Despite the relatively high cost of running a laboratory service and the low per capita healthcare budget in under-resourced countries, there are few data available on which to base rational decisions about ‘essential’ laboratory tests.^10,¹¹ In many of these countries, decisions are made at central level by healthcare planners without adequate consultation with laboratory managers and experts. The selection of ‘essential’ tests at each level should be based on the clinical and public health needs of the local community and the availability of qualified clinical and laboratory staff, as well as on the availability of funds. Essential test packages are usually defined as part of national policy and standards, taking into consideration medium- and long-term trends and the requirements of disease control programmes.

To ensure cost-effectiveness of the laboratory service, tests with no proven value should be eliminated and new tests for which there is independent evidence of clinical usefulness should be introduced,¹² as described in Chapter 24, p. 567. Tests that provide objective qualitative or quantitative information are preferred. Although it is not possible to draw up a list of essential tests that will be applicable to all countries, or even to different regions within a country, the following aspects should be considered.

Cost per Test

Often the cost of a test is calculated from the price of reagents divided by the number of tests performed. However, this oversimplifies the situation and is not accurate enough to form the basis for national policy decisions and budget allocation.¹¹ The factors that need to be taken into account when calculating the total annual costs for a laboratory are given in Chapter 24.

Diagnostic Reliability

It is important to know the sensitivity and specificity of a test and its predictive values (as calculated on p. 567) when selecting a laboratory test for clinical use. This information may be provided by manufacturers but it may not be locally applicable and, in many under-resourced countries, local test evaluation data are not available. In some countries, the ‘gold standard’ diagnostic services needed to determine ‘true positive’ and ‘true negative’ data in the local context are also lacking.

The quality of all tests carried out by a laboratory should be regularly monitored; systems for doing this are well-established (see Chapter 24) but are not easily implemented in under-resourced settings. The quality of a test influences its usefulness as well as its utilization by clinicians and community members. For example, if the result of a test in routine practice is correct only 80% of the time, then one in five tests will be wasted, reducing the effectiveness of the test by 20%. Furthermore, the inaccurate test may result in a patient receiving inappropriate treatment. Clinicians and the general public are increasingly aware of the need for accurate diagnosis. Clinicians may not order tests or use the results in patient management decisions if they do not trust the quality of the laboratory results¹³ and poor quality healthcare may deter patients from using health facilities.

Clinical Usefulness

An assessment of the clinical usefulness of a test should be carried out by an independent clinician who is familiar with local diseases and the diagnostic support services that are available. This assessor needs to compare actual clinical practice with locally agreed ‘best practice’ or, if available, local guidelines.¹² From observation of a range of clinical interactions, the percentage of times that ideal practice is followed can be calculated. For example, transfusion guidelines may recommend that transfusions are given routinely to children with an Hb of <5 g/dl. The assessor can record how many children with Hb below this level failed to receive a transfusion and how many transfusions were given without waiting for the Hb result or at an inappropriate Hb level. For each test, the assessor needs to judge whether the test has been appropriately requested and is used to influence patient management or public health decisions. The percentage of tests that are not used to guide clinical decisions will provide a figure for ‘clinical wastage’ of the test that can be entered into the formula (Chapter 24).

Maintaining quality and reliability of tests

Paradoxically, it is in under-resourced laboratories, where equipment and supplies are limited and training and supervision may be minimal, that the level of skills and motivation required to maintain a good-quality service need to be highest. Even the most basic of laboratories should ensure that procedures are in place to monitor quality (see Chapter 25). In addition to monitoring the technical quality of each test, the quality of the whole service must be ensured both within the laboratory and between laboratories. Standard operating procedures (see p. 575) should be drafted for every method performed in the laboratory; these can be adapted from existing standard operating procedure ‘models’. In addition to providing standardized techniques, standard operating procedures are excellent teaching resources and adherence to these procedures will minimize errors. Standard operating procedures need to be regularly reviewed and updated to keep pace with technical developments and changes in local circumstances (e.g. non-availability of reagents, technical limitations).

Quality Control of a Test Method (Technical Quality)

Methods for the control of various haematology tests are described in Chapter 25 but some may need to be adapted to specific local circumstances in resource-poor countries. For example, if commercial controls are not affordable, each batch of sickle cell screening tests should include known positive and negative samples from a previous batch of tests; for monitoring constancy of Hb estimations, a high and a low value sample can be re-tested several times during the day.

