Principle

The method to be described is based on that of Parpart et al . Small volumes of blood are mixed with a large excess of buffered saline solutions of varying concentration. The fraction of red cells lysed at each saline concentration is determined colorimetrically. The test is normally carried out at room temperature (15–25 °C).

Reagents

Prepare a stock solution of buffered sodium chloride, osmotically equivalent to 100 g/l (1.71 mol/l) NaCl, as follows: dissolve NaCl, 90 g; Na ₂ HPO ₄ , 13.65 g (or Na ₂ HPO ₄ 2H ₂ O, 17.115 g); and NaH ₂ PO ₄ .2H ₂ O, 2.34 g in water. Adjust the final volume to 1 litre. This solution will keep for months at 4 °C in a well-stoppered bottle. Salt crystals may form on storage and must be thoroughly redissolved before use.

In preparing hypotonic solutions for use, it is convenient to make first a 10 g/l solution from the 100 g/l NaCl stock solution by dilution with water. Dilutions equivalent to 9.0, 7.5, 6.5, 6.0, 5.5, 5.0, 4.0, 3.5, 3.0, 2.0 and 1.0 g/l are convenient concentrations. Intermediate concentrations such as 4.75 and 5.25 g/l are useful in critical work and an additional 12.0 g/l dilution should be used for incubated samples.

It is convenient to make up 50 ml of each dilution. The solutions keep well at 4 °C if sterile, but should be inspected for moulds before use and discarded if moulds develop.

Method

Heparinised venous blood or defibrinated blood may be used; oxalated or citrated blood is not suitable because of the additional salts added to it. The test should be carried out within 2 h of collection with blood stored at room temperature or within 6 h if the blood has been kept at 4 °C.

• 1.
  

  Deliver 5.0 ml of each of the 11 saline solutions into 12 × 75 mm test tubes. Add 5.0 ml of water to the 12th tube.

  
  
• 2.
  

  Add to each tube 50 μl of well-mixed blood and mix immediately by inverting the tubes several times, avoiding foam.

  
  
• 3.
  

  Leave the suspensions for 30 min at room temperature. Mix again and then centrifuge for 5 min at 1200 g .

  
  
• 4.
  

  Remove the supernatants and estimate the amount of lysis in each using a spectrometer at a wavelength setting of 540 nm or a photoelectric colorimeter provided with a yellow–green (e.g. Ilford 625) filter. Use as a blank the supernatant from tube 1 (osmotically equivalent to 9 g/l NaCl).

  
  
• 5.
  

  Assign a value of 100% lysis to the reading with the supernatant of tube 12 (water) and express the readings from the other tubes as a percentage of the value of tube 12. Plot the results against the NaCl concentration ( Fig. 12-1 ).

  
  

  
  

  
  

  

  
  

  
  

  Osmotic fragility curves.

  
  

  Osmotic fragility curves of patients suffering from the following: sickle cell anaemia, β thalassaemia major, hereditary spherocytosis and ‘idiopathic’ warm autoimmune haemolytic anaemia. The normal range is indicated by the unbroken lines.

  
  

Method

Defibrinated blood should be used, care being taken to ensure that sterility is maintained.

Incubate 1 ml or 2 ml volumes of blood in sterile 5 ml bottles. It is advisable to set up the samples in duplicate in case one has become infected, as indicated by gross lysis and change in colour.

After 24 h, if no infection is evident, pool the contents of the duplicate bottles after thoroughly mixing the sedimented red cells in the overlying serum and estimate the fragility as previously described.

Because the fragility may be markedly increased ( Fig. 12-2 ), set up additional hypotonic solutions containing 7.0 g/l and 8.0 g/l NaCl. In addition, use a solution equivalent to 12.0 g/l NaCl because sometimes, as in HS, lysis may take place in 9.0 g/l NaCl. In this case, use the supernatant of the tube containing 12.0 g/l NaCl as the blank in the colorimetric estimation.

Osmotic fragility curves before and after incubating blood at 37 °C for 24 h.

Results are from patients suffering from the following: hereditary spherocytosis, pyruvate kinase deficiency and hereditary nonspherocytic haemolytic anaemia of undiagnosed type. The normal range is indicated by the unbroken lines.

