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close this bookCommunicable Disease Control in Emergencies - A Field Manual (WHO - OMS, 2003, 223 p.)
View the document(introduction...)
View the documentINTRODUCTION
Open this folder and view contentsCHAPTER 1: RAPID ASSESSMENT
Open this folder and view contentsCHAPTER 2: PREVENTION
Open this folder and view contentsCHAPTER 3: SURVEILLANCE
Open this folder and view contentsCHAPTER 4: OUTBREAK CONTROL
Open this folder and view contentsCHAPTER 5: DISEASE PREVENTION AND CONTROL
View the documentANNEX 14: SAMPLE HEALTH CARD
View the documentANNEX 16: LIST OF PUBLISHERS




Blood and separated serum are the most common specimens taken in outbreaks of communicable disease. Venous blood can be used for direct isolation of the pathogen, or separated into serum for the detection of genetic material (e.g. by polymerase chain reaction), specific antibodies (by serology), antigens or toxins (e.g. by immunofluorescence). Serum is preferable to unseparated blood for the processing of most specimens for diagnosis of viral pathogens, except where otherwise directed. When specific antibodies are being assayed, it is often helpful to collect paired sera, i.e. an acute sample at the onset of illness and a convalescent sample 1-4 weeks later. Blood can also collected by finger prick for the preparation of slides for microscopy or for absorption on to special filter paper discs for analysis. Whenever possible, blood specimens for culture should be taken before antibiotics are administered to the patient.

Materials for collection of venous blood samples

· Skin disinfection: 70% alcohol (isopropyl alcohol, ethanol) or 10% povidone iodine, swabs, gauze pads, band-aids.

· Disposable latex or vinyl gloves.

· Tourniquet, Vacutainer® or similar vacuum blood collection devices, or disposable syringes and needles.

· Sterile screw cap tubes (or cryotubes if indicated), blood culture bottles (50 ml for adults, 25 ml for children) with appropriate media.

· Labels and indelible marker pen.

Method of collection

· Place a tourniquet above the venepuncture site. Disinfect the tops of blood culture bottles.

· Palpate and locate the vein. It is critical to disinfect the venepuncture site meticulously with 10% polyvidone iodine or 70% isopropyl alcohol by swabbing the skin concentrically from the centre of the venepuncture site outwards. Let the disinfectant evaporate. Do not repalpate the vein. Perform venepuncture.

· If withdrawing with conventional disposable syringes, withdraw 5-10 ml of whole blood from adults, 2-5 ml from children and 0.5-2 ml from infants. Under asepsis, transfer the specimen to appropriate transport tubes and culture bottles. Secure caps tightly.

· If withdrawing with vacuum systems, withdraw the desired amount of blood directly into each transport tube and culture bottle.

· Remove the tourniquet. Apply pressure to site until bleeding stops and apply band-aid.

· Label the tubes, including the unique patient identification number, using an indelible marker pen.

· Do not recap used sharps. Discard directly into the sharps disposal container.

· Complete the case investigation and the laboratory request forms using the same identification number.

Handling and transport

Blood specimen bottles and tubes should be transported upright and secured in a screw cap container or in a rack in a transport box. They should have enough absorbent paper around them to soak up all the liquid in case of spillage.

If the specimen will reach the laboratory within 24 hour, most pathogens can be recovered from blood cultures transported at ambient temperature. Keep at 4-8 °C for longer transit periods, unless a cold-sensitive bacterial pathogen is suspected such as meningococcus, pneumococcus, Shigella spp.


The specimen must be taken by a physician or a person experienced in the procedure. CSF is used in the diagnosis of viral, bacterial, parasitic and fungal meningitis/encephalitis.

Materials for collection

A lumbar puncture tray should be used that includes:

· sterile materials: gloves, cotton wool, towels or drapes
· local anaesthetic, needle, syringe
· skin disinfectant: 10% polyvidone iodine or 70% ethanol
· two lumbar puncture needles, small bore with stylet
· six small sterile screw-cap tubes and tube rack
· water manometer (optional)
· microscope slides and slide boxes.

Method of collection

As only experienced personnel should be involved in the collection of CSF samples, the method is not described here. CSF is collected directly into the separate screw-cap tubes. If the sample is not to be promptly transported, separate samples should be collected for bacterial and viral processing.

