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A collection of high quality essays on ‘Animal Tissue Culture’.
Essay on Animal Tissue Culture
Essay # 1. Introduction to Animal Tissue Culture:
The term tissue culture refers to the culture of whole organs, tissue fragments as well as dispersed cells on a suitable nutrient medium.
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It can be divided into two parts:
(1) Organ culture.
(2) Cell culture.
It is mainly on the basis of whether the tissue organisation is retained or not. In organ cultures whole embryonic organs or small tissue fragments are cultured in vitro in such a manner that they retain their tissue architecture.
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In contrast, cell cultures are obtained either by enzymatic or mechanical dispersal of tissues into individual cells or by spontaneous migration of cells from an explant; they are maintained as attached monolayers or as cell suspensions.
Freshly isolated cell cultures are called primary cultures, they are usually heterogeneous and slow growing, but are more representative of the tissue of their origin both in cell type and properties. Once a primary culture is sub-cultured, it gives rise to cell lines which may either die after several subcultures (such cell lines are known as finite cell lines) or may continue to grow indefinitely (these are called continuous cell lines).
Usually normal tissues give rise to finite cell lines, while tumours give rise to continuous cell lines. But there are several examples of continuous cell lines which were derived from normal tissues and are themselves non-tumorigenic, e.g., MDCK dog kidney, 3T3 fibroblasts etc. The evolution of continuous cell lines from primary cultures is supposed to involve a mutation which alters their properties as compared to those of finite lines.
Tissue culture refers to the ability to grow cells outside of the organism in a “synthetic” environment. Cells may be grown on a solid surface such as a dish that looks like a petri dish for growing bacteria (although cells of higher organisms will not grow on bacterial petri plates), in bottles, or in suspension.
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Cells growing on solid supports are bathed in highly complex liquid medium, in contrast to the simple defined medium for growing bacteria. Some of the components required for growth of some types of cells in culture are still undefined, and are supplied by the serum fraction of blood from e.g., fetal calves, sheep or horses. Growing cells in culture is quite expensive, mainly due to the cost of the serum which is added at a concentration of 5-10% depending on the cell type.
Cells are grown at (usually) 37 C in an incubator whose gaseous environment is maintained at 5% CO2, in order to maintain the correct pH in the medium.
Developing the techniques for in vitro (in glass) cell cultivation began in the beginning of this century. The first successes were with chunks of organs grown in lymph. Eventually, techniques were developed to separate tissues into individual cells.
This work was so important that the investigator who developed the conditions the tissue culture growth of cells in which polio virus was produced, necessary for the development of the polio vaccine, was awarded a share of the Nobel Prize.
In order to put cells from an organism into culture, the tissue is disaggregated by the use of an enzyme called trypsin; trypsin, a protease, works by digesting protein on the cell surface – don’t worry, they eventually are replaced. Single cells are deposited on a solid surface, covered with medium plus serum, and they then proceed to grow and divide.
Eventually, they fill the area and touch each other; such cells are said to be confluent. Confluent cells stop growing, a phenomenon called contact inhibition. Cancerous cells when grown in culture do not exhibit contact inhibition, reflecting their unregulated growth in the organism. Instead of remaining as a flat sheet (monolayer) on the surface of the growth vessel, they mound up and grow on top of each other).
If a virus can replicate with the same characteristics in a cancerous cell, it is advantageous to use the cancerous cell, for the simple reason that many more cells can be obtained, and therefore, a greater mass of viral constituents.
Essay # 2. Types of Tissue Culture Process:
1. Batch and Continuous Culture:
In standard culture, known as batch culture, cells are inoculated into a fixed volume of medium and, as they grow, nutrients are consumed and metabolites accumulate. The environment is therefore continuously accumulating the toxic waste products, or is density-dependent (imitation of growth in monolayer cultures).
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There are means of prolonging the life of a batch culture, and thus increasing the yield, by various substrates feed methods. Gradual addition of fresh medium, so increasing the volume of the culture (fed batch).
