tube and centrifuge 10 min at 400 × g, room temperature or 37◦ C. Remove supernatant. Resuspend cells in Medium C for a total volume of 10 ml. It is critical that the total volume in this step not be greater than 10 ml. Include any additional cytokines or growth factors in this 10-ml volume prior to adding to Medium D.
18. Transfer the 10 ml of cell suspension into the 90 ml of Medium D. Mix thoroughly by gently inverting the bottle. Once mixed, let sit for 15 min at room temperature or 37◦ C to allow the bubbles to rise to the top. It is not advisable to warm the bottle in a 37◦ C water bath. Instead, place the bottle in a 37◦ C incubator.
Induction of Immune Responses
2.5.13 Current Protocols in Immunology
19. Using a 12-ml syringe and 16-G blunt-end needle, aseptically plate out 9.5 ml of the cell suspension from step 15 into each of ten 100-mm petri dishes. Tilt the plates to level the mixture and try not to introduce air bubbles. Incubate dishes at 37◦ C in a 5% CO2 atmosphere. Do not disturb plates for 10 to 14 days. Methylcellulose (present in Medium D) is a viscous solution and cannot be accurately dispensed using pipets due to adherence of the medium to pipet walls. The incubator must be well humidified. It is advisable to put the plates in a separate plastic container together with an open 100-mm petri dish containing 10 ml sterile distilled water. Open and close the incubator door carefully to avoid shaking. IMPORTANT NOTE: Handling plates prior to the recommended 10- to 14-day incubation time may result in “runny” colonies. Identical daughter cells may break apart and spread across the plate, rather than remain in a tight colony. This will make it more difficult to identify single colonies to pick for cloning.
Harvest clones 20. At a time point 10 to 14 days after plating, examine the plates for the presence of colonies visible to the naked eye (a typical fusion will produce 1000 or more colonies over the ten plates). Recloning in semisolid medium (Support Protocol 4) is recommended if colonies are not well distributed in the semisolid Medium D at this step, or if it is difficult to distinguish between individual colonies. This should be done after cells have been harvested and established in Medium E (see subsequent steps).
21. Remove isolated colonies (usually 500 to 1000 colonies are picked) from the plates with sterile pipet tips using a micropipettor set to 10 μl. Pipet each clone into an individual well of a 96-well tissue culture plate containing 200 μl Medium E (Growth Medium containing HT). Incubate the plates at 37◦ C in a 5% CO2 atmosphere 1 to 4 days without feeding. Usually by the fourth day, each well has a high cell density and the medium has begun to turn yellow. It is a good idea to pick clones of different sizes, as slower-growing clones (i.e., smaller colonies) are often very good antibody producers. Such slow-growing hybridomas are often missed in other hybridoma screening procedures.
22. Transfer 150 μl of the supernatant from each hybridoma well to a separate well on a new 96-well plate and analyze by an assay system appropriate for the antigen involved; e.g., ELISA (UNIT 2.1), flow cytometry (Chapter 5), or immunoblotting (UNIT 8.10). 23. Add 150 μl of fresh Medium E to each hybridoma-containing well of the original 96-well plate. 24. Gently resuspend the hybridomas that showed a positive response in step 22. Transfer 100 μl of the resuspended positive hybridoma to each of two wells of a 24-well plate containing 1 ml of Medium E.
Production of Monoclonal Antibodies
25. When cells have grown to a suitable density (∼4 × 105 cells/ml), freeze the cells from one well by centrifuging the culture at 400 × g, room temperature, remove the medium using a pipet, and add ice-cold FBS/20% DMSO for a concentration of 4 × 105 cells/ml. Transfer cells and medium to a cryotube, and place at −70◦ C, −135◦ C, or in liquid nitrogen (also see APPENDIX 3G). Expand the remaining positive clones (duplicate, identical wells) in 25-cm2 tissue culture flasks containing 5 ml of Medium A and 5 ml of Medium B. Individual hybridomas may differ in their optimal cell density for growth. The culture should not be too sparse, or there will be insufficient numbers for recovery of frozen cells.
