- Email: [email protected]
Production of Microcell Hybrids
METHODS: A Companion to Methods in Enzymology 9, 3–11 (1996) Article No. 0002
Production of Microcell Hybrids Ann McNeill Killary1 and Steven T. Lott Division of Laboratory Medicine, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030
Microcell fusion is a technology that involves the transfer of single or small clusters of intact chromosomes from one cell to another. The transferred chromosome can be stably retained in the recipient cell background under dominant selective pressure. Hybrid clones generated by this method result in karyotypically simple and homogeneous populations that are excellent resources for physical mapping. This article will describe a general strategy for the efficient micronucleation of human and rodent cell populations and their use as donors for microcell fusion into recipient cell lines. q 1996 Academic Press, Inc.
Parasexual approaches for gene mapping were greatly enhanced with the development of the technology of microcell fusion. In microcell fusion experiments, single, intact chromosomes may be transferred from one cell to another, resulting in karyotypically simple somatic cell hybrids for genetic analysis. Five essential steps are involved in this technology; they include: (1) micronucleation of donor cells; (2) enucleation of micronucleate cell populations; (3) microcell isolation; (4) fusion of microcells to recipient cells; and (5) selection of viable microcell hybrid clones (diagrammed in Fig. 1). In outline, donor cells are exposed to the mitotic inhibitor Colcemid for a prolonged period. Colcemid-induced mitotic arrest results in the formation of micronuclei containing single or small numbers of chromosomes. Micronuclei are then extruded from the donor cell population by exposure to cytochalasin B in the presence of centrifugal force. The cytochalasin B-induced enucleation of micronucleate populations results in the isolation of an assortment of cell particles including microcells, enucleated whole nuclei (karyoplasts), whole cells, and cytoplasmic vesicles. Following enucleation, microcells (usually containing 1–5 interphase chromosomes) are purified and fused to recipient cells. Viable 1 To whom correspondence and reprint request should be addressed at Division of Laboratory Medicine, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Boulevard, Box 72, Houston, TX 77030-4095. Fax: (713) 794-1800.
microcell hybrid clones are then isolated using a variety of selection schemes. This article will describe a general strategy for the efficient micronucleation of human and rodent cell populations and their use as donors for microcell fusion into recipient cell lines.
DONOR CELL MICRONUCLEATION The first step in the technology of microcell fusion involves the prolonged treatment of growing cells with the mitotic inhibitor Colcemid (1–3). Colcemid interferes with the polymerization of microtubules that are necessary for the formation of a functional spindle during metaphase. In the absence of a spindle, chromosomes appear scattered throughout mitotic cells. Prolonged Colcemid arrest results in a fraction of cells in the population that attempt to escape the mitotic block in order to divide. The nuclear membrane, rather than surrounding the entire chromosome complement, reforms around single or small clusters of chromosomes, forming micronuclei (2, 4). A general protocol for the micronucleation of rodent and human cell lines is described below. Rodent Cell Micronucleation by Prolonged Mitotic Arrest Micronucleation of many rodent lines can be accomplished by exposure of growing cells to a prolonged mitotic block (48 h) using Colcemid concentrations in the range 0.01–0.1 mg/ml (5). Optimal concentrations of Colcemid, however, should be determined for each donor cell line. Conditions have been determined for murine L cell lines, including A9, and for microcell hybrids containing single human chromosomes in the A9 cell background (6). A standard protocol for micronucleation of A9-based monochromosomal hybrids involves the plating of donor cells onto thin plastic sheets, termed ‘‘bullets,’’ for micronucleation and subsequent enucleation. 3
1046-2023/96 $18.00 Copyright q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
/ m4819$131d
02-19-96 08:01:24
metha
AP: Methods
KILLARY AND LOTT
4
for 3–4 h to overnight to allow for cell attachment prior to Colcemid addition. Colcemid (Demecolcine, Sigma) is then added to each dish (0.06 mg/ml) for 48 h to induce micronucleation. The percentage of micronuclei obtained may be quantitated using several protocols (outlined in a later section of this chapter).
FIG. 1. The technology of microcell fusion.
Preparation of Bullets for Micronucleation and Enucleation Bullets, which resemble microscope slides with one rounded end, can be cut from the bottoms and sides of T75 or T150 tissue culture flasks using a hot wire cutter. The optimal size of each bullet is such that it fits snugly inside a 50-ml conical centrifuge tube (approximately 25 1 90 mm). Before cell plating, bullets should be washed in glassware detergent, rinsed thoroughly in water, and placed in 95% ethanol for 17–24 h. Bullets are then removed from ethanol using sterile forceps, placed in 150-mm tissue culture dishes (four bullets per dish), and left in the tissue culture hood to dry. Following enucleation, bullets can be soaked in a 3% solution of bleach overnight, washed with detergent, and rinsed in water. Micronucleation on Bullets For micronucleation of A9 cells or A9-based microcell hybrids, one T150 tissue culture flask of logarithmically growing cells at 80–90% confluence is used per experiment. The flask is trypsinized and replated into four 150-mm tissue culture dishes, each containing 4 bullets, in complete medium containing 50 mg/ml gentamycin. The number of bullets used per experiment is dependent on the percentage of micronuclei formed. A9 cells consistently micronucleate at 90–95%. Therefore, we routinely use 16 bullets per experiment for A9 or A9-based hybrids. Tissue culture dishes are left at 377C
/ m4819$131d
02-19-96 08:01:24
metha
Human Cell Micronucleation by Prolonged Mitotic Arrest The most efficient donor cells for human micronucleation are early passage human foreskin fibroblasts (7). Human diploid fibroblasts are also attractive donors for microcell fusion in that they represent early passage cultures with a normal diploid chromosome complement in contrast to established, transformed cell lines that may have undergone many chromosomal rearrangements. In addition, micronucleation of many human cell lines has proved to be difficult (7). Micronucleation of human lymphoblastoid cells has been reported; however, the percentage of micronucleation is low, which necessitates the use of large numbers of donor cells (8). Therefore, for the purposes of this article, the protocol described will be for use with human foreskin fibroblasts. For human cell micronucleation, four T150 tissue culture flasks containing logarithmically growing human foreskin fibroblasts at 80–90% confluence are seeded (1:2) into eight 150-mm tissue culture dishes, each containing four bullets in complete medium with 50 mg/ml gentamycin. Following cell attachment (3–4 h), Colcemid (10 mg/ml) is added to each dish, and cells are incubated at 377C for 48 h to induce micronucleation. Induction of micronucleation in human fibroblasts is critically dependent on the concentration of mitotic inhibitor and the absolute passage number of the donor cells (7). Colcemid concentrations required to induce micronucleation are in the range 10–20 mg/ml for a 48h arrest. Under these conditions, 40–60% of the cell population becomes micronucleate (Fig. 2). Increasing concentrations of Colcemid may result in increased cell toxicity and a decreased yield of micronucleated cells (7). By examination of the number of micronuclei per cell as a function of increasing Colcemid concentration and length of mitotic arrest, these conditions have proved optimal (7). Antikinetochore antibody staining was previously used to examine the numbers of kinetochores per micronucleus (A. McNeill Killary, R. L. Brown, S. Brinner, B. Brinkley, and E. Stubblefield, unpublished results). Results indicated that the smallest of micronuclei contain single chromosomes. Another critical variable for human cell micronucleation is the age of the culture (7). Early passage human foreskin fibroblasts micronucleate optimally; however, after a population doubling of 17–18, the percentage of micronucleate cells decreases dramatically (7). Also, as indicated previously, the length of time in the mi-
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
totic inhibitor is an important consideration for experimental design (6). The length of mitotic arrest is dependent on the growth rate of the donor population cell. Slower growing cell lines may require longer arrest times. We have also observed that the recovery of viable microcell hybrid clones containing intact donor chromosomes is greater in incubations of 48 h compared with a 72-h arrest time. In general, we have also observed that the longer the incubation time in Colcemid, the higher the percentage of microcell hybrids recovered containing fragmented donor chromosomes.
