Clonal Cultures in vitro for Haemopoietic Cells Using Semisolid Agar Medium

I. INTRODUCTION
An exponential increase in knowledge of the regulation of haemopoiesis has been witnessed since the mid-1980s. Over 20 cytokines (colony-stimulating factors, erythropoietin, thrombopoietin, and interleukins) have now been identified, molecularly cloned, and expressed. Many of these recombinant haemopoietic growth factors are now being used in clinical situations where they have been found to be useful in correcting anaemia and white cell deficiencies either in chronic-inherited disease or following acute treatment (e.g., chemotherapy, bone marrow transplantation) (Atkinson, 1993; Clark and Kamen, 1987; Mertelsmann et al., 1990; Sheridan et al., 1989).

The discovery of haemopoietic growth factors was facilitated greatly by the ability to grow haemopoietic cells in vitro. These culture systems enable the undifferentiated haemopoietic precursors to proliferate and differentiate into various haemopoietic cell lineages. Especially valuable has been the development of clonal cultures using semisolid agar or methylcellulose culture medium for haemopoietic precursor cells. In the presence of appropriate growth factors, these precursor cells proliferate and produce a clonal colony of differentiated cells. This will allow biological, viral, biochemical, or molecular studies to be performed on individual cell clones. In addition, by counting colonies, it is also possible to infer the number of precursor cells in the starting cell population. This is possible because a linear relationship exists between the number of colonies formed and the number of cells cultured. By comparison, liquid suspension cultures of primary haemopoietic cells do not allow enumeration of precursor cell numbers as the progeny are intermingled in the culture dish. The second feature of clonal cultures is the dose-response relationship that exists between the amount of growth factor and the number of colonies stimulated. This dose-response relationship is sigmoid, having a linear phase and a plateau phase. The linear portion of the curve can be used to determine the amount of growth factor activity; in cultures described here, 50 units of growth factor activity correspond to the amount of activity stimulating 50% of maximal colony numbers. More detailed information on haemopoietic colony formation and cytokines can be found in Metcalf (1984, 1985, 1986, 1991) and Nicola (1991).

II. MATERIALS AND INSTRUMENTATION
A. Semisolid Agar Medium Cultures
Iscove's modified Dulbecco's medium powder, with 4mM L-glutamine and 25 mM HEPES buffer (IMDM, Cat. No. 12200-085 for a 5-1itre batch), is from GIBCOInvitrogen. DEAE-dextran hydrochloride (Cat. No. D9885), L-asparagine monohydrate (Cat. No. A4284), 2-mercaptoethanol (Cat. No. M7522), penicillin G (Cat. No. P7794), and streptomycin (Cat. No. $9137) are from Sigma-Aldrich. Bacto agar is from Bacto Laboratories (Cat. No. 0140-17). Recombinant haemopoietic growth factors and cytokines can be purchased from a number of commercial suppliers (e.g., PeproTech EC Ltd, R&D Systems). For routine cultures, conditioned medium can be prepared from a number of tissues (see later). Foetal calf serum (FCS) can be obtained from a number of suppliers, but requires pretesting (see later). Bacteriological graded petri dishes (35-36 and 100mm diameter) can be obtained from a number of suppliers as a tissue culture grade dish is not required. For viable cell count, eosin Y from Sigma-Aldrich (Cat. No. E4009) is required. Distilled Milli-Q endotoxin-free water (H₂0) is routinely used.

B. Conditioned Medium
IMDM (Cat. No. 12200-085) and pokeweed mitogen (PWM, Cat. No. 15360-019) are from GIBCOInvitrogen. 2-Mercaptoethanol (2ME, Cat. No. M7522), sodium bicarbonate (NaHCO₃, Cat. No. $6014), penicillin G (Cat. No. P7794), and streptomycin (Cat. No. $9137) are from Sigma-Aldrich. FCS can be obtained from various suppliers, and batches can only be selected by prior testing. If a batch is available that is known to support good colony formation, then this can be used to prepare conditioned media. A variety of flasks can be used to prepare PWM-stimulated spleen cell-conditioned media (PWM-SCM) for murine cultures, although we routinely use 1- to 2-1itre glass flasks fitted with a cotton plug to allow gas diffusion. For preparation of human placental conditioned media, a variety of flasks can also be used, but we routinely use disposable 75-cm² tissue culture flasks (Cat. No. 25110-75, Corning).

