Cover: From top, Escherichia co//O157:H7 on sorbitol MacConkey agar, Vibrio cholerae O1 on TCBS agar, and Shige/la flexneri on xylose lysine desoxycholate agar. Funding for the development of this manual was provided by the U.S. Agency for International Development, Bureau for Africa, Office of Sustainable Development. This manual was developed as a result of a joint effort by the World Health Organization Regional Office for Africa, WHO Headquarters, and the Centers for Disease Control and Prevention as part of the activities of the WHO Global Task Force on Cholera Control. In particular, the staff of the project for Improving Preparedness and Response to Cholera and Other Epidemic Diarrhoeal Diseases in Southern Africa have worked closely with many laboratorians and epidemiologists in southern Africa to develop an integrated approach to the laboratory diagnosis of cholera and dysentery upon which this manual is based. We also appreciate the valuable assistance of Ms.# Introduction
Cholera and dysentery have afflicted humankind for centuries. The epidemics they cause have affected the outcome of wars and the fates of countries. In much of the world, epidemic cholera and dysentery are uncom mon, but during the past decade these two diseases have re-emerged as causes of significant morbidity and mortality in many developing countries.
Only a few pathogens cause epidemic diarrhea, although there are many that cause sporadic diarrhea. In developing countries, two etiologic agents are responsible for most epidemic diarrhea: toxigenic Vibrio choJerae serogroup O1, which causes watery diarrhea, and ShigeJJa dysenteriae serotype 1, which causes bloody diarrhea. Recently, two additional organisms have emerged to cause epidemic diarrhea, Vibrio choJerae serogroup O139, which causes watery diarrhea, and E scherichia coJi serotype O157:H7, which causes bloody diarrhea. The latter is a common agent of diarrhea only in developed countries.
This manual focuses on the epidemiology of these four organisms and the laboratory methods used to identify them and to test their susceptibility to antimicrobial agents in the epidemic setting. The laboratory techniques and study methodology described provide accurate and useful information for the control of epidemics using a minimum of resources. The manual emphasizes coordination of the activities of the microbiologist and the epidemiologist in order to obtain information that can be generalized to develop effective treatment policies for these epidemic diarrheal diseases. It encourages focused studies to determine the organisms causing epidemics and their antimicrobial susceptibility patterns rather than relying on random information that may not accurately represent a situation.
Often the countries that face the challenge of responding to an epidemic are those with the least resources. Therefore, the microbiology laboratory must use its resources wisely in order to have the greatest impact on reducing morbidity and mortality during an epidemic. There may be several ways to reach the end result of identifying the organism causing the outbreak or the epidemic. Often, however, a small added benefit requires a much larger expenditure of materials and time. In this manual this problem is addressed specifically. The procedures described are not new; most have been used for a number of years. However, these procedures were specifically selected for testing specimens from outbreaks rather than for general use in a clinical microbiology laboratory. The selected procedures minimize the materials needed by the laboratory while deriving the most useful information.
The two most common types of epidemic diarrhea in developing countries are watery diarrhea caused by Vibrio cholerae serogroup O1 and bloody diarrhea caused by Shigella dysenteriae serotype 1 (Sd1). This chapter presents an overview of these and other organisms that cause epidemic dysentery and cholera. Knowing the epidemiology and clinical presentation of these organisms will aid in understanding the procedures presented in the following chapters.

# Epidemic cholera
Cholera is a secretory diarrheal disease caused by enterotoxin-producing strains of V. cholerae. Although over 150 serogroups of V. cholerae have been identified, for decades toxigenic V. cholerae serogroup O1 was the only known cause of epidemic cholera. After a large epidemic in Asia in 1992 and 1993, it became clear that toxigenic V. cholerae serogroup O139 also could cause epidemics very similar to those caused by V. cholerae O1. According to World Health Organization (WHO) guidelines, both V. cholerae O1 and O139 are now recognized causes of cholera and should be reported the same way. Isolates of non-O1 and non-O139 V. cholerae can cause illness, but they do not pose the public health threat of the O1 and O139 serogroups.
Additional details on the epidemiology, historical background, clinical manifes tations and treatment of cholera are presented in Chapter 5.

# Epidemic dysentery
Dysentery, defined as diarrhea with visible blood, can be caused by many different organisms, including Shigella spp., enterohemorrhagic E scherichia coli serotype O157:H7, C am pylobacter je ju n i, enteroinvasive E. coli, Salm onella spp. and, infrequently, Entam oeba histolytica. Of these organisms, the only ones known to cause large epidemics are Shigella dysenteriae serotype 1 (Sd1), and much less frequently, E. coli O157:H7. Additional details on the epidemiol ogy, historical background, clinical manifestations and treatment of Sd1 infec tion are presented in Chapter 3.
Although uncommon, a species of parasitic ameba, E. histolytica, deserves mention. This organism is an occasional cause of dysentery, especially in young adults, but does not cause epidemic disease. Asymptomatic infection with E. histolytica, however, is frequent in developing countries, being present in up to 10% of healthy persons. Examination of specimens should be done by a trained microscopist since the organism must be differentiated from nonpathogenic amebae and from white blood cells, which are often mistaken for amebic trophozoites. In some epidemics of dysentery due to Sd1, E. histolytica was also identified and initially thought to be the cause. Because of this incorrect diagnosis, persons with dysentery were treated with anti-amebic drugs, resulting in continued transmission of Sd1 and excess preventable mortality. Finding E. histolytica in a bloody stool during an epidemic of dysentery does not indicate that it is the cause of the epidemic, or even that it is the cause of dysentery in an individual patient.
E. coli O157:H7 has caused at least one large outbreak of dysentery in southern Africa. It is suspected to have caused additional outbreaks, but these were not confirmed by microbiologic culture. E. coli O157:H7 is included in this manual so that laboratory workers will be familiar with the organism and will be able to identify it if necessary. It may return in the future to cause additional epidemics; laboratories must be prepared to identify it.
Additional details on the epidemiology, historical background, clinical manifestations and treatment of E. coli O157:H7 are presented in Chapter 7.

# B. Public Health Role of the Laboratory
Clinical laboratories play an especially crucial public health role during epidemics. A laboratory may be the only one in a country that can quickly provide the information needed to develop appropriate treatment policy during an epidemic. In countries with scarce resources, the role of the laboratory is to use those resources to provide the best information for developing treatment policy, rather than to focus on the diagnosis of individual patients. During an epidemic of cholera or dysentery, the laboratory has four primary roles:
- Initial identification of the organism causing the epidemic - Initial determination of the antimicrobial susceptibility patterns - Monitoring for changes in antimicrobial susceptibility patterns - Defining the duration and geographic extent of the epidemic The World Health Organization (WHO) recommends that countries at risk for epidemics establish an epidemic control committee. Since the laboratory plays an important role in the identification and control of epidemics, a microbiologist should be a part of the epidemic control committee.

# Initial identification of the organism causing the epidemic

# Preparation/laboratory network
In countries at risk for epidemics of dysentery or cholera, the laboratory's first role is to be prepared for an epidemic. This means having the supplies (or ready access to supplies) necessary to identify V. cholerae O1/O139 and Shigella. Annexes A and B in this manual list laboratory supplies required for isolation, identification, and antimicrobial susceptibility testing. A country-wide public health laboratory network should be established (see Annex C). All countries should have at least one national or central laboratory capable of identifying V. cholerae O1/O139 and Shigella, determining antimicrobial suscep tibility, and sending isolates to an international reference laboratory (Annex D).
To maintain a laboratory's capability to determine the antimicrobial suscepti bility patterns of bacterial pathogens accurately and reproducibly, investments must be made in the infrastructure of the laboratory. These investments include a steady supply of the material resources needed to perform appropriate testing; a trained staff with expertise to conduct the laboratory tests and sufficient time, materials, and supplies to maintain this expertise; and quality control of the staff, supplies, and reagents. Because antimicrobial susceptibility testing is so resource intensive, WHO recommends that this testing be carried out at only one or two laboratories in the country. Peripheral laboratories may perform initial isolation of Vibrio spp. or Shigella spp., and then refer isolates to the central or national reference laboratory for final confirmation and determination of antimicrobial susceptibility. Peripheral laboratories may also be the sites of focused studies to determine etiologic agents causing an outbreak. First-level laboratories should be supplied with transport medium and the means of sending the specimens to the next level laboratory or to the central laboratory.

# Diagnosing epidemics
During a suspected epidemic, the laboratory will determine the organism causing the epidemic and its antimicrobial susceptibilities. An epidemic may be suspected on clinical grounds: for instance, a surveillance system based on clinical diagnosis may note an increase in the number of cases of diarrhea. The laboratory should become involved as soon as possible to identify the causative agent. This underscores the need for good communication between the labora tory, the epidemiologists, and clinicians and other health care workers.
At times, the laboratory may be the first to suspect an epidemic. Laboratory workers may note an increase in the number of stool specimens submitted, an increase in the proportion of stool specimens with blood, or the appearance of a new organism. When a laboratory worker suspects an outbreak or epidemic, he or she should contact the appropriate clinicians and public health authorities as soon as possible.
Once the organism causing the epidemic is identified, it is not necessary to examine a large number of stool specimens. Patients can be treated on the basis of their syndrome.

# D iag n o sin g d ysen tery epidem ics
If an epidemic of dysentery is suspected, the most common cause in most parts of the world is Sd1. During an outbreak or epidemic, Sd1 is likely to be isolated much more frequently than the other organisms that cause dysentery. Therefore, a laboratory should conserve its resources and, according to WHO guidelines, once Sd1 has been confirmed as the cause of an epidemic, patients presenting with dysentery should initially be treated as if they are infected with Sd1. There is no need for the laboratory to examine the stools of all patients. Rather, it is better to take specimens from a small number of patients during an outbreak or to conduct periodic surveillance for organisms causing dysentery (see below).
If Sd1 is not isolated during a suspected outbreak, the laboratory should test for E. coli O157:H7. If neither of these organisms is isolated, arrangements should be made to send specimens to a reference laboratory.
Besides Sd1 and E. coli O157:H7, a number of organisms contribute in various proportions to the burden of dysentery in a country. The predominant causes of dysentery will vary by geographic location and time of year. Seasonal peaks occur and may reflect changes in the proportions of the various causative organisms. Laboratories should conduct periodic surveys of the organisms causing dysentery in order to monitor antimicrobial susceptibility patterns and to help clinicians and public health authorities develop rational guidelines for empiric treatment. Procedures for conducting such surveys are described in Annex E.

# D iag n o sin g cholera epidem ics
If an epidemic of cholera is suspected, the most common cause is V cholerae O1. If V cholerae O1 is not isolated, the laboratory should test for V. cholerae O139. If neither of these organisms is isolated, arrangements should be made to send stool specimens to a reference laboratory.
Infection with V. cholerae O139 should be handled and reported in the same manner as that caused by V. cholerae O1. The associated diarrheal illness should be called cholera and should be reported as a case of cholera to the appropriate public health authorities.

# Determining antimicrobial susceptibility patterns of epidemic organisms
Antimicrobial susceptibilities should be determined for the first 30 to 50 isolates identified by the laboratory at the beginning of an epidemic. That number will provide sufficient information to develop antimicrobial treatment policy for the organism. After that, the laboratory should conduct periodic surveys to detect any changes in antimicrobial susceptibility patterns (see Annex E).
The laboratory should not routinely test antimicrobial agents that are not available in the country or antimicrobial agents that are not recommended by WHO as efficacious in the treatment of cholera or dysentery (see Chapters 3 and 5). In addition, if all isolates are resistant to a particular antimicrobial agent during the first round of testing (for example, Sd1 resistance to ampicillin or trimethoprim-sulfamethoxazole), it is probably not useful to test against those agents during future surveys.
Once the organisms are isolated and the antimicrobial susceptibility patterns determined, these results should be transmitted as quickly as possible to the national epidemiologist and to other public health officials. They can then be used to make rational choices for antimicrobial treatment policy.
It is useful to send 10 to 20 of the initial isolates to an international reference laboratory for confirmation of the identification and antimicrobial susceptibility pattern (Annex D).

# Monitoring for changes in antimicrobial susceptibility
As the epidemic progresses, periodic surveys of 30 to 50 isolates of the epi demic organism should be carried out to detect any changes in the antimicrobial susceptibility pattern of the organism causing the epidemic. These should be conducted every 2 to 6 months, depending on conditions and resources. Any changes should be reported to the national epidemiologist and to other public health officials to modify the antimicrobial treatment policy. If any major changes are noted, it is useful to send isolates to an international reference laboratory for confirmation (Annex D).

# Defining the duration of the epidemic
The laboratory can help define the end of the epidemic, especially with cholera epidemics. In the course of an epidemic, the number of cases may decrease for several reasons: seasonal variation, transition to an endemic state, or disappear ance of cholera from an area. Cholera may nearly disappear in cool seasons, only to reappear in the summer months. The laboratory can assist in determining if the epidemic has actually ended by periodically analyzing stool specimens from patients with acute watery diarrhea. In order for an area to be declared cholera free by WHO, twice the incubation period (a total of 10 days) must pass without evidence of V. cholerae O1/O139. However, because of seasonal variation, surveillance should be maintained for at least 12 months.
Similarly, seasonal variation is seen with epidemic dysentery. The laboratory can periodically analyze stool specimens from patients with dysentery to see if Sd1 is still present in a particular area.

