1.1. Cautions

Bacterial spores are designed by nature to survive in the face of adverse conditions (i.e. levels of heat, radiation, desiccation, acidity, alkalinity and other chemical and physical conditions) that would rapidly kill other forms of life. It follows, therefore, that chemicals and procedures which can kill spores are necessarily highly lethal to less hardy cells, including those in human, animal and plant tissues.

The most widely used sporicides are chlorine (as in hypochlorite solutions or “bleach”) and formaldehyde, with some use being made of hydrogen peroxide and other oxidizing agents, or glutaraldehyde. At the concentrations necessary to be effective as sporicides, these are potentially hazardous to human health if handled incorrectly.

Precautions, therefore, should be taken not to get these on skin or into the eyes or, especially with the aldehydes, not to inhale them. In the case of fumigation, the work should only be carried out by trained professionals with appropriate protective clothing and breathing apparatus.

Attention is drawn to the importance of handling the concentrated liquid disinfectants referred to with caution, using gloves and aprons or overalls and goggles or eye shields to prevent contact with skin or eyes. Clean water should be at hand for immediate washing or showering in the event of an accident while handling concentrated disinfectants. All containers of disinfectants should be properly and accurately labelled as to their contents. Peroxides may be explosive under certain circumstances.

Appropriate (chemical) respirators should be worn by personnel disinfecting or fumigating closed spaces (rooms, stables, etc.) and when opening up such places to ventilate them at the end of the disinfection or fumigation procedure. Respirators should be fitted and tested by qualified personnel, and users of respirators should be trained in their correct use by qualified personnel.

Irradiation by gamma ray or particle bombardment should only be done by properly trained persons in properly monitored facilities. In the case of UV irradiation, care should be taken to protect the eyes and not to expose eyes or skin to direct UV light sources.

Further cautions are given as appropriate in the sections that follow.

1.2. Choice of disinfectants, fumigants or procedures

Probably because B. anthracis is essentially an obligate pathogen and depends for the continued existence of the species on the survival of the spores in the environment between infections of successive hosts, it appears to produce particularly tenaceous spores. The results are that (i) only a few fairly formidable chemicals and procedures are capable of reliably killing anthrax spores, and (ii) information on the sporicidal activities of disinfectants, fumigants, and disinfection and fumigation procedures based on other Bacillus species should be viewed with caution.

If heat treatment or incineration of the contaminated material is possible, this should be done in preference to chemical decontamination and disinfection. For certain materials or animal by-products, irradiation with gamma rays or particle bombardment may be appropriate (see 6.6 below). As noted in Annex 1, section 7.9, UV irradiation should not be relied on alone for decontamination, but should be used in conjunction with wiping down items to be decontaminated with hypochlorite or possibly formalin.

2.1. Titration of available chlorine in hypochlorite solutions

Hypochlorite is a strong oxidizing agent and will oxidize iodide ions to form elemental iodine. The iodine so formed may be titrated with standard sodium thiosulphate using starch solution as an indicator. The starch solution can be made by making a paste of 0.1 g of soluble starch with a little water and transferring the paste to 100 ml of boiling water. Boil for one minute. Allow the solution to cool and add 2–3 g of potassium iodide. The solution should be kept in a stoppered bottle.

1. The chlorine solution to be tested should be diluted to an estimated 10 000 ppm.
2. Fill a clean 50 ml burette with 0.1M sodium thiosulphate solution.
3. Accurately pipette 5 ml of the solution being tested into a clean flask and acidify with 5 ml of glacial acetic acid. Then add approximately 0.2 to 0.3 g of potassium iodide to the solution which now becomes orange in colour.
4. Titrate the mixture by adding the sodium thiosulphate from the burette until the colour is pale yellow.
5. Add 5 drops of the starch solution and continue the titration until the blue colour of the starch is just detectable (it will look slightly pink now).
6. Note the burette reading and then continue to add the sodium thiosulphate dropwise. The burette reading immediately preceding the observation of a colourless solution is the end point. Note the volume of sodium thiosulphate added from the burette and calculate the available chlorine in the test solution from the expression:

   1 ml 0.1M sodium thiosulphate ≡ 0.00355 g chlorine.

Correcting for the original dilution of the concentrated sample and converting to a percentage:

3.1. Fumigation of rooms

(Caution: this should only be done by trained professionals using PPE that includes a full-face respirator, fitted with a chemical filter and pretested for effectiveness. A formaldehyde dosimeter should be available also.)