Handling and transport

In general, specimens should be delivered to the laboratory and processed as soon as possible.

CSF specimens for bacteriology are transported at ambient temperature, generally without transport media. They must never be refrigerated, as these pathogens do not survive well at low temperatures.

CSF specimens for virology do not need a transport medium. They may be transported at 4-8 °C for up to 48 hours, or at -70 °C for longer periods.

Rapid diagnostic tests

Several commercial kits are available, based on the direct detection of N. meningitidis antigens in CSF by latex agglutination tests. Follow the manufacturer's instructions precisely when using these tests. For best results, test the supernatant of the centrifuged CSF sample as soon as possible. If immediate testing is not possible, the sample can be refrigerated (between 2 °C and 8 °C) up to several hours, or frozen at -20 °C for longer periods. Reagents should be kept refrigerated between at 2 °C and 8 °C when not in use. Product deterioration occurs at higher temperatures, especially in tropical climates, and test results may become unreliable before the expiration date of the kit. Latex suspensions should never be frozen. Note that some kits have a working temperature range and tropical temperatures may be above the recommended upper limit.


Stool specimens are most useful for microbiological diagnosis if collected soon after onset of diarrhoea (for viruses < 48 hours and for bacteria < 4 days) and preferably before the initiation of antibiotic therapy. If required, two or three specimens may be collected on separate days. Stool is the preferred specimen for culture of bacterial and viral diarrhoeal pathogens. Rectal swabs from faeces of infants may also be used for bacterial culture, but they are not useful for the diagnosis of viruses and of little value for the diagnosis of parasites.

Materials for collection

· Clean, dry, leak-proof screw cap container and adhesive tape.
· Appropriate bacterial transport media for transport of rectal swabs from infants.
· Parasitology transport pack: 10% formalin in water, polyvinyl isopropyl alcohol (PVA).

Method of collecting a stool specimen

· Collect freshly passed stool, 5-ml liquid or 5 g solid (pea-size), in a container.
· Label the container.

Method of collecting a rectal swab from infants

· Moisten a swab in sterile saline.

· Insert the swab tip just past the anal sphincter and rotate gently.

· Withdraw the swab and examine to ensure that the cotton tip is stained with faeces.

· Place the swab in a sterile tube/container containing the appropriate transport medium.

· If necessary, break off the top part of the stick without touching the internal part of the tube and tighten the screw cap firmly.

· Label the specimen tube.

Handling and transport

Stool specimens should be transported at 4 - 8°C. Bacterial yields may fall significantly if specimens are not processed within 1-2 days of collection. Shigella spp. are particularly sensitive to elevated temperatures.

Specimens to be examined for parasites should be mixed with 10% formalin or PVA, three parts stool to one part preservative. They should be transported at ambient temperature in containers sealed in plastic bags.

Fig. A8.1. Stool sampling and transport procedures for cholera and Shigella spp.


Specimens are collected from the upper or lower respiratory tract, depending on the site of infection. Upper respiratory tract pathogens (viral and bacterial) are found in throat and nasopharyngeal specimens. Lower respiratory tract pathogens are found in sputum specimens. For organisms such as Legionella, culture is difficult, and diagnosis is best based on the detection of antigen excreted in the urine.

When acute epiglottitis is suspected, no attempt should be made to take throat or pharyngeal specimens since these procedures may precipitate respiratory obstruction. Epiglottitis is generally confirmed by lateral neck X-ray, but the etiologic agent may be isolated on blood culture.

Materials for collection

· Transport media - bacterial and viral.
· Dacron and cotton swabs.
· Tongue depressor.
· Flexible wire calcium alginate tipped swab (for suspected pertussis).
· Nasal speculum (for suspected pertussis - not essential).
· Suction apparatus or 20-50-ml syringe.
· Sterile screw-cap tubes and wide-mouthed clean sterile jars (minimum volume 25 ml).


Method of collecting a throat swab

· Hold the tongue down with the depressor. Use a strong light source to locate areas of inflammation and exudate in the posterior pharynx and the tonsillar region of the throat behind the uvula.

