Annex B
Cleaning, Decontamination
and Waste Disposal
General
1. The agent thought to be responsible for CJD is recognised as being particularly resistant to standard physical and chemical methods of inactivation and decontamination. The standard autoclave regimen of 121°C for 15 minutes is ineffective, and autoclaving at 134°C for 3 minutes cannot be relied upon for equipment decontamination. Gases such as ethylene oxide and formaldehyde are ineffective, as are some chemical disinfectants such as alcohols, formalin, aldehydes (such as glutaraldehyde), ß propiolactone, hydrogen peroxide, iodophors, peracetic acid and phenolics. Sodium hypochlorite has been shown to be effective but at concentrations that pose certain practical constraints. Sodium hydroxide has a substantial effect but is not completely inactivating. Ionising or UV irradiation at conventional doses and dry heat are also not effective. The effectiveness of other processes and agents such as gas plasma have yet to be fully evaluated. Table B.1 gives a summary of recommended processes and agents, and Table B.2 those shown to be ineffective.
2. As many of the standard methods of decontamination cannot ensure complete inactivation of the agent, the emphasis must be on removal of the agents by thorough cleaning. This should be followed by an appropriate autoclaving or liquid chemical treatment as described below. Table B.3 gives a summary of the basic precautions for decontamination.
3. It is important to note that the advice here for the decontamination of instruments refers only to instruments that have been used on at risk patients where there has not been involvement of the brain, spinal cord or eye. All instruments used on known or suspect patients, and those used on at risk patients where there has been exposure to brain, spinal cord or eye must be disposed of by incineration. For suspect patients quarantine procedures can be used to hold the instruments pending confirmation of the diagnosis (see paragraph 4.28 in the main text).
4. Manufacturers of CE-marked re-usable medical devices are required to supply information on the appropriate processes to allow re-use (Medical Devices Regulations, 1994 - see list of references at the end of this Annex). Users should consult this information to ensure that the instruments are able to withstand the required decontamination processes, which are more rigorous than the processes normally used for re-processing. If there is any doubt, the manufacturer of the instrument or equipment should be contacted for further advice.
Table B.1
Chemicals & process RECOMMENDED for use against TSE agents |
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| Chemical disinfectants | Gaseous disinfectants | Physical processes | |
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20,000ppm available chlorine of sodium hypochlorite for 1 hour
2M sodium hydroxide for 1 hour* | none | porous load steam steriliser 134-137oC for a single cycle of 18 minutes, or 6 successive cycles of 3 minutes each*.; |
For histological samples only, 96% formic acid for 1 hour |
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| * but known not to be completely effective.
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Table B.2
Chemicals & process INEFFECTIVE against TSE agents |
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| Chemical disinfectants | Gaseous disinfectants | Physical processes | |
|
| alcohols | ethylene oxide | dry heat |
| ammonia | formaldehyde | |
| ß-propiolactone | | |
| chlorine dioxide | | ionising, UV or microwave |
| formalin | | radiation |
| glutaraldehyde |
| hydrochloric acid | | moist heat at 121oC |
| hydrogen peroxide | | for 15 minutes |
| iodophors |
| peracetic acid |
| phenolics |
| sodium dichloroisocyanurate |
| (e.g. ‘Presept’)** |
| 10,000ppm sodium hypochlorite | |
|
| ** the rate of release of chlorine from this product is insufficient to ensure complete inactivation of the agent. |
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Table B.3
Basic precautions for disinfection and decontamination |
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- Clean instruments thoroughly at least twice to remove body fluids prior to disinfection.
- Use automated decontamination processes where possible, and avoid mixing routine instruments with those used in TSE-related work in the same cycle.
- Recycle durable items for re use only after appropriate decontamination use only stringent autoclaving procedures or recommended chemical disinfection methods.
- Where possible, cover surfaces with disposable material, which can then be removed and incinerated; otherwise clean and decontaminate surfaces thoroughly use only recommended decontamination procedures.
- Use absorbent material to soak up spillages, which can then be contained and incinerated.