Note: formaldehyde is a gas which is soluble in water. The solution of formaldehyde in water is named “formalin”. Fully saturated (100%) formalin has a concentration of approximately 37% formaldehyde. For simplicity, concentrations of formalin are used where possible below. So, for example, 10% formalin would be a 3.7% formaldehyde solution.

Rooms where surfaces cannot be cleared before decontamination and disinfection, such as laboratories, can be fumigated by boiling off (for rooms up to 25–30 m³) 4 litres of 10% formalin in an electric kettle (fitted with a timing or other device to cut off the electricity when the fluid level has reached the element) and leaving overnight (or no less than 4 hours from the point in time when the boiling process has been completed) before venting.

Alternatively, paraformaldehyde can be vaporized in a pan on an electric element on the basis of 12 g per m³ with simultaneous evaporation of 4 litres of water to supply the necessary humidity. For formaldehyde fumigation, room temperature should be > 15 °C.

(Caution: vaporization of formalin or paraformaldehyde should not be done with gas or other naked flame heaters; formaldehyde is flammable. Avoid skin contact with formaldehyde solution or inhalation of formaldehyde vapour.)

Neutralization of formaldehyde can be carried out by vaporizing 15.5 g of ammonium bicarbonate per m³ or 13 g of ammonium carbonate per m³ in a second pan on an electrically heated element. An electric fan will assist in circulating the ammonia, but it may still be 24–48 hours before the room can be entered without a respirator.

(Note: ammonium carbonate and bicarbonate are hygroscopic. If they have become damp, greater weights than those given above should be vaporized to compensate.)

The presence of absorbent material in the room (paper, cardboard, fabric, etc.) reduces the rate of clearance and, indeed, can reduce the effectiveness of the fumigation process. Where there is extensive absorbent material present, the exposure time and possibly the starting concentration of the formalin or paraformaldehyde should be raised to compensate.

Before fumigation commences, all windows, doors and other vents to the outside should be sealed with heavy adhesive tape. Hazard warning notices should be posted on the door(s) and, if appropriate, windows. To ensure complete access of the fumigant, items of equipment should be held above bench or floor surfaces by racks or by tilting to allow the fumigant to penetrate underneath. Proper chemical respirators should be on hand and at least one nitrocellulose disk or filter paper which has, beforehand, been dipped in a spore suspension and dried should be placed at some point in the room distant from the kettle. Preferably the spore preparation should be an accepted standard, such as B. subtilis var globigii (NCTC 10073) or B. cereus (ATCC 12826), but failing the availability of these, the spores of the Sterne vaccine strain (34F₂) of B. anthracis would do.

At the end of the fumigation, the spore disc(s) should be retrieved into a sterile petri dish and the windows or vents to the outside air should be opened up. (Caution: a chemical respirator should be used for this. Respirators should be fitted and tested by qualified personnel and users of respirators should be trained in their correct use by qualified personnel.)

A fan, or fans, assists the extraction. Doors into the room should be kept closed and other personnel prevented from passing near or through them until venting is complete. If a formaldehyde meter is available, venting should not be considered complete until levels of less than 2 ppm have been reached. In the absence of a meter, the odour of formaldehyde should have become almost undetectable before entry into the room without a respirator is allowed.

The effectiveness of the fumigation procedure is checked by placing the spore disc(s) on plates of nutrient agar. In the case of formaldehyde fumigation, the nutrient agar should contain 0.1% histidine final concentration and be added as a filter-sterilized solution after the agar has been autoclaved and cooled to 50 °C. After overnight incubation at 37 °C, if fumigation was properly effective, the discs should show no bacterial growth.

Formaldehyde is becoming regarded as unacceptable in some places, at least for fumigation of large spaces, in particular in the USA on the basis of potential carcinogenicity. Following a peer-review process involving representatives of industry, academia and government, chlorine dioxide gas was identified by the United States Environmental Protection Agency as the best option for fumigation of buildings contaminated by the deliberate release events of October–November 2001 in the USA.

This requires the mixture of two solid precursor chemicals, sodium chlorite (NaClO) and sodium chlorate (NaClO₃), to produce chlorine dioxide (ClO₂) gas. The gas is an unstable compound and potentially explosive in air concentrations > 10 % v/v. It should therefore only be used as a fumigant by qualified personnel using the appropriate personal protective equipment, and generating equipment and a method based on liquid starting chemicals which cannot generate explosive concentrations (> 10% v/v).