Internal Quality Control

Internal quality control is a system within an individual laboratory for ensuring that the technical elements of the test are of acceptable quality. Monitoring of quality by processing control samples and plotting a control chart (see p. 593) will highlight major problems with the system. For example, an inaccurate differential white cell count may be due to problems with sample collection and handling, slide preparation, fixing and staining, morphological interpretation and microscope quality as well as inadequate microscopy technique. Measures such as the introduction of standard operating procedures, in-service training and equipment maintenance schedules can help to improve performance and reduce inaccuracies.

External Quality Assessment

The principles of external quality assessment (EQA) are described on p. 595. Poor communications and transport facilities make it difficult, but not impossible, to establish EQA in under-resourced laboratories. Among its other functions, EQA is used to identify poorly performing laboratories that require assistance and to detect the use of inaccurate techniques. Although participation in international, national or local external quality assessment schemes may be beyond the resources of a small rural laboratory, it should be possible for them to exchange samples and results with neighbouring facilities. Rural laboratories can also take advantage of supervisory visits by district officers to exchange samples, including external quality assessment materials and results with other laboratories. Accreditation schemes (see p. 577), either national or local, can be set up to formally recognize laboratories that are performing well and to assist those that are not.¹⁴ Step-wise, rather than pass/fail, accreditation schemes provide a better measure of performance and motivate laboratories to improve their services.

Basic haematology tests

Haemoglobinometry

Various methods for the measurement of Hb are given in Chapter 3. The most accurate method that may be available in under-resourced laboratories is the hemoglobin-cyanide (cyanmethaemoglobin) method. However, this requires a power source and considerable technical expertise to carry out accurate dilutions and to prepare the standard curve. Use of pre-set haemoglobinometers removes the need for calibration curves, but the machine may still need to be set correctly.

Methods for measuring Hb that are robust, accurate and can be used by health workers in peripheral settings are described below.

Direct Reading Haemoglobinometers

HemoCue Blood Hemoglobin System *

HemoCue Blood Hemoglobin System (see Chapter 3) is a battery- or mains-operated portable, direct read-out machine that uses disposable dry chemistry cuvettes. Measurements are precise and accurate (provided that only the specified cuvettes are used and the reading surfaces are kept clean) and, unlike most other systems, it does not require pre-dilution of the sample. Although the use of the unique disposable cuvettes makes this method relatively expensive, the cuvettes for the new Hb 301 version are cheaper than those for the Hb 201 and are designed for adverse climatic conditions. It is fast and very simple to use so that the cost may be offset by savings on training and supervision time.

Haemoglobin Colour Scale ^†

Many colour comparison methods have been developed in the past but these have become obsolete because the colours were not sufficiently comparable with blood or were not durable. The haemoglobin colour scale has been developed for anaemia screening where there is no laboratory.¹⁵ It consists of a set of printed colour shades representing haemoglobin levels between 4 and 14 g/dl (i.e. 40 and 140 g/l). The colour of a drop of blood collected onto a specific type of paper matrix is compared with that on the chart (Fig. 26.2). It is intended for detecting the presence of anaemia and estimating its severity in 2 g/dl (20 g/l) increments. The scale has been tested in resource-poor settings but more studies are needed to demonstrate whether it is superior to clinical diagnosis.¹⁶^–¹⁸ The instructions should be followed carefully because poor lighting, allowing the blood spot to dry out, and using the incorrect type of paper matrix for the test strips, can have detrimental effects on the results.

Figure 26.2 Haemoglobin colour scale. The stained test-strip being read on the right indicates a severe anaemia with a haemoglobin value of about 60 g/l (6 g/dl).

Packed Cell Volume

Although the packed cell volume (PCV) can be used as a simple screening test for anaemia and as a rough guide to the accuracy of haemoglobin measurements, it is not a substitute for a well-performed haemoglobin measurement. In addition to the technical problems (outlined on p. 28) there are particular problems with this method in resource-poor settings that may lead to errors in estimating the PCV. The lack of a mechanical mixing device means that specimens may not be adequately mixed before being introduced into the microhaematocrit tube. Poor-quality sealant means that the tubes often leak during centrifugation. Because the microhaematocrit tubes are difficult to label, samples may get mixed up in the centrifuge, especially when pressure of work is high and supervision is poor. Erratic power supplies, lack of devices for measuring g forces and poor equipment maintenance result in inadequate centrifugation with incomplete packing of the red cells.

