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1 The disease and its importance

Anthrax is primarily a disease of herbivores although few, if any, warm blooded species are entirely immune to it. From earliest historical records until the development of an effective veterinary vaccine mid–way through the present century (Sterne, 1937; Sterne et al., 1939), together with the subsequent advent of antibiotics, the disease was one of the foremost causes of uncontrolled mortality in cattle, sheep, goats, horses and pigs worldwide. Humans almost invariably contract anthrax directly or indirectly from animals. Today the Office international des épizooties, Paris, France (OIE), reports (OIE, 1997a) show that the disease is still enzootic in most countries of Africa and Asia, a number of European countries and countries/areas of the American continent and certain areas of Australia; it still occurs sporadically in many other countries.

2 Etiology and ecology

Anthrax is a bacterial disease caused by the spore–forming Bacillus anthracis, a Gram positive, rod–shaped bacterium (see section 6).

2.1 Spores and vegetative forms

When conditions are not conducive to growth and multiplication of the bacilli, they tend to form spores. Sporulation requires the presence of free oxygen; within the anaerobic environment of the infected host the organism is in the vegetative form.

Although the vegetative forms of B. anthracis grow and multiply readily on or in normal laboratory nutrient agars or broths, the evidence is that they are more "fragile" than the vegetative forms of other Bacillus species, dying more spontaneously in simple environments such as water or milk, and more dependent on sporulation for species survival (Turnbull et al., 1991; Bowen et al. 1992; Lindeque 1994). B. anthracis is, to all intents and purposes, an obligate pathogen.

The spore forms are markedly resistant to biological extremes of heat, cold, pH, desiccation, chemicals (and thus to disinfection), irradiation and other such adverse conditions. Therefore, the spore forms are the predominant phase in the environment and it is very largely through the uptake of spores that anthrax is contracted.

Within the infected host the spores germinate to produce the vegetative forms which multiply, eventually killing the host. A proportion of the bacilli released by the dying or dead animal (see Section 5) into the environment (usually soil under the carcass) sporulate, ready to be taken up by another animal. This cycle of infection is illustrated in Figure 1.

 

Figure 1. Cycle of infection in anthrax. The spore is central to the cycle, although infection
can also be acquired through uptake of the vegetative forms when, for example, humans or carnivores eat meat from an animal that died of anthrax or when biting flies transmit the disease.

The rate and extent of sporulation by vegetative cells shed from infected animals is affected in a complex manner by the environmental conditions into which they fall. Temperature, pH, oxygen availability and the presence of certain cations such as Mn? ? are the principal influencing factors.

Spores will germinate outside an animal if they fall into appropriate conditions, i.e. a temperature between about 8^o and 45^oC, a pH between about 5 and 9, a relative humidity greater than 95% and the presence of adequate nutrients. The extent to which subsidiary cycles of germination, multiplication and resporulation occur in the environment remains a topic of debate, but research (Bowen et al., 1992; Turnbull et al., 1991; Lindeque, 1994) suggests that the level of nutrient required for this to become possible is unlikely to occur very frequently under natural conditions. In fact, it seems that the fragile vegetative forms die rapidly in most environmental conditions, depending on sporulation in a proportion of the population for passing the infection to the next host (Bowen et al., 1992; Lindeque, 1994).

2.2 Climate and other influencing factors

There is little dispute that anthrax is a seasonal disease; its incidence in any one place is related to temperature, rains or drought; however, examination of the literature shows that the conditions which predispose to outbreaks differ widely from location to location. Climate probably acts directly or indirectly by influencing the way in which an animal comes into contact with the spores (for example, grazing closer to the soil in dry periods when grass is short or sparse, or movement of herds to restricted sites when water becomes scarce), or by affecting the general state of health of the hosts and thereby affecting their level of resistance to infection.

Much has been written and hypothesized about the effects of season, rainfall, temperature, soil, vegetation, host condition and population density on the epidemiology of anthrax, but little agreement exists on the roles played by these factors in the incidence of the disease. Most of the theories are based on concepts of conditions under which B. anthracis may germinate and multiply in the environment, but hard scientific supportive data are not readily available and, as said in 2.1, the conditions under which multiplication in the environment could take place probably only occur in exceptional circumstances.

No one model satisfactorily explains the varying observations on the relationship between the factors listed above and the incidence and persistence of anthrax in a locality. This is an area worthy of further structured research.

2.3 Strains

B. anthracis appears to be one of the most monomorphic species known. That is to say, isolates from whatever type of source or geographical location are almost identical phenotypically and genotypically. Phenotypically, strain differences are only apparent in non–quantifiable or semi–quantifiable characteristics, such as colonial morphology, flocculation in broth culture, cell size, LD₅₀ in animal tests, and so on. The biochemical, serological or phage typing methods available in the case of other pathogens have proved of no value for identifying different strains of B. anthracis. At the molecular level, genomic differences have also proved hard to detect, although, some progress is being made in attempts to devise chromosomally–based strain differentiation systems Henderson 1996; Anderson et al. 1996; Keim et al. 1997; Jackson et al. 1997.

It seems reasonable to attribute this exceptional degree of species monomorphism to the fact that B. anthracis encounters opportunities to multiply less often than most other bacterial and pathogenic species. Given the truth of the statements in 2.1 and 2.2 that opportunities to multiply in the environment are rare, multiplication cycles depend almost exclusively on infections in animal hosts – which, in turn, may only occur following considerable time intervals between sequential hosts. Furthermore, since multiplication occurs almost exclusively in the animal host, the vegetative form of the organism is rarely exposed to the mutagens, phages or other environmental factors responsible for strain variation in other bacterial species.

3 Anthrax in animals

3.1 Host range and susceptibility

As said at the outset (Section 1), anthrax is primarily a disease of herbivores. However, reports of its occurrence in dogs scavenging anthrax carcasses and in carnivorous animals in zoological gardens and wildlife sanctuaries or parks are not entirely uncommon, though outbreaks affecting large numbers of carnivorous animals are very rare. Published LD₅₀s for anthrax by the parenteral route range from <10 for a guinea pig through 3 x 10³ for the rhesus monkey, 10⁶ for the rat, 10⁹ for the pig and 5 x 10¹⁰ for the dog (Watson and Keir, 1994). Minimum infectious dose (MID) estimates are only rarely available, but an aerosol MID for sheep of 35 000 spores has been recorded (Fildes, 1943). De Vos and Scheepers (1996) record that 100 spores of a strain isolated from a kudu in the Kruger National Park consistently resulted in death from anthrax when administered parenterally in impala. In another recent study on 50 pigs given doses of 10⁷ to 10¹⁰ spores in feed containing grit (Redmond et al., 1997), the majority showed clinical illness with recovery, and just two died with confirmed anthrax 6 and 8 days respectively after ingestion of the spores; these were estimated to have received 1.6 x 10⁷ and 7.8 x 10⁷ spores respectively.

Since it is thought that animals generally acquire anthrax by ingestion of spores, and that some sort of lesion is necessary for the establishment of infection (see 3.3), LD₅₀s, particularly parenterally determined, only provide a rudimentary guide to natural susceptibility. Extrapolating experimental findings to the natural situation must also be done cautiously, taking into consideration the many factors influencing infectivity, such as the strain of B. anthracis, the route of infection, the specie, breed or strain and state of health of the animal tested, the times and sites at which tests are done and so on.

3.2 Incidence of anthrax in animals

The following information on the occurrence of anthrax in animals has been derived from official reports (OIE, 1997) as well as from data collected from other sources (Hugh-Jones; personal communication).

During the past three decades, there has been a progressive global reduction in livestock cases in response to national programmes. As a result, the disease in animals is now truly absent or only sporadic in the middle and higher latitudes of Europe; it is still common in the European countries adjoining the Mediterranean (Greece, Italy, Spain, Turkey and Yugoslavia). In Canada, apart from its continued incidence in bison in the MacKenzie Bison Range, North West Territory, and in the Wood Buffalo National Park, northern Alberta, it is sporadic in southern Alberta and Saskachewan, with a recent singular outbreak in Ontario. In the USA it is confined to a few persistent pockets with sporadic cases in South Dakota, Nebraska, and Oklahoma but probably a hyperendemic situation persists throughout the south western quadrant of Texas.

The true situation in Latin America awaits definition; under–reporting and failure to diagnose unexpected livestock deaths certainly occurs, especially in small ruminants; it is enzootic in Central America, Mexico and Guatemala, with decreasing occurrence as one goes farther south—it is absent in Belize, as it is throughout the Caribbean with the exception of Haiti. Anthrax is enzootic in Peru, Bolivia and Venezuela.

In South Africa, the annual number of outbreaks is less than 5 and occasionally zero, despite the continued occurrence of the disease in wildlife in the various parks. Good control programmes have been established in Botswana, Zimbabwe and Zambia, but the disease remains well–known in the latter two countries at least. In equatorial African countries, anthrax is enzootic or hyperenzootic.

The Middle Eastern and adjoining countries of former USSR republics continue to experience hyperendemic anthrax. Anthrax is enzootic in southern India but is less frequent to absent in the northern Indian states where the soil is more acid, while in Nepal it is endemic.

Some south–east Asian countries are severely affected—Myanmar, Vietnam, Cambodia, western China—while Thailand used to be free, now it is endemic and continually threatened and afflicted by its neighbour. Malaysia and Taiwan are free, and the disease is limited to single regions of the Philippines and Indonesia. All these countries differ from the rest of the world in that pigs and water buffalo, as opposed to ruminants, are the animals commonly affected.