Of the other effective sporicidal fumigants (vaporized hydrogen peroxide, methyl bromide, ethylene oxide), probably only vaporized hydrogen peroxide would be appropriate for attempts at room fumigation. It is by far the most ecologically acceptable, with the degradation products being oxygen and water. However it can be anticipated that more than one fumigation session may be needed before spore-strip tests pass completely. The process again requires the appropriate generating and personal protective equipment and should only be carried out by professionally qualified personnel.

3.2. Disinfection in rooms, animal houses, vehicles, etc

Where fumigation is not an option, or following fumigation of a facility, such as an animal room, containing extensive soiled matter, disinfection should be carried out in a three-step process aimed at (i) preliminary disinfection, (ii) cleaning, and (iii) final disinfection. (Caution: protective clothing, including eye cover and, at least with formalin, a combination chemical and biological respirator should be worn. Respirators should be fitted and tested by qualified personnel and users of respirators should be trained in their correct use by qualified personnel. Skin and eye contact with the disinfectants listed below or inhalation of their vapours should be avoided.)

3.3. Fumigation of safety cabinets; fumigation chambers

(Caution: a full-face respirator fitted with a chemical filter and a formaldehyde dosimeter should be on hand for this procedure. Respirators should be fitted and tested by qualified personnel and users of respirators should be trained in their correct use by qualified personnel.)

4.1. Chlorine solutions

Commercially-prepared hypochlorite as supplied to laboratories, hospitals, etc. frequently takes the form of stock solutions having approximately 10% available chlorine (100 000 ppm). Thus, what is familiarly referred to in laboratories as “10% hypochlorite solutions” is a 1:10 dilution of the stock solution containing 10 000 ppm available chlorine. (Note: “bleach” as sold in stores and supermarkets is frequently less concentrated, usually with 3%–5% available chlorine. This needs to be taken into account when making up daily working solutions.) If solid precursors of hypochlorous acid is available, stock solutions containing 100 000 ppm available chlorine should be prepared and the required dilutions made from this.

Unless a stabilizer such as 0.1% sodium carbonate is included, chlorine solutions are not highly stable and stock solutions should be titrated periodically to ensure that the correct level of available chlorine is present (see 2.1 above). Since stability is affected by concentration (and also by temperature and pH), subsequent dilutions should be made only as needed and these solutions should be changed frequently (preferably each day, but at least weekly). It should be remembered that chlorine solutions corrode metals and perish rubber, and that chlorine is rapidly neutralized by organic materials, including wood (as in wooden benches), soil, or specimens of blood or tissues.

Simple chlorine solutions are slow to kill spores (Jones & Turnbull, 1996). The sporicidal rate can be increased by using 50% methanol or ethanol to make the dilutions of the stock solution. However, the stability of these mixtures has not been established and, if used, these solutions should be made up fresh each day.

Simple garden-type spray bottles can be used for delivering hypochlorite solutions to surfaces prior to wiping down, although the chlorine will corrode the spring mechanism quite quickly and these spray bottles will need to be acquired and used on a semidisposable basis.

4.2. Rapid turnover items

Pipettes, disposable loops, microscope slides, sampling spoons, etc. may be immersed overnight in hypochlorite solutions with 10 000 ppm available chlorine. Small plastic items (loops, spoons, etc.) should then be transferred to an autoclave bin or bag for autoclaving, or to a bag for incineration. Glass items should be transferred to a sharps container for autoclaving and/or incineration. It is recommended that long plastic pipettes (1, 5, 10, 25 ml, etc.) are also discarded into sharps containers since they readily perforate autoclave bags.

4.3. Benches

Benches should be wiped down after use with hypochlorite solutions containing 10 000 ppm available chlorine. Because of their neutralizing effect on chlorine, wooden benches should be replaced by more suitable materials or covered with plastic or laminated sheeting, or with a proprietary covering designed for the purpose, such as Benchcote^T.¹

4.4. Spills and splashes on surfaces

Some thought should be given to the nature of the material spilled. For example, freshly growing B. anthracis cultures will have few, if any, spores and these will be incompletely dormant and more susceptible to disinfection procedures than, at the opposite extreme, purposely prepared spore suspensions.