· Rub the area back and forth with a Dacron or calcium alginate swab. Withdraw the swab without touching the cheeks, teeth or gums and insert into a screw-cap tube containing transport medium.

· If necessary, break off the top part of the stick without touching the tube and tighten the screw cap firmly.

· Label the specimen containers.

· Complete the laboratory request form.

Method of collecting nasopharyngeal swabs (for suspected pertussis)

· Seat the patient comfortably, tilt the head back and insert the nasal speculum.

· Insert a flexible calcium alginate/Dacron swab through the speculum parallel to the floor of the nose without pointing upwards. Alternatively, bend the swab and insert it into the throat and move the swab upwards into the nasopharyngeal space.

· Rotate the swab on the nasopharyngeal membrane a few times, remove it carefully and insert it into a screw-cap tube containing transport medium.

· If necessary, break off the top part of the swab without touching the inside of the tube and tighten the screw cap firmly.

· Label the specimen tube, indicating left or right side.

· Complete the laboratory request form.

· Repeat on the other side.


Method of collecting sputum

· Instruct the patient to take a deep breath and cough up sputum directly into a wide-mouth sterile container.

· Avoid saliva or postnasal discharge. The minimum volume should be about 1 ml. Label the specimen containers.

· Complete the laboratory request form

· Always label the jar NOT the lid.

Handling and transport

· All respiratory specimens, except sputum, are transported in appropriate bacterial/viral media.

· Transport as quickly as possible to the laboratory to reduce overgrowth by commensal oral flora.

· For transit periods up to 24 hours, transport bacterial specimens at ambient temperature and viruses at 4-8 °C in appropriate media.


Materials for collection

· Sterile plastic cup with lid (50 ml or more).
· Clean, screw-top specimen transport containers ("universal" containers are often used).
· Gauze pads.
· Soap and clean water (or normal saline) if possible.

Method of collection

· Give the patient clear instructions to pass urine for a few seconds, and then to hold the cup in the urine stream for a few seconds to catch a mid-stream urine sample. This should decrease the risk of contamination from organisms living in the urethra.

· To decrease the risk of contamination from skin organisms, the patient should be directed to avoid touching the inside or rim of the plastic cup with the skin of the hands, legs or external genitalia. Tighten the cap firmly when finished.

· For hospitalized or debilitated patients, it may be necessary to wash the external genitalia with soapy water to reduce the risk of contamination. If soap and clean water are not available, the area may be rinsed with normal saline. Dry the area thoroughly with gauze pads before collecting the urine.

· Urine collection bags may be necessary for infants. If used, transfer urine from the urine bag to specimen containers as soon as possible to prevent contamination with skin bacteria. Use a disposable transfer pipette to transfer the urine.

· Label the specimen containers.

Handling and transport

· Transport to the laboratory within 2-3 hours of collection. If this is not possible, do not freeze but keep the specimen refrigerated at 4-8 °C to reduce the risk of overgrowth of contaminating organisms.

· Ensure that transport containers are leak-proof and tightly sealed.


Chlorine is the recommended disinfectant for use in field laboratories. An all-purpose disinfectant should have a working concentration of 0.1% (= 1 g/litre = 1000 ppm) of available chlorine. A stronger solution of 0.5% (= 5 g/litre = 5000 ppm) available chlorine should be used in situations such as suspected viral haemorrhagic fever outbreaks.

In preparing appropriate dilutions, one must keep in mind that different products have different concentrations of available chlorine. To prepare solutions with the above concentrations, the manufacturer may provide appropriate instructions. Otherwise, use the guidelines provided below. Chlorine solutions gradually lose strength, and freshly diluted solutions must therefore be prepared daily. Clear water should be used because organic matter destroys chlorine.

Commonly used chlorine-based disinfectants include:

· sodium hypochlorite;

· commercial liquid bleaches such as household bleach (e.g. Chlorox®, Eau-de-Javel), which generally contain 5% (50 g/litre or 50 000 ppm) available chlorine.

However, the latter preparations lose a proportion of their chlorine content over time. Thick bleach solutions should never be used directly for disinfection purposes in disasters as they contain potentially poisonous additives.

To prepare a 0.1% chlorine solution with commercial bleach, make a 1 in 50 dilution, i.e. 1 part bleach in 49 parts water to give final concentrations of available chlorine of 0.1%. (For example, this could entail adding 20 ml of bleach to approximately 1 litre of water.)