As countries become free of anthrax or the annual incidence of outbreaks approaches unity, the numbers of animals affected in an outbreak increase. This seems to be due to the decreasing veterinary experience in recognizing cases and in dealing appropriately with outbreaks. The mere absence of reported livestock anthrax does not mean that a country is free of the disease. Reporting deficiencies and insufficient examination of unexpected livestock deaths are common throughout the world. Also, while civil unrest in northern latitudes can be expected to result in a fall in anthrax cases, in warmer countries, especially in Africa and Asia, civil wars invariably result in anthrax epidemics in livestock, which then readily spill over into human populations. The reason for this is that the northern countries import meat and bone meal and other commonly contaminated materials but imports are cut off in times of war. For example, Norway, which was occupied and pillaged in World War II, quickly ceased to experience outbreaks of anthrax; meanwhile the disease continued unabated in neutral Sweden. In contrast, during the civil war in Southern Rhodesia/Zimbabwe at the end of the 1970s, livestock vaccination ceased and by the end of the war there had been some 10 000 human cases, as compared with the previous normal annual rate of a dozen cases or less (Turner, 1980; Davies, 1982).

3.3 Transmission in animals

In economic and public health terms, the importance of the disease lies in its ability to affect large numbers of livestock at one time. Carcasses then pose a hazard to humans and other animals both in the vicinity and at a distance through their meat, hides, hair, wool or bones. Hides, hair, wool and bones may be transported large distances for use in industries, feedstuffs or handicrafts. Livestock may acquire the disease through contaminated feedstuffs or from spores that have reached fields in sewage sludge.

Although anthrax has been recognized for centuries, much remains unknown about the disease, including the precise manner by which grazing animals acquire it. The sporulated forms shed by an animal dying or dead from anthrax generally provide the source of infection of other animals (Figure 1). It is generally assumed that ingestion of the spores while grazing is a frequent mode of uptake; since B. anthracis is apparently non–invasive, it is believed a lesion is necessary for the initiation of infection. In view of associations between times of higher incidence and dry, hot conditions, theories have arisen that, at such times, the animal is forced to graze dry, spiky grass close to the soil. The spiky grass and grit produce the gastrointestinal lesions and if the soil is contaminated with anthrax spores, there is a high chance of infection occurring.

In countries with advanced agriculture, contaminated feedstuffs are or have been a common and significant source of infection, especially for dairy cows. The source can either be improperly treated locally produced meat and bone meals salvaged from moribund stock or imported infected bones/contaminated meat and bone meals. Cross–contamination can occur during shipping if the hulls of cargo ships and other containers are not cleaned out between shipments or if infected dry hides are placed on top of feed; this practice has in the past produced contaminated vegetable high–protein feeds which normally would not be expected to contain B. anthracis spores. Similar cross–contamination following the re–use of feed sacks has resulted in outbreaks of anthrax.

In England, Germany, Italy and Canada, pastures flooded by tannery waste water historically have posed a long–recognized hazard to grazing stock (Figure 2).

The examination of associations between climatic conditions and peak anthrax periods around the world have resulted in an number of other theories. Some contribute to the hypothesis that an animal can harbour the spores for long periods only manifesting the disease when stressed or compromised immunologically; seasonal stress may play a role in this regard. Others believe that acquisition of the disease by inhalation of spore–laden dust is not infrequent. The extent to which this might occur would vary with season. It is widely accepted that, in some regions, certain types of flies transmit anthrax; again this would be associated with season.

 Figure 2. Importation from endemic to other countries and cycles within the importing country.

 

 

3.4 Pathogenesis and symptoms

Except when taken up by the pulmonary route, B. anthracis needs a lesion through which to enter the body. Following entry, the spores, which may have commenced germination, are carried to the lymphatics where they multiply and, in terms used by workers in the 1950s, continuously feed the blood stream with the vegetative bacilli in a manner analogous to continuous culture. Initially, during the incubation period, the bacteria are filtered out by the spleen and other parts of the reticuloendothelial system. However, the system finally breaks down due to toxin action and, during the last few hours of life (fulminant systemic phase), the bacteria build up rapidly in the blood (doubling time approximately 0.75 to 2 hours depending on host species) to levels of >10⁸/ml together with massive toxaemia at the time of death. The action of the toxin on the endothelial cell lining of the blood vessels results in their breakdown, internal bleeding and the characteristic terminal haemorrhage to the exterior which is an essential part of the organism's cycle of infection (Figure 1 and front cover).

The incubation period in the susceptible herbivore ranges from about 36 to 72 hours and leads into the hyperacute systemic phase, usually without easily discernible prior symptoms. The first signs of an anthrax outbreak are one or more sudden deaths in the affected livestock, although farmers may reflect retrospectively that the animals had shown signs such as having been off their food or having produced less milk than usual. During the systemic phase, the animals become distressed, appear to have difficulty breathing and cease eating and drinking. Swellings in the submandibular fossa may be apparent; temperatures may remain normal for most of the period or may rise. The animal can remain responsive to treatment well into this period but if treatment fails it lapses into coma followed by death from shock. In highly susceptible species, the period between onset of visible symptoms and death may be just a few hours; the course of these events is more protracted in more resistant species.

3.5 Diagnosis in animals

The history is of major importance in the diagnosis of anthrax. Clinical manifestations to look for are:

Ruminants: Sudden death, bleeding from orifices, subcutaneous haemorrhage, without prior symptoms or following a brief period of fever and disorientation should lead to suspicion of anthrax.

Equines and some wild herbivores: Some transient symptoms?? fever, restlessness, dyspnoea, agitation?? may be apparent.

Pigs, carnivores, primates: Local oedemas and swelling of face and neck or of lymph nodes, particularly mandibular and pharyngeal and/or mesenteric.

At death in most susceptible species (the pig being a notable exception), the blood contains 10⁷ to 10⁸ bacilli per ml provided the animal has not been treated (numbers may also be lower in immunized animals which succumb to the disease). For reasons unknown, numbers of B. anthracis at death are very low in pigs (hundreds per millilitre or less).

A blood smear should be obtained with a swab from a small incision in the ear or from an ear clipping (the ear is usually recommended as being accessible, supplied with an extensive capillary network), or by means of a syringe from an appropriately accessible vein (the blood characteristically clots poorly or not at all upon death in anthrax victims and is dark and haemolysed). The smear is dried, fixed and stained with polychrome methylene blue (M'Fadyean stain) as described in Appendix I (A.I.2.1.2). Large numbers of the capsulated bacilli will be seen in smears from relatively fresh carcasses of most species (see preceding paragraph). A Gram stain will not reveal the capsule and may result in mistaken diagnosis, particularly if the carcass is not very fresh. B. anthracis does not compete well with putrefactive bacteria and, with increasing age of the carcass, the capsulated bacilli become harder to visualize (see also four paragraphs below).

The bacterium can also be cultured from the blood or from a swab of a ear clipping or other appropriate specimen on blood agar or other nutrient agar (see Appendix I). The haemorrhagic nasal, buccal or anal exudate will also carry large numbers of B. anthracis which can be cultured from swabs or from samples contaminated with the exudate.

If anthrax is suspected the carcass should not be opened; contamination of the environment by spilled body fluids with subsequent spore formation, is thereby avoided. If, mistakenly, the carcass has been opened, the dark unclotted blood and markedly enlarged haemorrhagic spleen are immediately apparent. The mesentery may be thickened and oedematous and peritoneal fluid may be excessive. Petechial haemorrhages may be visible on many of the organs and the intestinal mucosa may be dark red and oedematous with areas of necrosis. Where anthrax has been diagnosed after a carcass has been opened, special attention should be paid to decontamination of the site at which the postmortem was carried out and of the tools and materials that were used (see section 8).

In horses, the intestine and parenchymatous organs may be less affected than in sheep and cattle and the subcutaneous and intramuscular tissues may be oedematous. In pigs, as indicated above, blood smears may not reveal the capsulated bacilli and, if cervical oedema is present, smears and cultures should be made from fluid from the enlarged mandibular and suprapharyngeal lymph nodes. If intestinal anthrax is responsible for death in pigs, this may only become apparent at necropsy; smears and cultures should be made from the mesenteric fluid and lymph nodes. In other animals, in addition to any haemorrhagic exudates that may be observed, severe inflammation and oedematous swelling of the lips, tongue, gums, jowls and throat may be diagnostic indicators.

In the unopened carcass, the bacilli, unable to sporulate in the absence of oxygen, are destroyed by the putrefactive processes. Smears, as a diagnostic procedure, become unreliable about 24 hours after death although "shadons" (capsular material) may still be observed some time after the bacilli themselves can no longer be seen. B. anthracis can often be cultured from carcass skin residues for some days after death but, with increasing length of time between death and examination, this becomes progessively less easy. Diagnosis then becomes increasingly dependent on isolation of spores from soil or other environmental samples contaminated by the oral, buccal or anal exudates.

As mentioned in Appendix I (A.I.1.2), it may not be possible to find the bacilli in smears or to isolate B. anthracis from animals that were treated before death; treatment can sterilize the blood and tissues but, if sufficient toxin has been formed, the animal may still go on to die.

The thermostable antigen precipitin test devised by Ascoli (1911) (see Appendix I [A.I.3.1]) is still used in several countries of Europe and the Far East for detecting residual antigens in tissue in which it is no longer possible to demonstrate B. anthracis microscopically or by culture. However, it should be borne in mind that the antigens being detected are shared by other Bacillus species and that the test relies on the fact that, if Bacillus antigens are present in the tissues, this probably represents B. anthracis infection since infections with other Bacillus species are rare. Care has to be taken if the tissue being examined has been grossly contaminated with environmental materials (soil, sand, etc.) which frequently harbour large numbers of other Bacillus species.

A simple, rapid and highly sensitive and specific chromatographic device, utilising a monoclonal capture antibody detecting the anthrax–specific protective antigen, has now been designed (Burans et al., 1996) and hopefully will become available in the near future as a more reliable and more sensitive alternative to the Ascoli test.

3.5.1 Differential diagnosis

For differential diagnosis, other causes of sudden death should be considered. Amongst these are botulism, blackleg (Clostridium chauvoei), peracute babesiosis, chemical poisoning (heavy metals, other poisons), ingestion of toxic plants, snake bite, lightning strike or metabolic disorders such as lactic acidosis, magnesium deficiency, bloat. A recent outbreak of Rift Valley Fever in Kenya was initially thought to be anthrax (WHO, 1997a).