Intermittently, by replacing a constant fraction of the culture with an equal volume of fresh medium (semi-continuous batch), all batch culture systems retain the accumulating waste products, to some degree, and have a fluctuating environment. All are suitable for both monolayer and suspension cells.
2. Perfusion:
By the continuous addition of medium to the culture and the withdrawal of an equal volume of used (cellfree) medium, perfusion can be open, with complete removal of medium from the system, or closed, with recirculation of the medium, usually via a secondary vessel, back into the culture vessel. The secondary vessel is used to ‘regenerate’ the medium by gassing and pH correction.
3. Continuous-Flow Culture:
Continuous-flow Culture gives the true homeostatic conditions with no fluctuations of nutrients, metabolites, or cell numbers. It depends upon medium entering the culture with a corresponding withdrawal of medium plus cells. It is thus only suitable for suspension culture cells, or monolayer cells growing on micro carriers. Continuous-flow culture is described more fully.
Comparison of Batch, Perfusion, and Continuous-Flow Culture:
Continuous-flow culture is the only system in which the cellular content is homogeneous, and can be kept homogeneous for long periods of time (months). This can be vital for physiological studies, but may not be the most economical method for product generation. Production economics are calculated in terms of staff time, medium, equipment, and downstream processing costs.
Also taken into account are the complexity and sophistication of the equipment and process, as this affects the calibre of the staff required and the reliability of the production process. Batch culture is more expensive on staff time and culture ingredients, because for every single harvest a sequence of inoculum build-up steps and then growth in the final vessel has to be carried out, add there is also downtime whilst the culture is prepared for its next run.
Batch cultures can give repeated but smaller harvests, and the longer a culture can be maintained in a productive state then the more economical the whole process becomes.
Continuous-flow culture in the chemostat implies that cell yields are never maximal because a limiting growth factor is used to control the growth rate. If maximum yields are desired in this type of culture then the turbidostat option has to be used.
Some applications, such as the production of a cytopathic virus, leave no choice other than batch culture. Maintenance of high yields, and therefore high product concentration, may be necessary to reduce downstream processing costs and these could outweigh medium expenses.
For this purpose perfusion has to be used. Although for many processes this is more economical than batch culture, it does add to the complexity of the equipment and process, and increases the risk of a mechanical or electrical failure or microbial contamination prematurely ending the production run.
There is no clear-cut answer to which type of culture process should be used-it depends upon the cell and product, the quantity of product, downstream processing problems, and product licensing regulations (batch definition of product, cell stability, and generation number). However, a relative ratio of unit costs for perfusion, continuous-flow, and batch culture in the production of monoclonal antibody is 1:2:3.5.
4. Monolayer Culture:
Tissue culture flasks and tubes giving surface areas of 5-200 cm2 are familiar to all tissue culturists. The largest stationary flask routinely used in laboratories is the Roux bottle (or disposable plastic equivalent) which gives a surface area for cell attachment of 175-200 cm2 (depending upon type), needs 100-150 ml medium, and utilises 750-1000 cm2 of storage space.
This vessel will yield 2 x 107 diploid cells and up to 108 heteroploid cells. If one has to produce, for example, 1010 cells, then over 100 replicate cultures are needed (i.e. manipulations have to be repeated 100 times). In addition the cubic capacity of incubator space needed is over 100 litres.
Clearly there comes a time in the scale-up of cell production when one has to use a more efficient culture system. Scale-up of anchorage-dependent ceils reduces the number of cultures, is more efficient in the use of staff, and increases significantly the surface area/volume ratio. In order to do this a very wide and versatile range of tissue culture vessels and systems has been developed.
The methods with the most potential are those based on modifications to suspension culture systems because they allow a truly homogeneous unit process with enormous scale-up potential to be used. However, these systems should be attempted only if time and resources allow a lengthy development period.
Although suspension culture is the preferred method for increasing capacity, monolayer culture has the following advantages- It is very easy to change the medium completely and to wash the cell sheet before adding fresh medium. This is important in many applications when the growth is carried out in one set of conditions and product generation in another.
A common requirement of a medium change is the transfer of cells from serum to serum-free conditions. The efficiency of medium changing in monolayer cultures is such that a total removal of the unwanted compound can be achieved. If artificially high cell densities are needed then these can be supported by using perfusion techniques.