C. In Situ Colony Staining
Glutaraldehyde solution (grade II, 25% in water, Cat. No. G6257), ethanol (Cat. No. 02862), urea (Cat. No. U6504), acetylthiocholine iodide (Cat. No. A5751), sodium phosphate dibasic (Na₂HPO₄, Cat. No. 71629), sodium phosphate monobasic (NaH₂PO₄, Cat. No. 71492), sodium citrate (Cat. No. $1804), copper(II) sulphate (CuSO₄, Cat. No. C1297), potassium ferricyanide [K₃Fe(CN)₆, Cat. No. 60310], and Mayer's haematoxylin staining solution (Cat. No. MHS-16) are from Sigma-Aldrich. Luxol fast blue MBS (Code Pack No. 10732) is from NBS Biologicals. Hardened ashless 541 filter paper of 5.5-cm diameter (Cat. No. 1541 055) is from Whatman, and DePex (D.P.X) neutral mounting medium (Cat. No. 3197) is from Bacto Laboratories. Plain 2 × 2-in. microscope slides (Cat. No. 5075) and 45 × 50-mm cover glasses (Cat. No. 4550-1) can be obtained from Brain Research Laboratories.

D. Instrumentation
For colony examination and counting, a zoom stereomicroscope (Model SZ4045) fitted with a clear stage plate and a base illuminator from Olympus is used routinely. Many brands of CO₂ incubators are available and it is recommended that one with stainless-steel water jackets be obtained. In addition, to minimize the desiccation of cultures, an incubator without an inbuilt fan is preferred. A standard light microscope equipped with 10 and 40× objectives will be sufficient for routine cell counting and in situ colony typing. Graduated, glass blow-out pipettes in volumes of 1, 5, 10, and 25 ml are routinely used, although they can be replaced by disposable pipettes. For the concentration of conditioned media, we use an Amicon hollow-fibre concentrator (Model DC2A) fitted with a HIP10 membrane.

III. PROCEDURES
A. Method for Establishing and Scoring Agar-Medium Cultures
The fundamental steps in establishing agar-medium cultures are fourfold: (1) mix equal volumes of doublestrength medium and double-strength agar solution, (2) add the cells to be cultured and mix, (3) pipette the cell suspension onto the culture dishes, and (4) after gelling the culture dishes are put in an incubator for colony formation.

Solutions

1. IMDM for agar cultures (AIMDM): 88.3 g IMDM, 0.6g penicillin G, 0.375g streptomycin, 7.5 ml DEAEdextran (50mg/ml solution), 1.0g L-asparagine, 24.5g NaHCO3, and 29.5 µl 2-mercaptoethanol. Dissolve the contents of a 5-1itre package IMDM (88.3 g) in I litre of H₂O using a magnetic stirrer for mixing. Rinse the inside of the package to remove all traces of powder. Add afore-listed reagents while stirring continuously. Add H₂O to a final volume of 1.95 litres and gas medium with 100% CO₂ until it is a yellow-orange colour. The prepared endotoxin-free medium should then be filter sterilized and distributed in 100- or 250- ml aliquots, which are then tightly capped, stored at 4°C and protected from light. Each preparation of AIMDM should be batch tested against one that is currently in use if applicable.
2. Agar: 0.6 g Bacto-agar and 100ml H₂O. Weigh agar into a 100-ml flask, add 100ml H₂O, and plug flask loosely. Bring to boiling for 2min over a gas flame. Prepare immediately before setting up cultures and maintain at 45°C in a water bath. Each new lot of Bacto-agar should be batch tested against one that is currently in use if applicable.
3. Foetal calf serum: FCS is used as a source of nutrients in cell cultures. Batches of FCS should be tested extensively prior to purchase, for optimal colony formation in colony number and colony size, in semisolid cultures. They should also be titrated to determine the optimal concentration (final concentration is usually 5-20%). The storage/shelf life of FCS at -20°C is at least 2 years and at-70°C it as long as 10 years. Centrifugation may be necessary to remove any sediment that forms after thawing. It is otherwise ready for use. Heat inactivation is optional but not necessary.
4. Eosin for viable cell counts: Prepare stock solution [10% eosin-yellow powder (w/v)] in normal saline and keep at 4°C. Mix 0.2 ml of stock eosin solution with 8.6 ml normal saline and 1.5 ml FCS to prepare working solution. Aliquots of the working solution should be store frozen.
5. Hemopoietic growth factors: If purchased commercially, they should be pretested to determine the amount required for optimal colony formation. If conditioned media are prepared (see later), they also require titration to determine the optimal concentration for maximal colony formation without evidence of high-dose inhibition. Stimuli should be divided into aliquots at a concentration at least 10-fold higher than that to be used finally in the culture dish. They can be stored frozen, but once thawed, they should not be frozen again as this can result in a loss of activity.
6. Single-strength Iscove's modified Dulbecco's medium: Dissolve the entire contents of a 5-1itre package IMDM (88.3g) with 4.8 litres of H₂O and mix with gentle stirring. Rinse the inside of the package to remove all traces of powder. Add 15.12g NaHCO₃, 29.5H µl 2- mercaptoethanol, 150 mg penicillin G, and 100 mg streptomycin while stirring continuously. Add H₂O to a final volume of 5 litres and gas medium with 100% CO₂ until it is a yellow-orange colour. The prepared endotoxin- free medium should then be filter sterilized and distributed in 100- or 500-ml aliquots, which are then capped tightly, stored at 4°C and protected from light.
7. Pokeweed mitogen: This should be prepared immediately prior to use. Any material not used should be discarded. Make up powder with 5ml of doubledistilled, deionized water. Remove from the vial and dilute 1 : 15 (v/v) with H₂O.