The incubation fragility test is conveniently combined with the estimation of the amount of spontaneous autohaemolysis (p. 235).

Factors affecting osmotic fragility tests

In carrying out osmotic fragility tests by any method, three variables capable of markedly affecting the results must be controlled, quite apart from the accuracy with which the saline solutions have been made up. These are as follows:

• 1.
  

  The relative volumes of blood and saline

  
  
• 2.
  

  The final pH of the blood in saline suspension

  
  
• 3.
  

  The temperature at which the tests are carried out.

A proportion of 1 volume of blood to 100 volumes of saline is chosen because the concentration of blood is so small that the effect of the plasma on the final tonicity of the suspension is negligible. When weak suspensions of blood in saline are used, it is necessary to control the pH of the hypotonic solutions and it is for this reason that phosphate buffer is added to the saline. Even so, small differences will be found between the fragility of venous blood and maximally aerated (i.e. oxygenated) blood. For the most accurate results, it is recommended that the blood is mixed until bright red. Finally, it is ideal for tests to be carried out always at the same temperature, although for most purposes room temperature is sufficiently constant.

The extent of the effect of pH and temperature on osmotic fragility was well illustrated in the paper of Parpart et al . The effect of pH is more important: a shift of 0.1 of a pH unit is equivalent to altering the saline concentration by 0.1 g/l, the fragility of the red cells being increased by a decrease in pH. An increase in temperature decreases the fragility, an increase of 5 °C being equivalent to an increase in saline concentration of about 0.1 g/l.

Lysis is virtually complete at the end of 30 min at 20 °C and the hypotonic solutions may be centrifuged at the end of this time.

Further details of the factors that affect and control haemolysis of red cells in hypotonic solutions were given by Murphy.

Recording the results of osmotic fragility tests

In the past, osmotic fragility most often has been expressed in terms of the highest concentration of saline at which lysis is just detectable (initial lysis or minimum resistance) and the highest concentration of saline in which lysis appears to be complete (complete lysis or maximum resistance). It is, however, useful also to record the concentration of saline causing 50% lysis (i.e. the median corpuscular fragility, MCF) and to inspect the entire fragility curve ( Fig. 12-1 ). The findings in health are summarised in Table 12-1 .

Interpretation of results

The osmotic fragility of freshly taken red cells reflects their ability to take up a certain amount of water before lysing. This is determined by their volume-to-surface area ratio. The ability of the normal red cell to withstand hypotonicity results from its biconcave shape, which allows the cell to increase its volume by about 70% before the surface membrane is stretched; once this limit is reached lysis occurs. Spherocytes have an increased volume-to-surface area ratio; their ability to take in water before stretching the surface membrane is thus more limited than normal, and they are therefore particularly susceptible to osmotic lysis. The increase in osmotic fragility is a property of the spheroidal shape of the cell and is independent of the cause of the spherocytosis. Characteristically, osmotic fragility curves from patients with HS who have not been splenectomised show a ‘tail’ of very fragile cells ( Fig. 12-3 ). When plotted on probability paper, the graph indicates two populations of cells: the very fragile and the normal or slightly fragile. After splenectomy the red cells are more homogeneous, the osmotic fragility curve indicating a more continuous spectrum of cells, from fragile to normal.

Osmotic fragility curves of three patients suffering from hereditary spherocytosis belonging to the same family (brother, sister and uncle).

The area between the unbroken lines represents the normal range.

Decreased osmotic fragility indicates the presence of unusually flattened red cells (leptocytes) in which the volume-to-surface area ratio is decreased. Such a change occurs in iron deficiency anaemia and thalassaemia in which the red cells with a low mean cell haemoglobin (MCH) and mean cell volume (MCV) are unusually resistant to osmotic lysis ( Fig. 12-1 ). A simple one-tube osmotic fragility is a useful screening test for β thalassaemia and some haemoglobinopathies in countries with a high incidence of these abnormalities (p. 556). Reticulocytes and red cells from patients who have been splenectomised also tend to have a greater amount of membrane compared with normal cells and are osmotically resistant. In liver disease, target cells may be produced by passive accumulation of lipid, and these cells, too, are resistant to osmotic lysis.

