Plastics sterilisation

Smaller instruments are packed in sterilisation bags, which are made of sterilisation paper and transparent plastic. Sterilisation bags are sealed with equipment designed for the purpose. 

There are several ways to handle Petri dishes. The standard dishes are 15-20 mm deep and 12-15 cm in diameter. They are normally available in glass and also transparent plastic. Petri dishes are individually wrapped. The media is separately sterilised while we add the sterilised media to the Petri dishes in front of a flame. It is recommended to use 20-25 ml... 

It is common to sterilise the media and Petri dishes separately. When the medium is cooled to about 55 °C, in front of a flame or in a laminar flow chamber, lift the lid of the dish enough to pour about 25 ml of the medium to the desired depth and lower the lid in place. It is best to gently move the Petri dish in way that spreads a thin layer of agar uniformly without any ah bubbles. Distribution of media in the Petri dishes should be done in front of a flame. Most plastic Petri dishes are made of polystyrene and are not autoclaveable. Plastic Petri dishes are easily deformed during sterilisation at high temperature. Some plastic dishes can be autoclaved, but they ate more expensive. Please follow the instructions given by the manufacturer or obtain information from catalogues. 

Normal laboratory glassware must first be washed and cleaned. It has to be rinsed with deionised water. The clean glassware is sterilised in an oven set at 200 °C for 1 1 hours. It is suitable to cover glassware with aluminum foil to maintain aseptic conditions after removing the glassware from the oven. If aluminum foil is not available, special heat-resistant wrap paper can be used. The sterile glassware must be protected from the air, which has micro-flora, or any contaminants. Avoid the use of any plastic caps and papers. Detach any labelling tape or other flammable materials, as they are fire hazards. 

The non-technical nature of the problem becomes apparent when we consider a specific example. For instance, plastic bottles, which are tighter and cheaper than those made from glass, have superseded the traditional material in all sectors of the modern drinks industry. In Britain five billion plastic bottles are used a year, which leads to serious environmental problems. They are difficult to recycle or reuse and expensive to dispose of. They cannot be reused because of the need for sterility. Sterilising is done using high temperatures, which would cause softening or even melting if applied to plastics. 

The intensity of ionising radiation at the earth s surface is not high enough to significantly affect plastics, hence radiation exposure tests are only required in connection with applications in nuclear plant and possibly where radiation is used for sterilisation or to induce crosslinking. 

Fermenters with a capacity of over about 10 litres are too heavy to sterilise in autoclaves. Whilst they may still be laboratory-sized, they have to be constructed so that they may be sterilised in situ. They become, as a consequence, pressure vessels and the extensive use of glass becomes impractical and the preferred material of construction is a stainless steel. Seals are typically of silicone or other synthetic rubber or fluorinated plastics, with borosilicate glass being retained for sighting windows. This format is retained for vessels which are far larger than the laboratory scale, and Fig. 5.80 outlines the construction of a typical industrial deep tank fermenter. 

There are a number of well-established systems for the aseptic packaging of liquids. Notable among these are those packs constructed, box form, in situ on the filling line from a cardboard, aluminium, plastic laminate sheet, such as TetraPak or Combi-box. In the TetraPak system, the packing material enters the filling machine from a feed roll the sheet contact surface is sterilised with warm hydrogen peroxide solution it is formed into a tube, and its lower end is heat-sealed across the width the tube is filled, sealed at the upper end, cut and then folded into a box shape. This produces a continuous output of filled cartons with premium utilisation of bulk storage capacity. 

Bottle systems are more varied, whether for glass, polyethylene terephtha-late (PET) or other plastic. Bottles are rinsed with oxonia solution and then sterile water prior to filling. The filler is generally of a non-contact type (it does not touch the bottles) and product is either weighed in or measured volumetrically. Caps are also chemically sterilised (unless a foil closure is used) and applied on a capper monoblocked with the filler, enclosed in a high efficiency pure air (HEPA) filtered enclosure. The filler and final rinser are in a class 100 room and file operator wears full protective clothing to prevent infection of the product. 

There are several machine systems available for aseptically filling glass and plastic bottles for still juices. (Aseptic filling of drink cartons is covered in Section 9.7.) Carbonated drinks are not aseptically filled. There are two main aseptic filling workflows, with a third workflow used less frequently. The first system sterilises the container, fills and seals it the second takes a sealed, precleaned bottle, removes the seal in a sterile environment, fills and re-seals the container. The third system blows a bottle and while it is still sterile fills it and then seals it, all within the same machine, this is known as a form-fill-seaF (FFS) system. 

