Systems for Attached Cells 10311 Roller bottles

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The use of rotating bottles for large-scale cell culture was first described in 1933 (Gey 1933), since which time roller systems have been used for culturing a great many different types of attached cell. Clean glass bottles, e.g. 2.5-litre Winchester bottles, were used for many years, but purpose-designed reusable glass bottles or disposable plastic vessels are now available. The inner surface of the bottle or vessel is used as a cylindrical growth surface. Cells are introduced in a limited volume of medium, and the vessel is then placed in a horizontal position on or in an apparatus that will rotate it slowly around an axis parallel to that of the cylindrical growth surface. The cells attach to the vessel wall as it rotates, and are then cyclically immersed in and then removed from the bulk of the medium as the vessel rotates. When not immersed, there is a thin film of medium covering the cells, and the rotation rate must be such that this does not dry out to any significant extent before the cells are re-immersed in the bulk of the medium. Once cell attachment is complete, the amount of medium in the vessel is often increased, typically to around 8-15 % of the vessel's nominal capacity. Rotation rates will depend on the cell line in use and its attachment efficiency. A typical rate would be 0.5-1.0 r.p.m., perhaps with a slower rate (down to 0.1 r.p.m.) used immediately after seeding in order to facilitate cell adhesion. Some workers using cells that attach efficiently have recommended a different approach, using 0.2-0.4 r.p.m. during the attachment phase, then 0.08-0.16 r.p.m. once attachment is complete (Griffiths 1995). Systematic studies have permitted the mixing patterns in roller bottles to be characterized and modeled mathematically (Muzzio et al. 1999; Unger et al. 2000). From these studies and those of others (Tsao et al. 1992), it is clear that for best results optimization of rotation rates should be performed with the vessel/cell/medium combination actually in use. Periodic reversal of the direction of rotation may be useful in facilitating cell adhesion to the vessel walls (Muzzio et al. 1999). In addition, the introduction of a vertical rocking motion can improve mixing within the bulk of the medium (Unger et al. 2000).

Oxygen transfer is relatively efficient in this system, as the cells spend a large proportion of the time with only a thin film of medium separating them from the gas phase within the vessel. However, in order to avoid oxygen limitation in sealed vessels, it has been recommended that the gas-volume/medium ratio should not be less than 5:1 (Griffiths 1995). Alternatively, vented roller bottles are available. Another approach to improving oxygen transfer has been to attach an external recirculation loop to a roller bottle (Berson et al. 2002), but although highly effective this greatly increases the complexity of the system, negating one of the roller bottle's greatest assets: its simplicity.

A single flat-surfaced roller bottle will usually have a surface area of between 490 cm2 and 1800 cm2. This area may be increased by using a design with a ribbed culture surface, which can increase the surface area two to 2.5-fold without increasing the external dimensions of the vessel. A further increase in surface area has been achieved without increasing the dimensions of the vessel by adding concentric cylinders (Knight 1977) or plastic spiral films (Griffiths 2001), but such vessels do not seem to be widely available. Whilst longer vessels can be used, further scale-up is usually achieved simply by increasing the number of units used. This makes scale-up from the laboratory to initial production scale and beyond (as the market, or market penetration, increases) relatively simple, as there is no change in the unit process. However, large numbers of units have to be handled, and thus economies of scale can only be realized through automation of the various steps involved in roller bottle handling. The degree of automation can vary, but the largest-scale industrial systems are almost completely automated and have a capacity of tens of thousands of roller vessels. Such industrial systems have been in use in the vaccine industry since at least the mid-1960s (Nardelli & Panina 1976; Panina 1985). The emphasis now seems to be either on the replacement of human operatives with robots in order to reduce labour costs further and decrease the potential for contamination (Kunitake et al. 1997) or to convert these processes so that they can be run in fermenters.