3.5.2. Retrospective diagnosis (seroconversion)

Historically, there has been little need for serological support for the diagnosis of anthrax in animals. Either the animal had anthrax, recognized from the recent history of the herd or site, and was treated accordingly, or it died. Most of the interest in developing serological testing has been for research on humoral responses in humans, and—to a lesser extent—animals, to vaccines and for epidemiological studies involving naturally acquired seroconversion in humans, livestock and wild mammals.

Currently accepted as the best serological procedure is the ELISA in microtitre plates coated with the Protective Antigen (PA) component of the anthrax toxin at 3 –5 ?g/ml in high pH (9.5) carbonate coating buffer. The toxin antigens appear to be truly specific for B. anthracis, although there is at present no commercial source of these. This tends to mean that anthrax serology is currently confined to a few specialist laboratories. Various versions of the ELISA exist and can be found in standard laboratory manuals; any version will do for anthrax serology, although certain sera appear to be more "sticky" than others. A useful tip appears to be to use reconstituted dried milk as the blocking agent and to raise its concentration until control negative sera are giving reliable negative results. For bovine sera, this may be a 10% suspension or higher.

Examples of the successful field application of anthrax serology are given elsewhere (Turnbull et al., 1992; Redmond et al., 1997).

4 Anthrax in humans

4.1 Incidence

The major sources of human anthrax infection are direct or indirect contact with infected animals, or occupational exposure to infected or contaminated animal products. Other possible sources are rare and epidemiologically trivial. Human anthrax incidence is dependent on the level of exposure to affected animals and national incidence data for non–industrial cases reflect the national livestock situation. Historical analysis of epidemiological data globally reveals the following approximate ratios: (a) one human cutaneous anthrax case to ten anthrax livestock carcasses; (b) one incident of enteric human anthrax to 30–60 anthrax–infected animals eaten; (c) in humans, 100–200 cutaneous cases for each enteric case that occurs.

Industrial anthrax incidence data can be inferred from the volume and weight of potentially affected materials handled or imported, taking into account the quality of prevention, such as vaccination of personnel and forced ventilation of the workplace. These relationships are essentially all that can be used for many countries where human anthrax is infrequently, erratically or incompletely reported. In addition, certain countries suppress anthrax reporting at the local or national levels.

Human case rates for anthrax are highest in Africa, the Middle East and central and southern Asia. Where the disease is infrequent or rare in livestock, it is rarely seen in humans.

4.2 Susceptibility. Data for risk assessments

4.2.1 Historical information

Circumstantial evidence indicates that man is moderately resistant to anthrax. Before vaccines and antibiotics became available, and at a time when understanding of industrial hygiene was relatively basic, workers in at–risk industrial occupations processing animal products were exposed to significant numbers of anthrax spores on a daily basis. In Britain, 354 cases of anthrax in such industries were notified during the 13–year period 1899–1912 (Anon, 1918). Although the numbers of persons exposed is not known, it must have been many thousands, and the number of cases clearly represented only a very small proportion of the number exposed.

In 4 mills in the USA, in which unvaccinated workforces, varying in size from 148 to 655, were "chronically exposed to anthrax", annual case rates were only 0.6 to 1.4% (Brachman et al., 1962). In one mill, workers were found to be inhaling 600 to 1300 anthrax spores over an 8–hour shift without ill effect (Dahlgren et al., 1960) and in two goat–hair mills, B. anthracis was recovered from the nose and pharynx of 14 of 101 healthy persons. Despite extensive exposure to anthrax, cases among workers in wildlife reserves are exceedingly rare (Quinn and Turnbull, 1998).

Nevertheless, outbreaks and epidemics do occur in humans; sometimes these are sizeable, such as the epidemic in Zimbabwe which began in 1979, was still smouldering in 1984–5 and had by that time affected many thousands of persons, albeit with a low case fatality rate (Turner, 1980; Davies 1982; Kobuch et al. 1990). Occasionally, the case fatality rates are substantial, such as in the Sverdlovsk incident in Russia in 1979 (Abramova et al., 1993; Meselson et al., 1994). The outbreak in a mill in New Hampshire, USA, in 1957 was not associated with any unusual change in occupational exposure but seems to have been an isolated event within a prolonged period of exposure (Brachman et al., 1960).

4.2.2 Infectious dose

Infectious doses, which have not been established for man, and the severity of the resulting infection clearly depend on several factors such as route of infection, nutritional and other states of health on the part of the infected person, and probably on the relative virulence of the infecting strain. For the purpose of risk assessments, dependency on information from animal tests is unavoidable. The published data on infectious and lethal doses in animals have been collated elsewhere (Watson and Keir, 1994).

Cutaneous infections. It probably does not take many spores to initiate a cutaneous infection, but it is generally accepted that the spores must gain access to subepidermal tissue through a cut or abrasion before this can occur and risk of infection reflects the chance of this happening. This risk is greatly reduced in at–risk occupations by appropriate clothing and gloves, dressing of wounds, and other hygienic practices.

Pulmonary (inhalation) infections. Recorded inhalation LD₅₀s in non–human primates range from 2500 to 760 000 spores (Meselson et al., 1994; Watson and Keir, 1994). The US Department of Defence bases its strategies on an estimate that the LD50 for humans is 8000 to 10 000 spores (Meselson et al., 1994). However the only hard data on inhalation infectious doses in humans come from the studies in goat hair processing mills referred to in 4.2.1. In any event, substantial exposure is evidently necessary before the risk of inhalation anthrax becomes significant. In a recent study (Turnbull et al., 1998) the highest levels found in air sampled 3 to 9 m downwind from disturbed dry, dusty anthrax carcass sites in Namibia were 20 to 40 colony–forming units of spores per cubic metre. This corresponds to the conservative estimate that it would require about 2.5 minutes for an average human undergoing moderate activity to inhale 1 spore. It is, furthermore, well established that, at sizes above 5 µm, particles face increasing difficulty in reaching the alveoli of the lungs. The likelihood of inhaled spores penetrating far enough to induce inhalation anthrax therefore depends greatly on the size of the particles to which they are attached.

The overall conclusion from the available evidence is that the risks of pulmonary anthrax outside industrial situations are very low.

Oral route infections. There is very little information on infectious doses by the oral route, but what is true for the skin is probably largely true for the oropharyngeal and gastrointestinal epithelium. The chance of infection is likely to be enhanced by, if not dependent on, the existence of a lesion in the epithelium through which spores can gain entry and establish an infection.

Treatability. The fact that anthrax is readily treated if diagnosed at a sufficiently early stage of infection also needs to be taken into account when assessing risks. Awareness of the likelihood of exposure having taken place is clearly an important part of the equation.

4.2.3 Biological warfare associations

In the developed parts of the world where it is now seen rarely, anthrax has developed something of a "doomsday bug" status in the mind of the public, and the name frequently engenders unnecessary fear, for example, in relation to contaminated burial or industrial (e.g. tannery) sites. This anxiety results from the association of anthrax with the topic of biological warfare.

There is, in fact, no conflict between the statements and evidence given in 4.2.1 and 4.2.2 that humans are fairly resistant to fatal anthrax infection and the possible aggressive use of anthrax spores. The "worst case" natural contamination in the environment is found at the carcass sites of animals that have died of anthrax. In a study in Namibia of 106 such sites, the highest contamination level found was just over 1 million anthrax spores per gram of soil, but 79% had less than 1000 per gram and 25% less than 10 per gram (Lindeque and Turnbull, 1994). Levels in other types of inadvertently contaminated environments (soils at tannery sites, horsehair plaster, etc.) rarely exceed a few units or tens of spores per gram (Turnbull, 1996). Natural environmental exposure to infectious doses in the normal course of human life and endeavour is, therefore, a fairly unlikely event.

Aggressive scenarios, in contrast, envisage exposures to overwhelmingly massive doses (many millions of spores) which can only be created artificially. The Figure of 100 kg of dried anthrax spores, given in one article (Taylor, 1996) on the subject as technically feasible for aggressive delivery, represents dose levels in the order of 10¹³ human LD50s. It must be supposed that this could cause substantial devastation to human and animal communities within selectively targeted areas. The public health implication of deliberately induced anthrax outbreaks and its use as a biological weapon have been reviewed elsewhere (WHO, 1970).

4.3 Epidemiology and transmission: the forms of anthrax

Anthrax in humans is classically divided in two ways. The first type of classification, which reflects how the occupation of the individual led to exposure, differentiates between non–industrial anthrax, occurring in farmers, butchers, knackers, veterinarians and so on, and industrial anthrax, occurring in those employed in the processing of bones, hides, wool and other animal products. The second type of classification, reflecting the route by which the disease was acquired, distinguishes between cutaneous anthrax acquired through a skin lesion, gastrointestinal tract anthrax contracted from ingestion of contaminated food, primarily meat from an animal that died of the disease, or conceivably from ingestion of contaminated water and pulmonary (inhalation) anthrax from breathing in airborne anthrax spores.

Non–industrial anthrax, resulting from handling infected carcasses, usually manifests itself as the cutaneous form; it tends to be seasonal and parallels the seasonal incidence in the animals from which it is contracted. Cutaneous anthrax transmitted by insect bites and intestinal anthrax from eating infected meat are also non–industrial forms of the disease. Industrial anthrax also usually takes the cutaneous form but has a far higher probability than non–industrial anthrax of taking the pulmonary form through inhalation of spore–laden dust.

Humans almost invariably contract anthrax directly or indirectly from infected animals. Records of person–to–person spread or laboratory–acquired anthrax are rare (Heyworth et al., 1975; Collins 1988; Lalitha et al., 1988; Quinn and Turnbull, 1998).

It is generally believed that B. anthracis is non–invasive and cutaneous and gastrointestinal tract anthrax infection require entry through a small cut, abrasion or other lesion (insect bite, ulcer, etc.). Thus anthrax eschars are generally seen on exposed regions of the body, mostly on the face, neck, hands and wrists.