Similarly, to make a 0.5% chlorine solution, make a 1 in 10 dilution, i.e. 1 part bleach in 9 parts water to give final concentrations of available chlorine of 0.5%.(e.g. add 100 ml of bleach to 900 ml water.)

Chloramine powder

While the above-described bleach solution may satisfy all disinfection needs, chloramine powder may prove convenient for the disinfection of spills of blood and other potentially infectious body fluids. It may also prove useful under field conditions because of ease of transport. It contains approximately 25% available chlorine. In addition to its use as a powder on spills, chloramine powder may be used to prepare liquid chlorine solutions. The recommended formula is 20 g of chloramine powder to 1 litre of clean water.

Decontamination of surfaces

Wear an apron, heavy-duty gloves and other barrier protection if needed. Disinfect surfaces by wiping clean with 0.1% chlorine solution, then incinerate all absorbent material in heavy-duty garbage bags.

Decontamination of blood or body fluid spills

For spills, chloramine granules should be very liberally sprinkled to absorb the spill and left for at least 30 minutes. If chloramine powder is not available, one may use 0.5% chlorine solution to inactivate pathogens before soaking up the fluid with absorbent materials. These absorbent materials must then be incinerated.

Sterilization and reuse of instruments and materials

In field outbreak situations, it is not advisable to consider sterilization and reuse of any instruments or materials. Sterilization techniques are therefore not required, and are not described here

Disinfection of hands

The principal means for disinfecting hands is by washing with soap and water. If available, one may also use commercial hand disinfectants containing chlorhexidine or polyvidone iodine.


Any laboratory facility must fulfil four criteria:
It must be able to undertake the types of test required.
It must be able to handle the specimen load.
It must be safe and comfortable for the staff to work in.
If it is established in a community (rather than in a temporary camp) it should be sustainable in the long term.

To meet these needs it must have:
Adequate numbers of staff who have been trained in the tests to be undertaken.
Defined standard operating procedures covering the tests to be undertaken
Internal and external quality control to ensure consistency and accuracy of output.
A safety policy based on the tests undertaken and the risks posed by the organisms present in the area.
The appropriate equipment, reagents, media, glassware and disposables
A suitable building or room(s) appropriately laid out and furnished
Technical, engineering and logistic support.
Good access and external communications


In the early phases of a disaster only well trained staff should be employed for technical work. There will not be sufficient time to give to training laboratory assistants. The technical staff employed should be experienced in the relevant fields (particularly parasitology, and haematology although some knowledge of biochemistry, bacteriology and virology may be needed). At least one should have had experience of running a laboratory, preferably in field conditions, and be able to undertake additional duties such as laboratory management, ordering stock etc.

As the situation stabilises, the opportunity to expand the numbers of staff and to begin training or retraining local staff can begin.


Merely meeting the basic design and safety criteria for diagnostic laboratories is not enough. The staff must be able to undertake the required tests effectively and accurately. The output of the laboratory must be validated and a quality control system is essential. The way in which all diagnostic procedures undertaken in a laboratory should be performed should be laid down in "Standard Operating Procedures" (SOPs). These should include reference to full risk and hazard assessments and safety procedures. Protocols for internal and external quality assessment should also be laid down in these SOPs.

Internal quality control

All procedures undertaken in the laboratory must be measured against recognised standards. New batches of stains or reagents must be validated against the old. The work of the laboratory staff should be validated regularly by the blind inclusion of known positive and negative specimens in the routine diagnostic work.

External quality control

A suitable body to undertake external quality control should be identified as soon as possible after the establishment of the laboratory. This agency should provide known positive and negative specimens for assessment of the work of the laboratory.


Any agency intending to undertake medical work in an area should obtain detailed information as to the spectrum of diseases that it will have to deal with. This is essential both for the design of the laboratory "package" required and will help determine the microbiological risks that may face its staff (all its staff, not just laboratory staff members). The risks posed by any organism likely to be encountered can be classified according to the following criteria:


Mode of transmission and host range.