It is much easier to perfuse monolayer cultures because they are immobilised and a fine filter system (to withhold cells) is not required. Many cells will express a required product more efficiently when attached to a substrate.
The same apparatus can be used with different media/cell ratios which, of course, can be easily changed during the course of an experiment. Monolayer cultures are more flexible because they can be used for all cell types. If a variety of cell types are to be used then a monolayer system might be a better investment. It should be noted that the microcarrier system confers some of the advantages of a suspension culture system.
There are four main disadvantages of monolayer compared to suspension systems:
a. They are difficult and expensive to scale-up.
b. They require far more space.
c. Cell growth cannot be monitored as effectively, because of the difficulty of sampling and counting an aliquot of cells.
d. It is more difficult to measure and control parameters such as pH and oxygen, and to achieve homogeneity.
Micro carrier culture eliminates or at least reduces many of these disadvantages.
Essay # 3. Cell Attachment Used in Tissue Culture:
Animal cell surfaces and the traditional glass and plastic culture surfaces are negatively charged, so for cell attachment to occur, crosslinking with glyco-proteins and/or divalent cations (Ca2+, Mg2+) is required. The glycoprotein most studied in this respect is fibronectin, a compound of high molecular mass (220000) synthesised by many cells and present in serum and other physiological fluids.
Although cells can presumably attach by electrostatic forces alone, it has been found that the mechanism of attachment is similar, whatever the substrate charge. The important factor is the net negative charge, and surfaces such as glass and metal which have high surface energies are very suitable for cell attachment.
Organic surfaces need to be wettable and negative, and this can be achieved by chemical treatment (e.g. oxidising agents, strong acids) or physical treatment (e.g. high voltage discharge, UV light, high energy electron bombardment). One or more of these methods are used by manufacturers of tissue culture grade plastics.
The result is to increase the net negative charge of the surface (for example by forming negative carboxyl groups) for electrostatic attachment. Surfaces may also be coated to make them suitable for cell attachment.
A tissue culture grade of collagen can be purchased which saves a tedious preparation procedure using rat tails. The usefulness of collagen as a growth surface is also demonstrated by the availability of collagen- coated micro carrier beads Cytodex-3, Pharmacia).
Surfaces for Cell Attachment:
Schematic of a cell attachment blocking strategy using a surface impregnated with a molecular “blocking” analogue (blue circles). This analogue of the naturally produced “attachment blocking” biofilm signal of most bacteria in a mixed population would keep the surface clean (cells at left), but species or mutants that were not affected (at right) would colonise the surface/reactor.
i. Glass:
Alum-borosilicate glass (e.g. Pyrex) is preferred because soda lime glass releases alkali into the medium and needs to be detoxified (by boiling in weak acid) before use. After repeated use glassware can become less efficient for cell attachment, but efficiency can be regained by treatment with 1 mM magnesium acetate. After several hours soaking at room temperature the acetate is poured away and the glassware is rinsed with distilled water and autoclaved.
ii. Plastics:
Polystyrene is the most used plastic for cell culture, but polyethylene, polycarbonate, Perspex, PVC, Teflon, cellophane, and cellulose acetate are all suitable when pre-treated correctly.
iii. Metals:
Stainless steel and titanium are both suitable for cell growth because they are relatively chemically inert, but have a suitable high negative energy. There are many grades of stainless steel, and care has to be taken in choosing those which do not leak toxic metallic ions.
The most common grade to use for culture applications is 316, but 321 and 304 may also be suitable. Stainless steel should be acid washed (10% nitric acid, 3.5% hydrofluoric acid, 86.5% water) to remove surface impurities and inclusions acquired during cutting.
Scaling-up:
Step 1:
Roller Bottle:
The aims of scaling-up are to maximise the available surface area for growth and to minimise the volume of medium and headspace, while optimising cell numbers and productivity. Stationary cultures have only one surface available for attachment and growth, and consequently they need a large medium volume.