- Use secure leak-proof containers, e.g. double bagging, for the safe handling of clinical waste.
- Avoid external contamination of the waste container.
- Wear protective clothing at all times.
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Cleaning contaminated instruments
5. Manual handling of contaminated instruments should be kept to a minimum and automated decontamination processes, as described below, should be used wherever possible.
6. The cleaning of contaminated items to remove all body fluids and tissues in which a transmissible agent may be present is critical in ensuring the effectiveness of the decontamination regime. All items should be cleaned at least twice before treatment with moist heat or liquid chemicals. The first clean should be carried out in an ultrasonic cleaner; the second in an automated thermal washer/disinfector. Neutral or enzymatic detergent suitable for use with this processing equipment should be used.
7. Contaminated instruments should be processed through a covered ultrasonic bath and automated washer/disinfector in which no other instruments are being cleaned.
8. Items should be cleaned as soon as possible after use to minimise drying of blood and body fluids onto the item, which may then be more difficult to remove. Items should not be soaked in disinfectants prior to cleaning. Further details on the use of ultrasonic cleaners, thermal washer/disinfectors and manual cleaning are contained in the guidance from the Microbiology Advisory Committee on decontamination (see list of references at the end of this Annex).
9. Staff carrying out the cleaning and subsequent processing of instruments and equipment should follow standard basic precautions for avoiding exposure to infectious material (e.g. use protective clothing, cover abrasions with waterproof dressings, avoid use of sharps).
Decontaminating the cleaning equipment
10. Following processing of instruments, the ultrasonic bath and automated washer/disinfector should be run through an empty cycle. Any solid waste/tissue should be disposed of by incineration. Liquid waste should be disposed of safely, either by normal direct discharge from automated washers, or by collection and inactivation from equipment such as ultrasonic baths. Any cleaning aids such as brushes, if used, should be disposed of by incineration.
Autoclaving
11. After cleaning, the items should be processed in a porous load (high vacuum) steam sterilizer using one of the following cycles. Downward or upward displacement autoclaves must not be used:
- a single cycle of 134-137°C for a minimum holding time of 18 minutes; or
- 6 successive cycles of 134-137°C for a minimum holding time of 3 minutes for each cycle.
Other cycles are not recommended.
12. The sterilizer should have been validated and routinely tested in accordance with HTM 2010 (see list of references at the end of this Annex).
13. Whilst it has been suggested that the above cycles may not be entirely effective (Taylor et al 1994 - see list of references at the end of this Annex), it is considered that a substantial reduction in the level of contamination, i.e. the two-step process of cleaning to remove most of the body fluids or tissues in which the transmissible agent may be contained and protected, followed by processing through one of the above moist heat cycles, will be sufficient.
14. Recent work has suggested that a combination of autoclaving and chemical treatment may be effective (Taylor et al 1997 - see list of references at the end of this Annex). Further work is required to support this approach before it can be recommended.
Treatment of instruments with liquid chemicals
15. If the instrument is unable to tolerate the moist heat porous load cycles specified above, then liquid chemical treatment may be considered.
16. Chemical agents and contact times that have been found to be most effective include:
- 20,000ppm available chlorine of sodium hypochlorite for 1 hour;
- 2M sodium hydroxide for 1 hour.
Notes:
- 10,000ppm hypochlorite must not be used as it is ineffective against the agents of TSE at this concentration.
- Sodium dichloroisocyanurate (commonly used as 'Presept' tablets) has been shown also to be ineffective.
17. However, these chemical agents at the concentrations and contact times specified may have a detrimental effect on clinical instruments and equipment, and should only be used after seeking advice from the manufacturer of the instrument to ensure the item will withstand these corrosive processes (see paragraph 4 of this Annex).
Surface decontamination and the management of spillages
18. The above disinfectants should be used for cleaning surfaces. For the decontamination of surfaces, repeated wetting with the disinfectant is necessary over the treatment period. As this concentration of hypochlorite can be corrosive for some commonly used surface finishes, work that involves the handling of infected material should be conducted only on resistant surfaces or work benches shielded by disposable absorbent plastic-backed coverings. The use of enamel, heat-stable plastic or disposable trays is recommended to confine contamination. These should be autoclaved, and the disposable items incinerated after use.