ENUCLEATION OF MICRONUCLEATE CELL POPULATIONS Large-scale enucleation of micronucleate cell populations is accomplished by centrifugation of cells containing micronuclei in the presence of cytochalasin B. Cytochalasin B is a fungal metabolite that interferes with microfilament attachment to the cell membrane (9). High concentrations of cytochalasin B (10 mg/ml) result in nuclear extrusion in a fraction of cells in a population. Mass enucleation of cells was first demon-
strated by incubation in cytochalasin B combined with centrifugation (10–12). During the enucleation process, nuclei are extruded from the cell on thin cytoplasmic stalks and pellet at the bottom of the centrifuge tube (Fig. 3). The cytoplasm of enucleate cells remains attached to the bullets on which the cells are plated. Enucleation of micronucleate cells results in the formation of microcells, consisting of a single micronucleus containing approximately one to five chromosomes surrounded by a small rim of cytoplasm and an intact plasma membrane (13). After enucleation, the pellet in each centrifuge tube contains microcells of various sizes, enucleated whole cells, whole cells that were stripped off the bullet prior to enucleation, and cytoplasmic particles. Following are enucleation conditions for rodent and human cells in culture. Large-Scale Enucleation of Mouse A9 Cells or A9-based Hybrids Using the outlined protocol for the enucleation of A9 cells or A9-based hybrid cells, it is possible to achieve enucleation of ú90–95% of the cell population. A stock solution of cytochalasin B (Sigma) at 1–2 mg/ml is prepared by dissolving the powder in dimethyl sulfoxide (Sigma). The stock solution is then trans-
FIG. 2. Micronucleation by prolonged mitotic arrest. Acridine-orange-stained micronuclei from human diploid foreskin fibroblasts.
/ m4819$131d
02-19-96 08:01:24
metha
5
AP: Methods
KILLARY AND LOTT
6
ferred to sterile conical tubes and wrapped in foil to protect from light. For enucleation, eight 50-ml polycarbonate centrifuge tubes with lids (Nalgene) are autoclaved. To each tube, 38.5 ml of serum-free medium containing 10 mg/ml cytochalasin B is added and allowed to equilibrate at 377C. Aseptically, each of 16 bullets containing micronucleate A9 cells is removed from the tissue culture dish and placed inside a centrifuge tube. Bullets are placed such that 2 bullets are back-to-back inside each 50-ml tube. Lids are secured on tubes with tape. Tubes containing bullets are immediately placed inside a prewarmed (28–337C) superspeed centrifuge with a fixed angle rotor (for example, Sorvall RC-5B centrifuge and SS-34 rotor) such that bullets are parallel to rotor radii. Cells containing micronuclei are then centrifuged at 27,000g (15,000 rpm) for 70 min. Following centrifugation, bullets are removed from tubes and examined (as described in the next section) for the efficiency of enucleation. Bullets are then placed in a 3% bleach solution overnight, washed, and reused for subsequent experiments. Medium from each tube is carefully decanted, and pellets are resuspended. Pellets are redistributed in 1–2 ml serum-free medium and combined into a 15-ml conical tube. Following resuspension, an aliquot is saved for hemocytometer counts and quantification of yield. The tube is then incubated at 377C until fusion. Enucleation of Human Fibroblasts The conditions for human fibroblast enucleation are essentially the same as those outlined for A9 enucle-
ation, with the following exceptions. Optimal micronucleation for early passage human fibroblasts is 40– 60%, as opposed to approximately 95% for A9-based hybrids or A9 cells. We therefore increase the number of bullets used per experiment from 16 bullets as previously described for A9 enucleation to 32 bullets for human fibroblast enucleation. Bullets are centrifuged at 18,800g for 35 min in serum-free medium containing 5 mg/ml cytochalasin B. The remainder of the steps for large-scale enucleation of human fibroblasts are identical to those previously described for A9 cells. Enucleation of Nonadherent Cells on Bullets Procedures have been described for the efficient enucleation of cells that fail to adhere well to the substratum or for cells that grow in suspension (14). In this situation, bullets can still be used for enucleation if they have been pretreated with concanavalin A (6, 14). Concanavalin A (Con A) (Sigma) (15 mg/ml solution in 0.9% saline) is cross-linked to bullets using a crosslinking agent [1-cyclohexyl-3-(2 morpholinoethyl) carbodiimide metho-p-toluene sulfonate, water-soluble carbodiimide (WSC; Sigma)]. WSC solution is prepared at 75 mg/ml in 0.9% saline. For this procedure, bullets are removed from 95% ethanol and placed in tissue culture dishes to dry. Each of the bullets is layered with 0.6 ml of WSC solution followed by 0.6 ml of Con A solution, and the mixture is spread evenly. Bullets are incubated at room temperature for 1 to 2 h. Next, the Con A–WSC solution is aspirated off the bullets, and bullets are then rinsed twice with sterile phos-
FIG. 3. Electron micrograph of an enucleating cell (courtesy of Dr. Ronald L. Brown).
/ m4819$131d
02-19-96 08:01:24
metha
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
phate-buffered saline (PBS). Donor cells are induced to micronucleate in suspension by prolonged Colcemid arrest, as previously described in T75 or T150 flasks. Flasks are harvested by trypsinization, counted, and resuspended in PBS at 106 cells/ml. Then 1.2 ml of cell suspension is added to each bullet, and the cells are allowed to attach for 10–15 min at room temperature. Complete medium is added, and the cells are incubated for 30–60 min at 377C to allow for cell spreading. Con A bullets containing micronucleate cells can then be processed for enucleation as described in the preceding section. Suspension Enucleation Enucleation of suspension cultures can also be accomplished using gradient separation through Ficoll (15) and Percoll (8) gradients. Gradient separation is based on differences in buoyant densities of the nucleoplasm versus the cytoplasm under centrifugal force. Suspension enucleation is especially applicable to cell lines that fail to micronucleate efficiently. Lymphoblastoid cells have been used successfully as donors for microcell fusion despite their poor micronucleation efficiency because large numbers of cells can be used in the gradient. Enucleation in gradients, however, is often incomplete, and many partially enucleated cells will be lost because they partition away from isolated microcells. This approach is, nevertheless, the method of choice for cell lines that cannot be enucleated using bullets. The main variables associated with enucleation are (1) the concentration of cytochalasin B; (2) centrifugation speed; and (3) centrifugation time. The concentration of cytochalasin B is usually in the range 5–10 mg/ ml for both human and rodent lines. The variables that may need to be adjusted for each cell line are centrifugation time and speed. Careful monitoring of the steps in enucleation is imperative to achieve the optimal yield of microcells. Enucleation should result in virtually 99% nuclear extrusion. If incomplete enucleation occurs, then incremental increases in either time or speed can be monitored for complete enucleation. However, it should be noted that excess centrifugal force or time can result in the complete stripping of cytoplasm from the bullets.
QUANTITATION OF MICROCELLS This section will describe a general protocol for quantitation of microcell yield following enucleation. This procedure should also be used to stain bullets to determine the percentage of micronucleate cells in the population as well as to determine the efficiency of enucleation.
/ m4819$131d
02-19-96 08:01:24
metha
7
Following enucleation, the pellets that are recovered consist of a variety of types and sizes of cell particles, including microcells, intact nuclei, whole cells that stripped off the bullets prior to enucleation, and cytoplasmic particles. Hemocytometer counts can be used to determine the total number of particles per experiment. However, to determine the microcell particle yield, staining of the preparation is necessary to discriminate between nuclear and cytoplasmic particles. Aceto-orcein will differentially stain the nucleus from the cytoplasm and is used in conjunction with bright-field microscopy. Using this stain, nuclei appear red and the cytoplasm appears pink. An aceto-orcein solution is prepared by mixing 0.5% orcein solution (Sigma) in 50% acetic acid followed by refluxing this mixture for several hours. To quantitate microcell yield following enucleation, one drop of microcell preparation is mixed with one drop of aceto-orcein stain on a microscope slide. The preparation is coverslipped, stained for 2–5 min, and examined by bright-field microscopy. The staining time is critical for accurate determinations of nuclear versus cytoplasmic fragments. After an incubation time of approximately 5–10 min, all particles stain red. Thus, the interpretation of aceto-orceinstained material is sometimes difficult because discrimination between shades of the same color is required rather than visualization using fluorochromes of contrasting colors. The advantage of this staining protocol is its simplicity; visualization does not require a fluorescence microscope. A similar approach for the quantitation of microcell yield involves the use of the fluorochrome acridine orange. Acridine orange stains the nucleus and cytoplasm differentially in that nuclei and micronuclei fluoresce green, whereas cytoplasmic particles appear red using fluorescence microscopy. This stain is performed essentially as described for the aceto-orcein protocol and is an excellent method for quantitation of micronucleation, enucleation efficiency, or microcell yield. For this method, a stock solution of acridine orange is prepared by dissolving powder in 2 mg/ml sterile water. Acridine orange is a carcinogen; therefore, care should be taken in its handling. The stock solution is stored in a foil-wrapped conical tube at 47C. The working solution of acridine orange is 10 mg/ml prepared in water or normal saline. A foil-wrapped coplin jar is suitable for processing several bullets or slides for staining. To determine the efficiency of micronucleation or enucleation, bullets are removed from medium and placed in a coplin jar containing a 1:1 mixture of 95% methanol and serum-free medium for 10 min. Bullets are then transferred to a second coplin jar containing 95% methanol for 10 min and allowed to dry. Each bullet is then placed in the acridine orange stain for 30–60 s. Bullets are then rinsed twice with water and
AP: Methods
KILLARY AND LOTT
8
coverslipped using 0.05 M citrate phosphate mounting buffer, pH 4.1. The accurate discrimination of nucleus from cytoplasm is accomplished by screening bullets using fluorescence microscopy.
