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Metaphase Chromosome Preparation from Cultured Peripheral Blood Cells

UNIT 4.1

Chromosome preparations currently provide the only direct view of the genome as a whole. Although molecular methods allow a more detailed analysis of specific regions of the genome, the study of genetics is not complete without an appreciation of the metaphase cell. Peripheral blood provides a simple and reproducible source for large quantities of mitotic cells that may be used for clinical diagnosis (see UNITS 4.2 & 8.1) or studying chromosome structure and organization (see UNIT 4.3). From a clinical point of view, the metaphase cell reveals not only numerical and structural abnormalities, but also provides some biochemical information useful in diagnosing genetic disorders. For instance, the DNA instability associated with Fanconi anemia (UNIT 8.7) appears as increased chromosome breakage in metaphase cells; Bloom’s syndrome (UNIT 8.6) is manifested as increased sister chromatid exchange; and ataxia telangiectasia may be spotted in metaphase chromosome preparations as infidelity in somatic recombination. Likewise, other syndromes associated with abnormal chromosome behavior include Roberts/SC phocomelia syndrome (OMIM #269000), in which centromeric heterochromatin prematurely separates, and the immunodeficiency–centromere instability– facial anomalies syndrome (OMIM #242860), which also includes centromeric puffing and endoreduplication with breakage in centric heterochromatin. Chromosome preparations are essential for localizing genes and other DNA sequences on the physical map. This mapping is done directly by chromosomal in situ hybridization or indirectly by correlating the presence or absence of a marker (such as an enzyme, a particular RNA, or a DNA sequence) with the presence or absence of a chromosomal region. Chromosomal translocations have proven to be valuable in identifying the locations of genes for rare genetic syndromes. The identification of recurring chromosome abnormalities also plays a key role in diagnosing hematologic leukemia and lymphomas in both bone marrow and peripheral blood. The stimulated T cell system described here is the most widely used means of obtaining large numbers of mitotic cells for genetic analyses. Synchronization of the cell cycle in culture (Basic Protocol), combined with direct inhibition of chromosome condensation (Alternate Protocol), yield longer high-resolution prophase or prometaphase preparations. Such preparations are used for detailed analysis of microdeletions (UNIT 8.10) or subtle rearrangements, fine breakpoint analysis, and refined mapping. Although chromosomes may be obtained from other cells, human peripheral blood leukocytes are most amenable to synchronization, and thus to high-resolution analysis of their chromosomes. Microscope slide preparation of mitotic chromosomes from harvested cell culture suspensions is also explained in the Support Protocol. NOTE: All reagents and equipment coming into contact with live cells must be sterile.

CULTURE AND METAPHASE HARVEST OF PERIPHERAL BLOOD T lymphocytes in whole blood are stimulated with the mitogenic plant lectin phytohemagglutinin (PHA). The T lymphocytes “activate” to blast-like cells within 12 to 24 hr and continue to proliferate for 2 to 4 days. An optional cell-cycle synchronization step can be performed (see Fig. 4.1.1), which increases the number of cells harvested in early mitosis. (This synchronization is not as involved as the one described in the alternate protocol, which results in longer chromosomes and higher resolution). Metaphase

BASIC PROTOCOL

Cytogenetics Contributed by Charles D. Bangs and Timothy A. Donlon Current Protocols in Human Genetics (2005) 4.1.1-4.1.19 C 2005 by John Wiley & Sons, Inc. Copyright

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cells are obtained by treating cultures with Colcemid, a colchicine analog that disrupts the centriole/spindle-fiber complex by interfering with microtubule formation. This treatment results in mitotic arrest, which in turn leads to an accumulation of cells in metaphase. Mitotic arrest is followed by treatment with a hypotonic KCl solution (hypotonic “shock”) to increase cellular volume. The cells are then fixed with methanol/acetic acid to remove water and disrupt cell membranes before being spread onto slides (Support Protocol).