The medium volume can be reduced by rocking the culture or, more usually, by rolling a cylindrical vessel. The roller bottle has nearly all its internal surface available for cell growth, although only 15-20% is covered by medium at any one time. Plastic disposable bottles are available in c. 750 cm2 and c. 1500 cm2 (1400-1750 cm2) sizes.
Rotation of the bottle subjects the cells to medium and air alternately, as compared with the near anaerobic conditions in a stationary culture. This method reduces the volume of medium required, but still requires a considerable headspace volume to maintain adequate oxygen and pH levels.
The scale-up of a roller bottle requires that the diameter is kept as small as possible. The surface area can be doubled by doubling the diameter or the length. The first option increases the volume (medium and headspace) fourfold, the second option only twofold.
The only means of increasing the productivity of a roller bottle and decreasing its volume is by using a perfusion system, originally developed by Kruse et al., and marketed by New Brunswick and Bellco (Autoharvester). This is an expensive option, as an intricate revolving connection has to be made for the supply lines to pass into the bottle. However, cell yields are considerably increased and extensive multilayering takes place.
Protocol:
Use of standard disposable roller cultures.
Equipment:
I. 1400 cm2 (23×12 cm) plastic disposable.
II. Inverted microscope bottle (e.g. Costar, Bibby Sterilin)
Method:
1. Add 300 ml of growth medium.
2. Add 1.5 X 107 cells.
3. Roll at 12 r.p.h. at 37 °C for 2 h, to allow an even distribution of cells during the attachment phase.
4. Decrease the revolution rate to 5 r.p.h., and continue incubation.
5. Examine cells under an inverted microscope using an objective with a long working distance.
6. Harvest cells when visibly confluent (five to six days) by removing the medium, adding trypsin (0.25%), and rolling.
Yields will be very similar to those obtained in flasks, assuming enough medium was added. This method has the advantage of allowing the medium volume/surface area ratio to be altered easily. Thus after a growth phase the medium volume can be reduced to get a higher product concentration.
Step 2:
Roller Bottle Modifications:
The roller bottle system is still a multiple process, and thus inefficient in terms of staff resources and materials. To increase the surface area within the volume of a roller bottle, the following vessels have been developed.
i. SpiraCel:
Bibby Sterilin have replaced their bulk cell culture vessel with a SpiraCel roller bottle. This is available with a spiral polystyrene cartridge in three sizes, 3000, 4500, and 6000 cm2. It is crucial to get an even distribution of the cell inoculum throughout the spiral, otherwise very uneven growth and low yields are achieved. Cell growth can be visualised only on the outside of the spiral, and this can be misleading if the cell distribution is uneven.
ii. Glass Tubes:
A small scale example is the Bellco-Corbeil Culture System (Bellco). A roller bottle is packed with a parallel cluster of small glass tubes, (separated by silicone spacer rings). Three versions are available giving surface areas of 5 x 103, 1 x 104, and 1.5 x 104 cm2.
Medium is perfused through the vessel from a reservoir. The method is ingenious in that it alternately rotates the bottle 360 ° clockwise and then 360° anticlockwise. This avoids the use of special caps for the supply of perfused medium.
An example of its use is the production of 3.2 x 109 Vero cells (2.3 x 105/cm2) over six days using 6.5 litres of medium (perfused at 50 ml/ min) in the 10000.
iii. Increased Surface Area Roller Bottles:
In place of the smooth surface in standard roller bottles the surface is ‘corrugated’, thus doubling the surface area within the same bottle dimensions; e.g. extended surface area roller bottle (ESRB) available from Bibby Sterilin (Corning), or the ImmobaSil surface which is a textured silicone rubber matrix surface (Ashby Scientific Ltd. or Integra Biosciences).
Step 3:
Large Capacity Stationary Cultures:
The standard cell factory unit (Nunc) comprises ten chambers, each having a surface area of 600 cm2, fixed together vertically and supplied with interconnecting channels. This enables all operations to be carried out once only for all chambers.
It can thus be thought of as a flask with a 6000 cm2 surface area using 2 litres of medium and taking up a total volume of 12 500 ml. In practice this unit is convenient to use and produces good results, similar to plastic flasks. It is made of tissue culture grade polystyrene and is disposable.