Manual Cell Counts Using Counting Chambers

Visual counting of blood cells is an acceptable alternative to electronic counting for white blood cells but it is not recommended for routine red blood cell counts because the number of cells that can be counted within a reasonable time in the routine laboratory (e.g. about 400) will be too few to ensure a sufficiently precise result (see below). Although manual counting is recommended for platelet counts,¹⁹ in practice, it is often easier to detect gross reductions or increases in platelet counts from examination of the peripheral blood film.

Counting Chambers

The visibility of the rulings in the counting chamber ¹⁹ is as important as the accuracy of calibration, so that chambers with a ‘metallized’ surface and Neubauer or Improved Neubauer rulings are recommended. These have nine 1 × 1 mm ruled areas, which, when covered correctly with the special thick coverglass, each contain a volume of 0.1 μl diluted blood (Figs 26.3, 26.4). Coverslips designed for mounting of microscopy preparations must not be used with counting chambers. The sample is introduced between the chamber and the coverglass using a pipette or capillary tube and the preparation is viewed using ×40 objective and ×6 or ×10 eyepieces. With Neubauer and Improved Neubauer chambers, the cells in 4 or 8 horizontal rectangles of 1 × 0.05 mm (80 or 160 small squares) or in 5 or 10 groups of 16 small squares are counted, including the cells that touch the bottom and left-hand margins of the small squares.

Figure 26.3 Neubauer counting chamber. The total ruled area is 3 × 3 mm; the central ruled area is 1 × 1 mm. In the central area, 16 groups of 16 small squares are separated by triple rulings.

Figure 26.4 Improved Neubauer counting chamber. The central area consists of 25 groups of 16 small squares separated by closely ruled triple lines (which appear as thick black lines in the illustration).

Total White Blood Cell Count

To make the counting of white cells easier, diluted whole blood is mixed with a fluid to lyse the red cells and stain the white cell nuclei deep violet-black.

Diluent

The diluent is 2% (20 ml/l) acetic acid coloured pale violet with gentian violet.

Method

Make a 1 in 20 dilution of blood by adding 0.1 ml of well-mixed blood (lack of adequate mixing is a major source of error) to 1.9 ml of diluent * in a 75 × 10 mm plastic (or glass) tube. After sealing the tube with a lid or tightly fitting bung, mix the diluted blood in a mechanical mixer or by hand for at least 2 min by tilting the tube to an angle of about 120° combined with rotation, thus allowing the air bubble to mix the suspension. Fill a clean dry counting chamber, with its coverglass already in position, without delay. This is simply accomplished with the aid of a plastic Pasteur pipette or a length of stout capillary glass tubing that has been allowed to take up the suspension by capillarity. Take care that the counting chamber is filled in one action and that no fluid flows into the surrounding moat.

Leave the chamber undisturbed on a bench for at least 2 min for the cells to settle, but not much longer, because drying at the edges of the preparation initiates currents that cause movement of the cells after they have settled. The bench must be free of vibrations and the chamber must not be exposed to draughts or to direct sunlight or other sources of heat. It is important that the coverglass should be of a special thick glass and perfectly flat, so that when laid on the counting chamber, diffraction rings are seen. The coverglass should be of such a size that when placed correctly on the counting chamber, the central ruled areas lie in the centre of the rectangle to be filled with the cell suspension.

If any of the following filling defects occur, the preparation must be discarded and the filling procedure must be repeated using another clean dry chamber:

• Overflow into moat

• Chamber area incompletely filled

• Air bubbles anywhere in chamber area

• Any debris in chamber area.

To obtain a coefficient of variation of 5%, it is necessary to count about 400 cells (Table 26.1); in practice, it is reasonable to count 100 white cells. To minimize distribution errors, count the cells in the entire ruled area (i.e. 9 × 0.1 μl areas in an Improved Neubauer counting chamber). In practice, the 100 cell count can be achieved by counting the cells in the four larger corner squares.¹⁹

Table 26.1 Variance of haemocytometry count

Calculation

White blood cell count per litre (WBC/l)

Thus, if N cells are counted in 0.1 μl, then the WBC/l is as follows:

(e.g. if 115 cells are counted, the WBC is 115 × 200 × 10⁶/l = 23 × 10⁹/l).