In general, spills and splashes of cultures, or of materials known to be, or suspected of being contaminated with B. anthracis on floor, bench or apparatus should be covered with towelling and the towelling saturated with a hypochlorite solution containing 10 000 ppm available chlorine. The towelling should be left in place for at least 30 min before being transferred to an autoclave bin or bag and autoclaved, or to a bag for incineration. Vertical surfaces should be washed or wiped down thoroughly with cloths soaked in this solution. (Caution: the operator should wear gloves and safety spectacles or goggles while doing this.) In the event of substantial spills or splashes of spore suspensions, fumigation would be advisable after the initial hypochlorite decontamination. This would apply to the safety cabinet if the accident occurred within the cabinet, or the room if the accident occurred outside the cabinet.

Solutions of 10% formalin, 4% glutaraldehyde, 3% hydrogen peroxide or 1% peracetic acid are possible alternatives to hypochlorite, but the choice must be weighed against the greater personal protection needed when using these.

4.5. Biosafety cabinets

Decontamination of cabinets has been covered in Annex 1, section 7.9 and in section 3.3 above.

5.1. Spills and splashes on clothing

Rear-fastening laboratory gowns (surgical type) are the best type of overclothes to wear in laboratories working with B. anthracis, and disposable versions are available. In their absence, a plastic apron should be worn over the laboratory coat. Contaminated gowns/aprons/coats should be removed immediately and placed in autoclave bins or bags and autoclaved. Personal clothing that may still be contaminated – shoes, socks/stockings/upper garments if sleeves or collars are contaminated – should be removed as soon as possible and, if possible, autoclaved. Alternatively, they may be fumigated in a cabinet or fumigation chamber (section 3.3.2 above). Ideally, there should be an emergency shower and emergency clothing in the exit area that will allow the individual to put the contaminated clothes into an autoclave bag or bin, shower and dress to leave the area.

5.2. Spills and splashes on skin or in eyes

In case of contact (biological or chemical agents) with eyes, the eyes must be flushed out with copious quantities of water immediately for at least one full minute, preferably with running water. Ideally an eye-wash station should be included in the laboratory design. Avoid rubbing the eyes. The appropriate medical officer should be informed and the affected person kept under observation for at least a week.

In case of skin contact, the gross contamination should be washed off with water into a bowl and the washings subsequently neutralized by adding bleach and autoclaving. The skin should then receive a thorough soap and water wash (at least 2 minutes). The value of washing the skin itself with bleach is debatable since the contact time is too short to be effective. Bleach certainly should not be used on broken skin as it is likely to do more harm than good. Where the skin is broken (including needle-stick punctures), bleeding should be encouraged and the injury washed with copious amounts of water. The appropriate medical officer should be informed and the affected person kept under observation for at least a week.

5.3. Contamination in the mouth

At the outset, laboratory workers should be reminded that mouth pipetting in a microbiology laboratory is unacceptable. For contamination of the mouth with known or possible anthrax organisms, the mouth contents should be immediately spat out followed by thorough mouth washes with water. The appropriate physician should again be informed and the affected person be kept under observation for a week.

5.4. Suspected inhalation

In the event of suspected inhalational exposure, exhalation should be performed as hard as possible. Others present in the laboratory should be informed and, if necessary advised to evacuate the laboratory. The appropriate supervisor, safety officer and medical officer must be notified immediately and decisions on actions made without delay.

5.5. Exposure through sharps accidents

For sharps punctures (e.g. broken glass), gloves should be peeled off immediately and the wound encouraged to bleed under running tap water for 2–5 minutes followed by a thorough soap and water wash (washing wounds with disinfectant is not recommended). See also section 5.2 above.

6.1. Manure, dung, bedding, feed, etc

Where possible, anthrax-contaminated materials to be disposed of, such as bedding, feedstuffs, manure, etc., should be incinerated or autoclaved (121 °C core temperature for 60 minutes). Immersion in 10% formalin for > 12 hours is an alternative, but full penetration of the fluids must be ensured and natural degradation of the fumigant to the point at which the material can be handled in some way will be slow (at least several weeks). Probably a way of neutralizing and degassing the fumigant should be worked out in advance of taking this route. (Caution: avoid skin contact with formaldehyde solutions or inhalation of formaldehyde vapour. See cautions in section 1.1 above.)

Slurry from livestock suffering outbreaks of anthrax may be also disinfected with formaldehyde by adding undiluted formalin with thorough stirring until a final concentration of 10% formalin is reached. The mixture should be left a minimum of four days with stirring for at least one hour each day before being further processed (Williams et al., 1992). (Caution: avoid skin contact with formaldehyde solution or inhalation of formaldehyde vapour. See cautions in section 1.1 above.)