Steps

1. Warm AIMDM to room temperature.
2. Prepare agar solution.
3. Count viable cells using a haemocytometer and eosin as dye.
4. Draw culture layout, in a book, showing culture number, stimuli for each culture dish, and the number of cells for each culture dish.
5. Place required number of culture dishes on incubator trays and number lids individually according to the culture book.
6. Add required stimuli to appropriate culture dishes as described in the culture book. For each culture, the required amount of stimulus is usually added in 0.1 ml per culture dish. This amount can be less but should not exceed 0.2ml as the agar may not gel properly.
7. For agar cultures with 20% FCS final concentration, mix AIMDM (three parts) and foetal calf serum (two parts) first and then add an equal volume of agar (five parts). Cells are added last to this single-strength agar medium, which should be now roughly at 36-37°C before plating. For each group of cultures, allow 1 ml of agar medium each per culture plus at least 1 ml extra for wastage in pipetting. If by pretesting, a lower concentration of FCS is sufficient for optimal colony formation, replace the leftover volume with H₂O so that the amount of AIMDM and FCS equals 0.5 ml for each 1 ml of culture.
8. Aliquot 1-ml volumes into petri dishes and swirl to mix stimuli and agar medium-containing cells.
9. Allow mixture to gel and place in a fully humidified containing 5-10% CO₂ in air.
10. After the required incubation period (normally 7 days for murine progenitor cells and 14 days for human progenitors and murine high-proliferative stem/progenitor cells), remove cultures from incubator and count colonies using an Olympus SZ stereomicroscope at 30-35× magnification. Murine haematopoietic colonies are defined routinely as clones greater than 50 cells and human colonies greater than 40 cells (some investigators count human colonies as having greater than 20 cells). Place the microscope on top of a black platform and adjust the concave side of the mirror until cells appear white against a black background (Fig. 1). To aid in the enumeration of colonies, we routinely put the 35-mm agar culture dishes on top of an inverted 60-mm culture dish marked with 6-mm² grids.

B. Preparation of Conditioned Media
Although specific stimuli may be required for many situations, and always for use as a positive control, a variety of conditioned media containing mixtures of haemopoietic growth factors can be prepared. The following examples provide descriptions for the preparation of two conditioned media: one suitable for human cultures and the other for murine cultures.