Range of white blood cell count in health

See Chapter 2, Tables 2.1, 2.2 and 2.3.

Platelet Count

Automated full blood counters produce platelet counts with a precision that is much superior to that of manual platelet counts. Manual platelet counts may occasionally be needed for blood samples with a significant proportion of giant platelets. Manual platelet counts are best performed on ethylenediaminetetra-acetic acid (EDTA) anticoagulated blood that has been obtained by clean venipuncture. Skin-prick samples are associated with platelet clumping so platelet counts are significantly lower and less constant than those performed on venous blood. Manual platelet counts are performed by visual examination of diluted, lysed whole blood using a Neubauer or Improved Neubauer counting chamber as for total white cell counts.^20,²¹

Method

The diluent consists of 1% aqueous ammonium oxalate in which the red cells are lysed. This method is recommended in preference to using formal-citrate as diluent, which leaves the red cells intact and is more likely to give incorrect results when the platelet count is low.

Before diluting the blood sample, examine it carefully for the presence of blood clots. If these are present, a fresh specimen should be requested because clots will cause the platelet count to be artificially low. Make a 1 in 20 dilution of well-mixed blood in the diluent by adding 0.1 ml of blood to 1.9 ml of ammonium oxalate diluent (10 g/l). Not more than 500 ml of diluent should be made at a time, using scrupulously clean glassware and fresh glass-distilled or deionized water. If possible, the solution should be filtered through a micropore filter (0.22 μm) and kept at 4°C. For use, a small part of the stock is refiltered and dispensed in 1.9 ml volumes in 75 × 12 mm tubes.

Mix the suspension on a mechanical mixer for 10–15 min. Fill a Neubauer counting chamber with the suspension, using a stout glass capillary or Pasteur pipette. Place the counting chamber in a moist Petri dish and leave untouched for at least 20 min to give time for the platelets to settle.

Examine the preparation with the ×40 objective and ×6 or ×10 eyepieces. The platelets appear under ordinary illumination as small (but not minute), highly refractile particles if viewed with the condenser racked down; they are usually well separated and clumps are rare if the blood sample has been properly collected. To avoid introducing dirt particles into the chamber which might be mistaken for platelets, all equipment must be scrupulously clean. Platelets are more easily seen with the phase contrast microscope. A special, thin-bottomed (1 μm) counting chamber is best for optimal phase contrast effect. The number of platelets in one or more areas of 1 mm ² should be counted. The total number of platelets counted should always exceed 200 to ensure a coefficient of variation of 8–10%.

Calculation

Thus, if N is the number of platelets counted in an area of 1 mm² (0.1 μl in volume), the number of platelets per litre of blood is:

Peripheral Blood Morphology

Examination of the peripheral blood film is one of the most important and useful examinations in the peripheral laboratory but it requires a great deal of skill and experience as well as a good-quality microscope (see Chapter 5). In addition to providing information about quantitative changes in blood cells, careful analysis of the qualitative changes may help in elucidating the underlying reasons for clinical problems. These observations may identify the cause of anaemia or undiagnosed fever or the presence of a haemoglobinopathy (see Chapter 14). Problems that might occur when preparing blood films relate to the slides themselves and to the quality of fixing of the films and the quality of staining reagents. When glass slides are in short supply, laboratories sometimes find it necessary to wash and reuse slides (see p. 623). Traces of detergent can result in misleading appearances of the red cells (Fig. 26.5), as can residual stain and scratches on the slide. In humid conditions, particularly during the rainy season, water may be absorbed by the methanol used for fixing slides and this can cause gross artefactual changes in red cell appearance (see Fig. 4.2). To avoid this problem, stock bottles of methanol should be kept tightly closed after use; small amounts should be aliquoted into a bottle with a tightly fitting cap for daily use and replaced regularly from the stock bottle.

Figure 26.5 Red cell appearances on detergent washed slide.

Modified (One-Tube) Osmotic Fragility Test

This simple and inexpensive test for screening for β thalassaemia trait is useful when quantification of haemoglobin A₂ is not possible and standardized automated analysers are not available for accurate measurement of MCV and MCH.