The osmotic fragility of red cells after incubation for 24 h at 37 °C is also a reflection of their volume-to-surface area ratio, but the factors that alter this ratio are more complicated than in fresh red cells. The increased osmotic fragility of normal red cells, which occurs after incubation ( Fig. 12-2 ), is mainly caused by swelling of the cells associated with an accumulation of sodium that exceeds loss of potassium. Such cation exchange is determined by the membrane properties of the red cell, which control the passive flux of ions, and the metabolic competence of the cell, which determines the active pumping of cations against concentration gradients. During incubation for 24 h, the metabolism of the red cell becomes stressed and the pumping mechanisms tend to fail, one factor being a relative lack of glucose in the medium.

The osmotic fragility of red cells that have an abnormal membrane, such as those of HS and hereditary elliptocytosis (HE), increases abnormally after incubation ( Fig. 12-2 ). Similar results occur in hereditary stomatocytosis. The results with red cells with a glycolytic deficiency, such as those of pyruvate kinase (PK) deficiency, are variable. In severe deficiencies, osmotic fragility may increase substantially ( Fig. 12-2 ), but in other cases, the fragility may decrease owing to a greater loss of potassium than gain of sodium. In thalassaemia major and minor, osmotic fragility is frequently markedly reduced after incubation, again owing to a marked loss of potassium. A similar, although usually less marked, change is seen in iron deficiency anaemia.

To summarise, measurement of red cell osmotic fragility provides a useful indication as to whether a patient’s red cells are normal because an abnormal result invariably indicates abnormality. The reverse is, however, not true (i.e. a result that is within the normal range does not mean that the red cells are normal). The findings in some important haemolytic anaemias are summarised in Table 12-2 .

Principle

The osmotic fragility test lacks specificity and sensitivity and may fail to detect atypical or mild HS. Moreover, it can be affected by factors unrelated to red cell cytoskeletal defects; for example, positive results may be obtained for red cells from patients who are pregnant or who have immune or other haemolytic anaemias or renal failure. The flow cytometric (dye-binding) test of King and colleagues measures the fluorescence intensity of intact red cells labelled with eosin-5-maleimide (EMA), which reacts covalently with Lys-430 on the first extracellular loop of Band 3 protein. The N-terminal cytoplasmic domain of Band 3 interacts with ankyrin and protein 4.2, which in turn crosslink with the spectrin-based cytoskeleton and stabilises the membrane lipid bilayer. Deficiency or abnormality of Band 3 may result in decreased fluorescence. This is seen in HS red cells but has also been observed in cases of hereditary pyropoikilocytosis (HPP), Southeast Asian ovalocytosis, congenital dyserythropoietic anaemia Type II and the stomatocytic variant, cryohydrocytosis. Blood samples in ethylenediaminetetra-acetic acid (EDTA) can be analysed for up to 48 h after collection provided they have been stored in the refrigerator.

Reagents

Method

Thaw a tube of stock EMA solution in the dark at room temperature and dilute with an equal volume of PBS to obtain a working solution of 0.5 mg/ml. Mix 5 μl of washed packed red cells with 25 μl of EMA working solution in a plastic microfuge tube. Set up control tubes from blood of normal individuals, and perform all tests in duplicate. Leave a rack with the tubes in a cupboard in the dark at room temperature for 30 min. Mix and return to the cupboard for a further 30 min.

Then, spin the tubes in a bench-top microfuge for 5–10 s and remove the supernatant dye carefully with a fine-tip pipette.

Wash the labelled red cells three times with 500 μl PBS containing 0.5% BSA. The third wash should be colourless. If it is still pink, suggesting that traces of dye particle remain in the tube, discard the sample and repeat the cell-labelling procedure.

Resuspend the packed red cells in 500 μl of the PBS–BSA wash solution. Transfer 100 μl of the cell suspension into a plastic flow cytometer tube and add 1.4 ml of wash solution. Keep the cell suspensions in the dark by wrapping in aluminium foil until use. Set the analyser thresholds (gate) for red blood cells and count each sample for a minimum of 15 000 events. Select the FITC (fluorescein isothiocyanate) channel and record the mean fluorescence intensity (MFI). Compare the test with the mean value of several control samples analysed at the same time.