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If the culture is allowed to grow to too high a density, the medium will become acidic and cell viability may decrease. Suggested density is 5 × 104 cells/ml, with a maximum density of 4 × 105 cells/ml.
26. Keep a sample of cells in Medium E, in case they do not adapt well to the mixed medium. 27. When cells have grown to a suitable density (∼4 × 105 cells/ml), pipet 5 to 10 ml of the cell culture into 30 ml Medium A. Adjust the volume to ensure that the final cell concentration is 1–4 × 104 cells/ml. 28. Maintain expanded hybridomas in 100% Medium A at a concentration of 5 × 104 to 5 × 105 cells/ml. More aliquots of cells can be frozen at this point in order to secure the supply of hybridoma. Recloning in semisolid medium (Support Protocol 4) is recommended if hybridomas have been in continuous culture for a long period of time and are secreting antibody, but monoclonality or stability of the hybridoma needs to be confirmed.
CELL FUSION AND SELECTION USING ELECTROFUSION An alternative to PEG-mediated fusion in the production of mouse hybridomas is an electrically induced fusion method known as electrofusion. Electric-field mediated cell fusion was first described by Zimmermann et al. (reviewed in Zimmermann et al., 1985). This method has since been modified and improved over the years. Today, a number of pulse-generating instruments are commercially available from various manufacturers including Eppendorf and Harvard Apparatus. While these instruments may differ in chamber design and the parameters of the electrical pulses they deliver, most of them are based on a two-step model in the most basic form. First, the two cell types—spleen and myeloma cells in this case—are aligned in a string adjacent to one another by an alternating current (AC) electrical field. Second, a transient direct current (DC) electrical pulse is introduced to induce reversible membrane poration and the formation of two-cell hybrids. Additional steps of electrical pulsing may be included at the end to stabilize the fused cells. Resulting hybridomas can then be selected from unfused parental myeloma cells through incubation in HAT medium over the course of 1 to 2 weeks in a way similar to the selection process that follows PEG-mediated fusion. Compared to PEG-mediated fusion, electrofusion methods typically have lower toxicity and yield higher fusion efficiency. Moreover, since the fusion process is electronically defined, consistency is often improved over PEG-mediated methods. This section describes an electrofusion method using the Harvard Apparatus BTX Hybrimune Hybridoma Production System. This system utilizes a three-step fusion process composed of an AC pre-fusion pulse, followed by a DC fusion pulse, and a final AC post-fusion pulse (adapted from manufacturer’s instrument manual).
ALTERNATE PROTOCOL 3
Materials Myeloma cell line (e.g., SP2/0, NS-1, P3.X63Ag8.653, or FOX-NY, available from ATCC) Myeloma growth medium for the myeloma cell line chosen (see instructions provided by ATCC) 4% (w/v) sodium hydroxide solution Sterile reagent-grade H2 O Spor-Klenz sterilizing solution (Steris Life Sciences, http://www.sterislifesciences .com/) Serum-free Dulbecco’s modified Eagle medium (DMEM) BTX Cytofusion medium (Harvard Apparatus)
Induction of Immune Responses
2.5.15 Current Protocols in Immunology
Hybridoma selection medium such as DMEM-20/HEPES/pyruvate/HAT (APPENDIX 2A) 70% ethanol 125-cm2 tissue culture flasks Hybrimune Hybridoma Production System with a fusion chamber appropriate for the scale of fusion Computer 50-ml conical polypropylene centrifuge tubes96-well tissue culture plates Additional reagents and equipment for counting cells (APPENDIX 3A) and determining cell viability by trypan blue exclusion (APPENDIX 3B) Prepare myeloma cells (10 days prior to fusion) 1. At least 10 days prior to fusion, thaw the mouse myeloma cells and culture in myeloma growth medium in a 125-cm2 tissue culture flask at 1 to 4 × 105 cells/ml. Maintain the cell density between 1 to 4 × 105 cells/ml by passaging the cells every 2 to 3 days. Boost primed animal(s) (3 days prior to fusion) 2. Three days before fusion, boost primed animal(s) according to step 7 of Basic Protocol 1. Expand myeloma cells (1 to 2 days prior to fusion) 3. One to two days before fusion, split the myeloma culture to achieve logarithmic growth on the day of fusion. Ideally, myeloma cells should be in logarithmic growth, between 1 to 1.5.× 106 cells/ml and >95% viable on the day of fusion. Prepare to have approximately 2 × 108 myeloma cells for each spleen to be fused.