# Other duties of the laboratory during an epidemic
In addition to the major duties outlined above, the laboratory can support other activities related to the epidemic.

# Epidemiologic studies
At times, the laboratory may be asked to provide laboratory support to an epidemiologic study. By combining epidemiologic and laboratory data, studies that examine modes of transmission or risk factors for illness can be more specific and provide better information for the control of the epidemic.

# Defining the magnitude of the epidemic and improving surveillance data
Cultures taken from a series of patients that meet the clinical case definition used during an epidemic can determine the predictive value of the definition. Such studies will confirm the accuracy of the case definition used for surveillance purposes and can provide a more accurate picture of the magnitude of the epidemic.
In addition, the laboratory may be called upon to support other activities such as environmental monitoring for V. cholerae O1/O139. These requests place additional demands on the resources of the laboratory. Therefore, the microbiolo gist must be part of the decision-making process to determine whether the laboratory has the capacity to support the particular request and whether it is an appropriate use of the laboratory resources. Fecal specimens should be collected in the early stages of any enteric illness, when pathogens are usually present in the stool in highest numbers, and before antibiotic therapy has been started (Table 2-1). Collect stools from patients in clean containers without disinfectant or detergent residue and with tight-fitting, leak-proof lids. Specimens should not be collected from bedpans, as they may contain residual disinfectant or other contaminants. Unpreserved stool should be refrigerated if possible and processed within a maximum of 2 hours after collection. Specimens that cannot be cultured within 2 hours of collection should be placed in transport medium and refrigerated immediately.

# References

# Global

# Placing stool in transport medium
A small amount of stool can be collected by inserting a sterile cotton-or polyester-tipped swab into the stool and rotating it. If mucus and shreds of intestinal epithelium are present, these should be sampled with the swab. Immedi ately insert the swab into transport medium. (The transport medium should have been chilled for 1 to 2 hours, if possible.) The swab should be pushed completely to the bottom of the tube of transport medium and the top portion of the stick touching the fingers should be broken off and discarded. Replace the screw cap and tighten firmly. Place the tube in a refrigerator or cold box.

# Collection of rectal swabs
Rectal swabs may be collected as follows: moisten the swab in sterile transport medium, insert through the rectal sphincter 2 to 3 cm (1 to 1.5 inches) and rotate, withdraw and examine to make sure there is some fecal material visible on the swab. Immediately insert the swab into cold transport medium as described in above paragraph. Place the tube in a refrigerator or cold box.
The number of swabs needed will depend on the number of plates to be inocu lated. In general, if specimens will be examined for more than one pathogen, at least two stool swabs or rectal swabs should be collected per patient, and both swabs should be inserted into the same tube of transport medium.

# Transport media

# Cary-Blair transport medium
Cary-Blair transport medium can be used to transport many enteric pathogens, including Shigella, Vibrio cholerae, and E scherichia coli O157:H7 (Figure 2-1). Cary-Blair's semisolid consistency provides for ease of transport, and the prepared medium can be stored after preparation at room temperature for up to 1 year. Because of its high pH (8.4), it is the medium of choice for transport and preservation of V cholerae.

# P reparation a n d q u a lity con tro l o f C ary-B lair
Prepare according to manufacturer's instructions.  When Cary-Blair is prepared, it should be dispensed into containers in sufficient volume so that swabs will be covered by at least 4 cm of medium. For example, 5-to 6-ml amounts may be dispensed into 13 x 100 mm screw cap tubes. With the caps loosened, sterilize by steaming (do not autoclave) at 100C for 15 minutes. Tighten the caps after sterilization. Cary-Blair is quite stable if stored in tightly sealed containers in a cool dark place so that the medium does not dry out. Cary-Blair may be used for up to 1 year as long as there is no loss of volume, contamination, or color change.

# Other transport media
Other transport media that are similar to Cary-Blair are Amies' and Stuart's transport media. Both of these are acceptable for Shigella and E. coli O157:H7, but they are inferior to Cary-Blair for transport of V. cholerae.
Alkaline peptone water (APW) may be used to transport V. cholerae, but this medium is inferior to Cary-Blair and should be used only when the latter medium is not available. APW should not be used if subculture will be delayed more than 6 hours from the time of collection because other organisms will overgrow vibrios after 6 hours.
Buffered glycerol saline (BGS), a transport medium that is used for Shigella, is unsuitable for transport of V. cholerae. Additional disadvantages of BGS are that it can be used for only 1 month after it is made and, being a liquid medium, is more likely to leak or spill during transport.

# Storage of specimens in transport medium
If transport medium has been stored at room temperature, it should be chilled, if possible, for 1 to 2 hours before use. Specimens preserved in transport medium should be refrigerated until processed. If specimens will be kept more than 2 to 3 days before being cultured, it is preferable to freeze them immediately at -70C. It may be possible to recover pathogens from refrigerated specimens up to 7 days after collection; however, the yield decreases after the first 1 or 2 days. Prompt plating, refrigeration, or freezing of specimens in Cary-Blair is particularly important for isolation of Shigella, which is more fragile than other enteric organisms. Fecal specimens in transport medium collected from patients with cholera need not be refrigerated unless they are likely to be exposed to elevated temperatures (>40C).

# Unpreserved specimens
When transport medium is not available, one option for suspect V. cholerae specimens is to soak a piece of filter paper, gauze, or cotton in liquid stool and place it into a plastic bag. The bag must be tightly sealed so that the specimen will remain moist and not dry out. Adding several drops of sterile saline to the bag may help prevent drying of the specimen. Refrigeration during transport is desirable but not necessary. This method is not suitable for transport of Shigella or E. coli O157:H7 specimens and is less effective than transport medium for preserving V. cholerae organisms.

# B. Preparing Specimens for Shipment
Specimen tubes should be clearly labeled with the specimen number, and if possible, the patient's name and date of collection. Write the numbers on the frosted portion of the specimen tube, using an indelible marker pen. If there is no frosted area, write the information on a piece of first-aid tape and fix this firmly on the specimen container. Patient information should be recorded on a data sheet; one copy should be sent with the specimens and another kept by the sender. A sample data sheet is provided in Annex F.
If a package is to be shipped by air, refer to packaging regulations presented in the publication, D angerous G oods R egulations (DGR). International A ir Transport A ssociation (IATA). These regulations are summarized in Chapter 13, "Packing and Shipping of Clinical Specimens and Etiologic Agents." Even if the package will be shipped by other means, these regulations are excellent guidelines for packing all infectious or potentially infectious materials.

# Refrigerated specimens
Refrigerated specimens should be transported to the laboratory in an insulated box with frozen refrigerant packs or ice. If wet ice is used, place the tubes or containers in waterproof containers such as plastic bags that can be tightly sealed to protect the specimens from the water formed by melting ice.

# Frozen specimens
Frozen specimens should be transported on dry ice. The following precautions should be observed:
- Place tubes in containers or wrap them in paper to protect them from dry ice. Direct contact with dry ice can crack glass tubes. - If the specimens are not in leakproof containers, protect them from exposure to carbon dioxide by sealing the screwcaps with tape or plastic film or by sealing the tubes in a plastic bag. Carbon dioxide will lower the pH of the transport medium and adversely affect the survival of organisms in the specimen.
- Ensure that the cool box is at least one-third full of dry ice. If the specimens are sent by air and more than 2 kg of dry ice is used, special arrangements may be necessary with the airlines. Airlines accept packages with less than 2 kg of dry ice. - Address the package clearly, including the name and telephone number of the receiving laboratory. Write in large letters: EMERGENCY MEDICAL SPECIMENS; CALL ADDRESSEE ON ARRIVAL; HOLD REFRIGER ATED (or "FROZEN" if applicable). Be sure that all applicable labels and forms, such as those required by IATA, are correctly fixed to the outside of the package.

# Chapter 3 Epidemiology of Dysentery Caused by Shigella
Epidemic dysentery in developing countries is usually caused by Shigella dysenteriae serotype 1 (Sd1). Sd1 is an unusually virulent enteric pathogen that causes endemic or epidemic dysentery with high death rates. It is the most common cause of large-scale, regional outbreaks of dysentery. In recent years, Sd1 has caused epidemic dysentery in Central America, south Asia and central and southern Africa. An epidemic in Central America from 1969 to 1973 was responsible for more than 500,000 cases and 20,000 deaths. The epidemic in central and southern Africa began in 1979, initially affecting eastern Zaire, Rwanda and Burundi. In the early 1990s, epidemic dysentery moved southward, affecting first Zambia, then Malawi, Mozambique, Zimbabwe and southern Africa. A large rise in the number of cases associated with refugee camps was seen in central Africa in 1994.

# A. Epidemiology of Shigella
The genus Shigella is divided into four species: S. dysenteriae, S. flexneri, S. boydii, and S. sonnei. Each of these species, with the exception of S. sonnei, has several serotypes (Table 3-1). In general, S. sonnei is more common in developed countries and S. flexn eri and S. dysenteriae are more frequent in developing countries. The proportions of each species vary from country to country. Sd1 differs from the other Shigella species in several ways:
- Only Sd1 causes large and prolonged epidemics of dysentery.
- Antimicrobial resistance develops more quickly and occurs more frequently in Sd1 than in other Shigella species. - Infection with Sd1 causes more severe, more prolonged, and more frequently fatal illness than does infection with other Shigella species.

# B. Clinical Manifestations
The hallmark of infection with Sd1 is diarrhea with blood (dysentery). Shigella causes dysentery by invading and destroying cells that line the large intestine, leading to mucosal ulceration, a hemorrhagic inflammatory exudate and bloody diarrhea. Apart from bloody stools, patients with dysentery often have fever, abdominal cramps and rectal pain. However, the clinical response to infection spans a wide range, from mild to severe diarrhea with or without blood. In almost half of cases, Shigella causes acute nonbloody diarrheas that cannot be distinguished clinically from diarrhea caused by other enteric patho gens. Severity of symptoms appears to be dose related. Asymptomatic infec tions may occur, but not to the extent that they do in Vibrio cholerae O1 infections. A chronic carrier state does not occur, although the organisms may be excreted for several weeks. Sd1 infections are most often severe or fatal in young children and in the elderly and malnourished. Although most patients recover without complications within 7 days, persistent diarrhea may occasion ally occur.
Infection with Sd1 can be complicated by seizures, sepsis, rectal prolapse, or toxic megacolon. A more frequent complication is the hemolytic-uremic syndrome (HUS), which is characterized by the classic triad of hemolytic anemia, thrombocytopenia and renal failure. HUS may be mild with rapid recovery, or severe, leading to kidney failure and death.

# C. Treatment
The mainstay of treatment for Sd1 infection is appropriate antimicrobial therapy, which lessens the risk of serious complications and death. Other supportive measures should be used as well.
The

# Chapter 4 Isolation and Identification of Shigella
Isolation and identification of Shigella can be greatly enhanced when optimal laboratory media and techniques are employed. The methods presented here are intended to be economical and to offer laboratorians some flexibility in choice of protocol and media. Laboratories that do not have sufficient resources to adopt the methods described in this chapter should consider sending specimens or isolates to other laboratory facilities that routinely perform these procedures. When selective or differential media are incorrectly prepared, the reactions of organisms on those media can be affected. Therefore, it would be helpful to refer to Section D, "Media for isolation and identification of Shigella," for a discussion of these media, their preparation, and appropriate quality control strains.

# A. Isolation Methods
There is no enrichment medium for Shigella that consistently provides a greater recovery rate than use of direct plating alone.

# Inoculation of selective agar
Fecal specimens should be plated as soon as possible after arrival in the laboratory. Selective media may be inoculated with a single drop of liquid stool or fecal suspension. Alternatively, a rectal swab or a fecal swab may be used. If a swab is used to inoculate selective media, an area approximately 2.5 cm (1 inch) in diameter is seeded on the agar plates, after which the plates are streaked for isolation (Figure 4-2). Media of high selectivity such as XLD require more overlapping when streaking than media of low selectivity. When inoculating specimens to a plate for isolation, it is important to use the entire plate to increase the chances of obtaining well-isolated colonies. Incubate the plates for 18 to 24 hours at 35 to 37C.  After incubation, record the amount and type of growth (e.g., lactose-ferment ing or lactose-nonfermenting) on each isolation medium for each patient specimen (a sample worksheet is presented in Figure 4-3). Colonies of Shigella on MAC appear as convex, colorless colonies about 2 to 3 mm in diameter. S. dysenteriae 1 colonies may be smaller (

# B. Biochemical Screening Tests
Identification of Shigella spp. involves both biochemical and serologic testing. The use of biochemical screening media is usually advisable to avoid wasting antisera. Most laboratories will find KIA (or TSI) to be the single most helpful medium for screening suspected Shigella isolates. If an additional test is desired, motility medium can be used to screen isolates before doing serologic testing. Section D in this chapter further describes these media.