Interpretation of results

Results are expressed as a ratio of mean value of the test to control samples. Each laboratory should set the reference range and cut-off values for its own instrument. In our laboratory, a range of 0.83 to 1.17 is considered normal. HS red cells give a ratio of less than 0.8. Values as low as 0.6 may be seen in classic HS. Similar or even lower values are seen in the much rarer HPP, but the latter is readily distinguished from HS on the basis of red cell morphology and a very low MCV. A reduced ratio may also be encountered in cases of Southeast Asian ovalocytosis, congenital dyserythropoietic anaemia Type II and cryohydrocytosis, but these are associated with different types of red cell atypia and may generally be ruled out by examination of a stained blood film. EMA binding has a high sensitivity in the diagnosis of HS, however in a very small number of cases of typical HS the MFI ratio is normal. If HS is suspected and EMA binding normal, a second screening test may be informative.

Principle

Glycerol present in a hypotonic buffered saline solution slows the rate of entry of water molecules into the red cells so that the time taken for lysis may be more conveniently measured. Like the osmotic fragility test, differentiation can be made between spherocytes and normal red cells.

Reagents

Method

Add 20 μl of whole blood, anticoagulated with EDTA, to 5.0 ml of PBS, pH 6.85. Mix the suspension carefully.

Transfer 1.0 ml to a standard 4 ml cuvette of a spectrometer equipped with a linear-logarithmic recorder. Fix the wavelength at 625 nm and start the recorder. Add 2.0 ml of the glycerol reagent rapidly to the cuvette with a 2 ml syringe or automatic pipette and mix well.

The rate of haemolysis is measured by the rate of fall of turbidity of the reaction mixture. The results are expressed as the time required for the optical density to fall to half the initial value (AGLT ₅₀ ). The test can also be carried out using a colorimeter and stopwatch.

Results

Normal blood takes more than 30 min (1800 s) to reach the AGLT ₅₀ . The time taken is similar for blood from normal adults, newborn infants and cord samples. In patients with HS, the range of the AGLT ₅₀ is 25–150 s. A short AGLT ₅₀ may also be found in chronic renal failure, chronic leukaemias and autoimmune haemolytic anaemia; it also may be found in some pregnant women.

Significance of the acidified glycerol lysis-time test

The same principles apply as with the osmotic fragility test. Cells with a high volume-to-surface area ratio resist swelling for a shorter time than normal cells. This applies to all spherocytes, whether the spherocytosis is caused by HS or other mechanisms. The test is particularly useful in screening family members of patients with HS where morphological changes are too minor to indicate clearly whether the disorder is present.

Principle

Whereas osmotic fragility may be abnormal in any condition where spherocytes occur, it has been suggested that cryohaemolysis is specific for HS. This appears to result from the fact that the latter is dependent on factors that are related to molecular defects of the red cell membrane rather than to changes in the surface area-to-volume ratio. The test can be carried out on EDTA blood up to 1 day old.

Reagent

Method

• 1.
  

  Centrifuge the blood and wash the red cells three times with cold (4 °C) 9 g/l NaCl. Make a suspension of 50–70% cells in the saline and keep on ice until tested.

  
  
• 2.
  

  Prepare 2 ml volumes of reagent, thawing if frozen, and stand for 10 min in a 37 °C water bath to equilibrate.

  
  
• 3.
  

  Pipette 50 μl of the cell suspension into each of 2 tubes of the warmed reagent, vortex immediately for a few seconds and then incubate for exactly 10 min at 37 °C.

  
  
• 4.
  

  Without delay, transfer the tubes to an ice bath for another 10 min, vortex for a few seconds and then centrifuge to sediment the remaining cells. Transfer some of the supernatant to a clean tube.

  
  
• 5.
  

  Prepare a 100% haemolysate solution by pipetting 50 μl of the original sample into 2 ml of water. Centrifuge and dilute 200 μl of the supernatant in 4 ml of water.

  
  
• 6.
  

  Read absorbance at 540 nm of the test and the 100% lysis samples.