A variety of concentrations of buffered saline have been used. A concentration of 0.36% in buffered saline (see p. 622 for preparation) is recommended to ensure a high sensitivity with an acceptable specificity.²³ Because the false-positive rate is around 10%, confirmation of a positive result requires referral of a sample to a laboratory able to quantitate haemoglobin A₂. The test can also be used to screen for α⁰ thalassaemia trait, with positive samples being referred to a reference centre for DNA analysis. About 50% of samples containing haemoglobin E also give a positive result; this is an advantage rather than a disadvantage because detection of haemoglobin E is important in predicting the possibility of thalassaemia major or intermedia in compound heterozygotes with β thalassaemia.

Haemoglobin E Screening Test

Ideally, the diagnosis of haemoglobin E heterozygosity or homozygosity should be by haemoglobin electrophoresis at alkaline pH or high-performance liquid chromatography with a second method being used to confirm the provisional identification (see Chapter 14). When these facilities are not available, a screening test using the blue dye 2,6-dichlorophenolindophenol (DCIP), can be used. Samples containing haemoglobin E become faintly turbid when incubated with DCIP.^24,²⁵

Tests for Paroxysmal Nocturnal Haemoglobinuria

When facilities permit, flow cytometry for glycosylphosphatidylinositol-anchored proteins is the test of choice to detect paroxysmal nocturnal haemoglobinuria (PNH) clone. If positive, a sample of urine should be examined for haemosiderin. The Ham test and sucrose lysis test (see Chapter 13) can be used as tests for PNH. They are less sensitive and less quantitative than flow cytometry, but nevertheless are clinically useful.

Reagents for Prothrombin Time (PT) and Activated Partial Thromboplastin Time (APTT)

Rabbit Brain Thromboplastin for Prothrombin Time

Freeze-dried rabbit brain thromboplastins for use in the PT are now widely available commercially, with a shelf-life of 2–5 years or longer. Usually, they are calibrated against the World Health Organization (WHO) International Reference Preparation of thromboplastin and are supplied with an International Sensitivity Index (ISI) and a table converting prothrombin times to International Normalized Ratios (INR).

If a commercial preparation is not available, it is possible to prepare a homemade substitute using rabbit brain that does not require freeze drying and that is relatively stable.

Acetone-dried brain powder

Strip the membrane off freshly collected rabbit brain, wash free from blood and place in about three times its volume of cold acetone. Macerate for 2–3 min and then filter through absorbent lint (BP or USP grade) on a Büchner funnel. Repeat the extraction seven times; after two extractions, increase the time of exposure to acetone to c 20 min for each subsequent extraction. The material should become ‘gritty’ by the fourth or fifth extraction. After the last extraction, spread the acetone-dried brain on a piece of paper and allow to dry in air for 30 min. Rub through a 1 mm mesh nylon sieve to produce a coarse powder. Dispense into a batch of screw-capped bottles and dry over phosphorus pentoxide in a vacuum desiccator. After drying, screw down the caps tightly and store at 4°C or −20°C. At −20°C, the material should be stable for at least 5 years. A total of 100 g of whole brain yields c 15 g of dried powder.

Preparation of liquid suspension

Dissolve 0.9 g of NaCl and 0.9 g of phenol in 100 ml of water. Suspend 3.6 g of the acetone-dried brain in 100 ml of the phenol–saline solution at 15–20°C and allow to stand at this temperature for 4–5 h, mixing at 30-min intervals. Transfer to a 4°C refrigerator for 24 h and occasionally mix it. Thereafter, leave undisturbed at 4°C for 3 h and then decant the supernatant carefully through fine muslin or similar material. The ISI should be not more than 1.4 (see p. 469) and the mean normal prothrombin time should be 12–13 s. Store the suspension at 4°C. At this temperature, it will be stable for at least 6 months; at room temperature it will be stable for at least 7 days. It must not be allowed to freeze because freezing results in flocculation of the smooth suspension with deterioration of thromboplastic activity.

Phospholipid Reagent for Activated Partial Thromboplastin Time

Acetone-dried rabbit brain is suitable for preparing an APTT reagent. Bovine brain may also be used.