One of the disadvantages of the system can be turned to good use. In practice it is difficult to wash out all the cells after harvesting with trypsin, etc. However, enough cells remain to inoculate a new culture when fresh medium is added.
Given good aseptic technique, this disposable unit can be used repeatedly. The system is used commercially for interferon production (by linking together multiples of these units) (16). In addition, units are available giving 1200 (2 tray) and 24000 cm2 (40 tray).
i. Costar Cell Cube:
The Cell Cube has parallel polystyrene trays in a modular closed-loop per-fusion system with an oxygenator, pumps, and a system controller (pH, 02, level control). The unit is compact, with the trays being only 1 mm apart; thus the smallest unit of 21250 cm2 is less than 5 litres total volume (1.25 litres medium). Additional units of 42500 cm2 (2.5 litres medium) and 85000 cm2 (5 litres medium) are available and four units can be run in parallel, giving 340000 cm2 growth area.
ii. Hollow Fibre Culture:
Bundles of synthetic hollow fibres offer a onatrix analogous to the vascular system, and allow cells to grow in tissue-like densities. Hollow fibres are usually used in ultrafiltration, selectively allowing passage of macromolecules through the spongy fibre wall while allowing a continuous flow of liquid through the lumen.
When these fibres are enclosed in a cartridge and encapsulated at both ends, medium can be pumped in and will then perfuse through the fibre walls, which provide a large surface area for cell attachment and growth. Culture chambers based on this principle are available from Amicon.
The capillary fibres, which are made of acrylic polymer, are 350 ^m in diameter with 75 (Am walls. The pores through the internal lumen lining are available with molecular mass cutoffs between 10000 and 100000.
It is difficult to calculate the total surface area available for growth but units are available in various sizes and these give a very high ratio of surface area to culture volume (in the region of 30 cm2/ml). Upto 108 cells/ml has been maintained in this system. These cultures are mainly used for suspension cells but are suitable for attached cells if the polysulfone type is used.
iii. Optic Cell Culture System:
This system (Cellex Biosciences Inc.) consists of a cylindrical ceramic cartridge (available in surface area between 0.4-12.5 m2) with 1 mm2 square channels running lengthways through the unit. A medium perfusion loop to a reservoir, in which environmental control is carried out, completes the system.
It provides a large surface area/volume ratio (40:1) and its suitability for virus, cell surface antigen, and monoclonal antibody production is documented, Scale-up to 210 m2 is possible with multiple cartridges arranged in parallel in a single controlled unit. Cartridges are available for both attached and suspension cells, which become entrapped in the rough porous ceramic texture.
iv. Heli-Cel (Bibby Sterilin):
Twisted helical ribbons of polystyrene (3 mm x 5-10 mm x 100^m) are used as packing material for the cultivation of anchorage-dependent cells. Medium is circulated through the bed by a pump, and the helical shape provides good hydrodynamic flow. The ribbons are transparent and therefore allow cell examination after removal from the bed.
Step 4:
Unit Process Systems:
There are basically three systems which fit into fermentation (suspension Culture) apparatus
I. Cells stationary, medium moves (e.g. glass bead reactor)
II. Heterogeneous mixing (e.g. stack plate reactor)
III. Homogeneous mixing (e.g. micro carrier)
Bead Bed Reactor:
The use of a packed bed of 3-5 mm glass beads, through which medium is continually perfused has been reported by a number of investigators since 1962. The potential of the system for scale-up was demonstrated by Whiteside and Spier (19) who used a 100 litre capacity system for the growth of BHK21 cells and systems can be commercially obtained (Meredos GmbH).
Essay # 4. Methods of Animal Tissue Culture:
Small scale culture of cells in flasks of up to 1 litre volume (175 cm2 surface area) is the best means of establishing new cell lines in culture, for studying cell morphology and for comparing the effects of agents on growth and metabolism.