Interpretation

Streichman et al . reported the range of cryohaemolysis in normal subjects to be 3–15%, whereas in HS there was > 20% lysis. However, it is recommended that individual laboratories establish their own reference values for the method. We have found that most normal samples give < 3% lysis. Increased lysis is not exclusive to HS and may be observed in hereditary stomatocytosis.

Principle

Aliquots of blood are incubated both with and without sterile glucose solution at 37 °C for 48 h. After this period, the amount of spontaneous haemolysis is measured colorimetrically.

Method

It is essential to use aseptic techniques in setting up the autohaemolysis test to maintain sterility throughout the incubation period.

Use blood collected into ACD or defibrinated blood (see previous editions). Deliver four 1 ml or 2 ml samples into sterile 5 ml capped bottles. Retain a portion of the original sample; separate and store this as the preincubation serum.

Add to two of the bottles 50 or 100 μl of sterile 100 g/l glucose solution, so as to provide a concentration of glucose in the blood of at least 30 mmol/l. Make sure that the caps of the bottles are tightly closed and place the series of bottles in the incubator at 37 °C. A sample from a known normal individual should be run in parallel as a control.

After 24 h, thoroughly mix the content by gentle swirling. After incubating for a further 24 h, inspect the samples for signs of infection, thoroughly mix again, then from each bottle remove a sample for the estimation of the packed cell volume (PCV) (by the microhaematocrit method) and haemoglobin concentration (Hb) and centrifuge the remainder to obtain the supernatant serum.

Estimate the spontaneous lysis by means of a colorimeter or a spectrometer at 540 nm.

As a rule, it is convenient to make a 1 in 10 dilution of the incubated serum in cyanide-ferricyanide (Drabkin) solution (p. 20), unless there is marked haemolysis, when a 1 in 25 or 1 in 50 dilution is more suitable. A corresponding dilution of the preincubation serum is used as a blank and a 1 in 100 or 1 in 200 dilution of the whole blood in Drabkin solution indicates the total amount of Hb present and serves as a standard.

Calculate the percentage lysis, allowing for the change in PCV resulting from the incubation as follows:

The reading at time t is multiplied by (1 − PCV _t ) so as to give the concentration that would be found if the liberated haemoglobin were dissolved in whole blood (i.e. in both plasma and red cell compartments), not in the plasma compartment alone.

Normal range of autohaemolysis

Lysis at 48 h: without added glucose, 0.2% to 2.0%; with added glucose, 0% to 0.9%.

The results obtained are sensitive to slight differences in technique and each laboratory should use a carefully standardised procedure and establish its own normal range. If the amount of liberated haemoglobin is small, it is more accurate (although more time consuming) to measure lysis by a chemical method rather than by a direct spectrometric method (p. 216). It can also be measured directly by a simple and rapid procedure with a HemoCue Plasma/Low Hb system ( www.hemocue.com ).

Significance of increased autohaemolysis

Little or no lysis takes place when normal blood is incubated for 24 h under sterile conditions and the amount present after 48 h is small. If glucose is added so that it is present throughout the incubation, the development of lysis is markedly slowed. The amount of autohaemolysis that occurs after 48 h with and without glucose is determined by the properties of the membrane and the metabolic competence of the red cell. In membrane disorders such as HS, the rate of glucose consumption is increased to compensate for an increased cation leak through the membrane. During the 48 h incubation, glucose is therefore used up relatively rapidly so that energy production fails more quickly than normal unless glucose is added. This is one factor that contributes to the increased rate of autohaemolysis in HS. Usually, but not always, the addition of glucose to the blood decreases the rate of autohaemolysis in HS. This was referred to as Type 1 autohaemolysis. When the utilisation of glucose via the glycolytic pathway is impaired, as in PK deficiency, the rate of autohaemolysis at 48 h is usually increased but glucose fails to correct or may even aggravate lysis (Type 2 autohaemolysis). Although a similar result may be seen in severe HS (Type B), in the absence of spherocytosis, failure of glucose to diminish autohaemolysis is a strong indication of a glycolytic block. Blood from patients with G6PD deficiency or other disorders of the pentose phosphate pathway may undergo a slight increase in autohaemolysis (without additional glucose), which is corrected by the addition of glucose. Commonly, the result is normal, but examination of the incubated blood may show an increase in methaemoglobin (Hi) (discussed later). Not all glycolytic enzyme deficiencies give a Type 2 reaction so that a Type 1 result does not exclude the possibility of such a defect.