Prepare acetone-dried brain powder as described above. Suspend 5 g of the powder in 20 ml of chloroform (analytic grade) in a covered beaker for 1–2 h. Filter through filter paper to obtain a clear filtrate. Wash the brain deposit on the filter paper with 20 ml of chloroform and pool the clear filtrate with the previous filtrate. Evaporate the filtrate to dryness in a beaker of known weight in a waterbath at 60–70°C and weigh the residual deposit: 5 g of dried brain should yield c 1.5 g of phospholipid deposit. Emulsify in saline to give a 5% emulsion; 1.5 g of deposit should provide 30 ml of emulsion. Distribute the emulsion in small volumes in stoppered tubes. At −20°C it should be stable for at least 1 year.

For use, dilute 1 in 100 in saline and mix with an equal volume of 2.5 mg/ml kaolin suspension in imidazole buffer.

Laboratory support for management of hiv/aids: cd4-positive t-cell counts

The WHO recommends that district hospitals and higher level facilities should be able to provide a CD4+ cell count to aid identification of patients who would benefit from antiretroviral drugs and to monitor their progress. Several methods are available but their effectiveness in developing country settings is very variable.²⁶ Some of the low-cost methods that have good correlation with flow cytometric enumeration of CD4+ cells include the FACS count system (Becton Dickinson Sciences, CA), the Guava Easy CD4 assay (Guava technologies, Hayward, CA), Cyflow (Partec, Germany) and the panleucogating (PLG) CD4 technique. Manual methods such as Dynabeads (Dynal) and Cytospheres (Coulter) can also be used to measure CD4+ T-cell count but the cost, time and skill required make them unsuitable in many settings when other options are available.²⁷ Dried spots of serum or blood can be used to assess HIV-1 viral load and for resistance genotyping.²⁸

Laboratory management

Interlaboratory Communication

A well-planned infrastructure is necessary to facilitate flow of communication (e.g. about specimens, results, patient management) between remote clinics and referral or central laboratories. Cell telephones and e-mail using mobile telephone networks are now widely available and are being used to transmit health-related information. In more remote stations, voice and e-mail messages can be transmitted using high-frequency (HF) or satellite radio systems. Alternative power sources such as generators, solar energy and batteries are used to support communication facilities in remote rural clinics.

Digital wireless data communications systems such as the Global System of Mobile (GSM) network are being increasingly used for routine laboratory reporting. The data volume required for text-based laboratory reports is extremely modest and their transmission is low cost. Result reporting can either be incorporated into the mainstream laboratory information system or by individual handsets for smaller independent laboratories.

Specimen Transport

The need to ensure appropriate conditions for keeping specimens after they have been collected and other aspects of specimen transport to the laboratory are described in Chapter 1. The special problem of transporting specimens from remote clinics to laboratories and reference centres in low-resource countries requires further consideration. Peripheral laboratories can assist in rural development initiatives by creating employment opportunities for bicycle riders, motorcyclists, taxi operators and formal and informal courier companies. Commercial or private air services can be used to deliver samples to referral centres. The location of clinics is helped if Global Positioning System (GPS) coordinates are available for each clinic.²⁹

Staff Training

In poorer countries, there is often no system for regular supervision of individual laboratory staff and many staff do not receive continuing professional development. Monitoring standards of practice should continue for the whole professional life of laboratory staff to ensure high-quality results. For under-resourced countries, training may be provided in association with vertical health programmes such as HIV or tuberculosis control, but the need for training in basic haematological techniques is often overlooked because tests such as haemoglobin estimation and white cell counts are usually not linked to specific diseases and are therefore not incorporated into vertical programmes. However, because anaemia is the most prevalent disorder worldwide and is often the first sign of underlying disease, the importance of reliable haemoglobin measurements cannot be overemphasized. The need for integrated training programmes is becoming increasingly recognized, especially for peripheral laboratories. Continuing education needs to include the whole range of tests offered by the laboratory and not only tests that are used to support the diagnosis of specific diseases.

Individuals need to keep their own training records, perhaps in the form of a log book and to have their training achievements and plans regularly reviewed by their line managers. Central records of all training should also be maintained for monitoring purposes and to ensure equitable and appropriate distribution of training between different levels of staff. One of the most effective forms of continuing staff development is through regular on-site visits by trained supervisors, who stay for a sufficient length of time to be able to work with the resident staff, and are able to check equipment, supplies, records and attend to other administrative issues. Supervisors themselves need to be trained and monitored to ensure their own proficiency.