Figure 9.1 Aseptically operated filling and closing lines for bottles and wide-mouth containers made of glass and plastics. (1) one-lane feed transfer (2) in-feed into bottle sterilisation unit (3) bottle sterilisation unit (4) discharge (5) two-lane transport of bottles to filler (6) aseptic 10-up inline filler (7) closing machine (8) discharge for closed bottles (9) fid punch and in-feed (10) aseptic module.

Cells may be grown in dishes or flasks where the initial inoculum varies from 0.2 X 106 up to 2 X 106. The containers may be glass or plastic. The plastic ware is obtained in sterile wraps from commercial suppliers and is specially prepared for use in cell culture (Fig. 3.2a) (see Appendix 3). The glass bottles are usually medical flat bottles but any bottle with a flat side will do provided it is washed correctly and sterilised before use (see Chapter 8). To a large extent the disposable plastic flask has now replaced the glass bottle. 

Glass-coated and plastic beads are available from Cellon (Bioglas beads), ICN (Rapid Cell G and Rapid Cell P) and Nunc (Biosilon) and do not swell in aqueous solutions. They are sterilised by autoclaving at 121° for 15 min in water. 

Heat-stable solutions, rubber bungs and liners, bottles with plastic caps, ultrafiltration apparatus etc. are all sterilised by steam treatment at elevated pressure. Although the time required to sterilise is usually only about 15 min at 15 lb pressure the cycle time for modem autoclaves is several hours. This is because of the safety precautions built into these machines to prevent the doors being opened until the temperature of liquid within bottles has fallen to 80°C. 

Empty bottles should have their plastic caps only loosely screwed on to allow penetration and escape of steam during the sterilisation cycle. 

For small volumes, 13 or 25 mm diameter, filtration membranes may be fitted into plastic or stainless steel holders (e.g. the Swinnex filter holder made by the Millipore Corp. Appendix 3) which, after autoclaving, are fitted onto a syringe containing the liquid to be sterilised. Care must be taken in the assembling of the membrane in the holder as incorrect assembly leads to the escape of the membrane from its retaining gaskets with subsequent failure of the filtration process. The plastic holders have a limited life time as they distort on autoclaving. 

The methods of preparation of glassware are indicated in Chapter 8, and if sterilisation is monitored as described the glassware should not be a source of contamination. Likewise plasticware is obtained from the manufacturer in a sterile condition. Usually sterilisation of plastic is achieved using ethylene oxide or irradiation procedures and vessels are supplied wrapped in cellophane. 

The solubility of sterilising gases in polymers is important in determining the retention of residues which may, as in the case of ethylene oxide residues, be toxic. The quality control problems of polymers and plastics are considerable. Both the chemical and physical nature of the material has to be taken into account, as well as purity. 

Packaging for sterile products must be effectively contained and sealed to prevent microbial contamination, and must be robust enough to withstand any sterilisation process required. The sterilisation process can affect the leaching of components from the container into the product or affect the physical properties of the container. For example, autoclaving can soften plastic containers, and gamma irradiation can cause certain polymers to cross link. 

The unit dose is either glass or plastic, with the use of plastic form-fill-seal equipment being dominant. However, glass is considered more elegant pharmaceutically in some markets, and can be terminally sterilised where this is a requirement. With form-fill-seal processes, a strip of ampoules is produced, with the neck being much easier to break than glass. The plastic is pervious to moisture transmission, which will occur on storage, especially in dry environments thus an aluminium overwrap may be necessary. 

Semi-solid products have been traditionally packed in collapsible tin tubes. Metal tubes are a potential source of metal particles in ophthalmic products, and so the tubes have to be cleaned carefully prior to sterilisation. Also, the final product must meet limits for the number of metal particles found. Plastic tubes are not suitable because of their non-collapsible nature, which causes air to enter the tube after withdrawal of each dose. However, collapsible tubes made from laminates of plastic, aluminium foil and paper are a good alternative to tin tubes. Laminated tubes fitted with polypropylene caps can be sterilised by autoclaving, whereas tubes fitted with polyethylene caps are sterilised by gamma irradiation. The tubes are usually filled aseptically, sealed with an adhesive and then crimped. 

For ophthalmic products, there is a dilemma because recent market trends show that flexible LDPE plastic dropper bottles are popular with users because they offer several advantages, including ease of administration, better control of drop delivery and lower risk of contamination during patient use plastic dropper bottles are lightweight, yet more robust than glass. However, one disadvantage of LDPE containers is that they cannot withstand terminal heat sterilisation using pharmacopoeial recommended heat cycles. 

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