In the acquired haemolytic anaemias, the results of the autohaemolysis test are variable and generally not very helpful in diagnosis. In the autoimmune haemolytic anaemias, lysis may be increased in the absence of additional glucose but the effect of added glucose is unpredictable. In paroxysmal nocturnal haemoglobinuria (PNH), the autohaemolysis of aerated defibrinated blood is usually normal.

Autohaemolysis may be increased in haemolytic anaemias caused by oxidant drugs or when there are defects in the reducing power of the red cell. Heinz bodies, Hi or both will be detectable at the end of incubation. Normally, red cells produce < 4% Hi after 48 h incubation and Heinz bodies are not seen. Red cells containing an unstable haemoglobin also contain Heinz bodies at the end of the incubation period and increased amounts of Hi.

The nucleosides adenosine, guanosine and inosine, like glucose, diminish the rate of autohaemolysis when added to blood. Remarkably, adenosine triphosphate (ATP) strikingly retards haemolysis in PK deficiency, although glucose itself is ineffective. ATP does not pass the red cell membrane.

The autohaemolysis test lacks specificity. This has drawn much criticism on the test, including the suggestion that it has no place in the screening of blood for inherited defects. The best way to detect metabolic defects in red cells is undoubtedly to measure glucose consumption, lactate production and the contribution to metabolism of the pentose phosphate pathway. These measurements are, unfortunately, difficult and are likely to be undertaken only by specialised laboratories. The autohaemolysis test does provide some information about the metabolic competence of the red cells and helps to distinguish membrane defects from enzyme defects.

In summary, we feel that the autohaemolysis test is still useful in the investigation of patients who have or who may have chronic haemolytic anaemia for the following reasons:

• 1.
  

  If the result is entirely normal, an intrinsic red cell abnormality is unlikely.

  
  
• 2.
  

  If abnormal haemolysis is fully corrected by glucose, a metabolic abnormality is unlikely and a membrane abnormality is likely.

  
  
• 3.
  

  If abnormal haemolysis shows little or no correction by glucose, a metabolic abnormality is likely, provided that obvious features of spherocytosis are not present on the blood film.

Thus, in our experience, a combination of red cell morphology with the results of the autohaemolysis tests makes it possible to differentiate membrane abnormalities from enzyme deficiencies in the vast majority of cases.

Principle

NADPH, generated by G6PD present in a lysate of blood cells, fluoresces under long-wave ultraviolet (UV) light. In G6PD deficiency, there is an inability to produce sufficient NADPH; this results in a lack of fluorescence.

Reagents

Method

Thaw out sufficient tubes to set up test and controls. Allow reagents to reach room temperature before use.

Add 10 μl of whole blood (in EDTA, heparin, ACD [acid–citrate–dextrose] or CPD [citrate–phosphate– dextrose]) to 100 μl volumes of the reagent mixture and keep at room temperature (15–25 °C).

Place 10 μl of the reaction mixture on a Whatman No. 1 filter paper at the beginning of the reaction and again after 5–10 min. A shorter interval may be appropriate at a high ambient temperature ( c . 25–30 °C). Allow to air dry thoroughly before examining the spots under UV light. Record whether fluorescence is present (+) or absent (−). Always set up samples of normal blood and known G6PD-deficient blood in parallel.

If the samples are to be collected away from the laboratory or where delay is envisaged (e.g. during population screening) place about 10 μl of blood on Whatman No. 1 filter paper and allow it to dry. Cut out the disc of dried blood in the laboratory and add it to the reaction mixture. A sample of normal blood should be tested as a positive fluorescence (i.e. normal) control.

The test can be carried out on blood stored in ACD (provided it is sterile) for up to 21 days at 4 °C and for about 5 days at room temperature.