ISOLATION OF MICROCELLS There are several different strategies for purification of the microcell population following enucleation. This step in the microcell fusion technique is important if the goal is to transfer single chromosomes. Nuclepore Filtration The simplest strategy to purify microcells is to filter the crude microcell preparation through nuclepore filters to selectively isolate the smallest microcells in the preparation and to exclude contaminating whole cells and nuclei (7). For filtration, various sized nuclepore filters (8, 5, or 3 mm) are placed inside autoclaved Swinnex filter units (Swinnex, 25 mm) attached to sterile 10-cc syringes. Following enucleation, the crude microcell preparation is resuspended in a total volume of 10–15 ml serumfree medium. One-half of the crude preparation is poured into the syringe containing a 5-mm nuclepore filter and allowed to filter through gravity with only slight pressure applied as needed on the plunger. This step is repeated with the remaining half of the microcell preparation, using a new 5-mm filter set. Nuclepore filtration should result in exclusion of most of the whole cells and nuclei from the crude preparation. Preparations containing large numbers of intact cells should be passed through 8-mm filters prior to 5-mm filtration. The remaining preparation should contain various sizes of microcells and cytoplasmic particles. To select for the smallest microcells, which likely contain single chromosomes, the preparation is filtered again through two 3-mm filter units exactly as described previously for 5-mm separation. Careful monitoring of the filtration is required to ensure that the filter remains intact during the procedure. Excess pressure on the plunger could result in cracking of the filter. To check for the efficiency of the filtration step, aliquots are taken before and after filtration for quantitation using acridine orange or aceto-orcein. Using this filtration step, the resultant purified preparation should contain only the smallest of microcells, with complete elimination of whole cells and 99% of karyoplasts. The efficiency of this procedure is very much dependent on the proper use of the filter. If the filter is overloaded with the crude preparation, then clogging of the filter occurs. This results in a decrease in the total particles processed through the filter. Hemocytometer counting prior to filtration will allow the
/ m4819$131d
02-19-96 08:01:24
metha
optimal number of total particles ((1–5 1 106)/ml) to be passed through each filter unit. Microcell Purification via Unit Gravity Sedimentation Microcells can also be isolated using unit gravity sedimentation on density gradients (5). For unit gravity separation, a linear gradient of 1–3% bovine serum albumin (BSA) is prepared using a gradient mixing device connected to a peristaltic pump. Tubing is connected from the gradient mixing device through the peristaltic pump to a three-way valve. One valve outlet is connected to a 5-cc syringe that functions as a bubble trap. The second valve is connected to a 50-cc syringe that functions as the gradient chamber. Three glass beads are placed over the inlet of the chamber. Next, a small piece of tubing is connected to the other outlet, which serves as the sampling tube. The entire apparatus can be autoclaved. Next, sterile PBS is added to the gradient mixing device and pumped in to fill the tubing just at the bottom of the gradient chamber. Excess PBS is pumped into the bubble trap. To one side of the mixing device, 25 ml of 3% BSA in PBS is added, and to the other side, 25 ml of 1% BSA in PBS (containing phenol red) is added. The connection between the chambers is then opened and the stirrer motor started. Following initiation of the motor, the microcell preparation is resuspended in 2 ml of 0.5% BSA in PBS and added to the gradient chamber from the top. The pump is started and PBS in the lines is pumped into the bubble trap. When the BSA solution (red) reaches the trap, the valve is turned to allow the solution to enter the gradient chamber. The pumping rate for the BSA gradient should be 2–3 ml/min. The microcell preparation is then allowed to settle through the gradient for 3–3.5 h, which separates the microcells, whole cells, nuclei, and cytoplasmic particles by size. Fractions are collected by dripping through the sample tube. This type of separation efficiently separates whole cells and nuclei in the bottom 15–20 ml of the 50-ml gradient. The purified microcell preparation is recovered from the top 20–25 ml. Unit gravity sedimentation is probably the most efficient method for purification of the crude microcell preparation. This technique virtually eliminates whole cells and nuclei from the crude preparation. It is, nevertheless, a time-consuming procedure and therefore may not be the method of choice unless absolute purification of the preparation is needed.
FUSION OF MICROCELLS TO RECIPIENT CELLS In initial strategies for microcell fusion, inactivated Sendai virus was used to fuse microcells to recipient
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
cells (5). Later, protocols proved to be more simple and efficient by using a combined treatment of cells with phytohemagglutinin-P (PHA-P) to adhere microcells to recipient cells, followed by fusion using polyethylene glycol (PEG) (7, 14). Following are two protocols for microcell fusion. The first protocol involves the agglutination of microcells to a recipient monolayer using PHA-P, followed by fusion using PEG. The second method is a suspension protocol for use with cells that do not adhere to monolayer culture or for use when PHA-P is toxic to recipient cells. Two important variables need to be determined for each recipient cell line prior to microcell fusion. The first is the concentration of fusogen. This can be determined by simply fusing recipient cells in culture in increasing concentrations of fusogen. The conditions that yield the highest percentage of binucleate heterokaryons postfusion with the least amount of cell toxicity are optimal. Optimal fusogen concentrations are usually in the range 44–50% PEG w/w (Koch Chemical, United, MW 1540). Higher concentrations of fusogen can result in large numbers of fused cells en masse, with increased cell toxicity. PHA-P toxicity experiments should also be performed in increasing concentrations of the agglutinin in the range 50–200 mg/ml. In this case, the optimal conditions for PHA-P are such that cell–cell agglutination occurs with the least amount of cell toxicity. Agglutination can be monitored by phase-contrast microscopy, and the degree of cell toxicity can be measured by trypan blue staining. The following is a general protocol for monolayer microcell fusion. PEG is prepared in serum-free medium. For fusion to a recipient monolayer of A9 cells, a concentration of 50% w/w is optimal. The PEG solution is incubated at 377C until PEG has dissolved and then filter sterilized. The pH of the PEG solution should be in the range 7.5–8.0. PHA-P (Difco) is a lyophilized powder that is 50% salt by weight. The concentration of PHA-P used routinely for microcell fusion to a monolayer of A9 recipients is 100 mg/ml in serum-free medium. This solution can also be filter sterilized. PHA-P/PEG Monolayer Fusion Monolayer cultures of recipient cells are prepared in 25-cm2 flasks. Routinely, two flasks are used for microcell fusion, and one flask is reserved as a control. Recipient cell cultures should be subcultured 4–24 h prior to fusion to allow for firm attachment to the substratum. Each flask should be at 70–80% confluence at the time of fusion. The purified microcell preparation is then centrifuged at 2000g for 10 min (377C). The microcell pellet is resuspended in 4 ml PHA-P solution (100 mg/ml). At this step, it is important to redistribute the solution well to disperse any clumps of microcells prior to layering the solution onto the monolayer. Medium is then
/ m4819$131d
02-19-96 08:01:24
metha
9
aspirated off the recipient cell monolayers, and 2 ml of microcell/PHA-P solution is added to each of two T25 fusion flasks. Flasks are incubated at 377C until microcells have agglutinated to the monolayer (usually 10– 20 min). Next, the excess medium is carefully aspirated off the fusion monolayers, 0.5 ml PEG (50% w/w) is added, and the flask is gently rocked for 1 min. Following the 1-min incubation, the PEG solution is quickly aspirated off the monolayer, which is next rinsed three times with 4 ml serum-free medium. Complete, nonselective medium is then added, and flasks are returned to 377C for 17–24 h prior to addition of selective medium. The control T25 is treated as above with the exception that it does not receive donor microcells. Following incubation, fusion flasks are trysinized and replated into 10–20 T25s per fusion. Selective medium is then added to each flask. Viable microcell hybrids should be visible within 2–4 weeks. Suspension Microcell Fusion Microcells can be fused to recipient cell lines in suspension. This protocol does not use PHA-P and is appropriate for cell lines that grow in suspension or for which PHA-P is cytotoxic. For suspension fusion, recipient cells are counted and resuspended at 1 1 106 cells/ml in 5 ml serumfree medium. To this suspension, the filtered microcell preparation is added in a 15-ml conical centrifuge tube. The microcell/cell suspension is then centrifuged at 2000g for 15 min at 377C. The resulting pellet should be well dispersed. To this pellet, 0.5 ml PEG (50% w/ w) is added dropwise while gently resuspending the pellet for 1 min. Immediately, 9.5 ml of serum-free medium is added over 1 min with gentle swirling. Next, 20 T25s/fusion are prepared in complete nonselective medium. Twenty flasks are also prepared for the control. Following fusion, 0.5 ml of fusion or control suspension is aliquoted into each T25. Flasks are then incubated overnight (17–24 h) at 377C. Fusion and control flasks are then refed with complete selective medium and allowed to incubate at 377C until clones become visible. Both the monolayer and the suspension protocols have successfully resulted in the generation of viable microcell hybrid clones. The monolayer protocol for fusion is much more efficient than the suspension protocol, with transfer frequencies in the range 0.5–2.0 1 1005. The suspension protocol is the method of choice for cell lines that grow in suspension or adhere poorly to the monolayer or for which PHA-P is cytotoxic.