Formalin degrades naturally to formic acid and thence to carbon dioxide and water (Goring & Hamaker, 1972). It is photooxidized by sunlight to carbon dioxide, and it reacts with nitrogen compounds in the air or soil to form formic acid which, in turn, degrades to carbon dioxide. Its half-life in air is generally less than one hour (WHO, 1991). Certain bacteria and yeast are also able to bring about this degradation by means of dehydrogenases. An alkaline pH neutralizes the formic acid and thereby increases the rate of degradation by pulling the equilibrium in that direction. Decomposition is most rapid at pH 7–8 and with added nitrogen (e.g. in ammonium carbonate). Buffering or addition of lime, to counteract the lowered pH as formic acid is produced, will aid degradation. The treated slurry can be spread on uncultivated land and ploughed in or otherwise buried.

6.2. Sewage sludge

Sewage sludge containing effluents from tanneries that process hides from enzootic areas may contain anthrax spores. Dewatered sewage sludge up to a dry-matter content of 8% should be disinfected by bringing to 10 % formalin and retaining for 10 hours or 3% peracetic acid for 30 minutes. The disinfection process is not affected by polyelectrolytes and may be enhanced by lime added for dewatering the sludge (Lindner et al., 1987). Formalin degrades fairly rapidly naturally (6.1 above); degradation in sewage sludge specifically is covered by Dickerson et al. (1954). (Caution: avoid skin contact with formaldehyde or peracetic acid solutions or inhalation of their vapours. See cautions in section 1.1 above.)

6.3. Water

It is difficult to give general advice on treatment of water. The approach chosen depends on what type of body of water is to be treated, the likely extent of the anthrax spore contamination, what volumes are involved and where the water is to go, and what it may be used for after treatment. However, the choices are much the same as with other materials covered within this annex.

Autoclaving is the surest way of killing spores but is only applicable to fairly small volumes of water. Boiling for 20–30 minutes is a generally effective option. As reviewed by Rice et al. (2004), many experimenters have concluded that boiling inactivates B. anthracis spores with the outside time in the order of 12 minutes. Rice et al. found that there was a critical difference in the effectiveness of boiling when there was a lid on the container in which the water was boiled.

Treatment by bringing to a concentration of 10% formalin and retaining for at least 10 hours is feasible for volumes up to about 100 000 litres, as may result from industrial wastes, but holding tanks must be available and methods of neutralization and discharge without danger to the environment must be established. The necessary safety cautions will apply (see cautions in section 1.1 above). Cost is a major factor in this approach also.

The merits of chlorination are debatable; the levels of chlorine necessary to ensure effective killing of spores may be hard to attain in large volumes and, if the body of water is on open ground, it is likely to contain organic matter which rapidly neutralizes the chlorine.

Filtration, as for water treatment, is probably effective as far as the emerging water is concerned, but leaves unsolved the problem of contaminated filter beds.

In general, each situation should be considered on an individual basis and the best solution worked out for the particular circumstances that exist.

6.4. Soil

If possible, soil at the site of an anthrax carcass should be removed up to a depth of 20 cm and incinerated or heat-treated (121 °C throughout for 60 minutes). If this is not possible, it should be disinfected with 10% formalin at 50 litres per m². Where it is necessary to decontaminate soil to greater depths, such as burial sites of anthrax carcasses, 10% formalin should be injected below the soil surface at a rate of 30 ml for every 10 cm of depth at 0.5 m horizontal intervals across the contaminated area. (Caution: avoid skin contact with formaldehyde solution or inhalation of the vapour. See cautions in section 1.1 above.)

It is sometimes not possible to achieve sufficient penetration of even small clods of soil by formaldehyde or other sporicide solutions to result in complete kill of anthrax spores (Turnbull et al., 1996), especially in the case of water–saturated or heavy soils. Decontamination failure may result when attempting chemical disinfection, and the effectiveness of any such attempt should be checked by subsequent culture.

The decision on the best approach to making a contaminated site safe depends substantially on what the site is to be used for in the future. Where it is not feasible to incinerate or chemically decontaminate the soil or to remove it to an incinerator, the alternative is to close or seal off the site. Covering with concrete or tarmac for, say, a car park, is an alternative used in industrialized countries; planting with thorny bushes surrounded by a secure fence can be an aesthetic approach.

Further guidelines are supplied elsewhere (Turnbull, 1996).