Materials Heparinized whole blood obtained via Vacutainer (Becton Dickinson) or syringe with preservative-free sodium heparin (25 U/ml) Complete RPMI/10% FBS medium (APPENDIX 3G) containing 50 µg/ml gentamycin sulfate in place of penicillin and streptomycin 100× phytohemagglutinin-M (PHA) stock (GIBCO/BRL), reconstituted in sterile deionized water (store at 4◦ C) 10 µM methotrexate (optional; see recipe) 1 mM thymidine (optional; see recipe) 10 µg/ml Colcemid (GIBCO/BRL) 75 mM KCl (0.56 g in 100 ml H2 O; store ≤2 weeks at room temperature) Fixative: 3:1 (v/v) HPLC-grade absolute methanol/glacial acetic acid, (prepare fresh) 15-ml sterile disposable conical polypropylene centrifuge tubes TB syringe equipped with 21-G needle (VWR Scientific) IEC HN-SII centrifuge with 958 rotor (or equivalent) CAUTION: Human blood and methotrexate are hazardous; see APPENDIX 2A for guidelines on handling storage, and disposal. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Collect sample and initiate cultures 1. Collect peripheral blood by venipuncture into a sodium heparin Vacutainer or a syringe with 25 U preservative-free sodium heparin per milliliter of blood. Other anticoagulants, such as lithium heparin or EDTA are toxic to cells and should never be used. Samples should be shipped at room temperature. Blood in sodium heparin can be held for ≤4 days and still be cultured successfully, but cultures are best initiated as soon as possible. If necessary, the specimen can be stored at 4◦ C.

2. Inoculate 0.25 ml of the whole blood obtained in step 1 (0.2 ml for newborns ≤3 weeks old) into a sterile 15-ml centrifuge tube containing 5 ml complete RPMI/10% FBS medium, using a TB syringe equipped with a 21-G needle. Add 0.05 ml of reconstituted 100× PHA solution. Always replace the 25-G needle supplied with most TB syringes with a 21-G needle. Forcing blood through the narrower needle can lyse leukocytes. A single culture typically yields three to five full-slide preparations, or more if only part of the slide is used. Multiple cultures may be set up to meet clinical or research needs.

3. Incubate 2 to 4 days with tubes tilted at 45◦ in order to promote air exchange.

Chromosome Preparation from Peripheral Blood

Three-day incubations are optimal, but 2- or 4-day cultures can be used to accommodate laboratory scheduling concerns. Cultures from newborns will usually work well at 2 days but may also be harvested either directly or following a 1-day culture. Older patients’ leukocytes require 3- or 4-day cultures because they do not seem to respond as quickly to PHA stimulation.

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Current Protocols in Human Genetics

Tilting culture tubes to 45◦ increases gas exchange and prevents cells from packing too densely at the bottom. For these reasons it is best not to add more than 0.5 ml blood to any tube. If larger cultures are desired, they can be grown in T25 flasks using a similar ratio of blood to media.

Synchronize the cell cycle (optional) Synchronization (Fig. 4.1.1) is optional for routine blood culture and harvest, but will result in longer chromosomes on average and more mitotic cells. The synchronization presented here (steps 4 and 5) will yield a greater number of mitotic cells; however, these are not as long as those obtained using the high-resolution method (Alternate Protocol). 4. On the day before harvest (e.g., after 4:00 p.m.), add 0.05 ml of 10 µM methotrexate (10−7 M final) to block DNA replication. Incubate 16 to 18 hr (overnight). 5. On the following day (e.g., at 8:30 a.m.), add 0.05 ml of 1 mM thymidine (10−5 M final) to release the methotrexate block. Incubate ∼4 hr (e.g., until 12:30 p.m.). The exact hour of synchronization and release in steps 4 and 5 are not critical. However, because of methotrexate toxicity, the culture should be blocked no longer than 18 hr. The exact interval between release of block and harvest is critical, because cells will progress quickly through G2, which lasts for 4 to 6 hr, to metaphase, which lasts ≤1 hr.