# Kligler iron agar and triple sugar iron agar
To obtain true reactions in KIA or TSI or other biochemical tests, it is neces sary to inoculate with a pure culture. Carefully select at least one of each type of well-isolated colony on each plate. Using an inoculating needle, lightly touch only the very center of the colony. Do not take the whole colony or go through the colony and touch the surface of the plate. This is to avoid picking up contami nants that may be on the surface of the agar. If there is doubt that a particular colony is sufficiently isolated from surrounding colonies, purify the suspicious colony by streaking on another agar plate, after which the KIA or TSI slant may be inoculated.
KIA and TSI are inoculated by stabbing the butt and streaking the surface of the slant. After incubation for 18 to 24 hours at 35 to 37C, the slants are observed for reactions typical of Shigella. When incubating most biochemicals, caps should be loosened before placement in the incubator. This is particularly important for KIA and TSI. If the caps are too tight and anaerobic conditions exist, the characteristic reactions of Shigella spp. may not occur and a misleading result could be exhibited. It is also important that KIA and TSI be prepared so that the tubes have a deep butt and a long slant (see Section D).
Shigella characteristically produces an alkaline (red) slant and an acid (yellow) butt, little or no gas, and no H2S (Table 4-2; Figure 4-8    b K = alkaline (purple); A = acid (yellow); an alkaline reaction (purple) in the butt of the medium indicates that lysine was decarboxylated. An acid reaction (yellow) in the butt of the medium indicates that lysine was not decarboxylated. Motility agar should be inoculated with a straight inoculating needle, making a single stab about 1 to 2 cm down into the medium. Motility agar may be inocu lated with growth from a KIA or TSI that shows a reaction typical of Shigella. Alternately, motility agar can be inoculated at the same time as the KIA or TSI slant by using the same inoculating needle without touching the colony again. The motility agar should be inoculated first, after which, the KIA or TSI is inoculated by stabbing the butt first and then streaking the surface of the slant. Do not select a second colony to inoculate the KIA or TSI after the motility agar has been inoculated since it may represent a different organism.

# ID E N T IF IC A T IO N M O T c U R E A LIA
A 1 B D XLD X 1 X 2 X 3 MAC M 1 M 2 M 3 XLD X 1 X 2 X 3 M AC M 1 M 2 M 3 XLD X 1 X 2 X 3 MAC M 1 M 2 M 3 XYL/LAC -=
Examine after overnight incubation at 35 to 37C. Motility is indicated by the presence of diffuse growth (appearing as clouding of the medium) away from the line of inoculation (Figure 4-9). Nonmotile organisms do not grow out from the line of inoculation. Motility reactions may be difficult for inexperienced laboratorians to read; therefore reactions should be compared with positive and negative control strains. Shigella spp. are always nonmotile (Table 4-2).
The surface of the motility agar should be dry when used. Moisture can cause a nonmotile organism to grow down the sides of the agar creating a haze of growth and appearing to be motile (see Section D).
Sulfide-indole-motility medium is a combination medium that is commercially available in dehydrated form (see Section D). It can be used in place of motility medium.

# Motility agar

# Additional biochemical screening tests
Other biochemical tests such as urea medium and lysine iron agar may be used for additional screening of isolates before testing with antisera. The value of these should be assessed before using them routinely (Table 4-2, Annex G). These media, their preparation, and suggested quality control strains are described in Section D.

# Urea m ed iu m
Urea medium screens out urease-producing organisms such as K lebsiella and Proteus. Urea agar is inoculated heavily over the entire surface of the slant. Loosen caps before incubating overnight at 35 to 37C. Urease positive cultures produce an alkaline reaction in the medium, evidenced by a pinkish-red color (Figure 4-10). Urease negative organisms do not change the color of the medium, which is a pale yellowish-pink. Shigella spp. are always urease negative (Table 4-2

# ). L ysin e iro n agar
Lysine iron agar (LIA) is helpful for screening out H afnia spp. and certain E. coli, P roteus, and P rovidencia strains. LIA should be inoculated by stabbing the butt and streaking the slant. After incubation for 18 to 24 hours at 35 to 37C, organisms that produce lysine decarboxylase in LIA cause an alkaline reaction (purple color) in the butt of the medium and also on the slant (Figure 4-11). H2S production is indicated by a blackening of the medium. Organisms lacking lysine decarboxylase, produce an alkaline slant (purple) and an acid butt (yellow), no gas, and no H2S. P roteus and P rovidencia spp. will often produce a red slant caused by deamination of the lysine. LIA must be prepared so that the tubes have a deep butt (see Section D).
Shigella spp. are lysine negative and characteristically produce an alkaline (purple) slant and an acid (yellow) butt, no gas, and no H2S (Table 4-2 3-1
Serologic identification is performed typically by slide agglutination with polyvalent somatic (O) antigen grouping sera, followed, in some cases, by testing with monovalent antisera for specific serotype identification. Monovalent antiserum to S. dysenteriae 1 is required to identify this serotype, which is the most frequent cause of severe epidemic dysentery. Once one colony from a plate has been identified as Shigella, no further colonies from the same specimen need to be tested. Laboratorians should be aware that some Shigella commercial antiserum is labeled or packaged differently; for example, Shigella polyvalent A, which includes antisera to serotypes 1 through 7, may also be labeled polyvalent A1. Also, monovalent antiserum may be labeled in a way that it may be confused with polyvalent antiserum; for example, monovalent antiserum to S. dysenteriae 1 may be labeled "Shigella A1" instead of "S. dysenteriae serotype 1". When using newly purchased antisera, the laboratorian should read the package insert or check with the manufacturer if the label is not self-explanatory.

# Slide agglutination
Because S. dysenteriae 1 (followed by S. flexneri and S. sonnei) is the most common agent of epidemic dysentery, isolates that react typically in the screening biochemicals should be screened first with monovalent A1 antiserum, then with polyvalent B antiserum, and finally in polyvalent D antiserum.
Agglutination tests may be carried out in a petri dish or on a clean glass slide. An inoculating loop or needle, sterile applicator stick or toothpick is used to remove a portion of the growth from the surface of KIA, TSI, heart infusion agar (HIA), or other nonselective agar medium. Serologic testing should not be done on growth from selective media such as MAC or XLD because this may give false-negative results. Emulsify the growth in two small drops of physiological saline and mix thoroughly. Add a small drop of antiserum to one of the suspen sions. Usually approximately equal volumes of antiserum and growth suspension are mixed, but the volume of suspension may be as much as double the volume of the antiserum. To conserve antiserum, volumes as small as 10 microliters can be used. An inoculating loop may be used to dispense small amounts of antisera if micropipettors are not available (Figure 4-12). Mix the suspension and antiserum well and then tilt the slide back and forth to observe for agglutination. If the reaction is positive, clumping will appear within 30 seconds to 1 minute (Figure 4-13). Examine the saline suspension carefully to ensure that it is even and does not show clumping due to autoagglutination. If autoagglutination occurs, the culture is termed "rough" and cannot be serotyped.
Cultures that react serologically and show no conflicting results in the bio chemical screening tests are reported as positive for Shigella. Serologically negative isolates that are biochemically identified as Shigella may be sent to a reference laboratory.

# Quality control of antisera
All lots of antisera should be quality controlled before use. Quality control of antisera is discussed in Chapter 11.

# D. Media for Isolation and Identification of Shigella
This section contains descriptions of all media mentioned in this chapter and discussions of their characteristics, preparation, and appropriate quality control strains. Each manufacturer's lot number of commercial dehydrated media or each

# P reparation a n d q u a lity con tro l
Prepare according to manufacturer's instructions.  DCA medium is very heat-sensitive, and overheating during boiling should be avoided. Do not autoclave. Plates can be stored at 4C for up to a week.
For quality control of DCA, the following organisms should be adequate for confirmation of selective and inhibitory growth characteristics: E. coli may be somewhat inhibited, depending on the particular formulation used, but will produce pink colonies surrounded by a zone of precipitated bile; S. flexn eri and S. dysenteriae 1 will produce fair to good growth of colorless colonies.

# Hektoen enteric agar
Hektoen enteric agar (HE) is a differential selective agar that is useful for isolation of Salm onella and Shigella. It has an H2S-indicator system for selecting H2S-producing Salm onella, which produce blue-green colonies with a black center. Shigella colonies are green while rapid lactose-fermenters such as E. coli are pink to orange with a zone of bile precipitation.

# P reparation a n d q u a lity con tro l
Prepare according to manufacturer's instructions.  Heat to boiling to dissolve, but avoid overheating. Do not autoclave. When cool enough to pour, dispense into plates. Plates can be stored at 4C for up to 1 week.
For quality control of HE, the following organisms should be adequate for confirmation of selective and inhibitory growth characteristics: E. coli should produce colonies that are pink to orange surrounded by a bile precipitate; S. flexn eri should produce fair to good growth of green colonies, but S. dysenteriae 1 colonies should be smaller.

# Kligler iron agar and triple sugar iron agar
Kligler iron agar (KIA) and triple sugar iron (TSI) agar are carbohydratecontaining screening media widely used for identification of enteric pathogens, including Shigella. Both media differentiate lactose fermenters from nonfermenters and have a hydrogen sulfide indicator. H2S-producing organisms will cause blackening of the medium in both KIA and TSI.
KIA contains glucose and lactose. Organisms which ferment glucose cause the butt of the tube to become acid (yellow); some also produce gas. Lactosefermenting organisms will produce an acid (yellow) slant; lactose-nonfermenting organisms will have an alkaline (red) slant.
TSI contains sucrose in addition to the ingredients in KIA. Organisms which ferment either lactose or sucrose will produce an acid (yellow) slant while organisms that ferment neither carbohydrate will have an alkaline (red) slant. As in KIA, glucose-fermenters produce an acid (yellow) reaction in the butt (some times with gas produced).

# P reparation a n d q u a lity con tro l
Prepare according to manufacturer's instructions.  Dispense a quantity of medium in appropriate containers such that the volume of medium is sufficient to give a deep butt and a long slant. For example, dispense 6.5 ml of medium into 16 x 125-mm screw-cap tubes (leave caps loose), and after autoclaving allow the slants to solidify in a manner such that the medium in the butt of the tube is about 3.5 cm deep and the slant is about 2.5 cm long. Tighten caps and store at 4C for up to 6 months.
For quality control of KIA or TSI, the following organisms should be adequate for confirmation of biochemical response characteristics: E. coli should give an acid slant and butt, with the production of gas but no H2S; S. flexn eri should give an alkaline slant, acid butt, without production of gas or H2S (Figure 4-8); an H2S-producing Salm onella may be used to control this reaction.

# Lysine iron agar
Organisms that produce lysine decarboxylase in LIA cause an alkaline reaction (purple color) in the butt of the medium and also on the slant (Figure 4-11). H2S production is indicated by a blackening of the medium. Organisms lacking lysine decarboxylase, such as Shigella, typically produce an alkaline slant (purple) and an acid butt (yellow) no gas, and no H2S (Table 4-2). P roteus and P rovidencia spp. will often produce a red slant caused by deamination of the lysine. LIA must be prepared so that the volume of medium in the tube is sufficient to give a deep butt. It is important for LIA tubes to have a deep butt because the decarboxyla tion reaction occurs only in anaerobic conditions.

# P reparation a n d q u a lity con tro l
Prepare medium according manufacturer's instructions on the bottle.  Dispense a quantity of medium in appropriate containers such that the volume of medium is sufficient to give a deep butt and a long slant. For example, dispense 6.5 ml of medium into 16 x 125-mm screw-cap tubes (leave caps loose), and after autoclaving allow the slants to solidify in a manner such that the medium in the butt of the tube is 3.5 cm deep and the slant is 2.5 cm long. Tighten caps and store at 4C for up to 6 months.
For quality control of LIA, the following organisms may be used: S. flexn eri should produce an alkaline slant and an acid butt without production of H2S; an H2S-producing Salm onella strain may be used to control the H2S reaction and will most likely be lysine-positive and give an alkaline reaction in the butt of the tube.

# MacConkey agar
MacConkey agar (MAC) is a differential plating medium recommended for use in the isolation and differentiation of lactose-nonfermenting, gram-negative enteric bacteria from lactose-fermenting organisms. Colonies of Shigella on MAC appear as convex, colorless colonies about 2 to 3 mm in diameter. S. dysenteriae 1 colonies may be smaller.
Several commercial brands of MAC are available. Most manufacturers prepare several formulations of MAC, which may vary in selectivity and thereby affect the isolation of Shigella. For example, some formulations of MAC do not contain crystal violet, a selective agent; these types are not as selective and should not be used for isolation of Shigella. Oxoid MacConkey Agar No. 3, Difco Bacto MacConkey Agar, and BBL MacConkey Agar are all suitable.

# P reparation a n d q u a lity con tro l
Prepare according to manufacturer's instructions.  Sterilize by autoclaving at 121C for 15 minutes. Cool to 50C and pour into petri plates. Leave lids ajar for about 20 minutes so that the surface of the agar will dry. Close lids and store at 4C for up to 1 month. If plates are to be stored for more than a few days, put them in a sealed plastic bag to prevent drying.
For quality control of MAC, the following organisms should be adequate for confirmation of selective and inhibitory growth characteristics: E. coli should produce pink to red colonies with good to excellent growth; S. flexn eri should produce colorless colonies with fair to good growth, but S. dysenteriae 1 colonies may be smaller.