6.5. Other materials – clothing, tools, etc.

Where possible, contaminated materials should be incinerated or autoclaved at 121 °C for 60 minutes. In the case of nondisposable items such as clothing, boots, tools, etc., excess dirt should be scraped off into incineration or autoclave bags and the items themselves should be soaked overnight (at least 8 hours) in 10% formalin. (Caution: avoid skin contact with formaldehyde or glutaraldehyde solutions or inhalation of their vapours.) Bleach is a possible alternative if discoloration or corrosion is not of consequence, and there is little organic material left on the items after scraping.

Decontamination and disposal procedures following collection and examination of clinical specimens are covered in Annex 1, sections 7.7 & 7.8.

6.6. Wool, hair or bristles

Disinfection stations exist in a number of countries which import wool, hair or bristles from endemic regions, and the names and addresses of these may be obtained from the relevant veterinary authorities that control imports and exports of animal products. The requirements of an importing country will be specified as part of the approval process for getting a permit. One established disinfecting protocol is the Duckering process. This involves five stages, each of 10 minutes duration at 40.5 °C: (i) immersion in 0.25%–0.3% soda liquor; (ii) immersion in soap liquor; (iii) two immersions in 2% formaldehyde solution (5% formalin); (iv) rinsing in water; and (v) the wool or hair is finally dried in hot air and baled.

In countries where irradiation facilities are available, the preferred approach is to test samples of wool and hair from a consignment and, if positive for B. anthracis, to sterilize the consignment by irradiation. The dose needed to guarantee freedom from viable spores in a contaminated lot is very high; the D₁₀₀ in spore suspensions of 10⁸ to 10¹⁰ per ml have been found to exceed 40 kGy (4 MRad) (Bowen et al., 1996). Calculations of exposure times need to take into account the size and density of the bales being irradiated.

6.7. Hides and skins

No hazard need be expected in situations where hides come from properly supervised slaughtering. Dry hides of uncertain origin within enzootic countries should, on the other hand, be regarded as being of high risk in terms of anthrax. Where possible these should be decontaminated by fumigation (formaldehyde or ethylene oxide) or by irradiation prior to processing. It is considered by some tannery experts, however, that no preprocessing disinfection protocol has been devised for hides and skins that does not damage them (Anon., 1959). However the dehairing stage, which involves sodium sulphide liming with a mixture of sodium sulphide and calcium hydroxide, exposes the skins to a significant level of sodium hydroxide at high pH which probably kills any B. anthracis spores present (Robertson, 1948; Lindner & Böhm, 1985).

Control processes in tanneries should therefore be primarily targeted at stages before dehairing, particularly dust control and treatment of effluent from initial washing and rehydrating stages. In tanneries processing raw hides from anthrax-endemic areas, these effluents should be treated by bringing to 10% formalin and holding for 10 hours, with adequate time being allowed for natural degradation of the formaldehyde before discharging to sewerage. Peracetic acid (3%) for > 30 minutes is an alternative but more expensive treatment.

Precautions should be taken to avoid cross-contamination of hides and skins pre- and post-treatment through appropriate controls on movement of personnel, equipment and the hides themselves.

6.8. Bone, bonemeal, hoof and horn

Feed ingredients of bone, hoof and horn origin imported from endemic countries are still the cause of incidents among livestock in nonendemic importing countries. Similarly, bonemeal in fertilizers is periodically suspected of being the source of anthrax infection in humans and animals. However, in many developed countries anthrax is becoming very infrequent and there has been a significant decline in the occurrence of anthrax from feed contamination following the BSE (bovine spongiform encephalopathy) restrictions on the feeding of animal or ruminant materials to ruminants.

It is considered in most importing countries that mandatory requirements for sterilization of products of animal origin for international commerce would raise the costs of these products disproportionately to the human and animal health risks involved. Consequently few such countries have statutory requirements of this nature. However, some manufacturers consider it standard good practice to sterilize such products before placing them on the market, and this is certainly to be encouraged, particularly if they are to be used as fertilizers on land on which animals will subsequently graze. Similarly it should be a reasonable policy aim in any country to collect and process separately those raw bone, hoof and horn products obtained from regular and supervised slaughtering and those obtained from sources of uncertain origin, which present a higher risk in terms of anthrax.

Long-term control will be dependent on improved and effective control measures in the exporting countries. In the interim, control should depend on close adherence to the Terrestial Animal Health Code (Annex 4).

7.1. In-place incineration

7.3. Commercial incinerators