Figure 4.1.1 Cell culture synchronization. Two- or three-day asynchronous lymphocyte cultures are blocked by addition of methotrexate, a folate antagonist that prevents thymidine synthesis. Depletion of the thymidine pool prevents the cells from completing replication, and cells accumulate in the synthetic, or S, phase of the cell cycle. Subsequent addition of thymidine releases a synchronized wave of cells to complete replication and proceed through G2 and into mitosis.

Cytogenetics

4.1.3 Current Protocols in Human Genetics

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Harvest culture 6. Initiate harvest by adding 25 µl of 10 µg/ml Colcemid (0.05 µg/ml final). Incubate 30 min. For a 5-ml culture, this approximates two drops from a TB syringe equipped with a 25-G needle. If synchronization steps 4 and 5 have been omitted, the harvest can be initiated at any time 3 to 4 days following the culture described in step 3.

7. Centrifuge 8 min at 180 × g (1000 rpm in IEC 958 rotor), room temperature. Discard supernatant. 8. Add 6 ml of 75 mM KCl at room temperature and gently resuspend cells. Let stand 15 min at room temperature. The amount of hypotonic solution to be added should be adjusted to the volume of the pellet. Some laboratories vary the length of hypotonic treatment. Increasing the time will increase chromosome spreading, but this treatment is a hypotonic “shock,” so that increasing the amount of hypotonic solution will have more impact than increasing the time of treatment.

9. Add 10 to 12 drops of fixative with a Pasteur pipet and mix well. Centrifuge as in step 7. This treatment serves to reduce the pH of the cells gradually to precondition them for the following fixation steps. It also lyses remaining red blood cells and begins the process of clearing resulting cellular debris.

10. Remove all but 0.5 ml of the supernatant and resuspend pellet in remaining supernatant by drawing it gently up and down with a Pasteur pipet. Add 1 ml fixative and immediately mix gently. Adjust volume to 5 ml with fixative and mix thoroughly. Centrifuge as in step 7. The pellet after step 9 will be brown and clumpy because of erythrocyte debris. Resuspend gently but thoroughly to avoid clumped lymphocytes which may complicate slide-making. Do not draw too much volume into the pipet while resuspending because the cells will stick permanently to glass. Do not press the pipet tip against the bottom of the tube when drawing and delivering the suspension, as this will lyse cells. The pellet after step 10 will be more homogeneous, and will usually have a light-brown to white color. It may be ≤0.1 ml in volume.

11. Aspirate supernatant, resuspend pellet in 5 ml fixative, and centrifuge as in step 7. 12. Remove supernatant and resuspend pellet in a volume of fixative sufficient to produce a light milky suspension. Allow to stand 30 min at room temperature or store overnight at 4◦ C. Longer fixation will often improve chromosome spreading in difficult harvests. Keeping the suspension overnight at 4◦ C can improve the quality of the preparation or can be done for scheduling reasons. Suspensions should be kept in polypropylene tubes containing plenty of fixative (e.g., 5 ml). Polystyrene tubes will react with fixative and should not be used.

13. Prepare slides and analyze chromosome spreads as described in the Support Protocol.


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CULTURE AND HARVEST FOR HIGH-RESOLUTION PROMETAPHASE CHROMOSOMES

ALTERNATE PROTOCOL

Longer chromosomes provide higher resolution in analyses based on chromosome banding. Chromosomes condense and bands coalesce as cells progress through metaphase (Fig. 4.1.2); chromosome length and hence resolution of chromosome analyses are therefore functions of how early in mitosis fixation is performed. It is possible to obtain high-resolution cell harvests (i.e., harvests enriched for mitotic cells at or above the level of 750 G-bands per haploid karyotype) by a combination of cell-cycle synchronization and inhibition of chromosome condensation. In this protocol, synchronization is accomplished by blocking mitosis in cultures during the synthetic, or S, phase of the cell cycle. Methotrexate (amethopterin), a competitive inhibitor of dihydrofolate reductase, is used for this purpose and acts to reduce the levels of available thymidine, which is required for DNA synthesis. The methotrexate block is released by addition of thymidine and a series of sequential harvests are performed at short intervals to capture the wave of late-prophase or prometaphase cells, which will have the longest chromosomes. A short (10-min) mitotic arrest averts excessive chromosome condensation. Condensation may be reduced further by addition of the DNA-intercalating fluorochrome ethidium bromide, just before harvest.