As inferred earlier, in some countries mechanical transmission by biting insects is believed to be at least an occasional mechanism by which anthrax is contracted by humans (Rao and Mohiyudeen, 1958; Davies, 1983); that this can happen has been demonstrated experimentally (Sen and Minett, 1944; Turell and Knudson, 1987).

4.4 The clinical disease

Cutaneous anthrax is said to account for 95% or more of human cases globally. All three forms, cutaneous, gastro–intestinal tract and pulmonary, are potentially fatal if untreated, but the cutaneous form is often self–limiting. Data from pre–antibiotic and vaccine days indicate that 10–20% of untreated cutaneous cases might be expected to result in death (Anon, 1918). With treatment, less than 1% are fatal.

Overt gastrointestinal tract and pulmonary cases are more often fatal, largely because they go unrecognized until it is too late for effective treatment. However, serological and epidemiological evidence suggests that undiagnosed low–grade gastrointestinal tract or pulmonary anthrax with recovery can also occur, and may not be infrequent, among exposed groups (Brachman et al., 1960; Norman et al., 1960; Sirisanthana et al., 1988; Van den Bosch, 1996).

Development of meningitis is a dangerous possibility in all three forms of anthrax.

4.4.1 Cutaneous anthrax

The incubation period ranges from as little as 9 hours to 2 weeks, mostly 2 to 6 or 7 days.

The general scenario is as follows:

Day 0 Entry of the infecting B. anthracis (usually as spores) through a skin lesion
(cut, abrasion, insect bite, etc.).

Days 2–3 A small pimple or papule appears (see Figures M–R).

Days 3–4 A ring of vesicles develops around the papule. Vesicular fluid may be exuded. Unless the patient was treated, capsulated B. anthracis can be identified in polychrome methylene blue–stained (M'Fadyean stain) smears of this fluid and isolated on conventional agars, preferably blood agar (see Appendix I). Marked oedema starts to develop. Unless there is secondary infection, there is no pus and the lesion is not painful, although painful lymphadenitis may occur in the regional lymph nodes.

Days 5–7 The original papule ulcerates to form the characteristic eschar. Topical swabs will not pick up B. anthracis. Detection by polychrome methylene blue–stained smears or isolation requires lifting the edge of the eschar with tweezers (this gives no pain unless there is secondary infection) and obtaining fluid from underneath. The fluid will probably be sterile if the patient has been treated appropriately. Oedema extends some distance from the lesion. Clinical symptoms may be more severe if the lesion is located in the face, neck or chest. In these more severe forms, clinical findings are high fever, toxaemia, regional painful adenomegaly and extensive oedema; shock and death may ensue.

Day 10 (approximately). The eschar begins to resolve; resolution takes almost six weeks and is not hastened by treatment. A small proportion of cases, if untreated, develop systemic anthrax with hyperacute symptoms.

4.4.1.1 Differential diagnosis

Boil (early lesion), orf, vaccinia, glanders, syphilitic chancre, erysipelas, ulcer (especially tropical). These lack the characteristic oedema of anthrax. The absence of pus, the lack of pain, and the patient's occupation may provide further diagnostic pointers. The outbreak of Rift Valley Fever, referred to in 3.5.1 and initially thought to be anthrax, also affected numerous humans.

In differential diagnosis of the severe forms, orbital cellulitis, dacrocystitis and deep tissue infection of the neck should be considered in the case of severe anthrax lesions involving the face, neck and anterior chest wall. Necrotising soft tissue infections, particularly group A streptococcal infections and gas gangrene, and severe cellulitis due to staphylococci, should also be considered in the differential diagnosis of severe forms of cutaneous anthrax.

4.4.1.2 Immunological tests

Subject to certain provisos, serology can, on occasion, be useful in supportive or retrospective diagnosis of anthrax (see A.I.5). The practical aspects have been covered in 3.5.2.

Similarly, in the Russian sphere of influence, a skin test utilising Anthraxin^T (Antiplague Research Institute, Sovetskaya St., 13/15, Stavropol, 355106 Russian Federation; Fax: +7 8652 260312), first licensed in the former USSR in 1962, has become widely used for retrospective diagnosis of human and animal anthrax and for vaccine evaluation (Shylakhov et al., 1997). This is a commercially produced heat–stable protein–polysaccharide–nucleic acid complex, derived from oedematous fluid of animals injected with the vaccine STI or the Zenkowsky strains of B. anthracis. The test involves intradermal injection of 0.1 ml of Anthraxin and inspection after 24 h for erythema and induration at the site lasting for 48 hours after the injection. This delayed type hypersensitivity is seen as reflecting anthrax cell mediated immunity and was reportedly able to diagnose anthrax retrospectively some 31 years after primary infection in up to 72 % of cases (Shlyakhov et al., 1997). It was used with success in a retrospective investigation of a series of cases occurring in a spinning mill in Switzerland where synthetic fibres were combined with goat hair from Pakistan (Pfisterer, 1990). The diagnostic reliability of Anthraxin, like Ascoli test antigen (A.I.3.1), depends on the nature of anthrax rather than on the specificity of the antigens involved.

4.4.1.3 Precautions

Surgical tools should be sterilized without delay after use, and dressings should be incinerated. The wearing of surgical gloves by medical staff and orderlies is recommended but risks to these staff are NOT high. Direct human–to–human transmission is exceedingly rare (see 4.3). Vaccination of medical staff and orderlies is not necessary.

4.4.2 Gastrointestinal anthrax

There are two clinical forms of gastrointestinal anthrax which may present following ingestion of B. anthracis in contaminated food or drink.

1. Intestinal anthrax: Symptoms include nausea, vomiting, fever, abdominal pain, haematemesis, bloody diarrhoea and massive ascites. Unless treatment commences early enough, toxaemia and shock develop, followed by death. There is evidence that mild, undiagnosed cases with recovery occur.
2. Oropharyngeal anthrax: The main clinical features are sore throat, dysphagia, fever, regional lymphadenopathy in the neck and toxaemia. Even with treatment, the mortality is about 50% (Doganay et al., 1986).

The suspicion of anthrax depends largely on awareness and alertness on the part of the physician as to the patient's history and to the likelihood that he/she had consumed contaminated food or drink.

4.4.2.1 Confirmation of diagnosis. See 4.4.3.1

4.4.2.2 Differential diagnosis (gastrointestinal anthrax)

The differential diagnosis in intestinal anthrax includes food poisoning (in the early stages of intestinal anthrax), acute abdomen due to other reasons, and haemhorragic gastroenteritis due to other microorganisms, particularly necrotising enteritis due to Clostridium perfringens.

In the differential diagnosis of oropharyngeal anthrax, streptococcal pharyngitis, Vincent's angina, Ludwig's angina, parapharyngeal abscess, and deep tissue infection of the neck should be considered.

4.4.3 Pulmonary (inhalation) anthrax

Symptoms prior to the onset of the final hyperacute phase are non–specific and suspicion of anthrax depends on the knowledge of the patient's history. In probably the best–documented set of five case reports of inhalation anthrax (Plotkin et al., 1960), the illnesses began insidiously with mild fever, fatigue and malaise lasting one to several days. Headache, muscle aches, chills and fever were recorded in all four patients with development of a cough in four and mild pain in the chest in one. This mild initial phase was followed by the sudden development of dyspnoea, cyanosis, disorientation with coma and death in four of the patients, in whom treatment was unsuccessful. Death occurred within 24 hours of onset of the hyperacute phase.

4.4.3.1 Confirmation of diagnosis (pulmonary and intestinal anthrax)

As indicated, clinical diagnosis is dependent on a knowledge of the patient's history; early symptoms are non–specific and "flu–like" with mild upper respiratory tract signs in pulmonary anthrax or resembling mild food poisoning in intestinal anthrax. In fact, in pulmonary anthrax, the X–ray picture of the lung is very characteristic, with extremely enlarged mediastinal lymph nodes. Frequently, however, confirmatory diagnosis of pulmonary or gastrointestinal anthrax will usually take place after the patient has died or, if correct treatment is initiated early enough, when he or she is well recovered.

The definitive diagnosis is made by the isolation of B. anthracis from sputum in pulmonary anthrax and from vomitus, faeces and ascites in intestinal anthrax. Blood cultures may be positive in either form of the disease.

Depending on the treatment administered and the stage the disease has reached at the time of collection of specimens, smears stained for demonstration of the capsule (Appendix I) may be positive, or the specimens may be positive by culture. B. anthracis may be visualized in or isolated from sputum (pulmonary anthrax) or faeces (intestinal anthrax) but this cannot be relied upon. Specialized laboratories may be able to demonstrate anthrax toxin in fluid specimens (serum or oedematous fluid) or, in the case of patients who survive, anti–toxin and anti–capsular antibodies may be demonstrable in convalescent sera (Appendix I [A.I.5]). The Anthraxin hypersensitivity test referred to in 4.4.1.2 may also be applicable.

Death being due to the toxin, belated treatment can sterilize the blood and tissue fluids while still not preventing death. If this sterilising effect has not occurred, the capsulated B. anthracis may be visible in capsule–stained smears of these fluids and should be easily isolated from them by bacteriological culture.

Where anthrax has not been suspected prior to postmortem, characteristic signs are dark haemolysed unclotting blood, enlarged haemorrhagic spleen, petechial haemorrhages throughout the organs, and a dark oedematous intestinal tract, ulcerated or with areas of necrosis. In pneumonic anthrax, the mediastinal lymph nodes are always affected with haemorrhagic necrotizing lymphadenitis. Nevertheless, it may be hard to differentiate between pulmonary and intestinal anthrax at autopsy and the decision as to how the disease was contracted may have to be based on the patient's history.

4.4.4 Anthrax meningitis

Meningitis due to anthrax is a serious clinical development which may follow any of the other three forms of anthrax. The case fatality rate is almost 100%; the clinical signs of meningitis with intense inflammation of the meninges, markedly elevated CSF pressure and the appearance of blood in the CSF (the meningitis of anthrax is a haemorrhagic meningitis) are followed rapidly by loss of consciousness and death (Levy et al., 1981; Koshi et al., 1981; Lalitha et al., 1990; George et al., 1994; Lalitha et al., 1996). Only a few instances of survival as a result of early recognition of the problem and prompt treatment are on record (Khanne et al., 1989; Lalitha et al., 1996).