# Motility medium
Because Shigella spp. are always nonmotile, motility medium is a useful biochemical screening test. Motility is indicated by the presence of diffuse growth (appearing as clouding of the medium) away from the line of inoculation (Figure 4-9). Nonmotile organisms do not grow out from the line of inoculation. P reparation a n d q u a lity con tro l Follow manufacturer's instructions to weigh out and suspend dehydrated medium.  Heat to boiling to make sure medium is completely dissolved. Dispense into tubes or other types of containers, leaving caps loose, and sterilize at 121C for 15 min. Allow to solidify upright, forming a deep butt with no slant (e.g., about 4 to 5 ml of medium per 13 x 100-mm screw-cap tube). When the medium is solidified and cooled, leave caps loose until the surface of the medium has dried. Tighten caps and store at 4C for up to 6 months.
For quality control of motility medium, the following organisms may be used: E. coli is motile, while Shigella spp. are nonmotile. The surface of the medium should be dry when used. If moisture has accumulated in the tube, carefully pour it out before inoculating the tube. Moisture can cause a nonmotile organism to grow down the sides of the agar creating a haze of growth and appearing to be motile.

# Sulfide-indole-motility medium
Sulfide-indole-motility medium (SIM) is a commercially available combination medium that combines three tests in a single tube: hydrogen sulfide (H2S) production, indole production, and motility. The indole reaction is not useful for screening suspected Shigella isolates because strains vary in their reactions in this test. It is inoculated in the same way as motility agar, by using a needle to stab about 1 to 2 cm down into the medium, and is incubated overnight at 35 to 37C. The motility reaction is read the same as for motility medium. As in KIA or TSI, H2S production is indicated by blackening of the medium. Indole production can be tested by either the filter paper method or by adding Kovac's reagent to the tube.

# P reparation a n d q u a lity con tro l
Follow manufacturer's instructions to weigh out and suspend dehydrated medium. Heat to boiling to make sure the medium is completely dissolved. Dispense into tubes and sterilize by autoclaving for 15 minutes at 121C.
For quality control of SIM medium, the following organisms may be used: E. coli is indole positive, H2S negative, and motility positive; an H2S-producing Salm onella strain may be used to control the H2S reaction and will most likely be motile and indole negative; Shigella spp. are motility negative and H2S negative but are variable for the indole reaction.

# Urea medium
Urease-positive cultures produce an alkaline reaction in the medium, evidenced by a pinkish-red color (Figure 4-10). Urease-negative organisms do not change the color of the medium, which is a pale yellowish-pink. Shigella spp. are always urease-negative (Table 4-2). P reparation a n d q u a lity con tro l Follow manufacturer's instructions for preparation.  Prepare urea agar base as directed on the bottle. Sterilize at 121C for 15 min. Cool to 50 to 55C, then add urea concentrate according to manufacturer's directions. Before adding the urea to the agar base, make sure the agar base is cool since the urea is heat labile. Mix and distribute in sterile tubes. Slant the medium so that a deep butt is formed.
For quality control of urea medium, the following organisms may be used: P roteus spp. produce urease; E. coli is urease negative.

# Xylose lysine desoxycholate agar
Xylose lysine desoxycholate agar (XLD) is a selective differential medium suitable for isolation of Shigella and Salm onella from stool specimens. Differentiation of these two species from nonpathogenic bacteria is accomplished by xylose and lactose fermentation, lysine decarboxylation, and hydrogen sulfide production.
Shigella colonies on XLD agar are transparent pink or red smooth colonies 1 to 2 mm in diameter (Figure 4-5). S. dysenteriae 1 colonies on XLD agar are frequently very tiny, unlike other Shigella species (Figure 4-4). Coliforms appear yellow (4-6). Salm onella colonies are usually red with black centers but may be yellow with black centers.

# P reparation a n d q u a lity con tro l
Prepare according to manufacturer's instructions.  Mix thoroughly. Heat with agitation just until the medium boils. Do not overheat; overheating when boiling XLD or allowing the medium to cool too long may cause the medium to precipitate. Cool flask under running water until just cool enough to pour. Avoid cooling the medium too long. Pour into petri plates, leaving the lids ajar for about 20 minutes so that the surface of the agar will dry. Plates can be stored at 4C for up to a week.
For quality control of XLD, the following organisms should be adequate for confirmation of selective and inhibitory growth characteristics: S. flexn eri should produce fair to good growth of transparent pink or red smooth colonies that are 1 to 2 mm in diameter; S. dysenteriae 1 may produce very small transparent or red colonies; E. coli should produce poor to fair growth of yellow colonies.
Isolates of Vibrio choJerae serogroup O1 are classified into two biotypes, El Tor and classical, on the basis of several phenotypic characteristics. Currently, the El Tor biotype is responsible for virtually all of the cholera cases throughout the world, and classical isolates are not encountered outside of Bangladesh. In addition V choJerae O1 is classified into two serotypes, Inaba and Ogawa, based on agglutination in antiserum. A possible third serotype, Hikojima, has been described, but it is very rare. During an outbreak or epidemic, it is worth docu menting the biotype and serotype of the isolate; however, it is not necessary to know this information to respond appropriately to the epidemic.
Within the O1 and O139 serogroups, the ability to produce cholera toxin (CT) is a major determinant of virulence. In general, isolates of V. choJerae O1 or O139 that produce CT are considered fully virulent and capable of causing epidemic cholera (Table 5-1). Most V. choJerae isolated during cholera outbreaks will be toxigenic serogroup O1 or O139. However, some isolates of V choJerae O1 do not produce CT and cannot cause epidemic cholera. When these isolates are encountered, they must be considered within their clinical and epide miologic context. Nontoxigenic isolates may be associated with sporadic diarrheal disease.

# A. Historical Background
Cholera is thought to have its ancestral home in the Ganges Delta of the Indian subcontinent. In the nineteenth century, pandemic waves of cholera spread to many parts of the world. In 1961, a massive epidemic began in Southeast Asia; this is now recognized as the beginning of the seventh cholera pandemic. This pandemic was caused by the El Tor biotype of toxigenic V. choJerae O1. It spread rapidly through south Asia, the Middle East, and southeastern Europe, reaching Africa by 1970. In January 1991, epidemic cholera appeared in South America in several coastal cities of Peru and spread rapidly to adjoining countries. By the end of 1996, cholera had spread to 21 countries in Latin America, causing over 1 million cases and nearly 12,000 deaths. The number of cholera cases reported elsewhere in the world has also increased in the 1990s. In Africa in the early 1990s, the primary focus of cholera was in southern Africa. However, in the latter part of the decade, the focus moved to west Africa. Overall, more cases were reported from Africa in the 1990s than in a similar time period in previous decades.

# Vibrio cholerae serogroup O 139
Vibrio choJerae serogroup O139 appeared in India in late 1992. It quickly spread to Bangladesh and other Asian countries, although the rate of spread has slowed after the initial outbreaks. Through 1998, 11 countries have officially reported transmission of V choJerae O139 to WHO. Imported cases have been reported from the United States and other countries. At this time, V. choJerae O139 appears to be confined to Asia. The epidemiologic characteristics of the O139 serogroup are similar to those of the O1 serogroup. The isolation and identification characteristics of the O139 serogroup are identical to those of the O1 serogroup except that O139 antiserum is needed for identification. Biotyping tests for V cholerae O1 are not valid for V cholerae O139 or any non-O1/O139 serogroup.

# B. Clinical Manifestations
Cholera is a secretory diarrheal disease. The enterotoxin produced by V. cholerae O1 and O139 causes a massive outpouring of fluid and electrolytes into the bowel. This rapidly leads to profuse watery diarrhea, loss of circulation and blood volume, metabolic acidosis, potassium depletion, and ultimately vascular collapse and death. In severe cases, purging diarrhea can rapidly cause the loss of 10% or more of the body's weight, with attendant hypovolemic shock and death; however, 75% or more of initial infections with V. cholerae O1 or O139 may be asymptomatic, depending on the infecting dose. Of the 25% of persons with symptomatic infections, most have mild illness. Approximately 5% of patients have moderate illness that requires medical attention but not hospital ization. In only about 2% of patients does the illness progress to life-threatening "cholera gravis." Persons with blood type O are more likely to develop severe cholera than those with other blood types.

# C. Treatment
Successful treatment of cholera patients depends on rapid replacement of fluid and electrolyte losses. With proper treatment, mortality is less than 1% of reported cases. Fluids and electrolytes can be replaced rapidly through either oral or intravenous routes. Intravenous therapy is required for patients who are in profound shock or cannot drink.
Antimicrobial therapy is helpful, although not essential, in treating cholera patients. Antimicrobial agents reduce the duration of illness, the volume of stool, and the duration of shedding of vibrios in the feces. When antimicrobial agents are used, it is essential to choose one to which the organism is susceptible. Antimicrobial agents recommended by WHO for treating cholera patients include tetracycline, doxycycline, furazolidone, trimethoprim-sulfamethoxazole, erythro mycin, or chloramphenicol. Ciprofloxacin and norfloxacin are also effective. Because antimicrobial resistance has been a growing problem in many parts of the world, the susceptibility of V. choJerae O1 strains to antimicrobial agents should be determined at the beginning of an epidemic and be monitored periodically (see Annexes C and E).
For V. choJerae, the results of disk diffusion tests for ampicillin, sulfonamides, tetracycline, and trimethoprim-sulfamethoxazole (i.e., percentage of susceptible, intermediate, and resistant) correlate well with the minimum inhibitory concentra tion (MIC) results determined by broth microdilution. Disk diffusion tests should not be used for doxycycline and erythromycin because the results for these drugs are frequently inaccurate for V. choJerae O1 and O139 strains. However, the tetracycline disk test can be used to predict the likely susceptibility of isolates to doxycycline. Additional details on antimicrobial susceptibility testing are given in Chapter 9.

# D. Epidemiology
When cholera first appears in epidemic form in an unexposed population, it can affect all age groups. In contrast, in areas with high rates of endemic disease, most of the adult population have gained some degree of natural immunity because of illness or repeated asymptomatic infections. In this setting, the disease occurs primarily in young children, who are exposed to the organism for the first time, and in the elderly, who have lower gastric acid production and waning immunity. The poor are at greatest risk because they often lack safe water supplies, are unable to maintain proper hygiene within the home, and may depend on street vendors or other unregulated sources for food and drink.
Numerous investigations have linked cholera transmission to drinking water drawn from shallow wells, rivers or streams, and even to bottled water and ice. Food is the other important means of cholera transmission. Seafood has repeat edly been a source of cholera, particularly raw or undercooked shellfish harvested from sewage-contaminated beds or from environments where V. choJerae O1 occurs naturally. Although V. choJerae O1 and O139 are easily killed by drying, sunlight, and acidity, they grow well on a variety of moist alkaline foods from which other competing organisms have been eliminated by previous cooking. Cooked rice is an excellent growth medium, as are lentils, millet, and other cooked grains and legumes with neutral pH. Fruits and veg etables grown in sewage and eaten without cooking or other decontaminating procedures are potential vehicles of cholera transmission. Freezing foods or drinks does not prevent cholera transmission.
Person-to-person spread through direct contact, as by shaking hands or touch ing or by taking care of a patient, has not been shown to occur. Outbreaks on crowded hospital wards are likely to be due to contaminated food or water. Likewise, outbreaks following the funeral of a cholera patient have been caused by eating contaminated foods served at the wake, often prepared by the same persons who prepared the body for burial.

# E. Cholera Vaccine
During the past 15 years, considerable progress has been made in the develop ment of new oral vaccines against cholera. Two oral cholera vaccines, which have been evaluated with volunteers from industrialized countries and in regions with endemic cholera, are commercially available in several countries: a killed whole-cell V. choJerae O1 in combination with purified recombinant B subunit of cholera toxin (WC/rBS); and an attenuated live oral cholera vaccine, containing the genetically manipulated V. choJerae O1 strain CVD 103-HgR. The appear ance of V. choJerae O139 has redirected efforts to develop an effective and practical cholera vaccine. None of the currently available vaccines is effective against this strain.

# Chapter 6 Isolation and Identification of Vibrio cholerae Serogroups O1 and O139
Isolation and identification of V choJerae serogroups O1 and O139 can be greatly enhanced when optimal laboratory media and techniques are employed. The methods presented here are intended to be economical and to offer laboratorians some flexibility in choice of protocol and media. Laboratories that do not have sufficient resources to adopt the methods described in this chapter should consider sending the specimens or isolates to other laboratory facilities that routinely perform these procedures.