However, there are many applications in which large numbers of cells are required, for example extraction of a cellular constituent (109 cells can provide 7 mg DNA); to produce viruses for vaccine production (typically 5 X 1010 cells per batch) or other cell products (interferon, plasminogen activator, interleukins, hormones, enzymes, erythropoietin, and antibodies); and to produce inocula for even larger cultures.
Animal cell culture is a widely used production process in biotechnology, with systems in operation at scales over 10000 litres. This has been achieved by graduating from multiples of small cultures, an approach which is tedious, labour- intensive, and expensive, to the use of large ‘unit process’ systems.
Although unit processes are more cost-effective and efficient, achieving the necessary scale-up has required a series of modifications to overcome limiting factors such as oxygen limitation, shear damage, and metabolite toxicity. Another aspect of scale-up that will be discussed is increasing unit cell density 50- to 100-fold by the use of cell immobilisation and perfusion techniques.
Free suspension culture offers the easiest means of scale-up because a 1 litre vessel is conceptually very similar to a 1000 litre vessel. The changes concern -the degree of environmental control and the means of maintaining the correct physiological conditions for cell growth, rather than significantly altering vessel design.
Monolayer systems (for anchorage-dependent cells) are more difficult to scale-up in a single vessel, and consequently a wide range of diverse systems have evolved. The aim of increasing the surface area available to the cells and the total culture volume has been successfully achieved, and the most effective of these methods.
General Methods and Culture Parameters of Animal Tissue Culture:
Familiarity with certain biological concepts and methods is essential when understanding scale-up of a culture system. In small scale cultures there is leeway for some error. If the culture fails it is a nuisance, but not necessarily a disaster. Large scale culture failure is not only more serious in terms of cost, but also the system demands that conditions are more critically met. This article describes the factors that need to be considered as the culture size gets larger.
i. Cell Quantification:
Measuring total cell numbers (by haemocytometer counts of whole cells or stained nuclei) and total cell mass (by determining protein or dry weight) is easily achieved. It is far more difficult to get a reliable measure of cell viability because the methods employed either stress the cells or use a specific, and not necessarily typical, parameter of cell physiology. An additional difficulty is that in many culture systems the cells cannot be sampled (most anchorage-dependent cultures), or visually examined, and an indirect measurement has to be made.
ii. Cell Viability:
The dye exclusion test is based on the concept that viable cells do not take up certain dyes, whereas dead cells are permeable to these dyes. Trypan blue (0.4%) is the most commonly used dye, but has the disadvantage of staining soluble protein. In the presence of serum, therefore, erythrocin B (0.4%) is often preferred.
Cells are counted in the standard manner using a haemocytometer. Some caution should be used when interpreting results as the uptake of the dye is pH-and concentration-dependent, and there are situations in which misleading results can be obtained. Two relevant examples are membrane leakiness caused by recent trypsinisation and freezing and thawing in the presence of dimethyl sulfoxide.
A colorimetric method using the MTT assay can be used both to measure viability after release of cytoplasmic contents into the medium from artificially lysed cells, and for microscopic visualisation within the attached cell.
iii. Indirect Measurements:
Indirect measurements of viability are based on metabolic activity. The most commonly used parameter is glucose utilisation, but oxygen utilisation, lactic or pyruvic acid production, or carbon dioxide production can also be used, as can the expression of a product, such as an enzyme.
When cells are growing logarithmically, there is a very close correlation between nutrient utilisation and cell numbers. However, during other growth phases, utilisation rates, caused by maintenance rather than growth, can give misleading results.
The measurements obtained can be expressed as a growth yield (Y) or specific utilization/respiration rate (Q)-
Growth yield: Change in biomass concentration (dx)/Change in Substrate concentration (ds)
Specific utilisation/respiration rate (qa) = Change in substrate concentration (ds) time (dt) x cell mass/numbers (dx)
Typical values of growth yields for glucose (106 cells/g) are 385 (MRC-5), 620 (Vero), and 500 (BHK).
A method which is not so influenced by growth rate fluctuations is the lactate dehydrogenase (LDH) assay. LDH is measured in cell- free medium at 30° C by following the oxidation of NADH by the change in absorbance at 340 nm. The reaction is initiated by the addition of pyruvate (2).