Differential diagnosis should take into account acute meningitis of other bacterial aetiology. The definitive diagnosis is obtained by visualisation of the capsulated bacilli in the CSF and/or by culture.

4.4.5 Anthrax sepsis

Sepsis develops after the lymphohematogenous spread of B. anthracis from a primary lesion (cutaneous, gastrointestinal or pulmonary). Clinical features are high fever, toxaemia and shock, with death following in a short time.

In the differential diagnosis, sepsis due to other bacteria should be considered. Definitive diagnosis is made by the isolation of B. anthracis from the primary lesion and from blood cultures.

5 Pathogenesis and pathology

5.1 Toxin as the cause of death

The events occurring between entry of infecting B. anthracis into a lesion or uptake from the lungs and death were covered in 3.4. At one time it was held that death from anthrax was due to capillary blockage, hypoxia and depletion of nutrients by the exceedingly large numbers of bacilli. Subsequently it was shown that death is attributable to a toxin (Keppie et al., 1955).

5.2 The virulence factors of B. anthracis

The capsule and the toxin complex are the two known virulence factors of B. anthracis.

The poly–D–glutamic acid capsule is presumed to act by protecting the bacterium from phagocytosis.

The toxin complex, which consists of three synergistically acting proteins, Protective Antigen (PA, 83kDa), Lethal Factor (LF, 87 kDa) and Oedema Factor (EF, 89 kDa), is produced during the log phase of growth of B. anthracis. LF in combination with PA (lethal toxin) and EF in combination with PA (oedema toxin) are now regarded as responsible for the characteristics signs and symptoms of anthrax.

According to the currently accepted model, PA binds to receptors on the host's cells and is activated by a host protease which cleaves off a 20 kDa piece, thereby exposing a secondary receptor site for which LF and EF compete to bind. The PA+LF or PA+EF are then internalized and the LF and EF are released into the host cell cytosol.

EF is an adenylate cyclase which, by catalysing the abnormal production of cyclic–AMP (cAMP), produces the altered water and ion movements that lead to the characteristic oedema of anthrax. High intracellular cAMP concentrations are cytostatic but not lethal to host cells. EF is known to impair neutrophil function and its role in anthrax infection may be to prevent activation of the inflammatory process.

LF appears to be a calcium– and zinc–dependent metalloenzyme endopeptidase (Hammond and Hanna, 1998). It has recently been shown (Duesbery et al., 1998) that it cleaves the amino terminus of two mitogen–activated protein kinase kinases and thereby disrupts a pathway in the eukaryotic cell concerned with regulating the activity of other molecules by attaching phosphate groups to them. This signalling pathway is known to be involved in cell growth and maturation; the manner in which its disruption leads to the known effects of LF has yet to be elucidated. On the basis of mouse and tissue culture models, macrophages are a major target of lethal toxin which is cytolytic in these. The initial response of sensitive macrophages to lethal toxin is the synthesis of high levels of tumour necrosis factor and interleukin–1 cytokines and it seems probable that death in anthrax results from a septic shock type mechanism resulting from the release of these cytokines.

The endothelial cell linings of the capillary network may also be susceptible to lethal toxin and the resulting histologically visible necrosis of lymphatic elements and blood vessel walls is presumably responsible for systemic release of the bacilli and for the characteristic terminal haemorrhage from the nose, mouth and anus of the victim (see Figures on the front cover).

The detailed nature and mode of action of the toxin has been more thoroughly reviewed in various texts (Leppla, 1992; Quinn and Turnbull, 1998). Most of the recorded histopathological studies on anthrax were done between 1945 and 1970; these are reviewed elsewhere (Quinn and Turnbull, 1998).

6 Bacteriology

B. anthracis, the causative agent of anthrax, is a Gram positive, aerobic or facultatively anaerobic, endospore–forming, rod–shaped bacterium approximately 4 µm by 1 µm, although under the microscope, it frequently appears in chains of cells. In blood smears, smears of tissues or lesion fluid from diagnostic specimens, these chains are two to a few cells in length (see Figure B); in suspensions made from agar plate cultures, they can appear as endless strings of cells – responsible for the characteristic tackiness of the colonies (see Figure D). Also characteristic is the square–ended ("box–car" shaped) appearance traditionally associated with B. anthracis vegetative cells, although this is not always very clear. In the presence of oxygen, and towards the end of the exponential phase of growth, one ellipsoidal spore is formed in each cell; this does not swell the sporangium and is generally situated centrally, sometimes sub–terminally (see Figure A).

Under anaerobic conditions and in the presence of bicarbonate (HCO₃), the vegetative cell secretes a polypeptide (poly–( – D–glutamic acid) capsule. As covered in section 5, the capsule is formed in vivo and is one of the two virulence factors of B. anthracis. It is also a primary diagnostic aid (see 3.5 and 4.4.1, Appendix I [A.I.2] and Figure B).

6.1 Detection and isolation

In appropriate blood or tissue specimens collected within a few hours of death from animals (see 3.5) or humans with anthrax, B. anthracis is readily detected in capsule–stained (M'Fadyean–stained) smears and readily isolated in pure culture on blood or nutrient agar plates. The same applies to smears of fluid from cutaneous lesions of humans prior to treatment (see 4.4.1).

In old or decomposed animal specimens, or processed products from animals that have died of anthrax, or in environmental samples, detection is likely to involve a search for relatively few B. anthracis within a background flora of other bacteria, many of which will probably be other Bacillus species, in particular, the closely–related B. cereus. In this case, selective techniques are necessary. A procedure for the isolation of B. anthracis from such specimens is given in Appendix I (A.I.1).

Such is the nature of the properties of B. anthracis that few agents which differentially select between B. anthracis and other Bacillus species do so in favour of B. anthracis and those that do only do so unconvincingly. Of the selective media that have been proposed, the most successful is polymyxin–lysozyme–EDTA–thallous acetate (PLET) agar (Knisely, 1966), although care has to be taken to prepare this correctly. As yet no selective enrichment broth system has been devised for B. anthracis and, pending development of such a system, the sensitivity of in vitro detection of B. anthracis by conventional means in environmental samples or specimens from old or decomposed animals or from processed animal products is limited to approximately 5 cfu/g or /ml (Manchee et al., 1981).

As covered in Appendix I (A.I.1.4 and A.I.4), the most sensitive method for isolating the organism is inoculation into a guinea pig or mouse. Although, in line with increasing aversion to the use of animals for scientific tests, this should strictly be a last resort, there are times when this may still be the necessary approach, for example, when confirming anthrax in individuals or animals that have been treated with antibiotics, or in essential tests on environmental samples that contain sporostatic chemicals.

Polymerase chain reaction (PCR) detection systems have been developed for B. anthracis (Beyer et al. 1996; Patra et al. 1996; Sjöstedt et al. 1996; 1997), but it will probably be a few years before they become fully reliable, adequately sensitive and robust, and generally available for use in the non–specialist laboratory.

6.2 Identification and confirmation

With occasional exceptions, it is generally easy to identify B. anthracis and to distinguish it from other Bacillus species, including B. cereus. For all practical purposes, an isolate with the characteristic colonial morphology (Parry et al., 1983) on nutrient or blood agar (matt appearance, fairly flat, similar to B. cereus but generally rather smaller, more tacky, white or grey–white on blood agar, and often having curly tailing at the edges), and which is non–haemolytic or only weakly haemolytic, non–motile, sensitive to the gamma–phage and penicillin, and able to produce the capsule in blood or on anaerobic culture on bicarbonate media is B. anthracis (see Figures D and E, and Appendix I [A.I.2]) .

The PCR is becoming more widely available as a means of confirming the presence of the virulence factor (capsule and toxin) genes, and hence that an isolate is, or is not, virulent B. anthracis. For routine purposes, primers to one of the toxin genes (usually the Protective Antigen gene) and to one of the enzymes mediating capsule formation are adequate (Appendix I [A.I.6] and Figure F). In laboratories not equipped for PCR tests, if doubt remains at the end of the procedures given in Appendix I (A.I.1 and 2) as to the definitive identity of a suspect B. anthracis isolate, inoculation into a mouse or guinea pig may necessary (Appendix I [A.I.4]). However, as stated in 6.1, this should be a last resort procedure and confined to situations where a definitive identity is essential.

Movement of infectious or contaminated materials from the site of origin to a diagnostic or reference laboratory obviously presents a risk of spread of diseases if the materials inadvertently escape into the environment during transit. Attention is drawn to the recommendation of the United Nations Committee of Experts on Transport of Dangerous Goods for packaging, labelling and documentation in relation to transport of infectious specimens, detailed in other publications (WHO, 1997b), excerpts of which one is reported in Appendix VIII.

7 Treatment

Prompt and timely antibiotic therapy usually results in dramatic recovery of the individual or animal infected with anthrax. Almost all isolates of B. anthracis can be expected to be highly sensitive to penicillin and, being cheap and readily available in most parts of the world, this remains the basis of treatment schedules, particularly in animals and in humans in developing countries. The organism is also sensitive to numerous other broad spectrum antibiotics; should the use of penicillin be contraindicated, a wide range of alternative choices exist from among the aminoglycosides, macrolides, quinolones and tetracyclines. Chloramphenicol is also a satisfactory alternative.

We are only aware of four reports of the isolation of penicillin–resistant strains (Anon, 1996). The molecular basis of susceptibility and resistance is complex (Lightfoot et al., 1990). It looks as if it may prove to be the case that all B. anthracis strains carry the ? –lactamase gene(s) on their cromosomes but, apart from the exceptions mentioned, these are not expressed.

In pulmonary or gastrointestinal anthrax in humans, symptomatic treatment in an intensive care unit in addition to antibiotic therapy may save the patient's life; as referred to in 7.2.8, if available, plasmaphoresed hyperimmune serum or gamma–globulin from vaccinated persons may be considered in life–threatening situations.