SELECTION STRATEGIES FOR THE ISOLATION OF MICROCELL HYBRID CLONES Microcell hybrid clones containing single introduced chromosomes can be isolated under dominant selective
AP: Methods
KILLARY AND LOTT
10
pressure. Microcell hybrids have been used for complementation mapping, in which the recipient cells harbor a recessive, conditional lethal mutation that is complemented by the introduction of a wild-type copy of that gene on the introduced chromosome (16). The X chromosome, human chromosome 17, or mouse chromosome 11 can be selected using the nucleotide salvage pathway complementation called the HAT selective system (17). Also, adenine phosphoribosyl transferase (APRT) complementation via the AAT selective system can select for human chromosome 16 or mouse chromosome 8 (18). The most conceptually ideal system for microcell fusion would be one in which each donor chromosome contains a dominant selectable marker (7). Dominant selectable markers have been successfully transferred into donor chromosomes prior to micronucleation and then transferred into a variety of recipient lines (19– 22). The most popular vectors containing dominant selectable markers are pSV2neo, a plasmid vector containing the bacterial gene neo that confers resistance to the antibiotic G418 (23), and pSV2gpt, a similar vector containing the bacterial gene gpt that can be selected in HPRT0 cells using HAT selection or in wild-type back cells using mycophenolic acid (24). Retroviral vectors (pZIP-neo-SV(X1)) have also been used to efficiently tag donor chromosomes for microcell fusion (21). Retroviral transfer has an advantage in that the vector can be inserted into a single site per cell, and high frequency transduction is possible. Our laboratory has generated a standard protocol for the tagging of human chromosomes and subsequent transfer of marked human chromosomes into recipient A9 cells (6, 22). We use the technique of electroporation (25–27) as a simple, straightforward way to introduce pSV2neo into human diploid fibroblasts. Electroporation has distinct advantages over other systems in that, unlike calcium phosphate transfection, the DNA transferred is low copy number, the integration sites are limited, and the procedure can be used without the biosafety concerns associated with the use of retroviral vectors. For our experiments, the method of Potter (27) was modified for human foreskin fibroblasts and for use with a commercial electroporator (Bio-Rad gene pulser). Following is a general strategy for the tagging of normal diploid human fibroblasts with pSV2neo and subsequent selection in the antibiotic G418. Electroporation of pSV2neo Into Diploid Human Fibroblasts Human diploid fibroblasts (1 1 107 cells) are mixed with linearized plasmid DNA (10 mg/ml of pSV2neo) in PBS and incubated on ice for 10 min. A 2000-V electrical pulse is applied to the cell/DNA mixture, which is then incubated on ice for 10 min postelectroporation.
/ m4819$131d
02-19-96 08:01:24
metha
The suspension is replated in complete nonselective medium at 5 1 105 cells per T25. After a 48-h incubation at 377C, selective medium containing the antibiotic geneticin (G418) at 750 mg/ml is added to the medium to select for cells carrying tagged chromosomes. Clones can be observed at 10 days postelectroporation. Properties of Microcell Hybrid Clones Microcell hybrid clones can be isolated from flasks by first removing the tops of flasks with a soldering gun and then placing cloning rings over individual clones. Microcell hybrid clones, in general, have sizes and morphologies similar to that of the recipient cell line. They can be readily distinguished from whole-cell hybrid clones containing many introduced chromosomes, which are usually much larger. In general, microcell hybrids will stably retain the introduced chromosome as long as selective pressure is maintained. This results in a nearly homogeneous hybrid population for genetic studies. This property of microcell hybrids makes them ideal for quantitative somatic cell genetic studies that require high percentages of the introduced chromosome in the recipient cell population. Monochromosomal hybrids have a number of advantages over whole-cell hybrid clones because they necessarily contain only the introduced chromosome, as opposed to segregating whole-cell hybrids, in which other chromosomes, translocations, and chromosomal fragments could be a part of the hybrid genome. The stability of the introduced chromosome in the microcell hybrid is dependent to a certain extent on the individual recipient cells used (16). The isolation of microcell clones containing intact transferred chromosomes is much lower in certain recipient backgrounds, such as HT1080 and CHO cells (16). Microcell fusion is, to date, the only reliable strategy for the transfer of single, intact chromosomes. Microcell hybrids have also been extremely useful as a source of hybrids containing fragments of the introduced chromosome (16, 28). Together, monochromosomal microcell hybrids and fragment-containing hybrids are extremely valuable tools both for physical mapping in both interspecific and intraspecific crosses and for quantitative studies such as the identification of tumor suppressor genes and genes involved in tissue-specific gene regulation.
REFERENCES 1. Levan, A. (1954) Hereditas 40, 1–64. 2. Stubblefield, E. (1964) (Harris, R. J. C., Ed.), in Cytogenetics of Cells in Culture pp. 223–298, Academic Press, New York. 3. Phillips, S. G., and Phillips, D. M. (1969) J. Cell Biol. 40, 248. 4. Ege, T., Ringertz, N. R., Hamberg, H., and Sidebottom, E. (1977)
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.
in Methods in Cell Biology (Prescott, D. M., Ed.), Vol. XV, pp. 339–358, Academic Press, New York. Fournier, R. E. K., and Ruddle, F. H. (1977) Proc. Natl. Acad. Sci. USA 74, 319–323. Killary, A. M., and Fournier, R. E. K. (1995) in Methods in Enzymology, Academic Press, San Diego, in press. McNeill, C. A., and Brown, R. L. (1980) Proc. Natl. Acad. Sci. USA 77(9), 5394–5398. Sanford, J. A., and Stubblefield, E. (1987) Somat. Cell Mol. Genet. 13, 279. Carter, S. B. (1967) Nature 213, 261–264. Prescott, D. M., Myerson, D., and Wallace, J. (1972) Exp. Cell Res. 71, 480–485. Poste, G., and Reeve, P. (1972) Exp. Cell Res. 73, 287–294. Wright, W. E., and Hayflick, L. (1972) Exp. Cell Res. 74, 187– 194. Ege, T., and Ringertz, N. R. (1974) Exp. Cell Res. 87, 378–382. Fournier, R. E. K. (1981) Proc. Natl. Acad. Sci. USA 78(10), 6349–6353. Wigler, M. H., and Weinstein, I. B. (1975) Biochem. Biophys. Res. Commun. 63, 669–674. Fournier, R. E. K., and Moran, R. G. (1983) Somat. Cell Mol. Genet. 9, 69–84.
/ m4819$131d
02-19-96 08:01:24
metha
11
17. Littlefield, J. W. (1964) Science 145, 709–710. 18. Kusano, T., Long, C., and Green, H. (1971) Proc. Natl. Acad. Sci. USA 68, 82–86. 19. Athwal, R., Smarsh, M., Searle, B. M., and Surinder, S. D. (1985) Somat. Cell Mol. Genet. 11, 177–187. 20. Saxon, P. J., Srivatsan, E. S., Leipzig, G. V., Sameshima, J. H., and Stanbridge, E. J. (1985) Mol. Cell. Biol. 5, 140–146. 21. Lugo, T. G., Handelin, B., Killary, A. M., Houseman, D. E., and Fournier, R. E. K. (1987) Mol. Cell. Biol. 7, 2814–2820. 22. Killary, A. M., Wolf, M. E., Giambernardi, T. A., and Naylor, S. L. (1992) Proc. Natl. Acad. Sci. USA 89, 10877–10881. 23. Southern, P. J., and Berg, P. (1982) J. Mol. Appl. Genet. 1, 327– 341. 24. Mulligan, C., and Berg, P. (1980) Science 209, 1422–1427. 25. Zimmerman, U., and Vienken, J. (1982) J. Membr. Biol. 67, 165– 182. 26. Neumann, E., Schaefer-Ridder, M., Wang, Y., and Hofschneider, P. H. (1982) EMBO J. 1, 841–845. 27. Potter, H., Weir, L., and Leder, P. (1984) Proc. Natl. Acad. Sci. USA 81, 7161–7165. 28. Leach, R. J., Thayer, M. J., Schafer, A. J., and Fournier, R. E. K. (1989) Genomics 5, 167–176.