One unit of activity is defined as 1^mol/min NADH consumed. LDH is released by dead/dying cells and is therefore a quantitative measurement of loss of cell viability. To measure viable cells a reverse assay can be performed by controlled lysis of the cells and measuring the increase in LDH.
Equipment and Reagents of Animal Tissue Culture:
1. Culture Vessel and Growth Surfaces:
The standard non-disposable material for growth of animal cells is glass, although this is replaced by stainless steel in larger cultures. It is preferable to use borosilicate glass (e.g., Pyrex) because it is less alkaline than soda glass and withstands handling and autoclaving better.
Cells usually attach readily to glass but, if necessary, attachment may be augmented by various surface treatments. In suspension culture, cell attachment has to be discouraged, and this is achieved by treatment of the culture vessel with a proprietary sili-cone preparation (siliconisation).
Examples are Dow Corning 1107 (which has to be baked on) or dimethyldichlorosilane (Repelcote, Hopkins and Williams) which requires thorough washing of the vessel in distilled water to remove the trichloroethane solvent.
Complex systems use a combination of stainless steel and silicons tubing to connect various components of the system. Silicone tubing is very permeable to gases, and loss of dissolved carbon dioxide can be a problem. It is also liable to rapid wear when used in a peristaltic pump. Thick-walled tubing with additional strengthening (sleeve) should be used. Custom made connectors should be used to ensure good aseptic connection during process operation.
Safe removal of samples of the culture at frequent intervals is essential. An entry with a vaccine stopper through which a hypodermic syringe can be inserted provides a simple solution, but is only suitable for small cultures. Repeated piercing of the vaccine stopper can lead to a loss of culture integrity.
The use of specialised sampling devices, also available from fermented supply companies, is recommended. These automatically enable the line to be cleared of static medium containing dead cells, thus avoiding the necessity of taking small initial samples which are then discarded, and increasing the chances of retaining sterility.
Air filters are required for the entry and exit of gases. Even if continuous gassing is not used, one filter entry is usually needed to equilibrate pressure and for forced input or withdrawal of medium. The niters should have a 0.22 jim rating and be non-wet table.
2. Non-Nutritional Medium Supplements:
Sodium carboxymethylcellulose (15-20 centrepoise, units of viscosity) is often added to media (at 0.1%) to help minimise mechanical damage to cells caused by shear forces generated by the stirrer impeller, forced aeration, or perfusion. This compound is more soluble than methylcellulose, and has a higher solubility at 4°C than at 37 °C.
Pluronic F-68 (trade name for polyglycol) (BASF, Wyandot) is often added to media (at 0.1%) to reduce the amount of foaming that occurs in stirred and/or aerated cultures, especially when serum is present.
It is also helpful in reducing cell attachment to glass by suppressing the action of serum in the attachment process. However, it’s most beneficial action is to protect cells from shear stress and bubble damage in stirred and sparged cultures, and it is especially effective in low serum or serum-free media.
Practical Considerations of Animal Tissue Culture:
i. Temperature of Medium:
Always pre-warm the medium to the operating temperature (usually 37°C) and stabilise the pH before adding cells. Shifts in pH during the initial stages of a culture create many problems, including a long lag phase and reduced yield.
ii. Growth Phase of Cells:
Avoid using stationary phase cells as an inoculum since this will mean a long lag phase, or no growth at all. Ideally, cells in the late logarithmic phase should be used.
iii. Inoculation Density:
Always inoculate at a high enough cell density. There is no set rule as to the minimum inoculum level below which cells will not grow, as this varies between cell lines and depends on the complexity of the medium being used. As a guide, it may be between 5 x 104 and 2 x 105 cells/ml, or 5 x 103 and 2 x 104 cells/cm2.
iv. Stirring Rate:
Find empirically the optimum stirring rate for a given culture vessel and cell line. This could vary between 100-500 r.p.m. for suspension cells, but is usually in the range 200-350 r.p.m., and between 20-100 r.p.m. for micro carrier cultures.
v. Medium and Surface Area:
The productivity of the system depends upon the quality and quantity of the medium and, for anchorage-dependent cells.