Additional Materials (also see Basic Protocol) 1.25 mM ethidium bromide solution (see recipe) CAUTION: Ethidium bromide is hazardous; see APPENDIX 2A for guidelines on handling, storage, and disposal. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. 1. Prepare blood and initiate four cultures as in steps 1 and 2 of the basic protocol. Three cultures will be used for sequential harvest. The fourth culture is a backup in the event that the sequential harvests do not work well. All cultures will be treated with ethidium bromide in step 5. The backup culture will be treated with a 30-min Colcemid arrest as described in the basic protocol and step 6 below.

Figure 4.1.2 A partial karyotype of G-banded chromosome 7 from cells at successive stages of mitosis, illustrating the coalescing of subbands into larger, less defined, and less informative landmark bands. Cells captured in late prophase may show ≥850 bands per haploid karyotype, i.e., high resolution. By mid-metaphase, fine band detail is lost as chromosomes condense and bands fuse, and only 400 or fewer bands per haploid karyotype may be observed.

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Figure 4.1.3 High-resolution harvest. Prometaphase chromosomes are obtained by sequential harvests of synchronized cultures (1, 2, and 3), timed to capture a wave of cells early in mitosis, when the chromosomes are longest. Staggered 10-min mitotic arrests span a 30-min harvest window. The short mitotic arrest prevents excessive chromosome condensation. Addition of ethidium bromide ∼1 hr before mitotic arrest directly inhibits chromosome condensation. In a typical highresolution harvest, culture 1 may yield little or no mitoses, culture 2 may capture an early mitotic wave with long chromosomes, and culture 3 mitoses are shorter, having proceeded further into metaphase.

2. Incubate cultures 3 or 4 days with tubes tilted to 45◦ . 3. On the day prior to harvest (e.g., after 4:00 p.m.), add 0.05 ml of 10 µM methotrexate (10−7 M final) to block DNA replication. Incubate 16 to 18 hr.


The exact duration of the block is not critical but should be 16 to 18 hr to avoid methotrexate toxicity.

4. The next morning (e.g., at 8:30 a.m.), release the cultures from the methotrexate block by adding 0.05 ml of 1 mM thymidine (10−5 M final). Incubate 3 hr.

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5. Add 0.05 ml of 1.25 mM ethidium bromide solution (1.25 × 10−5 M final) to each culture. Incubate as described in steps 6 and 7. 6. After 50 min (e.g., at 12:20 p.m.) add 25 µl of 10 µg/ml Colcemid (0.05 µg/ml final) to the culture chosen for the first sequential harvest and the backup culture. Incubate the first harvest tube 10 min, then perform steps 7 through 12 of the basic protocol. 7. After an additional 10 min (e.g., at 12:30 p.m.), repeat step 6 using the second harvest culture. IMPORTANT NOTE: It is critical to stay on time! To maintain the 10-min intervals, it will be necessary to shorten all 8-min centrifugations described in the basic protocol (starting with that in step 7) to 6 min, and the 15-min hypotonic treatment in step 8 of the basic protocol to 12 min. By doing these sequential harvests, a 30-min period is spanned by three 10-min mitotic arrest intervals (Fig. 4.1.3). One of these arrest intervals should capture an early mitotic cell population with chromosomes that are less condensed than the others, providing optimal resolution. The backup culture spans the entire 30-min period. The optimum length of time between addition of thymidine (step 4) and initiation of harvest with Colcemid can vary between laboratories. It may be determined empirically by adding several extra 10-min mitotic arrest intervals so that the overall harvest period extends for a longer period (e.g., 50 min) and observing which interval results in the longest chromosomes.