AP: Methods
Production of Microcell Hybrids Ann McNeill Killary1 and Steven T. Lott Division of Laboratory Medicine, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030
Microcell fusion is a technology that involves the transfer of single or small clusters of intact chromosomes from one cell to another. The transferred chromosome can be stably retained in the recipient cell background under dominant selective pressure. Hybrid clones generated by this method result in karyotypically simple and homogeneous populations that are excellent resources for physical mapping. This article will describe a general strategy for the efficient micronucleation of human and rodent cell populations and their use as donors for microcell fusion into recipient cell lines. q 1996 Academic Press, Inc.
Parasexual approaches for gene mapping were greatly enhanced with the development of the technology of microcell fusion. In microcell fusion experiments, single, intact chromosomes may be transferred from one cell to another, resulting in karyotypically simple somatic cell hybrids for genetic analysis. Five essential steps are involved in this technology; they include: (1) micronucleation of donor cells; (2) enucleation of micronucleate cell populations; (3) microcell isolation; (4) fusion of microcells to recipient cells; and (5) selection of viable microcell hybrid clones (diagrammed in Fig. 1). In outline, donor cells are exposed to the mitotic inhibitor Colcemid for a prolonged period. Colcemid-induced mitotic arrest results in the formation of micronuclei containing single or small numbers of chromosomes. Micronuclei are then extruded from the donor cell population by exposure to cytochalasin B in the presence of centrifugal force. The cytochalasin B-induced enucleation of micronucleate populations results in the isolation of an assortment of cell particles including microcells, enucleated whole nuclei (karyoplasts), whole cells, and cytoplasmic vesicles. Following enucleation, microcells (usually containing 1–5 interphase chromosomes) are purified and fused to recipient cells. Viable 1 To whom correspondence and reprint request should be addressed at Division of Laboratory Medicine, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Boulevard, Box 72, Houston, TX 77030-4095. Fax: (713) 794-1800.
microcell hybrid clones are then isolated using a variety of selection schemes. This article will describe a general strategy for the efficient micronucleation of human and rodent cell populations and their use as donors for microcell fusion into recipient cell lines.
DONOR CELL MICRONUCLEATION The first step in the technology of microcell fusion involves the prolonged treatment of growing cells with the mitotic inhibitor Colcemid (1–3). Colcemid interferes with the polymerization of microtubules that are necessary for the formation of a functional spindle during metaphase. In the absence of a spindle, chromosomes appear scattered throughout mitotic cells. Prolonged Colcemid arrest results in a fraction of cells in the population that attempt to escape the mitotic block in order to divide. The nuclear membrane, rather than surrounding the entire chromosome complement, reforms around single or small clusters of chromosomes, forming micronuclei (2, 4). A general protocol for the micronucleation of rodent and human cell lines is described below. Rodent Cell Micronucleation by Prolonged Mitotic Arrest Micronucleation of many rodent lines can be accomplished by exposure of growing cells to a prolonged mitotic block (48 h) using Colcemid concentrations in the range 0.01–0.1 mg/ml (5). Optimal concentrations of Colcemid, however, should be determined for each donor cell line. Conditions have been determined for murine L cell lines, including A9, and for microcell hybrids containing single human chromosomes in the A9 cell background (6). A standard protocol for micronucleation of A9-based monochromosomal hybrids involves the plating of donor cells onto thin plastic sheets, termed ‘‘bullets,’’ for micronucleation and subsequent enucleation. 3
1046-2023/96 $18.00 Copyright q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
/ m4819$131d
02-19-96 08:01:24
metha
AP: Methods
KILLARY AND LOTT
4
for 3–4 h to overnight to allow for cell attachment prior to Colcemid addition. Colcemid (Demecolcine, Sigma) is then added to each dish (0.06 mg/ml) for 48 h to induce micronucleation. The percentage of micronuclei obtained may be quantitated using several protocols (outlined in a later section of this chapter).
FIG. 1. The technology of microcell fusion.
Preparation of Bullets for Micronucleation and Enucleation Bullets, which resemble microscope slides with one rounded end, can be cut from the bottoms and sides of T75 or T150 tissue culture flasks using a hot wire cutter. The optimal size of each bullet is such that it fits snugly inside a 50-ml conical centrifuge tube (approximately 25 1 90 mm). Before cell plating, bullets should be washed in glassware detergent, rinsed thoroughly in water, and placed in 95% ethanol for 17–24 h. Bullets are then removed from ethanol using sterile forceps, placed in 150-mm tissue culture dishes (four bullets per dish), and left in the tissue culture hood to dry. Following enucleation, bullets can be soaked in a 3% solution of bleach overnight, washed with detergent, and rinsed in water. Micronucleation on Bullets For micronucleation of A9 cells or A9-based microcell hybrids, one T150 tissue culture flask of logarithmically growing cells at 80–90% confluence is used per experiment. The flask is trypsinized and replated into four 150-mm tissue culture dishes, each containing 4 bullets, in complete medium containing 50 mg/ml gentamycin. The number of bullets used per experiment is dependent on the percentage of micronuclei formed. A9 cells consistently micronucleate at 90–95%. Therefore, we routinely use 16 bullets per experiment for A9 or A9-based hybrids. Tissue culture dishes are left at 377C
/ m4819$131d
02-19-96 08:01:24
metha
Human Cell Micronucleation by Prolonged Mitotic Arrest The most efficient donor cells for human micronucleation are early passage human foreskin fibroblasts (7). Human diploid fibroblasts are also attractive donors for microcell fusion in that they represent early passage cultures with a normal diploid chromosome complement in contrast to established, transformed cell lines that may have undergone many chromosomal rearrangements. In addition, micronucleation of many human cell lines has proved to be difficult (7). Micronucleation of human lymphoblastoid cells has been reported; however, the percentage of micronucleation is low, which necessitates the use of large numbers of donor cells (8). Therefore, for the purposes of this article, the protocol described will be for use with human foreskin fibroblasts. For human cell micronucleation, four T150 tissue culture flasks containing logarithmically growing human foreskin fibroblasts at 80–90% confluence are seeded (1:2) into eight 150-mm tissue culture dishes, each containing four bullets in complete medium with 50 mg/ml gentamycin. Following cell attachment (3–4 h), Colcemid (10 mg/ml) is added to each dish, and cells are incubated at 377C for 48 h to induce micronucleation. Induction of micronucleation in human fibroblasts is critically dependent on the concentration of mitotic inhibitor and the absolute passage number of the donor cells (7). Colcemid concentrations required to induce micronucleation are in the range 10–20 mg/ml for a 48h arrest. Under these conditions, 40–60% of the cell population becomes micronucleate (Fig. 2). Increasing concentrations of Colcemid may result in increased cell toxicity and a decreased yield of micronucleated cells (7). By examination of the number of micronuclei per cell as a function of increasing Colcemid concentration and length of mitotic arrest, these conditions have proved optimal (7). Antikinetochore antibody staining was previously used to examine the numbers of kinetochores per micronucleus (A. McNeill Killary, R. L. Brown, S. Brinner, B. Brinkley, and E. Stubblefield, unpublished results). Results indicated that the smallest of micronuclei contain single chromosomes. Another critical variable for human cell micronucleation is the age of the culture (7). Early passage human foreskin fibroblasts micronucleate optimally; however, after a population doubling of 17–18, the percentage of micronucleate cells decreases dramatically (7). Also, as indicated previously, the length of time in the mi-
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
totic inhibitor is an important consideration for experimental design (6). The length of mitotic arrest is dependent on the growth rate of the donor population cell. Slower growing cell lines may require longer arrest times. We have also observed that the recovery of viable microcell hybrid clones containing intact donor chromosomes is greater in incubations of 48 h compared with a 72-h arrest time. In general, we have also observed that the longer the incubation time in Colcemid, the higher the percentage of microcell hybrids recovered containing fragmented donor chromosomes.
ENUCLEATION OF MICRONUCLEATE CELL POPULATIONS Large-scale enucleation of micronucleate cell populations is accomplished by centrifugation of cells containing micronuclei in the presence of cytochalasin B. Cytochalasin B is a fungal metabolite that interferes with microfilament attachment to the cell membrane (9). High concentrations of cytochalasin B (10 mg/ml) result in nuclear extrusion in a fraction of cells in a population. Mass enucleation of cells was first demon-
strated by incubation in cytochalasin B combined with centrifugation (10–12). During the enucleation process, nuclei are extruded from the cell on thin cytoplasmic stalks and pellet at the bottom of the centrifuge tube (Fig. 3). The cytoplasm of enucleate cells remains attached to the bullets on which the cells are plated. Enucleation of micronucleate cells results in the formation of microcells, consisting of a single micronucleus containing approximately one to five chromosomes surrounded by a small rim of cytoplasm and an intact plasma membrane (13). After enucleation, the pellet in each centrifuge tube contains microcells of various sizes, enucleated whole cells, whole cells that were stripped off the bullet prior to enucleation, and cytoplasmic particles. Following are enucleation conditions for rodent and human cells in culture. Large-Scale Enucleation of Mouse A9 Cells or A9-based Hybrids Using the outlined protocol for the enucleation of A9 cells or A9-based hybrid cells, it is possible to achieve enucleation of ú90–95% of the cell population. A stock solution of cytochalasin B (Sigma) at 1–2 mg/ml is prepared by dissolving the powder in dimethyl sulfoxide (Sigma). The stock solution is then trans-
FIG. 2. Micronucleation by prolonged mitotic arrest. Acridine-orange-stained micronuclei from human diploid foreskin fibroblasts.