# A. Isolation Methods
Before 1992, of the more than 150 serogroups of Vibrio choJerae that have been reported, only the O1 serogroup was associated with epidemic and pandemic cholera. However in late 1992 and early 1993, large outbreaks of cholera due to a newly described serogroup, O139, were reported in India and Bangladesh. This strain, like serogroup O1 V. choJerae, produces cholera enterotoxin. Because the cultural and biochemical characteristics of these two serogroups are identical, the isolation and identification methods described below apply to both O1 and O139. Both serogroups must be identified using O-group-specific antisera. Annex A lists diagnostic supplies necessary for laboratory confirmation and antimicrobial susceptibility testing of V. choJerae.
Although V. choJerae will grow on a variety of commonly used agar media, isolation from fecal specimens is more easily accomplished with specialized media. Alkaline peptone water (APW) is recommended as an enrichment broth, and thiosulfate citrate bile salts sucrose agar (TCBS) is the selective agar medium of choice (Figure 6-1). In certain instances (for example, when the patient is in very early stages of illness), it may not be necessary to enrich specimens or use selective plating media. However, enrichment broth and a selective plating medium should always be used with convalescent patients, suspected asymptomatic infections, environmental specimens, and whenever high numbers of competing organisms are likely to be present in the specimen.
Refer to Section C, "Media and Reagents for V. choJerae," before preparing any of these media because incorrect preparation can affect the reactions of organisms in these tests. Chapter 11 discusses methods for quality control of selective media and antisera.

# Enrichment in alkaline peptone water
Enrichment in APW can enhance the isolation of V. choJerae when few organisms are present, as in specimens from convalescent patients and asymp tomatic carriers. Vibrio spp. grow very rapidly in APW, and at 6 to 8 hours will be present in greater numbers than non-Vibrio organisms. APW can be inoculated with liquid stool, fecal suspension, or a rectal swab. The stool inoculum should not exceed 10% of the volume of the broth. Incubate the tube with the cap loosened at 35 to 37C for 6 to 8 hours. After incubation, subculture to TCBS should be made with one to two loopfuls of APW from the surface and topmost portion of the broth, since vibrios preferentially grow in this area. Do not shake or mix the tube before subculturing. If the broth cannot be plated after 6 to 8 hours of incubation, subculture a loopful at 18 hours to a fresh tube of APW. Subculture this second tube to TCBS agar after 6 to 8 hours of incubation (Figure 6-1).

# Isolation from TCBS selective agar
TCBS agar is commercially available and easy to prepare, requires no auto claving, and is highly differential and selective (see Section C). Growth on this medium is not suitable for direct testing with V. choJerae antisera.

# Inoculation of TCBS
Figure 6-1 outlines the procedure for isolation of V. choJerae from fecal specimens. Inoculate the TCBS plate as described in Chapter 4 (Figure 4-2). After 18 to 24 hours' incubation at 35 to 37C, the amount and type of growth (e.g., sucrose-fermenting or sucrose-nonfermenting) on the TCBS plate should be recorded on data sheets (Figure 6-2). Colonies suspicious for V. choJerae will appear on TCBS agar as yellow, shiny colonies, 2 to 4 mm in diameter (Figure 6-3). The yellow color is caused by the fermentation of sucrose in the medium. Sucrose-nonfermenting organisms, such as V. parahaem oJyticus, produce green to blue-green colonies.

# Isolation of suspected V cholerae
Carefully select at least one of each type of sucrose-fermenting colony from the TCBS plate to inoculate a heart infusion agar (HIA) slant or another nonselective medium. Do not use nutrient agar because it has no added salt and does not allow optimal growth of V. choJerae. Using an inoculating needle, lightly touch only the very center of the colony. Do not take the whole colony or go through the colony and touch the surface of the plate. This is to avoid picking up contaminants that may be on the surface of the agar. If there is doubt that a particular colony is sufficiently isolated from surrounding colonies, purify the suspicious colony by streaking on another agar plate.
Incubate the HIA slants at 35 to 37C for up to 24 hours; however, there may be sufficient growth at 6 hours for serologic testing to be done. Slide serology with polyvalent O1 and O139 antisera is sufficient for a presumptive identification (see section B below for a description of serologic identification). F igu re 6-3. Growth of V. cholerae on TCBS F igu re 6-4. A positive oxidase test (shown here) results in the developm ent of a dark purple color within 10 seconds. V. cholerae is oxidase positive.

# Screening tests for suspected V. cholerae isolates
Generally for suspected V cholerae isolates from fecal specimens, screening with biochemical tests prior to testing with O1 and O139 antisera is not necessary. However, if the supply of antisera is limited, the oxidase test may be useful for additional screening of isolates before testing with antisera.

# O xidase test
Conduct the oxidase test with fresh growth from an HIA slant or any non carbohydrate-containing medium. Do not use growth from TCBS agar because it may yield either false-negative or false-positive results. Place 2 to 3 drops of oxidase reagent (1% N,N,N',N'-tetramethyl-p-phenylenediamine) on a piece of filter paper in a petri dish. Smear the culture across the wet paper with a platinum (not nichrome) loop, a sterile wooden applicator stick or toothpick. In a positive reaction, the bacterial growth becomes dark purple immediately (Figure 6-4). Oxidase-negative organisms will remain colorless or will turn purple after 10 seconds. Color development after 10 seconds should be disregarded. Positive and negative controls should be tested at the same time. Organisms of the genera Vibrio (including V. cholerae,  Small, curved rods with darting motility  K = alkaline; A = acid b An alkaline reaction (purple) in the butt of the medium indicates that lysine was decarboxylated. An acid reaction (yellow) in the butt of the medium indicates that lysine was not decarboxylated.

# Additional biochemical screening tests
The string reaction, Kligler iron agar (KIA) or triple sugar iron agar (TSI), lysine iron agar (LIA), Gram stain, and wet mount for motility are other possible tests that may be used for additional screening of isolates before testing with antisera (Table 6-1). The value of these tests should be assessed to determine their usefulness before they are applied routinely. See Section C for instructions on preparation of media and appropriate quality control strains.

# S trin g test
The string test, using fresh growth from nonselective agar, is useful for ruling out non-Vibrio spp., particularly A erom onas spp. The string test may be per formed on a glass microscope slide or plastic petri dish by suspending 18-to 24hour growth from HIA or other noninhibitory medium in a drop of 0.5% aqueous solution of sodium deoxycholate. If the result is positive, the bacterial cells will be lysed by the sodium deoxycholate, the suspension will lose turbidity, and DNA will be released from the lysed cells, causing the mixture to become viscous. A mucoid "string" is formed when an inoculating loop is drawn slowly away from the suspension (Figure 6-5). V. cholerae (

# K iig ier iro n ag ar a n d tripie su g a r iro n agar
Kligler iron agar (KIA) and triple sugar iron agar (TSI) can be used to rule out P seudom onas spp. and certain Enterobacteriaceae. The reactions of V. cholerae on KIA, which contains glucose and lactose, are similar to those of lactosenonfermenting E nterobacteriaceae (alkaline (red) slant, acid (yellow) butt, no gas, no H2S) (see Table 6-1 and Figure 6-6). However, on TSI, V. cholerae strains produce an acid (yellow) slant and acid (yellow) butt, no gas, and no H2S.
KIA or TSI slants are inoculated by stabbing the butt and streaking the surface of the medium. Incubate the slants at 35 to 37C and examine after 18 to 24 hours. Caps on all tubes of biochemicals should be loosened before incubation, but this is particularly important for KIA or TSI slants. If the caps are too tight and anaerobic conditions exist in the KIA or TSI tube, an inappropriate reaction will occur and the characteristic reactions of V cholerae may not be exhibited.
F igure 6-6. R eactions of V. cholerae in Kiigier iron ag ar (left) and triple sugar iron ag ar (right)

# L ysin e iro n agar
LIA is helpful for screening out A erom onas and certain Vibrio spp., which, unlike V. cholerae, do not decarboxylate lysine. LIA is inoculated by stabbing the butt and streaking the slant. After incubation for 18 to 24 hours at 35 to 37C, examine the LIA slants for reactions typical of V. cholerae. Organisms that produce lysine decarboxylase in LIA cause an alkaline reaction (purple color) in the butt of the tube (see Chapter 4, Figure 4-11). Organisms without the enzyme produce a yellow color (acid) in the butt portion of the tube. H2S production is indicated by a blackening of the medium. The LIA reaction for V cholerae is typically an alkaline slant (purple), alkaline butt (purple), no gas, and no H2S (Table 6-1). P roteus and Providencia spp. will often produce a red slant caused by deamination of the lysine.
It is important that KIA, TSI, and LIA be prepared so the tubes have a deep butt and a long slant. If the butt is not deep enough, misleading reactions may occur in these media. In LIA, the decarboxylation of lysine occurs only in anaerobic conditions and a false-negative reaction may result from insufficient medium in the tube (Section C).

# G ram sta in
Examining overnight growth from an HIA slant by Gram stain will demon strate typical small, curved gram-negative rods (Table 6-1

# ). Staining with crystal violet only is a more rapid technique and will still demonstrate the cell morphol ogy typical of Vibrio spp. W et m o u n t
Dark-field and phase-contrast microscopy have been used for screening suspected isolates of V. cholerae. With these techniques, saline suspensions are microscopically examined for the presence of organisms with typical small, curved rods and darting ("shooting star") motility (Table 6-1).

# B. Serologic Identification of V. cholerae O1 and O139

# Presumptive identification using O1 and O139 antisera
For slide agglutination testing with polyvalent O1 or O139 antisera, fresh growth of suspected V. cholerae from a nonselective agar medium should be used. Using growth from TCBS agar may result in false-negative reactions. Usually after 5 to 6 hours of incubation, growth on the surface of the slant is sufficient to perform slide serology with antisera; if not, incubate for a longer period. If the isolate does not agglutinate in O1 antiserum, test in O139 antiserum. If it is positive in the polyvalent O1 or in the O139 antiserum, it may be reported as presumptive V. cholerae O1 or O139. Presumptive V. cholerae O1 isolates should be tested in monovalent Ogawa and Inaba antisera (see below). Once one colony from a plate has been identified as V. cholerae O1 or O139, no further colonies from the same plate need to be tested.

# Confirmation of V cholerae O1 using Inaba and Ogawa antisera
The O1 serogroup of V. cholerae has been further divided into three serotypes,

# Inaba, Ogawa, and Hikojima (very rare). Serotype identification is based on agglutination in monovalent antisera to type-specific O antigens (see Table 6-2).
A positive reaction in either Inaba or Ogawa antiserum is sufficient to confirm the identification of a V. cholerae O1 isolate. Isolates that agglutinate weakly or slowly with serogroup O1 antiserum but do not agglutinate with either Inaba or Ogawa antiserum are not considered to be serogroup O1. Identifying these antigens is valid only with serogroup O1 isolates. For this reason, Inaba and Ogawa antisera should never be used with strains that are negative with poly valent O1 antiserum.
Strains of one serotype frequently produce slow or weak agglutination in antiserum to the other serotype, depending on how well the serotype-specific antisera have been absorbed. For this reason, agglutination reactions with Inaba and Ogawa antisera should be examined simultaneously, and the strongest and most rapid reaction should be used to identify the serotype. With adequately absorbed antisera, strains that agglutinate very strongly and equally with both the Ogawa and Inaba antisera are rarely, if ever, encountered. If such reactions are suspected, the strains should be referred to a reference laboratory for further examination and may be referred to as "possible serotype Hikojima."
Refer to Chapter 11 for a discussion on quality control of antisera.

# Slide agglutination procedures
Agglutination tests for V. cholerae somatic O antigens may be carried out in a petri dish or on a clean glass slide. Use an inoculating loop or needle, sterile applicator stick, or toothpick to remove a portion of the growth from the surface of HIA, KIA, TSI, or other nonselective agar medium. Emulsify the growth in two small drops of physiological saline and mix thoroughly. Add a small drop of antiserum to one of the suspensions. Usually approximately equal volumes of antiserum and growth suspension are mixed, but the volume of suspension may be as much as double the volume of the antiserum. To conserve antiserum, volumes as small as 10 microliters (0.01 ml) can be used. An inoculating loop may be used to dispense small amounts of antisera if micropipettors are not available (Figure 4-12). Mix the suspension and antiserum well and then tilt the slide back and forth to observe for agglutination. If the reaction is positive, clumping will appear within 30 seconds to 1 minute. Examine the saline suspension carefully to ensure that it does not show clumping due to autoagglutination. If autoagglutination occurs, the culture is termed "rough" and cannot be serotyped.

# Confirmation of V cholerae O139
A suspected V. cholerae isolate that reacts in O139 antiserum but not in polyvalent O1 antiserum should be sent to a reference laboratory. Confirmation of V. cholerae O139 includes testing for production of cholera enterotoxin and verification of the O139 antigen. No serotypes have been identified in the O139 serogroup. When inoculated into APW for quality control, V. cholerae O1 should show good growth at 6 to 8 hours.

# C. Media and Reagents for

# Kligler iron agar and triple sugar iron agar
Prepare according to manufacturer's instructions. Dispense a quantity of medium in appropriate containers such that the volume of medium is sufficient to give a deep butt and a long slant. For example, dispense 6.5 ml of medium into 16 x 125-mm screw-cap tubes (leave caps loose), and after autoclaving, allow the slants to solidify in a manner such that the medium in the butt of the tube is 3 cm deep and the slant is 2 cm long. Tighten caps and store at 4C for up to 6 months.
Each new lot should be quality controlled before use. E. coli should give an acid slant and butt, with production of gas but no H2S. S. flexneri should give an alkaline slant, acid butt, without production of gas or H2S .