8. Prepare slides and analyze chromosome spreads as described in the Support Protocol.

CHROMOSOME SLIDE PREPARATION Slide-making is the least standardized and understood of cytogenetic protocols, about which technologists have widely variable and sometimes contradictory ideas. In the end what really matters is that slide preparations are consistent and appropriate for the desired analysis. The protocol presented here is not the only approach to chromosome slide preparation but it works under varied physical conditions (slide-making is very climate-dependent) and for a wide range of cell cultures. It can be used for peripheral blood, bone marrow (UNIT 10.2), ascites and pleural effusions, amniotic fluid (UNIT 8.4) and tissue flask harvests (UNITS 8.3, 8.5 & 10.3), somatic cell or radiation hybrids (UNITS 3.2 & 3.3), lymphoblastoid cell lines, and nonhuman and hybridoma cultures—in short, any culture harvest that results in a fixed suspension of mitotic cells.

SUPPORT PROTOCOL

Harvested peripheral blood cultures suspended in methanol/acetic acid fixative (basic and alternate protocols) are applied to wet microscope slides, flooded with fixative, and air-dried. The drying process is adjusted according to ambient temperature and humidity to optimize spreading and morphology of chromosomes for subsequent banding and analysis. The protocol described here produces preparations that are particularly suitable for analysis by G-banding (UNIT 4.2) or in situ hybridization (UNIT 4.3), although many other staining techniques or procedures may be used.

Materials Fixed cultures (basic or alternate protocols) Fixative: 3:1 (v/v) methanol/acetic acid (use 100% methanol and glacial acetic acid, both AR grade, from J.T. Baker) Microscope slides (one end frosted) stored in 100% methanol (absolute AR grade, J.T. Baker) in Coplin jars Lint-free tissue (e.g., Kimwipe or gauze pad) Zeiss Standard phase-contrast microscope with 16× Ph2 objective and condenser ring (or equivalent)

Cytogenetics


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Figure 4.1.4 Chromosome slide preparation: (A) After blotting the long edge of the slide to obtain a thin uniform layer of water, the slide is tilted to ∼30◦ and 3 separate drops of fixed cell suspension are applied starting away from and proceeding toward the frosted end. This sequence allows excess fixative and water to flood onto the frosted end without pooling on the slide. Application of the drops 1/3 of the distance from the top of the slide (indicated by Xs) counteracts the downhill dispersal tendency of cells on the slide and promotes even dispersal across the slide width. (B) After application of the cell suspension, the slide is flooded with fixative across the top edge, again proceeding toward the frosted end. This displaces a front of remaining water across the slide and onto the frosted end. It is important to avoid pooling of excess fluid on the surface of the slide, and to obtain a thin, even film of fixative to ensure uniform drying.

1. Remove slide from methanol and polish with lint-free tissue, such as a folded Kimwipe or gauze pad. Dip slide once in methanol and then several times in deionized water until the methanol is gone and a thin, uniform film of water covers the slide. Good slides have few pits and imperfections and will hold a thin film of water across the entire slide, which reduces the surface tension prior to addition of the cell suspension. Cleaning each slide is essential, as few precleaned slides are truly clean enough for chromosome preparations.

2. Holding the frosted end between the thumb and finger, position the slide with the one long edge parallel to the bench top, and blot the lower long edge on a paper towel to draw off excess water. Keeping the lower long edge in contact with the paper towel, lower the opposite edge until the slide forms a 30◦ angle with the bench top, with the film of water facing up (Fig. 4.1.4).


3. From a Pasteur pipet held in a horizontal position 1 to 2 inches above the slide, place 3 drops of cell suspension, evenly spaced, onto the slide, moving successively toward the frosted end. Drops should strike the tilted slide one-third of its width from the elevated long edge (Fig. 4.1.4). The drops should burst on the water film and spread out evenly as they strike.

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Positioning and spacing of drops is critical. The goal is even dispersal of cells across the entire surface of the slide. This contributes to consistent and uniform slide-drying, which will optimize chromosome spreading. If discrete areas of cells are observed at the drop sites, surrounded by areas with few cells, the slide should be held at a lower angle (i.e.,