/ m4819$131d
02-19-96 08:01:24
metha
5
AP: Methods
KILLARY AND LOTT
6
ferred to sterile conical tubes and wrapped in foil to protect from light. For enucleation, eight 50-ml polycarbonate centrifuge tubes with lids (Nalgene) are autoclaved. To each tube, 38.5 ml of serum-free medium containing 10 mg/ml cytochalasin B is added and allowed to equilibrate at 377C. Aseptically, each of 16 bullets containing micronucleate A9 cells is removed from the tissue culture dish and placed inside a centrifuge tube. Bullets are placed such that 2 bullets are back-to-back inside each 50-ml tube. Lids are secured on tubes with tape. Tubes containing bullets are immediately placed inside a prewarmed (28–337C) superspeed centrifuge with a fixed angle rotor (for example, Sorvall RC-5B centrifuge and SS-34 rotor) such that bullets are parallel to rotor radii. Cells containing micronuclei are then centrifuged at 27,000g (15,000 rpm) for 70 min. Following centrifugation, bullets are removed from tubes and examined (as described in the next section) for the efficiency of enucleation. Bullets are then placed in a 3% bleach solution overnight, washed, and reused for subsequent experiments. Medium from each tube is carefully decanted, and pellets are resuspended. Pellets are redistributed in 1–2 ml serum-free medium and combined into a 15-ml conical tube. Following resuspension, an aliquot is saved for hemocytometer counts and quantification of yield. The tube is then incubated at 377C until fusion. Enucleation of Human Fibroblasts The conditions for human fibroblast enucleation are essentially the same as those outlined for A9 enucle-
ation, with the following exceptions. Optimal micronucleation for early passage human fibroblasts is 40– 60%, as opposed to approximately 95% for A9-based hybrids or A9 cells. We therefore increase the number of bullets used per experiment from 16 bullets as previously described for A9 enucleation to 32 bullets for human fibroblast enucleation. Bullets are centrifuged at 18,800g for 35 min in serum-free medium containing 5 mg/ml cytochalasin B. The remainder of the steps for large-scale enucleation of human fibroblasts are identical to those previously described for A9 cells. Enucleation of Nonadherent Cells on Bullets Procedures have been described for the efficient enucleation of cells that fail to adhere well to the substratum or for cells that grow in suspension (14). In this situation, bullets can still be used for enucleation if they have been pretreated with concanavalin A (6, 14). Concanavalin A (Con A) (Sigma) (15 mg/ml solution in 0.9% saline) is cross-linked to bullets using a crosslinking agent [1-cyclohexyl-3-(2 morpholinoethyl) carbodiimide metho-p-toluene sulfonate, water-soluble carbodiimide (WSC; Sigma)]. WSC solution is prepared at 75 mg/ml in 0.9% saline. For this procedure, bullets are removed from 95% ethanol and placed in tissue culture dishes to dry. Each of the bullets is layered with 0.6 ml of WSC solution followed by 0.6 ml of Con A solution, and the mixture is spread evenly. Bullets are incubated at room temperature for 1 to 2 h. Next, the Con A–WSC solution is aspirated off the bullets, and bullets are then rinsed twice with sterile phos-
FIG. 3. Electron micrograph of an enucleating cell (courtesy of Dr. Ronald L. Brown).
/ m4819$131d
02-19-96 08:01:24
metha
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
phate-buffered saline (PBS). Donor cells are induced to micronucleate in suspension by prolonged Colcemid arrest, as previously described in T75 or T150 flasks. Flasks are harvested by trypsinization, counted, and resuspended in PBS at 106 cells/ml. Then 1.2 ml of cell suspension is added to each bullet, and the cells are allowed to attach for 10–15 min at room temperature. Complete medium is added, and the cells are incubated for 30–60 min at 377C to allow for cell spreading. Con A bullets containing micronucleate cells can then be processed for enucleation as described in the preceding section. Suspension Enucleation Enucleation of suspension cultures can also be accomplished using gradient separation through Ficoll (15) and Percoll (8) gradients. Gradient separation is based on differences in buoyant densities of the nucleoplasm versus the cytoplasm under centrifugal force. Suspension enucleation is especially applicable to cell lines that fail to micronucleate efficiently. Lymphoblastoid cells have been used successfully as donors for microcell fusion despite their poor micronucleation efficiency because large numbers of cells can be used in the gradient. Enucleation in gradients, however, is often incomplete, and many partially enucleated cells will be lost because they partition away from isolated microcells. This approach is, nevertheless, the method of choice for cell lines that cannot be enucleated using bullets. The main variables associated with enucleation are (1) the concentration of cytochalasin B; (2) centrifugation speed; and (3) centrifugation time. The concentration of cytochalasin B is usually in the range 5–10 mg/ ml for both human and rodent lines. The variables that may need to be adjusted for each cell line are centrifugation time and speed. Careful monitoring of the steps in enucleation is imperative to achieve the optimal yield of microcells. Enucleation should result in virtually 99% nuclear extrusion. If incomplete enucleation occurs, then incremental increases in either time or speed can be monitored for complete enucleation. However, it should be noted that excess centrifugal force or time can result in the complete stripping of cytoplasm from the bullets.
QUANTITATION OF MICROCELLS This section will describe a general protocol for quantitation of microcell yield following enucleation. This procedure should also be used to stain bullets to determine the percentage of micronucleate cells in the population as well as to determine the efficiency of enucleation.
/ m4819$131d
02-19-96 08:01:24
metha
7
Following enucleation, the pellets that are recovered consist of a variety of types and sizes of cell particles, including microcells, intact nuclei, whole cells that stripped off the bullets prior to enucleation, and cytoplasmic particles. Hemocytometer counts can be used to determine the total number of particles per experiment. However, to determine the microcell particle yield, staining of the preparation is necessary to discriminate between nuclear and cytoplasmic particles. Aceto-orcein will differentially stain the nucleus from the cytoplasm and is used in conjunction with bright-field microscopy. Using this stain, nuclei appear red and the cytoplasm appears pink. An aceto-orcein solution is prepared by mixing 0.5% orcein solution (Sigma) in 50% acetic acid followed by refluxing this mixture for several hours. To quantitate microcell yield following enucleation, one drop of microcell preparation is mixed with one drop of aceto-orcein stain on a microscope slide. The preparation is coverslipped, stained for 2–5 min, and examined by bright-field microscopy. The staining time is critical for accurate determinations of nuclear versus cytoplasmic fragments. After an incubation time of approximately 5–10 min, all particles stain red. Thus, the interpretation of aceto-orceinstained material is sometimes difficult because discrimination between shades of the same color is required rather than visualization using fluorochromes of contrasting colors. The advantage of this staining protocol is its simplicity; visualization does not require a fluorescence microscope. A similar approach for the quantitation of microcell yield involves the use of the fluorochrome acridine orange. Acridine orange stains the nucleus and cytoplasm differentially in that nuclei and micronuclei fluoresce green, whereas cytoplasmic particles appear red using fluorescence microscopy. This stain is performed essentially as described for the aceto-orcein protocol and is an excellent method for quantitation of micronucleation, enucleation efficiency, or microcell yield. For this method, a stock solution of acridine orange is prepared by dissolving powder in 2 mg/ml sterile water. Acridine orange is a carcinogen; therefore, care should be taken in its handling. The stock solution is stored in a foil-wrapped conical tube at 47C. The working solution of acridine orange is 10 mg/ml prepared in water or normal saline. A foil-wrapped coplin jar is suitable for processing several bullets or slides for staining. To determine the efficiency of micronucleation or enucleation, bullets are removed from medium and placed in a coplin jar containing a 1:1 mixture of 95% methanol and serum-free medium for 10 min. Bullets are then transferred to a second coplin jar containing 95% methanol for 10 min and allowed to dry. Each bullet is then placed in the acridine orange stain for 30–60 s. Bullets are then rinsed twice with water and
AP: Methods
KILLARY AND LOTT
8
coverslipped using 0.05 M citrate phosphate mounting buffer, pH 4.1. The accurate discrimination of nucleus from cytoplasm is accomplished by screening bullets using fluorescence microscopy.