# Lysine iron agar
 Prepare medium according to manufacturer's instructions on the bottle. Dis pense a quantity of medium in appropriate containers such that the volume of medium is sufficient to give a deep butt and a long slant. For example, dispense 6.5 ml of medium into 16 x 125-mm screw-cap tubes (leave caps loose), and after autoclaving, allow the slants to solidify in a manner such that the medium in the butt of the tube is about 3 cm deep and the slant is about 2 cm long. When the agar is cooled and solidified, tighten caps and store at 4C for up to 6 months.
Each new lot of dehydrated medium should be quality controlled before use. S. flexn eri should produce an alkaline slant and an acid butt without production of H2S. H2S-producing Salm onella strains should produce an alkaline slant and an alkaline butt with blackening of the medium due to H2S. V. cholerae strains are lysine-positive and will produce an alkaline reaction in the butt of the LIA.

# Oxidase reagent
-Tetramethyl-p-phenylenediamine dihydrochloride 0.05 g Distilled water 5.0 ml Dissolve the reagent in purified water (do not heat to dissolve). Prepare fresh daily.
Positive and negative controls should be tested every time the reagent is prepared. V. cholerae is oxidase positive; E. coli is oxidase negative.

# Sodium deoxycholate reagent (0.5%) for string test

# Sodium deoxycholate 0.5 g Sterile distilled water
100.0 ml Add sterile distilled water to sodium deoxycholate and mix well. Store at room temperature for up to 6 months.
Each new batch should be quality controlled before use. A V. cholerae O1 strain should be used as positive control. E. coli may be used as a negative control.

# Thiosulfate citrate bile salts sucrose agar
 Follow manufacturer's instructions to weigh out and suspend the dehydrated medium. Heat with agitation. Medium should be completely dissolved. Cool agar in a water bath until cool enough to pour (50 to 55C). Pour into petri plates, leaving lids ajar about 20 minutes so that the surface of the agar will dry. Close lids and store at 4C for up to 1 week.
Each new lot should be quality controlled before use since TCBS is subject to lot-to-lot and brand-to-brand variations in selectivity. V. cholerae O1 should show good growth of yellow colonies. E. coli should have none to poor growth of translucent colonies.

# Chapter 7 Epidemiology of Escherichia coliSerotype O157:H7
E scherichia coli O157:H7 is a recently recognized pathogen that causes a dysentery-like illness. The disease is typically a bloody diarrhea, often without prominent fever, that can be complicated by hemolytic uremic syndrome. It is primarily found in developed countries. Only one confirmed outbreak has occurred in a developing country-in Swaziland in 1992 affecting 20,000 persons. Other outbreaks have been thought to occur but have not been confirmed.
The major modes of transmission are through undercooked beef, unpasteurized milk, and foods that have come in contact with materials of animal origin. Water borne outbreaks have been reported, as have outbreaks associated with swimming in contaminated lakes.
The organism produces toxins similar to those produced by Shigella dysenteriae serotype 1. Treatment with antimicrobial agents has not been demonstrated to be useful in improving the course or outcome of infection with E. coli O157:H7. In fact, treating with some agents may actually worsen the outcome. Since no treatment is recommended, it is not necessary to test the antimicrobial susceptibil ity of isolates of E. coli O157:H7.
Laboratories should be familiar with this organism and should periodically look for it in stools from patients with bloody diarrhea. It is not necessary to examine every stool submitted to the laboratory for E. coli O157:H7, but it should be considered in outbreaks of dysentery in which Shigella spp. are not isolated from the stools of patients with bloody diarrhea. Laboratory supplies required for diagnosis of E. coli O157:H7 are listed in Annex H.

# Chapter 8 Isolation and Identification of Escherichia coli Serotype O157:H7
Isolation and identification of E scherichia coli serotype O157:H7 can be greatly enhanced when optimal laboratory media and techniques are employed. The methods presented here are intended to be economical and to offer laboratorians some flexibility in choice of protocol and media. Laboratories that do not have sufficient resources to adopt the methods described below should consider sending specimens or isolates to other laboratory facilities that routinely perform these procedures. Laboratory supplies required for diagnosis of E. coli O157:H7 are listed in Annex H.

# A. Isolation and Identification Methods
E. coli O157:H7 rapidly ferments lactose and is indistinguishable from most other E. coli serotypes on traditional lactose-containing media. However, unlike approximately 80% of other E. coli, nearly all isolates of E. coli O157:H7 ferment D -sorbitol slowly, or not at all. Sorbitol-MacConkey (SMAC) agar was developed to take advantage of this characteristic by substituting the carbohydrate sorbitol for lactose in MacConkey agar, and it is the medium of choice for isolation of E. coli O157:H7. Sorbitol-negative colonies will appear colorless on SMAC (Figure 8-1).
F igu re 8-1. E. c o liO 157:H7 colonies are colorless on SMAC. Non-O157 E. coli colonies are pink.
Enrichment for E. coli O157:H7 is not usually necessary for isolation of the organism from acutely ill patients.
Figure 8-2 illustrates the procedure for recovery of E. coli O157:H7 from fecal specimens. SMAC is inoculated as described in Chapter 4 (Figure 4-2). Incubate 18 to 24 hours at 35 to 37C. After 18 to 24 hours' incubation, the amount and type of growth (e.g., sorbitol-positive or sorbitol-negative) on SMAC should be recorded on data sheets for each patient specimen (Figure 8-3). Colonies suspicious of E. coli O157:H7 will appear colorless and about 2 to 3 mm in diameter (Figure 8-1).
Test sorbitol-negative colonies selected from SMAC with E. coli O157 anti serum or latex reagents (O157 antibody-coated latex and control latex) according to the procedures recommended by the manufacturer. Suspected colonies may be tested with antisera directly from the SMAC plate or subcultured to a nonselective medium (HIA, for example) and tested the next day. This provides more growth on which to perform the agglutination assay (however, some manufacturers of O157 latex reagents recommend testing only colonies taken directly from the plate). If colonies are tested directly from the plate, a colony that is positive in O157 latex reagent should also be transferred to another medium for subsequent testing. Once one colony from a plate has been identified as O157-positive, no further colonies from the same plate need to be tested.  If O157 latex reagent is used, it is important to test any positive colonies in the latex control reagent also; this is because some sorbitol-nonfermenting organisms will react nonspecifically with latex. The manufacturers of these kits recommend that strains reacting in both the antigen-specific and control latex reagents be heated and retested. However, in a study that used this procedure, none of the nonspecifically reacting strains were subsequently identified as E. coli O157:H7.
Isolates that are O157 positive should be sent to a reference laboratory for confirmation. The reference laboratory should identify isolates biochemically as E. coli because strains of several species cross-react with O157 antiserum. Identification of the H7 flagellar antigen is also required for confirmation. Isolates that are nonmotile or that are negative for the H7 antigen should be tested for production of Shiga toxins to identify pathogenic strains.
It is not necessary to test E. coli O157:H7 isolates for susceptibility to antimicrobial agents (see Chapter 7).

# B. Preparation and Quality Control of Sorbitol-MacConkey agar
Prepare according to the manufacturer's instructions.  Sterilize by autoclaving at 121C for 15 minutes. Cool to 50C and pour into petri plates. Leave lids ajar for about 20 minutes so that the surface of the agar will dry. Close lids and store at 4C for up to 1 month. If medium is to be stored for more than a few days, put plates in a sealed plastic bag to prevent drying. Each new lot should be quality controlled before use.
E. coli should produce good to excellent growth of pink to red colonies. E. c o liO157:H7 should produce colorless colonies.

# Chapter 9 Antimicrobial Susceptibility Testing (Agar Disk Diffusion Method)
The disk diffusion method presented in this chapter has been carefully stan dardized by the National Committee for Clinical Laboratory Standards (NCCLS) and if performed precisely according to the protocol below, will provide data that can reliably predict the in vivo effectiveness of the drug in question. However, any deviation from the method may invalidate the results. For this reason, if laboratories lack the resources to perform the disk diffusion test exactly as described, they should forward isolates to other laboratories for susceptibility testing.

# A. Considerations for Antimicrobial Susceptibility Testing
As antimicrobial resistance increases in many parts of the world, it becomes increasingly important to monitor the antimicrobial susceptibility of Shigella and Vibrio cholerae O1 and O139. However, because antimicrobial therapy for E scherichia coli O157:H7 infection has not been demonstrated to be efficacious or safe, except for cases of cystitis and pyelonephritis, determination of the antimicrobial susceptibility pattern is usually not meaningful.
See Chapters 3 and 5 for a discussion of appropriate antimicrobial agents for treatment of dysentery and cholera. Testing Shigella, V. cholerae, and E. coli O157:H7 against certain drugs may yield misleading results when in vitro results do not correlate with in vivo activity. Shigella spp., for instance, are usually susceptible to aminoglycosides (e.g., gentamicin, kanamycin) in the disk diffusion test, but treatment with these drugs is often not effective. Some special consider ations for susceptibility testing of V cholerae are discussed in section B below. Antimicrobial agents suggested for use in susceptibility testing of Shigella and V cholerae are listed in Table 9-1.

# B. Procedure for Agar Disk Diffusion
Figure 9-1 summarizes the disk diffusion method of susceptibility testing. Laboratory supplies required for Shigella and V. cholerae disk diffusion testing are listed in Annexes A and B.

# Mueller-Hinton susceptibility test agar
Mueller-Hinton agar medium is the only susceptibility test medium that has been validated by NCCLS. Mueller-Hinton agar should always be used for disk diffusion susceptibility testing, according to NCCLS and international guidelines. Because the way Mueller-Hinton is prepared can affect disk diffusion test results, it is very important to refer to Section C below for instructions on preparation and quality control of this medium. a If resistant to nalidixic acid, test with ciprofloxacin. b The results from the tetracycline disk are used to predict susceptibility to doxycycline also.

# McFarland turbidity standard
A McFarland 0.5 standard should be prepared and quality controlled prior to beginning susceptibility testing (see Section C). If tightly sealed to prevent evaporation and stored in the dark, the standard can be stored for up to 6 months. The McFarland standard is used to adjust the turbidity of the inoculum for the susceptibility test.

# Preparation of inoculum
Each culture to be tested should be streaked onto a noninhibitory agar medium (blood agar, brain heart infusion agar, or tryptone soy agar) to obtain isolated colonies. After incubation at 35C overnight, select 4 or 5 well-isolated colonies with an inoculating needle or loop, and transfer the growth to a tube of sterile saline (see Section C) or nonselective broth (Mueller-Hinton broth, heart infusion broth, or tryptone soy broth) and vortex thoroughly. The bacterial suspension should then be compared to the 0.5 McFarland standard. This comparison can be made more easily if the tubes are viewed against a sheet of white paper on which sharp black lines are drawn (see Figures 9-2 and 9-3). The turbidity standard should be agitated on a vortex mixer immediately prior to use. If the bacterial suspension does not appear to be the same density as the McFarland 0.5, the turbidity can be reduced by adding sterile saline or broth or increased by adding more bacterial growth.
Alternatively, the growth method may be used to prepare the inoculum. Four or five colonies are picked from overnight growth on agar and inoculated into broth (Mueller-Hinton broth, heart infusion broth, or tryptone soy broth). Incubate the broth at 35C until turbid, and then adjust the turbidity to the proper density.

# A g en ts fo r Shigella
A g en ts fo r V. cholerae

# Inoculation procedure
Within 15 minutes after adjusting the turbidity of the inoculum suspension, dip a sterile cotton swab into the suspension. Pressing firmly against the inside wall of the tube just above the fluid level, rotate the swab to remove excess liquid. Streak the swab over the entire surface of the medium three times, rotating the plate approximately 60 degrees after each application to ensure an even distribution of the inoculum (Figure 9-4). Finally, swab all around the edge of the agar surface.
F igu re 9-4. The Mueller-Hinton plate should be swabbed over the entire surface of the medium three times, rotating the plate 60 degrees after each application.

# Antimicrobial disks
The working supply of antimicrobial disks should be stored in the refrigerator (4C). Upon removal of the disks from the refrigerator, the package containing the cartridges should be left unopened at room temperature for approximately 1 hour to allow the temperature to equilibrate. This reduces the amount of conden sation on the disks. If a disk-dispensing apparatus is used, it should have a tightfitting cover, be stored in the refrigerator, and be allowed to warm to room temperature before using.

# Antimicrobial Susceptibility Testing (Agar Disk Diffusion M ethod)
Apply the antimicrobial disks to the plates as soon as possible, but no longer than 15 minutes after inoculation. Place the disks individually with sterile forceps or with a mechanical dispensing apparatus, and then gently press down onto the agar. In general, place no more than 12 disks on a 150-mm plate and no more than 4 disks on a 100-mm plate. This prevents overlapping of the zones of inhibition and possible error in measurement. Diffusion of the drug in the disk begins immediately; therefore, once a disk contacts the agar surface, the disk should not be moved.