7.1 Treatment of animals

7.1.1 General principles and approaches

Following the first incident of anthrax in a herd, the remaining animals should be moved immediately from the field or area where the index case died and regularly checked at least three times a day for two weeks for signs of illness (rapid breathing, elevated body temperature) or of submandibular or other oedema. Any animal showing these signs should be separated from the herd and given immediate treatment. Clinical experience has frequently demonstrated that animals, especially cattle, will respond favourably to treatment even though apparently in the terminal stages of anthrax. Even if they go on to die (death in anthrax results from the effect of the toxin), the infecting load of B. anthracis will have been greatly reduced, if not entirely eliminated, thereby significantly reducing the chance of subsequent transmission from the carcass to other animals.

In endemic areas, or if there is concern that the outbreak may spread, the herd should be vaccinated (see Appendix V). Further anthrax deaths usually cease within 8 to 14 days of vaccination. Herd quarantine can be lifted 21 days after the last death. Decontamination of the site(s) where the index case or other case(s) died should be carried out (see Section 8 and Appendix III). [It should be remembered that vaccination and treatment should not be done simultaneously; the treatment will prevent the live vaccine taking effect].

In certain countries, treatment is not permitted and slaughter policies are in place (7.1.2.4).

7.1.2 Specific procedures

7.1.2.1 Antimicrobial therapy in animals

The recommended procedure for treating animals showing clinical illness in which anthrax is thought to be the likely or possible cause is immediate intravenous administration of sodium benzylpenicillin as directed by the manufacturer's instructions (usually in the range 12 000–22 000 units per kg of body weight) followed 6–8 hours later by intramuscular injection of long acting benethamine penicillin (manufacturers' instructions usually recommend dose within range 6000–12 000 units per kg of body weight) or other appropriate long–acting preparation such as Clamoxyl^R (15 mg/kg), a long–acting preparation of amoxycillin. If long–acting preparations are unavailable, procaine penicillin, (dose recommended by manufacturers is usually 6000–12 000 units/kg) can be used for intramuscular injection but should be administered again after 24 and 48 hours.

Streptomycin acts synergically with penicillin and penicillin/streptomycin mixtures are available commercially. Recommended doses of streptomycin to be administered together with penicillin intramuscularly are 5–10 mg per kg body weight in large animals and 25–100 mg per kg body weight in small animals.

Veterinary experience in Britain is that, in contrast to advice frequently found in textbooks, treatment with tetracyclines may not be fully effective.

Attention should be paid to manufacturers' recommendations regarding precautions and limitations of use and aspects relating to withdrawal periods after use in food animals.

Cost and availability are likely to be a major consideration in choice of treatment. For example, combined penicillin and streptomycin treatment can be expected to cost twice as much as penicillin alone.

7.1.2.2 Supportive therapy for animals

Symptomatic treatment may also be useful and a range of possible agents is available. Supportive therapy with an agent such as flunixin (an analgesic with anti–inflammatory, anti–pyretic and anti–endotoxic properties) may be advantageous although it will add significantly to the cost of the therapy.

7.1.2.3 Hyperimmune serum therapy for animals

Hyperimmune serum has been used in the past for treating anthrax cases (Sterne, 1959) and it was generally considered that homologous sera (e.g. serum prepared in cattle for bovine use, etc.) were more effective than heterologous sera. Serum treatment of livestock is apparently still practised in Russia.

The protective effect of immune serum administered therapeutically was demonstrated in monkeys (Henderson et al., 1956). However, these were challenged by the inhalation route and, once the effects of the immune serum wore off at about 20 days, the animals began to die of anthrax from continued uptake into the lymphatics of spores that still remained in the lungs. The possibility of relapse after the effect of this therapy has worn off should be borne in mind.

7.1.2.4 Countries prohibiting treatment in livestock

It should be noted that treatment of animals is forbidden in several countries. Veterinary requirements in these countries are that, in a herd that has experienced a case of anthrax, other animals showing signs of illness must be killed without spilling of blood or exsanguination and the unopened carcass must be heat–treated in a rendering plant. Certain countries require the slaughter of the entire herd following a case of anthrax; this draconian approach is unnecessary and wasteful.

7.1.2.5 Therapy in wildlife

While the use of antibiotics for controlling anthrax in wildlife is, generally speaking, unlikely to be feasible, it has reportedly been done with success in an outbreak in Tanzania where a single treatment of roan and sable antelope and kudu, 50 animals in total, by direct darting, appeared to arrest an outbreak (Dr SFH Jiwa, Faculty of Veterinary Medicine, Sokoine University of Agriculture, P.O. Box 3110, Morogoro, Tanzania, personal communication).

7.2 Treatment of humans

7.2.1 Mild uncomplicated cases

In mild uncomplicated cases of cutaneous anthrax, penicillin V, 500 mg, taken orally every 6 hours for 5–7 days is adequate, but the treatment usually recommended is 3 to 7 days of intramuscular procaine penicillin, 600 mg (1 million units), every 12–24 hours or intramuscular benzylpenicillin (penicillin G), 250 000 units at 6–hourly intervals. Cutaneous lesions usually become sterile within the first 24 hours of such regimens but, although early treatment will limit the size of the lesion, it will not alter the evolutionary stages it must go through (Kobuch et al., 1990).

7.2.2 Severe or potentially life–threatening cases

In severely affected patients or when pulmonary or gastrointestinal anthrax is suspected, the initial treatment is penicillin G, 1200 mg (2 million units) per day by infusion or by slow intravenous injection (<300 mg/min) every 4–6 hours until the patient's temperature returns to normal; at this point treatment should continue in the form of intramuscular procaine penicillin administered as described above. Streptomycin, 1–2 g per day intramuscularly, may act synergically with penicillin.

General measures for treatment of shock may be life saving since death is due, at least in part, to toxin–induced shock. Intubation, tracheotomy or ventilatory support may be required in the event of respiratory problems, and vasomotor support with dopamine may be necessary when there is haemodynamic instability. Primary haematological, renal or liver function disorders are not generally seen.

7.2.3 Supportive treatment for cutaneous anthrax

As covered in 5.2, the swelling seen in an anthrax infection is due to the action of oedema toxin and there is very little associated inflammation. In theory, therefore, steroids should be of little value. In practice, some reports indicate that these have been administered with evidence of benefit but others (Kobuch et al., 1990) have concluded that they were ineffective, discontinuing their use.

Early tracheotomy is advised if there is danger of tracheal obstruction; once oedema is extensive it can be difficult to find the trachea at operation.

7.2.4 Alternatives to penicillin

In the event of allergy to penicillin, several antibiotics are effective alternatives, including tetracyclines, chloramphenicol, gentamicin and erythromycin. Of the tetracycline family, tests in animals have indicated doxyxycline is very effective and that the quinolone, ciprofloxacin may also be suitable (Friedlander et al., 1991). Trimethoprim is not effective.

7.2.5 Adequate doses

In vitro tests (Lightfoot et al., 1990) have indicated that some strains of B. anthracis grown in the presence of sub–inhibitory concentrations of flucloxacillin become resistant to penicillin and amoxycillin either by induction of beta–lactamase or through some other, unidentified mechanism. It is therefore important to use adequate doses of penicillin when this is being used for treatment.

7.2.6 Duration of treatment

The appropriate duration of treatment is a subject for debate. B. anthracis cannot be isolated from cutaneous lesions 24 to 48 hours after commencement of antibiotic therapy and Kobuch et al (1990) could see no advantage to continuing treatment beyond 4 days. A report from Ethiopia (Martin, 1975) records that 100 patients with cutaneous anthrax were treated with a single intramuscular dose of procaine penicillin, 600 000 units, and 99 of these were sent home with the invitation to return if complications occurred. Only 5 returned on account of further developments. Physicians tend to continue treatment for 7 to 14 days, but it needs to be kept in mind that the lesion will continue to progress through its cycle of development and resolution regardless of the elimination of the infecting B. anthracis (7.2.1). Excessive treatment may be wasteful and counterproductive.

In cases of inhalation anthrax, anthrax spores have been shown to persist for many weeks in the lungs of monkeys infected experimentally by the aerosol route and kept on antibiotics (Henderson et al., 1956; Friedlander et al., 1991); the animals succumbed to the disease once the antibiotic treatment was stopped. Although this is unlikely to occur in the natural situation, in cases where known or suspected inhalation of anthrax spores has taken place, especially if this was likely to have been substantial, it may be prudent to consider the administration of a non–living vaccine simultaneously with treatment. The treatment should be continued for about 6 weeks to allow for development of adequate vaccine–induced immunity. (The issue of simultaneous treatment and vaccination in countries that administer live vaccines to humans is discussed elsewhere (Pomerantsev et al., 1996; Stepanov et al., 1996).

7.2.7 Treatment of anthrax meningoencephalitis

Lessons from the few recorded instances of survival in cases of anthrax meningoencephalitis suggest that penicillin (2 million units of crystalline penicillin intravenously every 2 hours initially) remains the antibiotic of choice because it diffuses into the CSF through highly inflamed meninges. Chloramphenicol (1 g intravenously every 4 hours) is a suitable alternative for hypersensitive patients. The effectiveness of newer antibiotics, known to achieve a satisfactory degree of penetration into the CSF, has not been determined.

Essential supportive therapy includes the early institution of anticerebral oedema measures, such as 100 ml of 20% mannitol intravenously every 8 hours and hydrocortisone, 100 mg every 6 hours. Useful references to anthrax meningoencephalitis are given in 4.4.4.

7.2.8 Hyperimmune serum therapy

The use of anti–anthrax serum was abandoned years ago in most western countries as of negligible value, but it is apparently still done in China (Dong, 1990) and the Russian Federation (Anon, 1996). With the availability of modern plasmaphoresis techniques, specific human gamma–globulin from vaccinated individuals could well be of life–saving value in an emergency. The disease is readily responsive to early antibiotic treatment but, as has been pointed out before, death is due to the toxin and delayed treatment may sterilize the blood and tissues while not preventing death. Thus, if initiated after 24–36 hours following clinical onset, antibiotic treatment may fail to save life in pulmonary, intestinal or complicated cutaneous cases. In these cases, specific gamma–globulin containing anti–toxin antibodies may be life–saving.

