|Methods for the Evaluation of the Impact of Food and Nutrition Programmes (UNU, 1984, 287 p.)|
|4. Measuring impact using laboratory methodologies|
Barbara A. Underwood and Abraham Stekel
Development of a primary nutritional deficiency
Choice of tissue for laboratory assessment
Selection of laboratory methodologies for nutritional impact evaluation
Laboratory methods for assessment of nutritional impact
Annex A. Laboratory evaluation of protein nutriture
Annex B. Laboratory evaluation of vitamin A nutriture
Annex C. Suggested methods for hematology
Prolonged dietary inadequacy alters the biochemical milieu of the body, and consequently enzymatic activities. in advance of the appearance of clinical symptoms and signs. Laboratory measurements of the nutrient adequacy of body fluids or tissues, therefore, can provide objective, specific, and sensitive indicators of nutriture.
These measurements, judiciously selected for some nutrients, can provide subclinical information useful in evaluating the nutritional impact of nutrition interventions.
Depending on the specific nutrient in question, adequacy may be measured biochemically by:
Which laboratory approach is appropriate for measuring adequacy for a particular nutrient will depend on an understanding of its basic biochemical role, the distribution among body compartments during periods of dietary lack and sufficiency, and how this distribution is influenced by short and long-term changes in the physiological environment, e.g.. acute and chronic infections, and hormonal imbalances and variations.
Figure 4.1. (see
It is important to keep in mind that in practice, the relative availability of some nutrients to support biochemical functions varies from day to day with fluctuations in intake. Hence, flow in figure 4.1 may be upward or downward and a single laboratory measurement or clinical observation will not reveal the direction of events. Furthermore, the rapidity with which the direction of flow responds to alterations in food or nutrient supply will vary. For example, anatomical signs, though the last to appear, may take substantially longer to disappear than will restoration of the activity of nutrient-dependent enzymes or the concentration of the nutrient in blood. Thus the biochemical assessment of nutritional status for a specific nutrient does not always directly correlate with the findings from dietary or clinical assessment, particularly when applied on an individual basis or among populations of small sample size. Generally, when applied to populations for assessment of nutritional status, the trend will be in the same direction for dietary, biochemical, and clinical findings.
To use cross-sectionally obtained measurements for purposes of evaluating the nutritional impact of nutrition interventions, a laboratory measurement should be chosen that represents the cumulative effects on nutriture. i.e., nutritional status, rather than immediate responses to dietary intake. On the other hand (as noted earlier) acute changes in the physiological environment, acute infection, hormonal balances etc., as well as diet, can shift the distribution of nutrients among compartments, thus affecting biochemical events. Therefore it is often also useful to have an indicator of the immediate situation, particularly when dealing with individuals or populations of small size. In all cases, an appropriate comparison group is necessary to evaluate associations among laboratory measurements with nutritional intervention.
Biochemical measurements with nutritional implications can be made quite non-invasively on tissues such as hair and nails, or at the extreme of invasiveness on liver and muscle. But in practice, blood and its cellular components and urine are the most readily available tissues and can be obtained with moderate evasiveness, for estimating status in surveys. For most nutrients, urine is unsuitable for nutritional status assessment unless a timed or 24-hour sample can be obtained to compensate for diurnal variations in nutrient excretion rates and in volume. Relating values to creatinine concentrations reduces but does not eliminate this problem. This criticism is most applicable when applied to individuals and to populations of small size. On the other hand, casual urine specimens can be useful if the purpose is to evaluate compliance to a food or nutrient supplementation programme rather than nutritional impact. For example, by including a nutrient marker in the supplement that normally is excreted in the urine, such as riboflavin, increased levels can be detected qualitatively when related to non-participant or non-complying comparison groups.
As already noted, choice of fluid or cellular component in blood that best reflects status rather than immediate dietary intake will vary in relation to how specific nutrients are distributed between extra and intracellular compartments, how responsive this distribution is to dietary change, and how it is influenced by altered physiological conditions. such as acute or chronic infections. drugs, and stressful) circumstances, which are unrelated to diet. For many nutrients, compensatory biochemical mechanisms exist to adjust for short-term fluctuations in dietary intake and to delay the onset of clinical signs of inadequacy. Body reserves of varying size and half-life exist for this purpose. Ideally, assessment of change in the magnitude of the reserve supply of a nutrient (step 2, fig. 4.1.) would be most useful for nutritional surveillance purposes and for measuring the nutritional impact of certain nutrient-specific interventions where homeostatic mechanisms maintain body fluid levels until reserves are depleted, such as for iron and vitamin A. This would represent the earliest stage of dietary inadequacy and indicate when preventive measure should be instituted. However, biochemical indicators of tissue reserves that are present in blood and accessible to laboratory evaluation in surveys are available for only some nutrients, e.g., serum ferritin as a reflector of tissue iron stores. There is no "true" reserve store of protein, only variation in active protein mass that is not easily measured in surveys by laboratory methods.