This can be influenced by:existing levels of immunity density of the host population host population movements

Vectors and reservoirs weather (ii) environmental factors (topography, plant species and distribution etc.). (iii) sanitation and environmental hygiene

· Availability of effective preventive measures including: Immunisation / preventive antisera

i. Sanitation
ii. Vector and reservoir control

· Availability of effective treatment


i. Passive immunisation
ii. Post-exposure vaccination
iii. Antimicrobials/chemotherapeutic agents (including any data on resistance patterns)


In the emergency phase it is likely that only basic laboratory facilities can be established and hence only a limited number of tests can be offered. As the situation stabilises it will be possible to establish a more sophisticated laboratory and hence to offer a wider range of tests.

1) Emergency phase

· Malaria - microscopy and/or spot tests

· Meningococcal meningitis - spot tests

· Stool examinations for ova and parasites

· If blood transfusions are being undertaken there must be a facility that can type and cross match blood and test for HIV.

· Culture and identification of Shigella dysenteriae

· Detection of blood borne agents other than malaria (trypanosomes, leishmanias, rickettsia)

· Haematocrit

· Differential white cell counts

· Sickle cell detection

· Clotting time

2) Post acute emergency

Sputum microscopy for the diagnosis of TB is only of value if the condition can be treated and this should only be done in the context of a properly designed and functioning DOTS programme.


The safety of the staff must be a prime consideration when a laboratory is set up. It is wholly unethical to expect staff to work in conditions where safety is ignored. The level of safety required has profound implications for the design and working of the laboratory.

Safe working in the laboratory depends on the observance of basic safety precautions (Table A9.1) and on good training of staff both in safety and in good bench work. The level of the safety precautions that may need to be established over and above the general safety principles (Table A9.1) will depend on:

· the types of tests to be done

· the types of organism present and the risks they pose due to their pathogenicity, mode of transmission etc.

· whether work higher risk work is appropriate at the local level

The safety levels that can be achieved in a laboratory (and hence the types of test that can be offered) also depend on the possibility of maintaining and sustaining safety equipment.

Infectious micro-organisms can be classified into four risk groups (Table A9.2) and the level of safety needed to handle them can then be determined (Table A9.3).

Table A8.1. General Safety Principles

1) The laboratory should have a written manual of safe practice and this should be followed at all times.

2) A first aid box must be provided and a staff member trained in first aid should be present at all times when the laboratory is working.

3) Eyewash facilities must be provided.

4) Only the laboratory staff should be permitted to enter the working area of the laboratory.

5) Laboratory staff should wear protective clothing, which should be removed when they leave the laboratory itself. It should not be worn in laboratory support areas such as offices, staff rooms etc.

6) Protective clothing should never be stored in the same lockers as street clothing. Appropriate shoes should be worn.

7) Open toed shoes (sandals) are not suitable for wear in the laboratory. Face protection (goggles/masks/eyeshields) should be provided and worn when procedures that may produce aerosols or splashes are undertaken.

8) Rubber gloves should always be worn when handling specimens and they or other appropriate protective gloves should be worn for other hazardous procedures.

9) Mouth pipetting should be absolutely forbidden.

10) Hypodermic syringes and needles should not be used as pipetting devices.

11) All contaminated material (specimens, glassware, sharps etc) should be decontaminated before disposal or cleaning for re-use.

12) To this end, appropriate containers (sharps bins, sealable plastic bags, disinfectant pots) and disinfectants must be provided. A predictable unidirectional airflow across and out of the laboratory should be maintained when the laboratory is in use (see "Ventilation" below).

13) Eating, drinking, smoking and applying cosmetics should be forbidden in the laboratory.

14) Laboratory staff should clean and disinfect all benches at the end of the working day or if infectious material is spilt.

15) Laboratory staff should always wash their hands when leaving the laboratory and facilities must be provided for this purpose.

16) All spills, accidents etc. should be reported to the laboratory supervisor.

Table A8.2. Infectious micro-organisms classified by risk group





No or very low individual and community risk

A micro-organism that is unlikely to cause animal or human disease


Moderate risk to individuals, low risk to the community

A pathogen that can cause human or animal disease but is unlikely to be a serious hazard to laboratory workers, the community, livestock or the environment. Laboratory exposures may cause serious infection but effective treatment and preventive measures are available and the risk of spread of infection is limited


High risk to individuals, low risk to the community

A pathogen that usually causes serious human or animal disease but does not ordinarily spread from one infected individual to another. Effective treatment and preventive measures are available.