7.2.9 Recurrence after treatment

Recurrence of disease on termination of treatment is very rare but convalescent cases should remain under observation for at least a week after treatment has ceased.

8 Control

Control measures are aimed at breaking the cycle of infection depicted in Figure 3. Each of the following actions must be rigorously implemented.

• correct disposal of anthrax carcasses (point X)
• correct disinfection, decontamination and disposal of contaminated materials (point Y)
• vaccination of exposed susceptible animals (point Z1) and humans in at–risk occupations (point Z2).

8.1 Disposal of anthrax carcasses (point X in Figure 3)

Because sporulation of B. anthracis requires oxygen and therefore does not occur inside a closed carcass, regulations in most countries forbid postmortem examination of animals when anthrax is suspected. Most, if not all, the vegetative B. anthracis cells in the carcass are killed in a few days by putrefactive processes. Nevertheless, with the characteristic terminal serosanguinous exudates from the nose, mouth and anus, some organisms may escape and sporulate. The precise length of time after which no viable B. anthracis remain within the carcass is unpredictable but depends greatly on climatic conditions, particularly temperature.

Figure 3. (see Figure 1 [section 2.1] for further notes).

8.1.1 Alternatives

In most countries, the preferred method of disposal of an anthrax carcass is incineration, although, in some countries, it is considered that the best approach to the disposal of anthrax carcasses is an effective controlled heat–treatment or "rendering". Where neither of these approaches is possible, for example due to lack of fuel, burial is the remaining best alternative. In some developing country situations, however burial, incineration or rendering may not be feasible. The last resort in such situations is to leave the carcass unmoved and adequately closed off from other animals, particularly scavengers, or people. The carcass should be fenced off and covered using branches of trees, corrugated iron or any other available materials, and hazard signs should be posted around the site. ???anthracis?within the animal carcass does not sporulate and is inactivated by the putrefactive process in a few days.

However, environmental contamination due to bloody exudate escaping from the mouth, nose and anus of the death animals may still occur. The resulting environmental contamination could be minimised by scorching the site with fire, after the carcass has effectively putrefies, though this may be many months later. Clearly, this is far less satisfactory than incineration, rendering or burial which should be carried out as the preferred option unless absolutely impossible.

8.1.2 Burial

Periodic reports of viable anthrax spores at burial sites of animals which died many years previously have testified to the unreliability of burial procedures for long term control of the disease. Disturbance of such sites, for example by ploughing, or laying drainage, brings the spores to the surface; even without site disturbance, spores can work their way up to the soil surface. In either case, this may result in new livestock cases.

Further disadvantages to burial sites are that scavengers may dig down to reach the carcass, and, in dry, dusty areas, the digging process can spread the contaminated soil extensively. In Wood Buffalo National Park, Canada, where burials of bison that had died of anthrax were carried out in the 1960s, the raised burial mounds became excavation sites for foxes and wolves building their dens and nesting sites for ants (D.C. Dragon, Department of Medical Microbiology and Immunology, University of Alberta, Edmonton, Canada, personal communication).

In summary, burial should be discouraged in favour of incineration or rendering where possible.

8.1.3 Incineration

Guidelines on incineration procedures are given in Appendix III (A.III.3). Ideally, the soil surrounding and under the carcass, particularly around the nasal and anal regions, should be decontaminated (see 8.3.8) and then incinerated with the carcass.

Incineration must be done with appropriate care to ensure complete burning from beneath. Usually this involves raising the carcass off the ground before the process is started (see Appendix III [A.III.3]). Commercial incinerators designed to ensure this are available (see

Figures G–J). It must be appreciated that spores that have soaked into the soil may survive the incineration process although isolation of B. anthracis from incineration sites is rare. The down–directed blow–torch shown in Figures K and L (see also Appendix III [A.III.3.3]), illustrates an alternative incineration procedure which ensures severe scorching of the soil to several centimetres of depth.

Comments are occasionally encountered opposing incineration on the basis that anthrax spores may survive the fire and become aerosolized in the updraft. Circumstantial evidence does not support the contention that incineration of anthrax carcasses results in the transmission of anthrax in this manner.

Blenkharn and Oakland (1989) were able to isolate Gram positive bacteria, predominantly Bacillus species, from the base of the exhaust stack of a hospital waste incinerator with design–specified operating temperatures of 800 °C and 1000 °C in the primary and secondary chambers respectively, thereby demonstrating that there is no room for complacency. However, numbers were very low, averaging 56 cfu per cubic metre (range 0–400 cfu per cubic metre) and would be subject to rapid further dilution on leaving the chimney. A badly constructed pyre producing smoke with little or no flame might result in a higher survival rate of organisms collected by the updraft. However, as discussed in relation to windborne spores from anthrax carcass sites (Turnbull et al., 1998), the dilution effect on anthrax spores dispersed in airborne form from a carcass is such that, for even the most susceptible animal species, the risks of infectious doses arising are absolutely minimal.

A further consideration is that the anthrax organisms in an unopened carcass are in the vegetative form (see 8.1) and are highly susceptible to heat and other adverse conditions. The spores will be confined to where the blood has been shed through the mouth, nose or anus, and will mostly be in the soil beneath these points. Consequently, relatively few spore forms will enter the fire and updraft; vegetative forms are very unlikely to survive.

8.1.4 Rendering

Rendering is essentially a cooking process which results in sterilisation of raw materials of animal origin such that parts of carcasses may be utilized safely for subsequent commercial purposes.

There are a number of variations of the rendering process, broadly divided into batch processes and continuous processes. In general, the raw materials are finely chopped and then passed into a steam–heated chamber and subjected to temperatures ranging from 100 o C to 150 o C for 10–60 minutes (this does not include the time taken to bring the material to the peak temperature or the subsequent cooling period time).

The rendering procedure involves correct performance at each of three stages: collection, transport and treatment of the carcass (Riedinger et al., 1975; Riedinger, 1980; Strauch, 1991). These should be supervised by veterinary authorities. The rendering plant must be properly divided into "dirty" and "clean" areas. The dirty side must be suitably equipped for disinfection of the transport vehicles and other equipment involved. Wastewater from the dirty side must be collected and autoclaved. Before the heat treatment, carcasses have to be broken down in a closed system into pieces not larger than 10 cm 3 . Controlled heat treatment is then carried out with temperature, pressure and time of sterilisation recorded.

The level of hygiene being maintained in the clean side of the rendering plant should be monitored at least twice yearly by the veterinary authorities.

8.2Infection control in the management of human anthrax cases

As said in 4.3, the risk of human–to–human transmission is not a serious one given that sensible precautions are taken. For example, cutaneous anthrax lesions should be dressed for the first 24–48 hours after treatment; disposable gloves should be worn, or gloves that can be sterilized, while applying the dressing and during subsequent disposal of specimens or sterilisation of materials and equipment (see 8.3.9).

Prophylactic antibiotics and vaccination are not necessary for health workers or family contacts, though the latter should be instructed to consult their physicians in the event of suspicious signs or lesions.

In fatal cases, postmortem should be discouraged; cremation is preferable to burial where local custom permits. It is advisable for the body to be placed in an impervious body bag for transport from the place of death and preferably the body should not be extracted from the bag. Where only burial is permitted, the bagged body should be placed in a hermetically sealed coffin and buried without re–opening. Useful guidelines are available elsewhere (Healing et al., 1995; Young and Healing, 1995).

Bedding and contaminated materials should be bagged and incinerated, autoclaved or fumigated as appropriate. Whether room fumigation is necessary will depend on the perceived level of contamination in the room where the patient died (see 8.2.3).

8.3 Disinfection, decontamination and disposal of infected/contaminatedmaterial(siteYinFigure3)

In addition to helping break the local cycle of anthrax infection, disinfection, decontamination and correct disposal of infected/contaminated material are of considerable importance in preventing international transmission of anthrax. In non–endemic countries, risks arise largely from animal products — wool, hair, hides, bone, etc. — imported from endemic regions. Regulations regarding importation of untreated animal products vary from country to country but several importing countries take the view that the financial costs that would be incurred from legislating for the sterilisation of such imports would be disproportionate to the benefits.

In many countries, some or all of the following requirements are in place to limit the risks of importation or dissemination of products contaminated with anthrax spores:

· products must be accompanied by a certificate signed by a veterinary official in the country of export certifying that they derive from anthrax–free sources (see Appendix IV)

· products regarded as having a high chance of containing anthrax spores are subject to some form of monitoring or control

· finished or raw materials from certain countries may be subject to import restrictions

· finished or raw materials from certain countries may have to undergo a general

sporicidal treatment before processing or distribution.

In the case of hides and skins (where no sterilisation procedure that does not damage the materials has been devised), the exporting countries nowadays often require the initial processing stages to be carried out before export for financial benefit. This has meant that far fewer anthrax–contaminated hides and skins now reach non–endemic countries than was the case a few years ago.

Sections 8.3.1–12 aim to supply information by which each country can formulate its own control programme according to its particular circumstances. However, it is clear that long–term global control depends almost entirely on the application of appropriate measures to prevent the disease among livestock in enzootic exporting countries. Ideally, national policies should ensure that materials known to be contaminated with anthrax spores are appropriately disposed of and are not included in any industrial process. The procedures in sections 8.3.10–12 are essentially designed to protect workers and the environment in situations where inadvertently contaminated materials may enter the processing line.

8.3.1 Decontamination of manure, dung, bedding, unused feed, etc.

Where possible, anthrax–contaminated materials to be disposed of, such as bedding, feedstuffs, manure, etc, should be incinerated or autoclaved (121+1 o C core temperature for 30 min). Immersion in 4% formaldehyde (10% formalin) for >12 hours is an alternative but full penetration of the fluids must be ensured. (Caution: avoid skin contact with formaldehyde solutions or inhalation of formaldehyde vapour).