Prepare Hybrimune waveform generator and fusion chamber (on the day of fusion) 4. Connect the computer to the waveform generator. Turn on both components and run the waveform generator interface software. Specify the pre-pulse, pulse, and post-pulse waveform parameters. A typical set for hybridoma electrofusion is as follows: Pre-pulse:
Start voltage = 60 V peak Stop voltage = 60 V peak Function = linear Duration = 15 sec Frequency = 1.4 MHz Amplitude = 800 V Width = 40 μsec Pulse number = 1 Start voltage = 60 V peak Stop voltage = 5 V peak Function = linear Duration = 30 sec Frequency = 1.4 MHz.
The values listed above are intended to be used as a guideline only. Optimal values for the various parameters of the waveforms may vary depending on the particular line of myeloma cells used. Production of Monoclonal Antibodies
5. Clean the fusion chamber as per manufacturer’s instruction.
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Typically, this involves a 5-min soak with 4% sodium hydroxide solution followed by 10 rinses with reagent-grade water and a final rinse with 70% (v/v) ethanol. The chamber can then be air dried until the fusion step.
Prepare spleen and myeloma cells 6. Harvest splenocytes according to Basic Protocol 2, steps 7 to 9, then fill up the tube with serum-free DMEM. Optionally, lyse red blood cells (Basic Protocol 1, steps 10 to 13). Count white blood cells (APPENDIX 3A) and determine viability (APPENDIX 3B). 7. Harvest myeloma cells and place into 50-ml conical tubes. Count the myeloma cells (APPENDIX 3A) and determine viability (APPENDIX 3B). Both cell populations should be >95% viable for efficient fusion.
8. Centrifuge both cell populations 8 at 400 × g, room temperature, and remove the supernatant. Resuspend each population in 20 ml of BTX Cytofusion medium. Repeat this step once more. 9. Perform a viable count on both cell populations (APPENDIX 3B). Combine equal numbers of splenocytes and myeloma cells. Centrifuge the mixture 8 min at 400 × g, room temperature, and remove the supernatant. Resuspend the mixture at 1–2 × 107 cells/ml in BTX Cytofusion medium and transfer the suspension to a pre-sterilized fusion chamber (see step 5 above).
Perform cell fusion 10. Initiate fusion protocol. After the post-pulse step is finished, allow the cells to sit undisturbed in the fusion chamber for 5 min. 11. Gently transfer the cells to a 50-ml conical centrifuge tube. Rinse the fusion chamber with BTX Cytofusion medium and transfer the wash to the same 50-ml conical tube, to minimize cell loss in the chamber. 12. Fill the 50-ml tube with hybridoma selection medium. 13. Incubate at 37◦ C for 10 to 20 min to allow fused cells to rest. 14. While waiting for the fused cells to stabilize, clean the fusion chamber as described in step 5, above. Let stand for 5 min, then empty the chamber and refill it with sterile reagent-grade water. Let stand for 10 sec, then empty the chamber. Repeat this procedure nine more times. Rinse with 70% ethanol and air dry. 15. After the fused cells have been stabilized, adjust the cell density to target plating density. Plate out the cell suspension in 96-well tissue culture plates. Incubate the plated cells at 37◦ C with 5% CO2 . Typically, cells can be plated at 5 to 10 × 103 cells/well in 200 μl/well. Plating density may be adjusted as desired.
16. Refeed cultured cells 3 and 5 days after plating as described in step 34 of Basic Protocol 2. Screen the supernatants 12 to 14 days after plating for reactivity against immunogen. Refeeding schedule can be varied according to growth conditions.
SCREENING PRIMARY HYBRIDOMA SUPERNATANTS The vast majority of wells will not contain the desired antibody, or may contain nonproducing hybridomas. The purpose of screening is to discover which wells (