High risk to individuals and the community

A pathogen that usually causes serious human or animal disease and that can be readily transmitted from one individual to another, directly or indirectly. Effective treatment and preventive measures are not usually available.

Table A8: Risk groups, biosafety levels, laboratory practice and safety equipment

Risk group

Biosafety level

Types of laboratories

Laboratory practice

Safety equipment


1 (Basic)

Basic teaching


None. Open bench work


2 (Basic)

Primary health services, primary level hospital diagnostic, teaching and public health services

GMBT + basic protective clothing; biohazard signs

Open bench + Class I or II BSC** for potential aerosols


3 (Containment)

Special diagnostic

As level 2 + special protective clothing, controlled access, directional air flow

Class I or II BSC and/or other primary containment for all work


4 (Maximum containment)

Dangerous pathogen units

As level 3 + airlock entry, shower exit, special waste disposal

Class III BSC or positive pressure suits, double ended autoclave, filtered air

* GMBT = Good microbiological bench technique
** BSC = Biological safety cabinet

Where toxic or corrosive chemicals are involved a suitable fume hood with an extractor fan will be required. Such equipment is not safe for work involving biological hazards. Where the work involves dangerous pathogens, (risk group 2 and higher - Tables1 & 2) properly designed safety cabinets will be required.

In general the type of laboratory that will be set up in the early stages of an emergency will be of basic Level 1. However, in many areas where disasters occur there is a risk of exposure to organisms in the higher risk groups. This must be taken into account when the laboratory is set up. The tests that can be done may be limited by the potential risk. It is rare that high biosafety levels will be appropriate for a local laboratory in an emergency but it may well be that as a result some tests simply cannot be undertaken locally because safe working cannot be guaranteed.


The ability of a laboratory to perform even the most basic tests depends on the quality of its equipment. The equipment must not only be suitable for the tests required, it must also be safe. It should be designed with certain general principles in mind:

1) It should prevent (or at least limit) contact between the operator and infectious material

2) It should be made from materials that are corrosion resistant, impermeable to liquids and sufficiently strong.

3) It must be free of sharp edges and moving parts must be protected.

4) It must be simple to install, operate, maintain, decontaminate and clean.

5) It must be electrically safe.


When considering where to buy microscopes consider buying locally if this does not compromise quality. It could greatly ease supply of spares, maintenance etc.

Centrifuges may be required for:

· Measurement of haematocrit (Packed Cell Volume - PCV)
· Separation of blood cells from plasma
· Concentration of casts and cells in urine
· Concentration of cells in CSF
· Concentration of stool samples


World Health Organisation publications:

Basic Laboratory Methods in Medical Parasitology. WHO, Geneva, 1991.

Bench Aids for the diagnosis of malaria infections. (2nd Edition). WHO Geneva, 2000.
Bench Aids for the diagnosis of filarial infections. WHO Geneva, 1997.
Bench Aids for the diagnosis of intestinal parasites. WHO,Geneva 1994.

Guidelines for the Collection of Clinical Specimens during Field Investigation of Outbreaks. WHO Department of Communicable Disease Surveillance and Response. Geneva, 2000.

Health Laboratory Facilities in Emergency and Disaster Situations. WHO Regional Office for the Eastern Mediterranean, Alexandria, 1994.

Laboratory Biosafety Manual. (2nd Edition). WHO, Geneva, 1993.

Maintenance and repair of laboratory Diagnostic, Imaging and Hospital Equipment. Geneva, 1994.

Safety in Health-Care Laboratories. Geneva, 1997.

Selection of Basic Laboratory Equipment for Laboratories with Limited Resources. WHO Regional Office for the Eastern Mediterranean, 2000.

2) Other relevant publications:

Cheesbrough M. Laboratory Practice in Tropical Countries.(Part 1). Cambridge University Press, Cambridge, 1998.

Cheesbrough M. Laboratory Practice in Tropical Countries. (Part 2) Cambridge University Press, Cambridge, 2000.

Medecins Sans Frontieres. Refugee Health. Macmillan, London, 1997.