Interpretation

Fluorescence is produced by NADPH formed from NADP ⁺ in the presence of G6PD. Some of the NADPH produced is oxidised by GSSG, but this reaction, catalysed by glutathione reductase, is normally slower than the rate of NADPH production. Red cells with < 20% of normal G6PD activity do not cause detectable fluorescence.

Like all screening tests, this method is useful when large numbers of samples are to be tested, but the result must be interpreted with caution in an individual patient. The main causes of erroneous interpretations are as follows:

False-normal . If there is reticulocytosis, a vivid fluorescence may be seen with a genetically G6PD-deficient blood sample because young red cells have more G6PD activity. If the test is carried out during an acute haemolytic episode, the patient’s blood should be retested when the reticulocyte count has returned to normal.

False-deficient . If the patient is anaemic, very little fluorescence may be seen despite the G6PD being genetically normal, simply because there are relatively few red cells in the 10 μl of blood used.

Although it is possible to correct for either or both of these contingencies, if in doubt, it is best to proceed directly to a quantitative enzyme assay (discussed later).

The test is meant to give only a + or − (normal or deficient) result by comparison with the controls, and it does not make sense to grade by eye the intensity of fluorescence. If a control G6PD-deficient sample is not available, the appearance of the ‘zero time’ spot can be used for reference. The threshold for a ‘deficient’ result can be worked out by making dilutions of a normal blood sample in saline and is best set by regarding as deficient the fluorescence obtained when G6PD activity is 20% of normal or less (corresponding to a 1 in 5 dilution of normal blood). This means that very mildly deficient variants and a substantial proportion of heterozygotes (see below), will be missed. However, clinically important haemolysis is unlikely to occur in subjects who have more than 20% G6PD activity and therefore this seems an appropriate (although arbitrary) threshold for a diagnostic laboratory. Because the test depends on visual inspection, it is best to select the time of incubation in relation to ambient temperature in preliminary trials. NADPH production is a cumulative process. Therefore, given enough time, a G6PD-deficient sample will fluoresce. The time allowed for the reaction should be one at which the contrast in fluorescence between a G6PD-normal and a G6PD-deficient sample is maximal.

Principle

Sodium nitrite converts haemoglobin to Hi. When no methylene blue is added, methaemoglobin persists, but incubation of the samples with methylene blue allows stimulation of the pentose phosphate pathway in subjects with normal G6PD levels. The Hi is reduced during the incubation period. In G6PD-deficient subjects, the block in the pentose phosphate pathway prevents this reduction.

Reagents

Method

Use anticoagulated blood (EDTA or ACD) and test the samples preferably within 1 h of collection if left on the bench or within 6 h if kept at 4 °C. Blood in ACD can be stored for up to 1 week but will be unsatisfactory if there is any haemolysis. With blood from patients who are severely anaemic, adjust the PCV to 0.40 ± 0.05.

Add 2 ml of blood to the tube containing 0.2 ml of the combined reagent either freshly prepared or dried. Close the tube with a stopper and gently mix the contents by inverting it 15 times.

Prepare control tubes by adding 2 ml of blood to a similar tube without reagents (normal reference tube) and to a tube containing 0.1 ml of sodium nitrite– dextrose mixture without methylene blue (‘deficient’ reference tube).

Incubate the samples at 37 °C for 90 min. If the blood has been heparinised, incubation should be continued for 3 h.

After the incubation, pipette 0.1 ml volumes from the test sample, the normal reference tube and the deficient reference tube into 10 ml of water in separate, clear glass test tubes of identical diameter. Mix the contents gently. Compare the colours in the different tubes (see below).

Interpretation

Normal blood yields a colour similar to that in the normal reference tube (i.e. a clear red). Blood from deficient subjects gives a brown colour similar to that in the deficient reference tube. Heterozygotes give intermediate reactions.

Although this method takes longer than the fluorescent test, its advantages include the fact that it is extremely inexpensive and that the only equipment required is a water bath. In addition, the test can be complemented by cytochemical analysis that lends itself to detecting G6PD deficiency in patients with reticulocytosis and in heterozygotes.

Test kits

Several commercial kits are available for detection of G6PD deficiency. A fluorescent spot test (Trinity Biotech 203-A) and a test based on reduction of the dye dichloroindophenol to a colourless state in the presence of phenazine methosulphate (Trinity Biotech 400) are available commercially ( www.trinitybiotech.com ).