# Recording and interpreting results
After the disks are placed on the plate, invert the plate and incubate at 35C for 16 to 18 hours. After incubation, measure the diameter of the zones of complete inhibition (including the diameter of the disk) (Figure 9-5) and record it in millimeters (Figures 9-6, 9-7). The measurements can be made with a ruler on the undersurface of the plate without opening the lid. With sulfonamides and trimethoprim-sulfamethoxazole, a slight amount of growth may occur within the inhibition zone. In this instance, slight growth (80% inhibition) should be ignored and the zone diameter should be measured to the margin of heavy growth. The zones of growth inhibition should be compared with the zone-size interpretative table (see Table 9-2), and recorded as susceptible, intermediate, or resistant to each drug tested.
Colonies growing within the clear zone of inhibition may represent resistant variants or a mixed inoculum. The distance from the colony(ies) closest to the disk to the center of the disk should be measured and then doubled to obtain a diameter. The diameter of the outer clear zone should be recorded as well and an interpretation recorded for each diameter. The colony(ies) inside the zone should be picked, re-isolated, re-identified, and retested in the disk diffusion test to confirm the previous results. The presence of colonies within a zone of inhibition may predict eventual resistance to that agent.

# Quality control
To verify that susceptibility test results are accurate, it is important to include at least one control organism (ATCC 25922 is the E. coli control strain used when testing E nterobacteriaceae and V cholerae) with each test. Zone diameters obtained for ATCC 25922 should be compared with NCCLS published limits (see Table 9-2 for diameters of the zones of inhibition for ATCC 25922). If zones produced by the control strain are out of the expected ranges, the laboratorian should consider possible sources of error.
Susceptibility tests are affected by variations in media, inoculum size, incuba tion time, temperature, and other factors. The medium used may be a source of error if it fails to conform to NCCLS recommended guidelines. For example, agar containing excessive amounts of thymidine or thymine can reverse the inhibitory effects of sulfonamides and trimethoprim, causing the zones of growth inhibition to be smaller or less distinct. Organisms may appear to be resistant to these drugs F igu re 9-5. R esults of the disk diffusion assay. This Shigella isolate is resistant to trim ethoprim -sulfam ethoxazole and is growing up to the disk (SXT), the zone of which is recorded as 6 mm. when in fact they are not. If the depth of the agar in the plate is not 3 to 4 mm or the pH is not between 7.2 and 7.4, the rate of diffusion of the antimicrobial agents or the activity of the drugs may be affected.
If the inoculum is not a pure culture or does not contain a concentration of bacteria that approximates the McFarland standard, the susceptibility test results will be affected. For instance, a resistant organism could appear to be susceptible if the inoculum is too light. Also, if colonies from blood agar medium are used to prepare a suspension by the direct inoculum method, trimethoprim or sulfonamide antagonists may be carried over and produce a haze of growth inside the zones of inhibition surrounding trimethoprim-sulfamethoxazole disks even when testing susceptible isolates.
If antimicrobial disks are not stored properly or are used beyond the stated expiration date, their potency may decrease; this will usually be indicated by a decrease in the size of the inhibition zone around the control strain.
As mentioned above, testing some bacteria against certain antimicrobial agents may yield misleading results because these in vitro results do not neces sarily correlate with in vivo activity. Examples include narrow-and expandedspectrum cephalosporins and aminoglycosides (e.g., gentamicin) tested against Shigella spp. (see Chapter 3), and erythromycin tested against V. cholerae (see section C below).

# C. Special Considerations for Susceptibility Testing of V. cholerae
Although the disk diffusion technique is the most commonly used method for antimicrobial susceptibility testing, zone size interpretive criteria for V cholerae O1 and O139 have been established only for ampicillin, chloramphenicol, sulfonamides, tetracycline and trimethoprim-sulfamethoxazole. It has been determined that disk diffusion results are not accurate for V. cholerae when testing erythromycin and doxycycline, and these agents should not be tested by this method. The results from the tetracycline disk should be used to predict suscepti bility to doxycycline. If susceptible to tetracycline, the strain will be susceptible to doxycycline. At this time there is no in vitro method to accurately determine susceptibility to erythromycin.
The reliability of disk diffusion results for other antimicrobial agents, including ciprofloxacin, furazolidone and nalidixic acid, has not been validated. Until interpretive criteria have been established for V. cholerae, disk diffusion may be used to screen for resistance to ciprofloxacin, using interpretive criteria for the E nterobacteriaceae as tentative zone size standards. Tentative breakpoints have been proposed for testing furazolidone and nalidixic acid with V. cholerae (see Table 9-2). When screening with the disk diffusion method for these agents, results should be interpreted with caution. If zone sizes for these drugs fall within the intermediate range, the organism should be considered possibly resistant.

# D. Preparation and Quality Control of Media and Reagents

# Mueller-Hinton agar
 Follow manufacturer's instructions to prepare medium. After autoclaving, cool medium to 50C. Measure 60 to 70 ml of medium per plate into 15 x 150-mm plates, or measure 25 to 30 ml per plate into 15 x 100-mm plates. Agar should be poured into flat-bottom glass or plastic petri dishes on a level pouring surface to a uniform depth of 4 mm. Using more or less agar will affect the susceptibility results. Agar deeper than 4 mm may cause false-resistance results, whereas agar less than 4 mm deep may be associated with a false-susceptibility report.
Freshly prepared plates may be used the same day or stored in a refrigerator (2 to 8C) for up to 2 weeks. If plates are not used within 7 days of preparation, they should be wrapped in plastic to minimize evaporation. Just before use, if excess moisture is on the surface, plates should be placed in an incubator (35 to 37C ) until the moisture evaporates (usually 10 to 30 min). Do not leave lids ajar because the medium is easily contaminated.
Each new lot should be quality controlled before use by testing the E. coli ATCC 25922 and/or Staphylococcus aureus ATCC 25923 standard strains. These standard strains are used with every test run for E nterobacteriaceae and gram-positive aerobes, respectively. The pH of each new lot of Mueller-Hinton should be between 7.2 to 7.4. If outside this range, the pH medium should not be adjusted by the addition of acid or base; the batch of plates should be discarded and a new batch of plates prepared. If the pH for every batch is too high or low, the entire lot of dehydrated medium may have to be returned to the manufacturer as unsatisfactory.

# Turbidity standards (McFarland)
McFarland 0.5 turbidity standards are available from various manufacturers. Alternately, the 0.5 McFarland may be prepared by adding 0.5 ml of a 1.175% (wt/vol) barium chloride dihydrate (BaCl2^2H2O) solution to 99.5 ml of 1% (vol/ vol) sulfuric acid. The turbidity standard is then aliquoted into test tubes identical to those used to prepare the inoculum suspension. Seal the McFarland standard tubes with wax, Parafilm, or some other means to prevent evaporation. McFarland standards may be stored for up to 6 months in the dark at room temperature (22 to 25C). Discard after 6 months or sooner if any volume is lost. Before each use, shake well, mixing the fine white precipitate of barium sulfate in the tube.
The accuracy of the density of a prepared McFarland standard should be checked by using a spectrophotometer with a 1-cm light path; for the 0.5 McFarland standard, the absorbance at a wavelength of 625 nm should be 0.08 to 0.1. Alternately, the accuracy of the McFarland standard may be verified by adjusting a suspension of a control strain (e.g., E. coli ATCC 25922) to the same turbidity, preparing serial 10-fold dilutions, and then performing plate counts (see Figure 9-8). The adjusted suspension should give a count of 108 colony forming units/ml. F igu re 9-8. Procedure for preparation and quality control of the M cFarland 0.5 stan dard

# Frozen storage (ultralow freezer, -70C; or liquid nitrogen freezer, -196C)
Isolates may be stored indefinitely if they are maintained frozen at -70C or below. Storage at -20C is not recommended because some organisms will lose viability at this temperature.
- give greater assurance that the medium is adequate for recovery of a small number of organisms from a clinical specimen. An example of a protocol for quality control of media is given here:
- The control strain is inoculated to nonselective broth (e.g., tryptone soy broth) and grown up overnight. - To prepare a standardized inoculum for testing selective and inhibitory media, make a 1:10 dilution of the overnight nonselective broth culture. If testing nonselective media, prepare an additional 1:10 dilution (to give a 1:100 dilution of the broth). - One tube or plate of each medium should be inoculated with the standardized inoculum of the control strain(s). When testing selective plating media, a nonselective plating medium such as heart infusion agar should be inoculated at the same time for comparison purposes. - Using a loopful of the 1:10 or 1:100 dilution prepared above (use a calibrated loop, if available) inoculate media to be tested, streaking for isolation on plating media. The same loop should be used for all quality control of all media; it is more important to use the same inoculating loop every time than it is to use a calibrated loop.

# Sources of quality control strains

# B. Quality Control of Reagents
As with all other products used in testing, reagents, either purchased or pre pared in the laboratory, should be clearly marked to indicate the date on which they were first opened and the expiration date, if appropriate. Each reagent should be tested to make sure the expected reactions are obtained. If the reagent is a rare, expensive, or difficult-to-obtain product such as diagnostic antiserum, it does not necessarily have to be discarded on the expiration date. If satisfactory sensitivity and specificity can still be verified by normal quality control procedures, the laboratory may indicate on the vial label the date of verification of quality of the reagent. All reagents should be tested for quality at intervals established by each laboratory to ensure that no deterioration has occurred.
For quality control of antiserum, two or more control strains (one positive and one negative) should be used to test its agglutination characteristics. The results of all reactions should be recorded. Following is an example of a typical quality control procedure.
- Place a drop (about 0.05 ml) of each antiserum on a slide or plate. Also, place a drop of 0.85% saline on each slide or plate to test each antigen for roughness or autoagglutination. - Prepare a densely turbid suspension (McFarland 2 or 3; see Corresponding approxim ate density o f bacteria/m l 0.5 0.5 ml 99.5 ml 1 x 108 1 0.1 ml 9.9 ml 3 x 108 0.2 ml 9.8 ml 6 x lO 8 0.3 ml 9.7 ml 9 x 10s 0.4 ml 9.6 ml 12 x lO 8 0.5 ml 9.5 ml 15 x 10s 0.6 ml 9.4 ml 18 x 108 0.7 ml 9.3 ml 21 x 108 0.8 ml 9.2 ml 24 x lO 8 0.9 ml 9.1 m l 27 x 10s 10 1.0 ml 9.0 ml 30 x lO 8

# M o u th p ip e ttin g
Mouth pipetting should be strictly prohibited in the laboratory. Rubber bulbs or mechanical devices should be used.

# S h a rp s
A high degree of precaution must always be taken with any contaminated sharp items, including needles and syringes, slides, pipettes, capillary tubes, and scalpels. Dispose of sharps in designated containers. To minimize finger sticks, used disposable needles must not be bent, sheared, broken, recapped, removed from disposable syringes, or otherwise manipulated by hand before disposal. Nondisposable sharps, including syringes, should be placed in a labeled discard pan for decontamination before cleaning. Broken glassware should not be handled directly by hand but should be removed by mechanical means such as a brush and dustpan, tongs, or forceps.

# A eroso ls
Perform all procedures carefully to minimize the creation of splashes or aerosols. Techniques that tend to produce aerosols should be avoided. Cool inoculating wires and loops by holding them still in the air for 5 to 10 seconds before touching colonies or clinical material. Loops containing infectious material should be dried in the hot air above the burner before flaming. Vortexing and centrifugation should be done in closed containers. Gauze should be used to remove the tops on blood specimens and should be placed around the top of blood culture bottles to minimize aerosol production during removal of the needle. Needles should never be cut or removed from the syringe before autoclaving. All body fluids should be centrifuged in carriers with safety caps only.
When procedures with a high potential for creating infectious aerosols are conducted or when there is a risk of splashing or spraying the face with infectious or other hazardous materials, laboratory work should be conducted in a safety cabinet or with face protection (goggles, mask, face shield or other splatter guards). Procedures that pose a risk may include centrifuging, grinding, blending, vigorous shaking or mixing, sonic disruption, opening containers of infectious materials whose internal pressures may be different from ambient pressures, inoculating animals intranasally, and harvesting infected tissues from animals or eggs. Face protection should also be used when working with high concentrations or large volumes of infectious agents.

# D eco n ta m in a tin g ben ch tops a n d o th er su rfa ces
Bench tops should be wiped with a disinfectant (a phenolic disinfectant, 1% sodium hypochlorite, or 70% alcohol) routinely after working with infectious agents or clinical specimens or after spills, splashes, or other contamination by infectious materials. Solutions of disinfectants should be maintained at the work station (see D isinfectants below).

# D isposal o f con ta m in ated m a teria ls
All discarded plates, tubes, clinical samples or other contaminated materials are to be placed in disposal containers at each bench. Special disposal boxes must be used for sharps such as syringes or broken glass to minimize the risk of injury. Avoid overfilling such containers. Containers of contaminated material should be carefully transported to the autoclave room and autoclaved before disposal.