A number of modern biological medicines have been produced in large-scale roller bottle facilities, including a number of vaccines as well as recombinant erythropoietin (by Amgen, Johnson & Johnson, and Janssen). However, many of these have now been converted to fermenter processes, either with or without microcarriers (see below). Stacked-plate systems NUNC 'cellfactory'/Corning 'CellSTACK

These culture systems contain a number of flat culture surfaces stacked in parallel one above the other within a single unit, in order to increase the culture surface area that can be handled in one operation. Unlike roller bottles, they require no agitation and can be thought of for most purposes as large culture flasks. Initial scale-up is by increasing the number of surfaces within a unit, and there may be up to 40 within a single unit (Figure 10.1). Media and other solutions are added through an access port and distributed between the layers by tilting the unit. However, the requirement to move the whole unit, complete with its contents, into different orientations means that the

Cell Factory Layers

Figure 10.1 A 40-layer cell factory. Photograph courtesy of NUNC.

Nunck Cell Factory
Figure 10.2 Part of a rig for handling four 40-layer cell factories simultaneously, shown with only one cell factory in place. Photograph courtesy of NUNC.

largest unit that can realistically be handled by an unaided operator is a 10-stack (10 parallel culture surfaces). Manual rigs are available to facilitate handling of a 40-stack. Beyond this, scale-up is by the use of multiple units, and larger, electrically operated rigs can be used to handle multiple 40-stacks simultaneously (Figure 10.2).

Although the culture environment in these units is very similar to that in a culture flask, there are a number of disadvantages to the system. Unlike in a culture flask or roller bottle, there is no means of direct access to the culture surface, so removal of cells by scraping, for example, is not possible. The units are made of rigid plastic, and are thus susceptible to damage if mishandled or hit by other objects (in contrast to modern plastic roller bottles, which are more resilient). They are also completely dependent on the integrity of the seals between the multiple components of the unit (again in contrast to roller bottles, which are nowadays made as a single, seamless moulding) and these seals are easy to break by simply adding liquids or gases to a unit too quickly. Until relatively recently, it was also difficult (for the above reason) to add a CO2-enriched atmosphere to the unit, and even more difficult to maintain such an atmosphere in an effective dynamic manner in order to ensure adequate buffering with CO2/HCO3~-buffered media as used with many mammalian cells. This problem has now been overcome by NUNC with the introduction of 'active gassed' cell factories that are engineered with a gas distribution system. A CO2/air mixture is pumped constantly or intermittently at a low flow rate (typically 20 ml per square cm of culture surface per hour) to this distribution system, which ensures even distribution both within and between layers, resulting in efficient buffering and gas exchange without the risk of damaging the unit. Such gas flow rates are easy provided by using a small pump such as would be used in a home aquarium.

'Cell factories' have been available for over 30 years, and have been used with a wide range of cell lines (see Siegl et al. 1984; Goetz et al. 1992; Bishop et al. 1994; Dickinson & Kohwi-Shigematsu 1995; Litjens et al. 1997; Nelson et al. 1997; Ohtaki et al. 1998; Lee et al. 1999; Lay et al. 2000; Suehiro et al. 2000; Loewen et al. 2002) and at the industrial scale for vaccine manufacture (Hagen et al. 1996). As with roller bottles, scale-up and adjustment to market demand has been relatively simple, being achieved by the use of multiple identical layers, and then multiple identical units. However, economies of scale are again difficult to realize other than by automation. Corning CellCube®

Superficially, the CellCube® is similar in principle to the 'cell factory', having at its heart a culture unit composed of multiple flat surfaces bonded together in parallel. However, unlike the 'cell factory', the CellCube® is a perfusion culture device, with medium being continuously circulated through the system. Also, cells attach to both faces of the flat culture surfaces and, once seeded with cells, these culture surfaces are held in a vertical orientation. There is no air space within the system (as there is between each layer of a 'cell factory'), oxygenation/gas exchange being achieved by the passage of the medium through an external oxygenator during recirculation of the medium through the system. It also requires a controller, and a circulation and medium pump. In fact, the true precursor of this system was not the cell factory, but a vertical-plate system using glass plates separated by PTFE spacers that was first described by Mann (1977) and could be scaled up to a surface area of 10 m2. In the modern CellCube®, by using multiple modules the culture surface area can be increased from 8500 cm2 to 340 000 cm2 whilst still using a single controller. For exploratory work and initial process development, a simplified version called the 'E-cube' is available.