ISOLATION OF MICROCELLS There are several different strategies for purification of the microcell population following enucleation. This step in the microcell fusion technique is important if the goal is to transfer single chromosomes. Nuclepore Filtration The simplest strategy to purify microcells is to filter the crude microcell preparation through nuclepore filters to selectively isolate the smallest microcells in the preparation and to exclude contaminating whole cells and nuclei (7). For filtration, various sized nuclepore filters (8, 5, or 3 mm) are placed inside autoclaved Swinnex filter units (Swinnex, 25 mm) attached to sterile 10-cc syringes. Following enucleation, the crude microcell preparation is resuspended in a total volume of 10–15 ml serumfree medium. One-half of the crude preparation is poured into the syringe containing a 5-mm nuclepore filter and allowed to filter through gravity with only slight pressure applied as needed on the plunger. This step is repeated with the remaining half of the microcell preparation, using a new 5-mm filter set. Nuclepore filtration should result in exclusion of most of the whole cells and nuclei from the crude preparation. Preparations containing large numbers of intact cells should be passed through 8-mm filters prior to 5-mm filtration. The remaining preparation should contain various sizes of microcells and cytoplasmic particles. To select for the smallest microcells, which likely contain single chromosomes, the preparation is filtered again through two 3-mm filter units exactly as described previously for 5-mm separation. Careful monitoring of the filtration is required to ensure that the filter remains intact during the procedure. Excess pressure on the plunger could result in cracking of the filter. To check for the efficiency of the filtration step, aliquots are taken before and after filtration for quantitation using acridine orange or aceto-orcein. Using this filtration step, the resultant purified preparation should contain only the smallest of microcells, with complete elimination of whole cells and 99% of karyoplasts. The efficiency of this procedure is very much dependent on the proper use of the filter. If the filter is overloaded with the crude preparation, then clogging of the filter occurs. This results in a decrease in the total particles processed through the filter. Hemocytometer counting prior to filtration will allow the
/ m4819$131d
02-19-96 08:01:24
metha
optimal number of total particles ((1–5 1 106)/ml) to be passed through each filter unit. Microcell Purification via Unit Gravity Sedimentation Microcells can also be isolated using unit gravity sedimentation on density gradients (5). For unit gravity separation, a linear gradient of 1–3% bovine serum albumin (BSA) is prepared using a gradient mixing device connected to a peristaltic pump. Tubing is connected from the gradient mixing device through the peristaltic pump to a three-way valve. One valve outlet is connected to a 5-cc syringe that functions as a bubble trap. The second valve is connected to a 50-cc syringe that functions as the gradient chamber. Three glass beads are placed over the inlet of the chamber. Next, a small piece of tubing is connected to the other outlet, which serves as the sampling tube. The entire apparatus can be autoclaved. Next, sterile PBS is added to the gradient mixing device and pumped in to fill the tubing just at the bottom of the gradient chamber. Excess PBS is pumped into the bubble trap. To one side of the mixing device, 25 ml of 3% BSA in PBS is added, and to the other side, 25 ml of 1% BSA in PBS (containing phenol red) is added. The connection between the chambers is then opened and the stirrer motor started. Following initiation of the motor, the microcell preparation is resuspended in 2 ml of 0.5% BSA in PBS and added to the gradient chamber from the top. The pump is started and PBS in the lines is pumped into the bubble trap. When the BSA solution (red) reaches the trap, the valve is turned to allow the solution to enter the gradient chamber. The pumping rate for the BSA gradient should be 2–3 ml/min. The microcell preparation is then allowed to settle through the gradient for 3–3.5 h, which separates the microcells, whole cells, nuclei, and cytoplasmic particles by size. Fractions are collected by dripping through the sample tube. This type of separation efficiently separates whole cells and nuclei in the bottom 15–20 ml of the 50-ml gradient. The purified microcell preparation is recovered from the top 20–25 ml. Unit gravity sedimentation is probably the most efficient method for purification of the crude microcell preparation. This technique virtually eliminates whole cells and nuclei from the crude preparation. It is, nevertheless, a time-consuming procedure and therefore may not be the method of choice unless absolute purification of the preparation is needed.
FUSION OF MICROCELLS TO RECIPIENT CELLS In initial strategies for microcell fusion, inactivated Sendai virus was used to fuse microcells to recipient
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
cells (5). Later, protocols proved to be more simple and efficient by using a combined treatment of cells with phytohemagglutinin-P (PHA-P) to adhere microcells to recipient cells, followed by fusion using polyethylene glycol (PEG) (7, 14). Following are two protocols for microcell fusion. The first protocol involves the agglutination of microcells to a recipient monolayer using PHA-P, followed by fusion using PEG. The second method is a suspension protocol for use with cells that do not adhere to monolayer culture or for use when PHA-P is toxic to recipient cells. Two important variables need to be determined for each recipient cell line prior to microcell fusion. The first is the concentration of fusogen. This can be determined by simply fusing recipient cells in culture in increasing concentrations of fusogen. The conditions that yield the highest percentage of binucleate heterokaryons postfusion with the least amount of cell toxicity are optimal. Optimal fusogen concentrations are usually in the range 44–50% PEG w/w (Koch Chemical, United, MW 1540). Higher concentrations of fusogen can result in large numbers of fused cells en masse, with increased cell toxicity. PHA-P toxicity experiments should also be performed in increasing concentrations of the agglutinin in the range 50–200 mg/ml. In this case, the optimal conditions for PHA-P are such that cell–cell agglutination occurs with the least amount of cell toxicity. Agglutination can be monitored by phase-contrast microscopy, and the degree of cell toxicity can be measured by trypan blue staining. The following is a general protocol for monolayer microcell fusion. PEG is prepared in serum-free medium. For fusion to a recipient monolayer of A9 cells, a concentration of 50% w/w is optimal. The PEG solution is incubated at 377C until PEG has dissolved and then filter sterilized. The pH of the PEG solution should be in the range 7.5–8.0. PHA-P (Difco) is a lyophilized powder that is 50% salt by weight. The concentration of PHA-P used routinely for microcell fusion to a monolayer of A9 recipients is 100 mg/ml in serum-free medium. This solution can also be filter sterilized. PHA-P/PEG Monolayer Fusion Monolayer cultures of recipient cells are prepared in 25-cm2 flasks. Routinely, two flasks are used for microcell fusion, and one flask is reserved as a control. Recipient cell cultures should be subcultured 4–24 h prior to fusion to allow for firm attachment to the substratum. Each flask should be at 70–80% confluence at the time of fusion. The purified microcell preparation is then centrifuged at 2000g for 10 min (377C). The microcell pellet is resuspended in 4 ml PHA-P solution (100 mg/ml). At this step, it is important to redistribute the solution well to disperse any clumps of microcells prior to layering the solution onto the monolayer. Medium is then
/ m4819$131d
02-19-96 08:01:24
metha
9
aspirated off the recipient cell monolayers, and 2 ml of microcell/PHA-P solution is added to each of two T25 fusion flasks. Flasks are incubated at 377C until microcells have agglutinated to the monolayer (usually 10– 20 min). Next, the excess medium is carefully aspirated off the fusion monolayers, 0.5 ml PEG (50% w/w) is added, and the flask is gently rocked for 1 min. Following the 1-min incubation, the PEG solution is quickly aspirated off the monolayer, which is next rinsed three times with 4 ml serum-free medium. Complete, nonselective medium is then added, and flasks are returned to 377C for 17–24 h prior to addition of selective medium. The control T25 is treated as above with the exception that it does not receive donor microcells. Following incubation, fusion flasks are trysinized and replated into 10–20 T25s per fusion. Selective medium is then added to each flask. Viable microcell hybrids should be visible within 2–4 weeks. Suspension Microcell Fusion Microcells can be fused to recipient cell lines in suspension. This protocol does not use PHA-P and is appropriate for cell lines that grow in suspension or for which PHA-P is cytotoxic. For suspension fusion, recipient cells are counted and resuspended at 1 1 106 cells/ml in 5 ml serumfree medium. To this suspension, the filtered microcell preparation is added in a 15-ml conical centrifuge tube. The microcell/cell suspension is then centrifuged at 2000g for 15 min at 377C. The resulting pellet should be well dispersed. To this pellet, 0.5 ml PEG (50% w/ w) is added dropwise while gently resuspending the pellet for 1 min. Immediately, 9.5 ml of serum-free medium is added over 1 min with gentle swirling. Next, 20 T25s/fusion are prepared in complete nonselective medium. Twenty flasks are also prepared for the control. Following fusion, 0.5 ml of fusion or control suspension is aliquoted into each T25. Flasks are then incubated overnight (17–24 h) at 377C. Fusion and control flasks are then refed with complete selective medium and allowed to incubate at 377C until clones become visible. Both the monolayer and the suspension protocols have successfully resulted in the generation of viable microcell hybrid clones. The monolayer protocol for fusion is much more efficient than the suspension protocol, with transfer frequencies in the range 0.5–2.0 1 1005. The suspension protocol is the method of choice for cell lines that grow in suspension or adhere poorly to the monolayer or for which PHA-P is cytotoxic.