# A u to cla vin g
An autoclave must be available for the BSL-2/3 laboratory and must be operated only by personnel who have been properly trained in its use. To verify that each autoclave is working properly, spore strips or other biological indicators designed to test for efficiency of sterilization should be included in autoclave loads on a regular basis. Each autoclave load should be monitored with tempera ture-sensitive tape, thermograph, or other means (e.g., biological indicators).

# G eneral la bo rato ry p o licies
All areas of the laboratory must be kept clean and orderly. Dirt, dust, crowd ing, or clutter is a safety hazard and is not consistent with acceptable biological research. Floors should be kept clean and free of unnecessary clutter. They should be washed with a germicidal solution on a regular basis and after any spills of infectious material have occurred.

# R efrig era to rs a n d freezers
Refrigerators and freezers should be regularly inspected for the presence of broken vials or tubes containing infectious agents. Wear gloves and proper attire when removing and discarding broken material. Refrigerators and freezers should be regularly cleaned with a disinfectant and defrosted to prevent possible contami nation and temperature failure.

# F ire p reven tio n
Keep burners away from lamps and flammable materials. Bulk flammable material must be stored in the safety cabinet. Small amounts of these materials, such as ethyl acetate, ethyl alcohol, and methanol, can be stored in safety contain ers. Turn off burners when not in use. Know the location of fire extinguishers, fire blankets, and showers. Fire safety instructions and evacuation routes should be posted.

# B. Special Practices

# T ra n sp o rt o f biohazardous m a teria ls
Transport of biohazardous materials from one building to another increases the risk of breakage and spills. If transport is necessary, the primary infectious agent container (regardless of size) must be placed in an unbreakable second container that can be sealed (e.g., screw-top tube, plastic bag).

# D isin fecta n ts
Organisms may have different susceptibilities to various disinfectants. As a surface disinfectant, 70% alcohol is generally effective for the E nterobacteriaceae, but other organisms are more resistant. However, 70% alcohol is not the disinfectant of choice for decontaminating spills. Phenolic disinfectants, although expensive, are usually effective against many organisms. Always read disinfectant labels for manufacturers' recommendations for dilution and for exposure times for efficacy, especially before use on BSL-3 organisms such as M ycobacterium tuberculosis. A good general disinfectant is a 1:100 (1%) dilution of household bleach in water; at this dilution, bleach can be used for wiping surfaces of benches, hoods and other equipment. A 1:10 (10%) dilution of bleach is corro sive and will pit stainless steel and should not be used routinely; however, it may be used to clean up spills of cultured or concentrated infectious material where heavy contamination has occurred. Dilutions of sodium hypochlorite should be made daily from a stock solution.

# D econtam ina tio n o f sp ills
The following procedure is recommended for decontaminating spills. Isolate the area to prevent anyone from entering. Wear gloves and protective clothing (gown or lab coat; mask if the spill may contain a respiratory agent or if the agent is unknown). Absorb or cover the spill with disposable towels. Saturate the towels with an appropriately diluted intermediate or high level disinfectant (e.g., a phenolic formulation or household bleach). Place disinfectant-soaked towels over the area and leave them in place for at least 15 minutes before removing and discarding them. Wipe area using clean disinfectant-soaked towels and allow area to air dry. Place all disposable materials used to decontaminate the spill into a biohazard container. Handle the material in the same manner as other infectious waste.

# A ccid en ts
All injuries or unusual incidents should be reported immediately to the supervi sor. When cuts or puncture wounds from potentially infected needles or glassware occur, the affected area should be promptly washed with disinfectant soap and water. In the event of a centrifuge accident in which safety carriers have not been used, other personnel in the area should be warned immediately and the area isolated to prevent anyone from entering.

# C. Protective Clothing and Equipment

# L ab oratory coats
Protective coats, gowns, smocks, or uniforms designated for laboratory use must be worn while working in the laboratory. This protective clothing should be removed and left in the laboratory before leaving for non-laboratory areas. All protective clothing is either disposed of in the laboratory or laundered by the institution; it should never be taken home by personnel.

# G loves
Regardless of the type of infectious material, gloves should be worn when performing potentially hazardous procedures (e.g., slide agglutination) in which there is a risk of splashing or skin contamination or when the laboratory worker has cuts or broken skin on his or her hands. Gloves should always be worn when handling clinical specimens, body fluids, and tissues from humans and animals. These tissues should be assumed to be positive for hepatitis B virus, HIV, other bloodborne pathogens, or M ycobacterium tuberculosis. Gloves must be removed when contaminated by splashing or spills or when work with infectious materials is completed. Gloves should not be worn outside the laboratory. Do not use the telephone or open doors with gloves that have been used in laboratory procedures. Dispose of all used gloves by discarding them with other disposable materials and autoclaving. Hands should be washed immediately after removing gloves.

# B a rrier p reca u tio n s
Clinical specimens, body fluids, and tissues from humans and animals should be assumed to be positive for hepatitis B virus, HIV, other bloodborne pathogens, or M ycobacterium tuberculosis. These materials should be handled in a safety cabinet or using other barrier precautions such as goggles, mask, face shield or other splatter guards whenever there is a potential for creating an aerosol. infectious substances, anyone packaging isolates or infectious specimens should consult the appropriate national regulations and the current edition of D an gerous Goods R e g u la tio n s before packing and shipping infectious substances by any means of transport. D efin itio n o f in fe ctio u s sub stan ces Infectious substances are defined as substances known to contain, or reason ably expected to contain, pathogens. Pathogens are microorganisms (including bacteria, viruses, rickettsia, parasites, fungi) or recombinant microorganisms (hybrid or mutant) that are known or reasonably expected to cause infectious disease in humans or animals. C la ssifica tio n o f clinica l/dia gn ostic specim en s
- Specimens (human, animal, food, environmental, etc.) known or reasonably expected to contain pathogens are now to be classified as infectious sub stances. When these specimens are transported/shipped for any purpose, including initial or confirmatory testing for the presence of pathogens, they are to be packaged and shipped as infectious substances (see below). - Specimens that have a relatively low probability of containing pathogens are to be classified as clinical/diagnostic specimens. When these specimens are transported/shipped for the purpose of routine screening tests or initial diagnosis for other than the presence of pathogens, they are to be packaged and shipped as clinical/diagnostic specimens. - Those specimens known not to contain pathogens are to be packaged and shipped as nonrestricted, i.e., packaging and shipping is not regulated. They are to be packaged in watertight primary containers and leakproof secondary containers. Unless it has been specifically determined, i.e., by testing, that a clinical/ diagnostic specimen does not contain a pathogen(s), it is considered to fall within the categories either of those specimens known or reasonably expected to contain pathogens or those specimens that have a relatively low probability of containing pathogens.

# G u id elin es fo r packaging and la beling in fectio u s sub stan ces
Persons who ship infectious agents or diagnostic specimens must comply with all local and international regulations pertaining to the packaging and handling of these items. They must ensure that specimens arrive at their destination in good condition and that they present no hazard to persons or animals during shipment. The inner packaging must include the following:
- An inner watertight primary container that is glass, metal, or plastic and has a leakproof seal. Petri plates should not be shipped. - A watertight, impact-resistant secondary container - Absorbent material between the primary container and the secondary container. If multiple primary containers are placed in a single secondary packaging, they must be wrapped individually to ensure that contact between them is prevented. The absorbing material, such as cotton wool, must be sufficient to absorb the entire contents of all primary containers. - An itemized list of contents between the secondary packaging and the outer packaging Multiple primary receptacles placed in a single secondary packaging must be wrapped individually, or for infectious substances transported in liquid nitrogen, separated and supported to ensure that contact between them is prevented. The absorbing material must be sufficient to absorb the entire contents of all primary receptacles. The outer packaging must meet the following requirements:
- Be of sufficient strength to adequately protect and contain the contents - Be at least 100 mm (4 inches) in its smallest overall external dimension - Be durably and legibly marked on the outside with the address and tele phone number of the consignee. A biohazard warning label must be affixed to the outside of the outer container, and must bear the inscription, "Infec tious substance. In case of damage or leakage immediately notify public health authority." Packaging for infectious substances must be marked with United Nations specification markings denoting that the packaging has been tested and certified for shipping infectious substances. Figure 13-1 illustrates these packaging recommendations. G uidelines fo r p a cka g in g a n d la b elin g clinica l/dia gn ostic specim en s n o t fo r m icrob iolog ic exam in ation Clinical specimens with a low probability of containing an infectious agent that are not being transported for examination for the presence of pathogens must be packaged as follows:
- Be "triple packaged" as described above for infectious agents - Be in packaging that will not leak after a 4-foot drop test - Have a "Clinical Specimens" label affixed to the outside of the outer container - If being shipped by air, bear the following statement, "Contents not re stricted, packed in compliance with IATA packing instruction 650." Figure 13-2 illustrates these packaging recommendations. S u p p lies need ed b y each n a tio n a l referen ce labo rato ry fo r con firm ation 5 x 100 sterile cotton or polyester swabs 5 x 500 g Cary-Blair medium 5 x 500 g TCBS medium 5 x 25 g sodium desoxycholate Glass slides for serologic testing and string test 5 x 5 g A^ALrALftetramethyl-D-phenylenediamine dihydrochloride (oxidase reagent) Filter paper Sterile wooden sticks or platinum inoculating loops 5 x 500 g nonselective agar- (e.g., tryptone soy agar, heart infusion agar) 20 x 2 ml polyvalent V. cholerae O1 diagnostic antiserum 5 x 2 ml V. cholerae O139 diagnostic antiserum 5 x 2 ml V. cholerae O1 serotype Ogawa diagnostic antiserum 5 x 2 ml V. cholerae O1 serotype Inaba diagnostic antiserum 5 x 500 g Bacto-peptone medium 5 x 500 g NaCl NaOH pH paper or pH meter 5 x 500 petri dishes (9 cm) 5 x 1000 test tubes (e.g., 13 x 100 mm or 16 x 125 mm)

# National Reference Laboratory
S u p p lies n eed ed b y each n a tio n a l referen ce labo rato ry fo r con firm ation 500 sterile cotton or polyester swabs 5 x 500 g Cary-Blair medium or other transport medium 5 x 500 g XLD medium 5 x 500 g MacConkey medium 3 x 500 g Kligler iron agar 3 x 500 g motility agar 3 x 500 g nonselective agar (e.g., tryptone soy agar, heart infusion agar) Diagnostic antisera: 20 x 2 ml monovalent S. dysenteriae serotype 1 (not Group A polyvalent) 10 x 2 ml polyvalent S. flexn eri (Group B) 5 x 2 ml polyvalent S. sonnei (Group D) Glass slides for serologic testing 5 x 500 disposable petri dishes (9 cm) 5 x 1000 disposable test tubes (e.g., 13 x 100 mm or 16 x 125 mm) A n tim icro b ia l su scep tib ility te st su p p lies fo r 100 Shigella iso lates 2 x 500 g Mueller-Hinton Agar 200 disposable petri dishes ( 9 National Reference Laboratory (Materials to confirm 500 isolates)
- Heart infusion agar (HIA) or other nonselective medium - Petri dishes - Tubes for transport (HIA used as transport medium for isolates)

# Reference level
The regional laboratory sends isolates to a national reference laboratory for confirmation and antimicrobial susceptibility testing. Each reference laboratory will need sufficient materials to confirm at least 500 isolates sent from the regional level throughout the year. The basic materials needed are as follows:
- TCBS medium

# Referral to international reference laboratories
As part of the laboratory-based surveillance process, isolates should periodi cally be sent to an international reference laboratory for confirmation of the antimicrobial susceptibility patterns. This is especially important if strains exhibit a new or unusual antimicrobial susceptibility pattern. Arrangements can be made through WHO for sending such specimens to an international reference laboratory and to provide for the rapid feedback of the results.

# Feedback of Results
R eg io n a l la bo rato ry to district, co n firm a tio n o f outbreak When the regional laboratory confirms the presence of V. cholerae O1 in stool specimens received from the district, it should contact the district as quickly as possible to inform the health authorities that V. cholerae O1 has been identified from the district.

# R eferen ce laboratories to reg io n a l laboratories
The reference laboratory should regularly communicate the results of the studies carried out on isolates submitted from the regional laboratory. The results should be sent to the regional laboratory and to the Ministry of Health. This includes the results of isolates sent both for confirmation of outbreaks and for the routine surveillance for V. cholerae carried out every 3 months. These results can serve as an internal quality control for the regional laboratories. In addition, every 3 months summaries of the results from all national reference laboratories should be distributed to the regional laboratories and appropriate persons in the Ministry of Health for further distribution to all relevant parties. The international reference laboratory should provide timely feedback of results to the national reference laboratory that is coordinating the shipping of the specimens. These results will be shared with the other reference laboratories and will serve as an external quality control for identification of V. cholerae O1 and O139 and for determining the antimicrobial susceptibility of the these strains.

# Additional components for network
This system could be expanded to include other bacterial pathogens, such as those causing dysentery. For instance, periodic surveillance of isolates from patients presenting with bloody diarrhea could be done to determine the prevalence of various organisms causing dysentery and their antimicrobial susceptibility patterns.
Interna tion a l reference laboratory to reference laboratory