1. Preparation of Human Placenta-Conditioned Medium

Steps

1. Placenta should be obtained within 9 h of birth. Place it on a large sterile tray in a biological safety cabinet.
2. Using sterile instruments, remove outer layer of placenta. Assume that this portion is not sterile. Instruments can be kept sterile by periodically returning them to boiling water.
3. Cut a portion (1 cm³) of exposed placenta and place in a 100-mm petri dish containing 10ml IMDM. Limit each petri dish to 8-10 pieces.
4. Having removed sufficient pieces of placenta, rinse each piece through three changes of IMDM in 100-mm petri dishes to remove most of the blood.
5. Place 18-20 pieces of placenta into tissue culture flasks (75cm², Cat. No. 25110-75, Coming) in 60ml IMDM with 5% (v/v) FCS.
6. Place the placenta cultures, with the caps sealed loosely, in a 37°C fully humidified incubator containing 5-10% CO₂ in air.
7. After 5-7 days of incubation, harvest the medium free of placenta by pouring the contents of each flask through cotton gauze into a collection flask. Centrifuge the medium at 3000g and store the supernatant at -20°C until 4-5 litres has accumulated.
8. Concentrate the placenta-conditioned medium approximately 10-fold using a hollow fibre concentrator (this type of concentrator is preferred because of the relatively large volumes involved).
9. Filter sterilize the concentrate and test by titration using human bone marrow semisolid agar cultures. Select the batches of conditioned media that display a sigmoid dose-response relationship with the number of colonies formed and without evidence of a high-dose inhibition.

2. Preparation of Murine Pokeweed Mitogen- Stimulated Spleen Cell-Conditioned Medium

Steps

1. Prepare a single-cell suspension of murine spleen cells, either by teasing the spleen tissue with needle or by forcing it through a fine stainless steel mesh.
2. Place the spleen cells in a tube and allow them to stand for 5rain to allow larger tissue fragments to sediment. Remove the supernatant and determine viable cell numbers.
3. Make up the cells to 2 × 10⁶/ml in IMDM containing 10% FCS. (The concentration of FCS should be as low as possible and can be determined only by preliminary testing.) Add pokeweed mitogen (0.05 ml of a 1:15 dilution of freshly prepared stock is added for each millilitre of culture medium).
4. Incubate the cells in medium for 7 days at 37°C in a fully humidified incubator containing 5-10% CO₂ in air. The cells can be incubated in a variety of containers. We routinely use 2-1itre flasks, with cotton plugs, containing 250ml of medium.
5. After incubation, harvest the conditioned medium and centrifuge at 3000g to remove cellular debris. Concentrate the medium 10-fold as described earlier for human placenta-conditioned medium.
6. Titrate the concentrated PWM-SCM using cultures of routine bone marrow cells to determine the concentration required to give plateau numbers of colonies.
7. The conditioned medium can then be diluted to a concentration 10 times that is required for maximal colony formation, divided into 20-ml aliquots, and stored at-20°C until required. Once thawed the PWMSCM should be stored at 4°C.

III. COLONY TYPING
Colonies grown in agar cultures can be typed using an in situ whole plate staining sequentially with Luxol Fast Blue to detect eosinophil granules (Johnson & Metcalf, 1980), acetylcholinesterase to detect megakaryocytes (von Melchner and Lieschke, 1981) and Mayer's haematoxylin for nuclear morphology to detect neutrophils and monocyte/macrophages.

Preparation of Fixative and Staining Solutions
A 2.5% (v/v) glutaraldehyde in PBS is prepared by mixing 1 part of 25% glutaraldehyde with 9 parts of PBS. The luxol fast blue staining solution is prepared by dissolving 0.1 g powdered luxol fast blue MBS dye (NBS Biologicals Ltd., Hungtingdon, Cambs, England) in 100ml of 70% ethanol saturated with urea. The substrate solution for acetylcholinesterase staining should be prepared fresh each time by dissolving 10mg acetylthiocholine iodide (Sigma-Aldrich) in 15 ml of 100 mM sodium phosphate buffer, pH 6.0. One millilitre of 100mM sodium citrate, 2ml of 30mM copper sulphate, and 2ml of potassium ferricyanide solutions are then added sequentially with constant stirring.

Steps

1. Add 2ml 2.5% glutaraldehyde into individual culture dishes and leave at room temperature overnight.
2. Transfer the gel onto a 3 × 2-in. glass slide with the aid of a water bath and then cover the gel with a wet 5.5-cm Whatman 541 filter paper and allow drying at room temperature in a fume hood.
3. Remove the filter paper, leaving a thin film of gel containing compressed colonies on the glass slides.
4. Slides are first stained for acetylcholinesterase by incubating the slides in the dark with the substrate solution at room temperature for 3 h.
5. After washing under running tap water for 10- 15 min, transfer slides into the luxol fast blue staining solution and stain for 30min at room temperature.
6. After another washing under running tap water for 30min, counterstain slides with Mayer's haematoxylin for 1 min, wash, and "blue" in running tap water.
7. Dry slides at room temperature, mount with DePex, and examine under a light microscope after drying.