SELECTION STRATEGIES FOR THE ISOLATION OF MICROCELL HYBRID CLONES Microcell hybrid clones containing single introduced chromosomes can be isolated under dominant selective
AP: Methods
KILLARY AND LOTT
10
pressure. Microcell hybrids have been used for complementation mapping, in which the recipient cells harbor a recessive, conditional lethal mutation that is complemented by the introduction of a wild-type copy of that gene on the introduced chromosome (16). The X chromosome, human chromosome 17, or mouse chromosome 11 can be selected using the nucleotide salvage pathway complementation called the HAT selective system (17). Also, adenine phosphoribosyl transferase (APRT) complementation via the AAT selective system can select for human chromosome 16 or mouse chromosome 8 (18). The most conceptually ideal system for microcell fusion would be one in which each donor chromosome contains a dominant selectable marker (7). Dominant selectable markers have been successfully transferred into donor chromosomes prior to micronucleation and then transferred into a variety of recipient lines (19– 22). The most popular vectors containing dominant selectable markers are pSV2neo, a plasmid vector containing the bacterial gene neo that confers resistance to the antibiotic G418 (23), and pSV2gpt, a similar vector containing the bacterial gene gpt that can be selected in HPRT0 cells using HAT selection or in wild-type back cells using mycophenolic acid (24). Retroviral vectors (pZIP-neo-SV(X1)) have also been used to efficiently tag donor chromosomes for microcell fusion (21). Retroviral transfer has an advantage in that the vector can be inserted into a single site per cell, and high frequency transduction is possible. Our laboratory has generated a standard protocol for the tagging of human chromosomes and subsequent transfer of marked human chromosomes into recipient A9 cells (6, 22). We use the technique of electroporation (25–27) as a simple, straightforward way to introduce pSV2neo into human diploid fibroblasts. Electroporation has distinct advantages over other systems in that, unlike calcium phosphate transfection, the DNA transferred is low copy number, the integration sites are limited, and the procedure can be used without the biosafety concerns associated with the use of retroviral vectors. For our experiments, the method of Potter (27) was modified for human foreskin fibroblasts and for use with a commercial electroporator (Bio-Rad gene pulser). Following is a general strategy for the tagging of normal diploid human fibroblasts with pSV2neo and subsequent selection in the antibiotic G418. Electroporation of pSV2neo Into Diploid Human Fibroblasts Human diploid fibroblasts (1 1 107 cells) are mixed with linearized plasmid DNA (10 mg/ml of pSV2neo) in PBS and incubated on ice for 10 min. A 2000-V electrical pulse is applied to the cell/DNA mixture, which is then incubated on ice for 10 min postelectroporation.
/ m4819$131d
02-19-96 08:01:24
metha
The suspension is replated in complete nonselective medium at 5 1 105 cells per T25. After a 48-h incubation at 377C, selective medium containing the antibiotic geneticin (G418) at 750 mg/ml is added to the medium to select for cells carrying tagged chromosomes. Clones can be observed at 10 days postelectroporation. Properties of Microcell Hybrid Clones Microcell hybrid clones can be isolated from flasks by first removing the tops of flasks with a soldering gun and then placing cloning rings over individual clones. Microcell hybrid clones, in general, have sizes and morphologies similar to that of the recipient cell line. They can be readily distinguished from whole-cell hybrid clones containing many introduced chromosomes, which are usually much larger. In general, microcell hybrids will stably retain the introduced chromosome as long as selective pressure is maintained. This results in a nearly homogeneous hybrid population for genetic studies. This property of microcell hybrids makes them ideal for quantitative somatic cell genetic studies that require high percentages of the introduced chromosome in the recipient cell population. Monochromosomal hybrids have a number of advantages over whole-cell hybrid clones because they necessarily contain only the introduced chromosome, as opposed to segregating whole-cell hybrids, in which other chromosomes, translocations, and chromosomal fragments could be a part of the hybrid genome. The stability of the introduced chromosome in the microcell hybrid is dependent to a certain extent on the individual recipient cells used (16). The isolation of microcell clones containing intact transferred chromosomes is much lower in certain recipient backgrounds, such as HT1080 and CHO cells (16). Microcell fusion is, to date, the only reliable strategy for the transfer of single, intact chromosomes. Microcell hybrids have also been extremely useful as a source of hybrids containing fragments of the introduced chromosome (16, 28). Together, monochromosomal microcell hybrids and fragment-containing hybrids are extremely valuable tools both for physical mapping in both interspecific and intraspecific crosses and for quantitative studies such as the identification of tumor suppressor genes and genes involved in tissue-specific gene regulation.
REFERENCES 1. Levan, A. (1954) Hereditas 40, 1–64. 2. Stubblefield, E. (1964) (Harris, R. J. C., Ed.), in Cytogenetics of Cells in Culture pp. 223–298, Academic Press, New York. 3. Phillips, S. G., and Phillips, D. M. (1969) J. Cell Biol. 40, 248. 4. Ege, T., Ringertz, N. R., Hamberg, H., and Sidebottom, E. (1977)
AP: Methods
PRODUCTION OF MICROCELL HYBRIDS
5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.
in Methods in Cell Biology (Prescott, D. M., Ed.), Vol. XV, pp. 339–358, Academic Press, New York. Fournier, R. E. K., and Ruddle, F. H. (1977) Proc. Natl. Acad. Sci. USA 74, 319–323. Killary, A. M., and Fournier, R. E. K. (1995) in Methods in Enzymology, Academic Press, San Diego, in press. McNeill, C. A., and Brown, R. L. (1980) Proc. Natl. Acad. Sci. USA 77(9), 5394–5398. Sanford, J. A., and Stubblefield, E. (1987) Somat. Cell Mol. Genet. 13, 279. Carter, S. B. (1967) Nature 213, 261–264. Prescott, D. M., Myerson, D., and Wallace, J. (1972) Exp. Cell Res. 71, 480–485. Poste, G., and Reeve, P. (1972) Exp. Cell Res. 73, 287–294. Wright, W. E., and Hayflick, L. (1972) Exp. Cell Res. 74, 187– 194. Ege, T., and Ringertz, N. R. (1974) Exp. Cell Res. 87, 378–382. Fournier, R. E. K. (1981) Proc. Natl. Acad. Sci. USA 78(10), 6349–6353. Wigler, M. H., and Weinstein, I. B. (1975) Biochem. Biophys. Res. Commun. 63, 669–674. Fournier, R. E. K., and Moran, R. G. (1983) Somat. Cell Mol. Genet. 9, 69–84.
/ m4819$131d
02-19-96 08:01:24
metha
11
17. Littlefield, J. W. (1964) Science 145, 709–710. 18. Kusano, T., Long, C., and Green, H. (1971) Proc. Natl. Acad. Sci. USA 68, 82–86. 19. Athwal, R., Smarsh, M., Searle, B. M., and Surinder, S. D. (1985) Somat. Cell Mol. Genet. 11, 177–187. 20. Saxon, P. J., Srivatsan, E. S., Leipzig, G. V., Sameshima, J. H., and Stanbridge, E. J. (1985) Mol. Cell. Biol. 5, 140–146. 21. Lugo, T. G., Handelin, B., Killary, A. M., Houseman, D. E., and Fournier, R. E. K. (1987) Mol. Cell. Biol. 7, 2814–2820. 22. Killary, A. M., Wolf, M. E., Giambernardi, T. A., and Naylor, S. L. (1992) Proc. Natl. Acad. Sci. USA 89, 10877–10881. 23. Southern, P. J., and Berg, P. (1982) J. Mol. Appl. Genet. 1, 327– 341. 24. Mulligan, C., and Berg, P. (1980) Science 209, 1422–1427. 25. Zimmerman, U., and Vienken, J. (1982) J. Membr. Biol. 67, 165– 182. 26. Neumann, E., Schaefer-Ridder, M., Wang, Y., and Hofschneider, P. H. (1982) EMBO J. 1, 841–845. 27. Potter, H., Weir, L., and Leder, P. (1984) Proc. Natl. Acad. Sci. USA 81, 7161–7165. 28. Leach, R. J., Thayer, M. J., Schafer, A. J., and Fournier, R. E. K. (1989) Genomics 5, 167–176.
AP: Methods