The CellCube® has not as yet been widely used for the production of GMP-grade material, although GMP-grade retroviral vectors for gene therapy have been produced in the system (Wikstrom et al. 2004). Multidisc propagators

Industrial requirements, predominantly in the vaccine industry, for large surface-area single units in which cells could be cultured for weeks and from which multiple harvests of (virus-containing) medium could be removed, resulted in the development of the multidisc propagator (Molin & Heden, 1969; McAleer et al. 1975). This normally took the form of a stack of titanium discs, mounted on a spindle with a gap of 2-3 mm between the discs, the whole assembly rotating within a stainless steel vessel partly filled with medium such that the cells were periodically immersed in the medium in a manner analogous to a roller bottle. Typically, an assembly of 100 disks would be contained in a 10-litre vessel (Elliott 1990). However, problems of getting an even distribution of cells on the plates and of confluent sheets sliding off the (vertical) surface of the plates led to a change of configuration, so that the plates were mounted one above the other with their surfaces horizontal and medium being circulated by a pump (Schleicher & Weiss 1968). In this form the systems were scaled up to over 200 litres, but had to be tilted to remove the medium and could only use one of the two surfaces on each plate for cell growth. Thus potential for scale-up was limited. Nevertheless, these vessels in their various forms were used for many years for the production of a variety of vaccines, including those against measles, mumps, rubella and polio (McAleer et al. 1975; Elliott 1990). However, use outside the vaccine industry appears to have been limited, and many of the processes that formerly used this system have been converted to fermenter-based systems using microcarriers, in order to overcome the problems already described, to utilize some of the advantages of microcarriers detailed below, and to achieve further increases in the size of the unit process. Microcarriers

Microcarrier beads were conceived as a way of improving the volumetric efficiency (surface area per unit volume) of culture systems for attached cells. Using these tiny beads (typical diameter 200 |im), surface areas in excess of 30 cm2 per cm3 of culture medium are easily attainable for use in simple batch culture (van Wezel 1967), and higher values can be employed for more intensive

(fed-batch or perfused) cultures. This compares with values of around 3 cm2/cm3 for a T-flask or cell factory. However, if one compares surface area with total culture unit volume (a real measure of the amount of incubator/laboratory space required for scale up), then microcarriers in a spinner flask will attain around 10 cm2/cm3, whereas a 225 cm2 T-flask will give c.0.2 cm2/cm3, and a 10-stack cell factory c.0.55 cm2/cm3.

Originally DEAE-Sephadex A-50 beads were used (van Wezel, 1967), but the density of positive charges on these (equivalent to an exchange capacity of ca 4 meq/g) was too high for optimum cell attachment and growth. Reducing the charge density to about 2 meq/g overcame the problem (Levine et al. 1977, 1979), and nowadays negatively charged and amphoteric surfaces are also used. A wide range of microcarriers is now commercially available (see Table 10.1), made from a variety of materials. Some have special surface coatings to encourage the attachment of particular cell types, while others are made from materials that can be enzymatically digested in order to release the cells with minimum cellular damage.

A number of other physical properties are also important in defining the utility of microcarrier beads:

• Density The beads must be dense enough not to float, but not so dense that they are difficult to keep in suspension. Values between 1.02 and 1.04 g/cm3 are most frequently used (except in fluidized-bed applications).

• Transparency A highly transparent bead material aids microscopic examination of the attached cells.

Table 10.1

Some commercially available microcarriers.


Trade name




Cytodex 1

GE Healthcare

DEAE-substituted crosslinked dextran

Cytodex 3

GE Healthcare

Collagen-coated dextran


Cytopore 1

GE Healthcare

Macroporous DEAE-coupled cellulose

(1.1 meq/g)

Cytopore 2

GE Healthcare

Macroporous DEAE-coupled cellulose

(1.8 meq/g)


Cytoline 1

GE Healthcare

Macroporous, silica-weighted polyethylene,

density = 1.32 g/cm3

Cytoline 2

GE Healthcare

Macroporous, silica-weighted polyethylene,

density = 1.03 g/cm3



Gelatin-coated plastic beads

2D MicroHex


Flat polystyrene hexagons


Solohill Engineering

Collagen-coated polystyrene

Collagen (C)

Solohill Engineering

Collagen-coated polystyrene

Hillex (H)

Solohill Engineering

Trimethylammonium-modified polystyrene

Hillex II (H)

Solohill Engineering

Trimethylammonium-modified polystyrene

Glass (G)

Solohill Engineering

Silica glass-coated polystyrene

Plastic (P)

Solohill Engineering


Plastic Plus (PP)

Solohill Engineering

Polystyrene (cationic)

Pronectin® F (PF)

Solohill Engineering

Recombinant fibronectin-coated polystyrene



Percell Biolytica AB

Crosslinked gelatin


Percell Biolytica AB

Crosslinked gelatin (increased thermal


Pronectin® F is a registered trade mark of Sanyo Chemical Industries.