IV. PITFALLS

1. Numerous pitfalls are associated with these procedures. A major problem involves selection of a suitable batch of FCS. If possible, a known positive sample should be obtained from a colleague or a commercial source (e.g., StemCell Technologies) for use as a control when testing new batches.
2. The agar needs to be boiled to ensure that it is dissolved properly. For this reason, it is recommended to use a gas flame. The agar will initially bubble up when boiliing, and care must be taken to prevent it from overflowing. Once the bubbling has subsided, the agar should be boiled for another minute. An autoclave should not be used to prepare the agar solution.
3. It is imperative that the incubator being used is fully humidified, as desiccation of the cultures will prevent colony growth. With satisfactory cultures, a small volume of liquid will be evident at the edge of the agar medium when the cultures are tilted. If desiccation has occurred, the surface of the agar medium will display irregularities instead of being smooth and shiny. To prevent desiccation, the incubator should contain one or more large open trays containing H₂O. Humidity can be improved by pumping the air-gas mixture into the incubator via a tube immersed in one of the trays of water. Many incubators are fitted with a fan to produce a uniform atmosphere with the closed incubator. This can also cause the desiccation of cultures and the fan may have to be disconnected.
4. The possibility of cultures drying out is increased by extending the incubation time. For cultures in excess of 7 days it is a good policy to place culture dishes in a 100-cm petri dish (two cultures per dish) containing a third open-lid, 35-mm culture dish with H₂O. It is also a good policy to minimize the number of openings of the incubator.
5. When establishing the cultures, problems can arise due to the temperature of the agar-medium mixture. If too cold, it will gel prematurely and the cells will not be immobilized properly. If too hot, it will kill the cells. It is recommended that the agar be maintained at 45°C and that the AIMDM and FCS be allowed to warm to room temperature (18-20°C). When agar, medium, and FCS are mixed, the temperature of the solution will be about 37°C which will not kill the cells but will still be above the gelling temperature of the agar. Once cells have been added to the agar medium mixture and mixed, it should be dispensed to culture dishes as soon as possible. As 0.1 ml of stimulus is usually present in the culture dish, the mixture should be swirled gently to allow homogeneous mixing. All these actions should be executed before gelling starts to occur.

References
Atkinson, K. (1993). Cytokines in bone marrow transplantation. Today's Life Sci. 5, 28-38.

Clark, S. C., and Kamen, R. (1987). The human hematopoietic colony-stimulating factors. Science 236, 1229-1237.

Johnson, G. R., and Metcalf, D. (1980). Detection of a new type of mouse eosinophil colony by Luxol-Fast-Blue staining. Exp. Hematol. 8, 549-561.

Mertelsmann, R., Herrman, F., Hecht, T., and Schulz, G. (1990). Hematopoietic growth factors in bone marrow transplantation. Bone Marrow Transplant. 6, 73-77.

Metcalf, D. (1984). "The Hemopoietic Colony Stimulating Factors." Elsevier, Amsterdam. Metcalf, D. (1985). The granulocyte-macrophage colony-stimulating factors. Science 229, 16-22.

Metcalf, D. (1986) How reliable are in vitro clonal cultures? Some comments based on hemopoietic cultures. Int. J. Cell Cloning 4, 287-294.

Metcalf, D. (1991). Control of granulocytes and macrophages: Molecular, cellular, and clinical aspects. Science 254, 529- 533.

Nicola, N. A. (1991). Receptors for colony stimulating factors. Br. J. Haematol. 77, 133-138.

Sheridan, W. P., Morstyn, G., Wolf, M., Lusk, J., et al. (1989). Granulocyte colony-stimulating factor and neutrophil recovery after high dose chemotherapy and autologous bone marrow transplantation. Lancet 2, 891-895.

von Melchner, H., and Lieschke, G. J. (1981). Regeneration of hemopoietic precursor cells in spleen organ cultures from irradiated mice: Influence of genotype of cells injected and of the spleen microenvironment. Blood 57, 906-912.