Regular monitoring of the quality of results from individual laboratories enables specific problems to be identified. Integrated External Quality Assurance (EQA) programmes tailored for peripheral laboratories are an excellent way to monitor laboratory performance across a wide range of tests. In addition, this quality monitoring allows discrete training needs to be identified, enabling limited teaching resources to be specifically targeted. Systems need to be established to report issues such as equipment failures, discontinuity of supplies and communication breakdowns to the local or regional management teams.

Access to up-to-date information is important for adapting and enhancing laboratory performance. Many medical and technical journals now publish their articles in full on the internet and by an arrangement between the main international publishers and WHO, they can be accessed free of charge or at a significantly discounted cost in most under-resourced countries (see www.who.int/hinari/en).

Clinical Staff Interaction

A major delay in test turnaround time (see p. 574) and the failure of clinicians to use test results is often due to the slow delivery of reports within the health facility after the test has been performed. Appropriate clinical use of the laboratory has a direct impact on the cost-effectiveness of the service ¹². Laboratory tests may be initiated by nurses, health field workers and public health officers, as well as doctors. Many of these individuals have little or no training in how to request appropriate tests, how to provide timely and suitable samples and how to use the results for maximal benefit to patients. Training for laboratory users needs to be incorporated into laboratory training programmes and be closely monitored. In poorer countries, there is often a dearth of clinicians with both clinical and laboratory experience who are qualified to provide such training. However, the relationship between the laboratory and clinical staff can be enhanced and use of the laboratory can be optimized by using a clinician/scientist team to provide joint training in both disciplines. Various publications are now available to guide laboratory users on which test to select, how to interpret the results and how to select and collect the correct samples.

Health Management Teams

Local health management teams are often responsible for ensuring that their laboratories are provided with the necessary tools to deliver a high-quality service. As a rule, these teams do not include members of the laboratory profession, who are often represented by other allied professionals such as pharmacists. Staff in under-resourced laboratories are therefore not always responsible for purchasing supplies and equipment for the laboratory, and this can result in the purchase of inappropriate or poor-quality equipment and reagents. In addition to encouraging adequate representation of laboratory professionals at management level, it is important to ensure that non-laboratory personnel responsible for making procurement decisions for the laboratory are aware of the needs of their laboratory service and seek advice from laboratory professionals.

Health and Safety

Details of laboratory safety issues are described in Chapter 24. Awareness of these issues should be constantly promoted within all laboratories,³⁰ and the working environment needs to be made as safe as possible. A ‘Code of Safe Laboratory Practice’ should be prepared that is affordable and relevant to the local circumstances. It should include the following:

• Risk assessment to identify potential workplace hazards and the risk they pose to individuals working or visiting the laboratory

• Education on safe working practices

• Provision of appropriate and adequate safety items such as white coats, gloves and pipetting devices

• Monitoring of adherence to health and safety regulations

• Prompt reporting and investigation of laboratory accidents

• Addressing medical waste disposal practices including use of sharps containers and waste bags and disposal of items in incinerators or deep pits.

Disposable syringes (see p. 2) are intended for single-use only. They cannot withstand sterilization and should never be reused. Similarly, disposable lancets for skin puncture must never be reused. The practice of using a single lancet on several patients consecutively and cleaning it with alcohol between use is unacceptable.

References

1 WHO. Guidelines for the Treatment of Malaria, 2010. Available at: www.who.int/malaria/publications/atoz/9241546948/en/index.html Accessed 25 01

2 Akpek G., Lee S.M., Gagnon D.R., et al. Bone marrow aspiration, biopsy and culture in the evaluation of HIV-infected patients for invasive mycobacteria and histoplasma infections. Am J Hematol. 2001;67:100-106.

3 Larsen C. The fragile environment of inexpensive CD4+ T-cell enumeration in the least developed countries: strategies for accessible support. Clinical Cytometry Society. 2008;74B(Suppl 1):S1007-S1116.

4 WHO. Consultation on Laboratory Services for Monitoring HIV Anti-retroviral Therapy in Resource-limited Settings. Available at: www.who.int/diagnostics_laboratory/events_hivlab/en/, 2004. Accessed 25 01 2010