The Quantase kit ( www.bio-rad.com ) is a photometric method for use on whole blood or dried blood spots; NADPH produced by oxidation of G6P to 6PG is measured by an increase in absorption at 340 nm.

Each test or batch of tests should include a normal and a G6PD-deficient sample. Sheep blood is a useful source of naturally deficient blood. Where possible, participation in an external quality assessment (or proficiency testing) scheme is also recommended.

Reagents

Method

Venous blood collected into ACD should be used. The test should be carried out within 8 h of collection and the blood should be kept at 4 °C until it is tested. Centrifuge the blood at 4 °C for 20 min at 1200–1500 g .

Discard the supernatant and add 0.5 ml of the packed red cells to 9 ml of 9 g/l NaCl and 0.5 ml of sodium nitrite solution contained in a 15 ml glass centrifuge tube. Incubate at 37 °C for 20 min. Centrifuge at 4 °C for 15 min at approximately 500 g , then discard the supernatant fluid without disturbing the buffy coat and uppermost layer of red cells. Wash the cells three times in cold saline. After the last washing, remove the buffy coat, mix the packed cells well and transfer 50 μl to a glass tube containing 1 ml of the incubation medium. Incubate the suspension undisturbed at 37 °C for 30 min. Then add 0.2 ml of MTT solution, shake gently and incubate at 37 °C for 1 h. Resuspend the cells thoroughly. Place one drop adjacent to one drop of hypotonic saline on a glass slide, mix the drops thoroughly and cover with a coverslip.

Examine the red cells with an oil-immersion objective, noting the presence of formazan granules ( Fig. 12-5 ).

Cytochemical demonstration of G6PD (glucose- 6-phosphate dehydrogenase) activity.

Normal blood; positive reaction with formazan granules in the red cells.

Interpretation

When G6PD activity is normal, all the red cells are stained. In hemizygotes for G6PD deficiency, the majority of the red cells are unstained. In heterozygotes, mosaicism is usually seen; usually 40–60% of the cells are unstained, but the proportion may be much less and in extreme cases as few as 2–3% may be unstained.

Principle

The nucleotide pool of normal red cells consists largely (> 96%) of purine (adenine and small amounts of guanine) derivatives. The levels of cytidine and uridine are normally extremely low. However, in P5N-deficient cells, more than 50% of this pool consists of pyrimidine nucleotides.

In acidic solutions, cytidine nucleotides have an absorbance maximum at approximately 280 nm, whereas adenine, guanine and uridine nucleotides absorb maximally at 260 nm. The ratio of absorbance at 260 nm to absorbance at 280 nm reflects the relative abundance of cytidine nucleotides; the absorbance ratio is lower when pyrimidine derivatives are increased.

Reagents

Method

For sample preparation, centrifuge blood freshly collected in EDTA at 1200 g for 5 min, remove the plasma and wash the cells three times with ice-cold 9 g/l NaCl solution. Add 1 ml of a 50% suspension of the washed red cells to 4 ml of ice-cold 4% PCA solution and then shake vigorously for 30 s. Transfer the clear supernatant obtained after centrifugation at 1200 g for 15 min to a small test tube. Prepare a sham extract by adding 1 ml of 9 g/l NaCl to 4 ml of 4% PCA solution.

Add 500 μl of water and 300 μl of 1 mol/l glycine buffer to each of two cuvettes. To correct for optical differences between the cuvettes, read the sample cuvette against the blank at 260 and at 280 nm, giving readings B ²⁶⁰ and B ²⁸⁰ . Add 200 μl of the red cell extract to the sample cuvette and 200 μl of the sham extract to the blank cuvette. With the spectrometer zeroed at 260 nm on the blank cuvette, read the sample cuvette to obtain the value S ²⁶⁰ . Repeat the process at 280 nm to obtain the reading S ²⁸⁰ .

The A ²⁶⁰ /A ²⁸⁰ absorbance ratio (R) is calculated by subtracting the cuvette blank readings (positive or negative) at 260 and 280 nm from the readings obtained on the red cell extract when blanked against the sham extract:

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