8.3.2 Disinfection of surfaces in rooms, animal houses, vehicles, etc?

For general principles on disinfection and sporicidal disinfecting agents, see Appendix III. Disinfection of contaminated surfaces involves a three–step approach aimed at (i) preliminary disinfection, (ii) cleaning, and (iii) final disinfection. Effective disinfection of spores can be extremely difficult and, under some circumstances, it may not be possible to achieve this completely. It is important, therefore, to act promptly following cases of anthrax in order to prevent, as far as possible, the release and sporulation of vegetative cells from the dying or dead animal.

Stage 1: Preliminary disinfection

 

One of the following disinfectants may be used in amounts of 1–1.5 litres per square metre for an exposure time of 2 hours (Caution: avoid skin contact with the disinfectants listed below or inhalation of their vapours):

· 10% formaldehyde (approximately 30% formalin), or

· 4% glutaraldehyde (pH 8.0–8.5)

A high pressure cleaner may be used but, to avoid spreading the contamination, the pressure should not exceed 10 bar.

Stage 2: Cleaning

Where practical, cleaning of all surfaces should be done by straightforward washing and scrubbing using ample hot water. The operator should wear protective clothing, face and hands included. Cleaning should be continued till the original colours and surfaces are restored and the waste water is free of dirt particles. At the end of the process, residual water should be removed and the surfaces dried. High pressure cleaners are again discouraged because of the greater potential to spread the contamination through aerosols. If used, however, the water jet should be applied at a pressure of 80–100 bar delivering 13–15 litres/minute.

Stage 3: Final disinfection

For final disinfection, one of the following disinfectants should be applied at a rate of 0.4 litres per square metre for an exposure time of at least 2 hours:

· 10% formaldehyde (approximately 30% formalin)

· 4% glutaraldehyde (pH 8.0–8.5)

· 3% hydrogen peroxide

· 1% peracetic acid.

Hydrogen peroxide and peracetic acid are not appropriate if blood is present. When using glutaraldehyde, hydrogen peroxide or peracetic acid, the surface should be treated twice with an interval of at least one hour between applications. Formaldehyde and glutaraldehyde should not be used at temperatures below 10 o C. After the final disinfection, closed spaces such as rooms or animal houses should be well ventilated before re–commissioning.

The effectiveness of the disinfection procedure cannot be assumed and attempts should be made to confirm it has been adequate by means of swabs and culture.

In the case of surfaces within a room, it may be considered appropriate to finish the disinfection process by fumigating the room itself as described in 8.3.3.

8.3.3 Fumigation of closed spaces – cabinets, rooms, etc.

Safety cabinets should be treated as fumigation chambers and fumigated according to the procedure given in paragraph 2 of 8.3.9.

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 m 3 ) 4 litres of water containing 400 ml of concentrated formalin (37% w/v formaldehyde) 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. Room temperature should be >15 o C.

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. 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 (preferably 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) and dried should be placed at some point in the room distant from the kettle. (Caution: avoid skin contact with formaldehyde solution or inhalation of formaldehyde vapour).

At the end of the fumigation, the spore disk(s) should be retrieved into a sterile petri dish and the windows or vents to the outside air should be opened up. A chemical respirator should be used for this. A fan, or fans, assist 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 disk(s) on plates of nutrient agar containing 0.1% histidine (final concentration and added as a filter–sterilized solution after the agar has been autoclaved and cooled to 50 o C). After overnight incubation at 37 o C, if fumigation was properly effective, the disks should show no bacterial growth.

For health and environmental protection reasons, there are moves to replace formalin with hydrogen peroxide vapour as the recommended general purpose microbiological fumigant. Indeed, the use of formalin is being increasingly forbidden in some countries. However, the equipment needed for hydrogen peroxide fumigation is, at present, cumbersome, elaborate and expensive and is not universally available. It would not lend itself to routine fumigation of safety cabinets after every use. This section will be updated when the appropriate progress has been made.

8.3.4 In the laboratory - spills, splashes, accidents

Chlorine solutions. Commercially–prepared hypochlorite 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. 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.

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 Appendix III (A.III.2.1.4)). 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 (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 and Turnbull, 1996). The sporicidal rate can be increased by using 50% methanol or ethanol to make the dilutions of the stock solution.

Rapid turnover items such as pipettes, disposable loops, microscope slides, sampling spoons, etc., should be immersed overnight in hypochlorite solutions with 10 000 ppm available chlorine and then transferred to an autoclave bin or bag for autoclaving, or to a bag for incineration.

Benches should be wiped down after use with hypochlorite solutions containing 10 000 ppm available chlorine. Because of their neutralising 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 (Whatman International Ltd, Maidstone, UK).

Spills and splashes on surfaces. Some thought must 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 on floor, bench or apparatus should be flooded with hypochlorite solution containing 10 000 ppm available chlorine and vertical surfaces should be washed or wiped down thoroughly with cloths soaked in this solution (the operator should wear gloves and safety spectacles while doing this). Spills and splashes from fresh cultures can be mopped up with towelling after 5 minutes; the towelling should be placed in an autoclave bin or bag and autoclaved or in a bag for incineration. Spills or splashes of spore suspensions should be left for 30–60 minutes before mopping up unless the area can be sealed off and fumigated, in which case mopping up can be done after a few minutes and fumigation carried out immediately as in 8.3.3.

An alternative approach is to cover the contaminated area with absorbent material and wet this with an excess of disinfectant. Solutions of 10% formalin, 4% glutaraldehyde or 1% peracetic acid may be more appropriate than hypochlorite, but the choice must be weighed against the greater personal protection needed when using these. HAZ–TAB disinfectant granules (Guest Medical, Edenbridge, Kent TN8 6EW, UK) are commercially prepared for this purpose.

Spills and splashes on clothing. 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 (disposable gowns can be incinerated). 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 fumigated as in 8.3.3.

Spills and splashes on skin, in eyes. For skin contamination the area should be bathed in hypochlorite solution containing 5000 ppm available chlorine for one minute and then washed thoroughly with soap and water. Where the skin is broken (including needle–stick punctures), bleeding should be encouraged and the injury washed with copious amounts of water. The current Occupational Health wisdom is that immersing punctured skin in hypochlorite solutions does more harm than good. The appropriate medical officer should be informed and the affected person kept under observation for at least a week.

For splashes in the eye, the eye must be flushed out with copious quantities of water immediately. Avoid rubbing the eye. The appropriate medical officer should be informed and the affected person kept under observation for at least a week.

Contamination in the mouth. At the outset laboratory workers should be reminded that mouth pipetting in a microbiology laboratory is totally unacceptable. For contamination of the mouth with known or possible anthrax organisms, the mouth contents should be immediately spat out followed by a thorough mouth wash with hypochlorite solution containing 2000 ppm available chlorine and several subsequent mouth washes with water. The appropriate physician should again be informed and the affected person be kept under observation for a week.

8.3.5 Decontamination of liquid manure

Slurry from pigs and cattle suffering outbreaks of anthrax should be disinfected with formaldehyde added with thorough stirring to a final concentration of 2–4% depending on the content of dry matter. This is approximately equivalent to 50–100 kg of formalin (37% formaldehyde solution) per m 3 of slurry. The mixture should be left a minimum of 4 days with stirring for at least one hour each day before being removed elsewhere. ?Caution: avoid skin contact with formaldehyde solution or inhalation of formaldehyde vapour?.

The formalin degrades naturally and the treated slurry can be spread on uncultivated land and ploughed in or otherwise buried.

8.3.6 Decontamination of sewage sludge

Sewage sludge containing effluents from tanneries may contain anthrax spores. Dewatered sewage sludge up to a dry matter content of 8% should be disinfected with 5% formaldehyde for 10 hours or 3% peracetic acid for 30 minutes. The disinfection process is not affected by the use of lime or polyelectrolytes for dewatering the sludge (Lindner ?????., 1987). ? Caution: avoid skin contact with formaldehyde or peracetic acid solutions or inhalation of their vapours?.

8.3.7 Treatment of 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 section 8.3.

Autoclaving is the surest way of killing spores but is only applicable to fairly small volumes of water.

Treatment with formaldehyde (5–10% final concentration) 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 neutralisation and discharge without danger to the environment must be established. 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 must be considered on an individual basis and the best solution worked out for the particular circumstances that exist.

8.3.8 Treatment of 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 o C throughout for 20 min). If this is not possible, it should be disinfected with 5% formaldehyde solution at 50 litre per m 2 . Where it is necessary to decontaminate soil to greater depths, such as burial sites of anthrax carcasses, 5% formaldehyde solution 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).

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 can 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).

8.3.9 Other materials – clothing, tools, etc.

Where possible, contaminated materials should be incinerated or autoclaved at 121 ° C for 30 minutes. In the case of non–disposable 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 4% formaldehyde solution or 2% glutaraldehyde (pH 8.0–8.5). (Caution: avoid skin contact with formaldehyde or glutaraldehyde solutions or inhalation of their vapours).

Materials and equipment that cannot be autoclaved, boiled or immersed in formalin or other solutions may be fumigated in fumigation chambers of 1 to 3 m 3 using 15–50 ml of 37% formaldehyde solution diluted 2–3–fold with water and boiled off with an electric element.

The temperature should be ambient (>18 o C) and exposure time overnight (at least 4 hours, but preferably more than 12 hours, especially if contamination is likely to be heavy or penetration by the fumigant into the material being fumigated is likely to be slow). The chamber should be properly constructed, airtight with a system of venting to the outside away from places of human or animal movement at the end of the fumigation procedure. The relative humidity within the chamber should be >90% during the fumigation procedure.

Ethylene oxide may be used as an alternative to formaldehyde if the facilities are available. The gas is acutely toxic at concentrations of >50 ppm and can cause skin burns and blistering; it is also explosive under alkaline conditions or if exposed to certain other chemicals. Although highly effective, it is really only to be recommended whe