Pronectin® F is a registered trade mark of Sanyo Chemical Industries.

• Porosity Most microcarrier beads are designed such that cells will grow on their surface. However, some are specifically designed to be macroporous, so that the cells actually grow within the body of the bead. This may have advantages in terms of protecting the cells from bead/bead impacts and high liquid shear environments that can be damaging to cells (Cherry & Papoutsakis 1988), although these mechanisms may only become significant at high agitation rates (Croughan et al. 1987, 1988). However, removal of the cells from the beads by trypsiniza-tion may be more difficult.

• Diameter A large number of beads/cm3 is required, both to ensure that the suspension is essentially homogeneous, and to obtain the required volumetric efficiency. However, some cells find it difficult to migrate between beads, so each bead should have the ability to carry several hundred cells. Given these constraints, an optimum balance is often obtained with bead diameters in the 150-230 |im range. The size distribution should be as small as possible to ensure an even distribution of cells between the beads, otherwise smaller beads may be colonized at the expense of larger ones (van Wezel 1985).

Clearly, microcarrier beads must not be toxic to the cells, a problem during the early years of microcarrier use that has since been overcome (Giard et al. 1977). Similarly, the monomeric material from which they are made, and any other substance (such as surface coatings) liable to leach from the beads, must not be inhibitory to cell growth.

Although most microcarrier beads are spherical, this shape is not essential. Cylindrical cellulose carriers can be used (Reuveny 1990), and NUNC currently market flat, hexagonal polystyrene carriers (see Table 10.1).

One of the advantages of using microcarrier beads is that the culture of attached cells can be carried out in the same type of equipment that is employed for the homogeneous stirred culture of cells growing in suspension (Reuveny 1990), usually with slight modifications to help keep the relatively rapidly sedimenting microcarriers suspended. This has numerous benefits in terms of scale-up and environmental control, and also permits direct sampling and observation of the cells, something that may be difficult if not impossible in some other systems for culturing attached cells. These benefits have led to the widespread use of microcarrier culture, particularly in the vaccine industry where this technology was first commercialized more than 20 years ago (Meignier et al. 1980; Montagnon et al. 1984).

Macroporous microcarriers have also been utilized in both packed-bed (Looby & Griffiths 1988) and fluidized-bed systems. Formerly, a fluidized-bed system using weighted collagen carriers was available from Verax Corp., but although this is no longer available, a similar unit (the Cytopilot) can now be obtained from GE Healthcare for use with their Cytoline carriers (see Table 10.1). Such systems increase volumetric efficiency (see above) beyond that attainable in a homogeneous suspension system (G.E.Healthcare 2002) Packed-bed systems

Packed-bed systems are used widely in the chemical and water treatment industries to obtain a large surface area in the minimum volume. In cell culture, a variety of packing materials has been investigated for attached cells (Spier 1985), with glass beads having been probably the most widely applied (Burbidge 1980; Robinson et al. 1980; Griffiths et al. 1982) and having been scaled up to at least 100 litres (Whiteside & Spier 1981). A simplified diagram of such a system is shown in Figure 10.3. However, packing and stability issues (grinding of the cells on the surface of the beads can occur if the bed is sub-optimally packed, or subject to vibration) and a complex relationship between bead diameter (and hence both surface area and inter-bead channel size), cell yield, uniformity of cell distribution, and medium flow rate (and direction) (Griffiths 1990, 2001) have

Roller Bottles Cell Culture
Figure 10.3 Highly simplified diagram of a glass bead packed-bed culture system.

meant that some of the theoretical advantages of the system have been difficult to realize. The system is best suited to the harvesting of a secreted product over a period of time, as it can be difficult to remove cells from the bed efficiently (Robinson et al. 1980; Griffiths 2001). For further discussion of packed beds, see Section

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