19. For minor spillages, the surface should be disinfected as above. For spillages of larger volumes of liquid, absorbent material should be used to absorb the spillage (a number of proprietary absorbent granules are available for such use). The surface should be disinfected as above, and all waste disposed of as clinical waste by incineration. Disposable gloves and an apron should be worn when removing any spillage and these should be disposed of by incineration.
Inactivation of samples
20. All tissues for histological examination should be immersed in 96% formic acid for 1 hour after routine fixation, unless they have been exposed to phenol. If samples have been exposed to phenol it is not considered safe to then expose them to formic acid. Such samples should therefore be handled as un-decontaminated tissue.
21. Paraffin sections from blocks of tissue not previously decontaminated should be immersed in 96% formic acid for 5 minutes after de-waxing.
22. Clinical samples, e.g. CSF, should be autoclaved or immersed in a solution of sodium hypochlorite resulting in 20,000ppm free chlorine for 1 hour before final disposal by incineration.
Decontamination of safety cabinets
23. Formalin, or rather, in this context, gaseous formaldehyde, which is the conventional medium for the fumigation of safety cabinets, is not effective against TSE agents. Nonetheless, fumigation will need to be carried out as a precaution against other infectious agents that may be impacted on the surface of the cabinet's HEPA filter. The unit should be decontaminated before changing filters.
24. Due to the difficulties associated with their decontamination, it is recommended that safety cabinets used for work with TSE agents should be of the type with the facility for removing HEPA filter units by bagging. Whether or not bagging of the filter as it is withdrawn is possible, spraying the filter face after fumigation and before removal with 'eg hair spray' will help to limit the shedding of particulate matter. Where a Class II cabinet (BS:5762:1992) is to be used, a model that has the main HEPA filter immediately below the work surface is preferred, as this will prevent contamination of the plenum of the cabinet. With the filter in this position, use may be made of liquid latex to seal the filter surface before removal. Pre filters (dust filters) are generally easily removed, and after immersion treatment with 2M sodium hydroxide solution (see above) to limit dust dispersal they should be contained securely for incineration or safe transport to the autoclave. If made of durable but not heat stable material, they may, alternatively, be treated with hypochlorite solution containing 20,000 ppm available chlorine.
25. Working in a shallow tray in the cabinet will limit dispersal onto work surfaces by splashing, but it is essential to ascertain, by testing the cabinet with the tray in situ, that containment for operator protection is not affected (see BS 5726:1992 for detail of containment testing). Another option is to tape disposable plastic-backed absorbent paper to the working surface in order to minimise contamination. The covering must be renewed regularly (preferably after each period of work) and incinerated.
Waste disposal
26. All material classified as clinical waste should be disposed of by incineration at an authorized incineration site. For the safe handling of clinical waste, use secure leak-proof containers, e.g. double bagging, where appropriate. Avoid external contamination of the container.
References in Annex B
| 1. | Medical Devices Regulations 1994 (SI 1994 No 3017). |
| 2. | Department of Health. Sterilization, Disinfection and Cleaning of Medical Equipment: guidance on decontamination. Microbiology Advisory Committee to the Department of Health Medical Devices Agency, Part 1: Principles, Part 2: Procedures. |
| 3. | Department of Health (1994). HTM 2010 Part 3: Validation and verification: Sterilization. HMSO. |
| 4. | Taylor DM, Fraser H, McConnell I, Brown KL, Lamza KA and Smith GRA. (1994). Decontamination studies with the agents of bovine spongiform encephalopathy and scrapie. Arch Virol 139 313-326. |
| 5. | Taylor DM, Fernie K and McConnell I. (1997). Inactivation of the 22A strain of scrapie agent by autoclaving in sodium hydroxide. Vet Microbiol 58 87-91. |
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