Blood samples obtained from fasting subjects are preferred to avoid fluctuations in some nutrients that reflect immediate dietary intake. However, under field conditions, especially among children, non-fasting specimens are often all that can be obtained practically. By selection of the appropriate parameter, non-fasting specimens can be used without prejudice in interpretation for estimation of protein and iron status, and for vitamin A status except following a meal that contains a concentrated source of the vitamin (e.g., animal liver). Since blood samples are usually obtained in the morning hours and rich sources of preformed vitamin A are unusual breakfast items in developing countries, this is an unlikely significant confounding variable (1). Fasting specimens may be more critical in the laboratory assessment of certain other nutrients, particularly water-soluble vitamins (2).
Caution must be used in interpreting blood data obtained from subjects with acute infections. These may cause a transient lowering of blood level of some nutrient-specific transport proteins, such as retinol-binding protein and transferrin. Chronic infections too can cause a lowering of circulating levels for some nutrients. These blood concentration changes may not reflect a depletion of the total available body pool, but a temporary redistribution of the nutrient that is without physiological significance.
The appropriateness of using laboratory measurement for nutritional impact evaluation will depend on the nature of the intervention programme and the kind, severity, and prevalence of nutritional problems in the recipient population (see TABLE 4.1. Summary of Laboratory Methodology (Nutritional Biochemistry and Hematology)). Laboratory measurements are most appropriately applied in tandem with the introduction of specific, population-based nutrient interventions, such as iron, iodine, or vitamin A food fortification programmes, or in interventions targeted to individuals who are given specific supplements for which specific before-and after-treatment effects can be measured.
TABLE 4.1. Summary of Laboratory Methodology (Nutritional Biochemistry and Hematology)
|Kind of Information||Kind of Intervention||Unit of Observation||Personnel |
|Resources Required||Time Level||Survey|
|Albumin/ Prealbumin||Protein-calorie supplement to malnourished vulnerable groups||Individual||Lab. tech. (2 weeks)||Radio-immuno diffusion kit, refrigeration||10-30 |
|Prevalence of low serum retinol levels||Vitamin A||Individual (1 month)||Lab. tech.||HPLC Spectrophotometric |
|20 specs./day||Sophisticated Appropriate for LDCs|
|Hemoglobin concentration or hematocrit||Iron||Individual||Lab. tech. (2 weeks)||Colorimetric||3 minutes||Minimum for difficult field conditions|
|Transferrin saturation||Iron||Individual||Lab. tech. (1 month)||Spectrophotometer||20 minutes||Sophisticated|
|Free erythrocyte||Iron||Individual||Lab. tech.
|Protoporphyrin||Lab. tech. (1 month)||Fluorometer or Spectrophotometer||20 minutes|
|Serum ferritin||Iron||Individual||Lab. tech. (1 month)||Refrigerator Centrifuge Gamma counter or Spectrophotometer Serum ferritin kit||20 minutes||Sophisticated|
Biochemical measurements are also appropriate where intervention programmes, even non-nutrient-specific ones, are clearly targeted to vulnerable population groups with known significant dietary inadequacies. One example would be preschool children from poor environments in which PEM is prevalent and among their pregnant and lactating mothers.
Biochemical methods may not be useful in evaluating general food aid programmes for adult workers with only marginally adequate diets. Under these circumstances, limitations in the magnitude of biochemical responsiveness to moderate dietary change that can be reliably detected by laboratory measurements could preclude usefulness. For example, clearly it would be inappropriate to apply laboratory assessment of iron and vitamin A status to evaluate a food aid programme such as a grain distribution in a food-for-work programme, that did not include significant amount of these nutrients. Furthermore. it is unlikely that significant alterations in protein status would be detected by laboratory methodology in this type of adult recipient population.
In contrast, however, a feeding programme including a protein ration for preschool, poor children is likely to show a shift upward in the distribution of albumin levels from the lower range of the distribution curve. Because laboratory measurements are costly with respect to employment of professional personnel and possible continued cooperation of programme recipients, the decision of evaluators to include laboratory assessment should be carefully matched to the specific programme being evaluated, to maximize the potential for obtaining interpretable data on nutritional impact.
The major nutrient deficiencies of public health significance which food or nutrient-specific distribution programmes have most often been designed to alleviate include inadequate food energy intake, protein-energy malnutrition, iron deficiency anaemia. vitamin A deficiency, and iodine deficiency. Evaluation by laboratory measurement of the nutritional impact of food programmes to combat these five nutrient-specific problems is possible using several methods that vary in degree of sophistication, reliability, and accuracy. Individual laboratories should evaluate their own resources and select the method best suited to their situation. We have chosen to include in this chapter those methods that are most practical and least costly, that required a moderate level of training of personnel, yet provide the degree of precision and accuracy required for programme evaluation. Other methods might be more appropriate for research purposes, or where laboratories are well-equipped and have a highly trained staff.
Protein-Energy Malnutrition (PEM)
Energy balance is best assessed by non-laboratory measurements (see chapter 3), while protein status can be reflected in biochemical measurements. The speed at which changes in protein status occur will vary according to the turnover rate of the protein in question. In contrast to iron and vitamin A, for which there are tissues that accumulate reserve stores (step 2, fig. 4.1.), there is no true storage tissue for protein, only variations in the amount of total active protein mass. Knowledge of turnover rates of various protein species found in blood is needed, therefore, to select the parameter appropriate for use in evaluating nutritional impact of nutritional interventions, i.e., long-term effects rather than short-term responses to the relative availability of a balanced amino acid supply for protein synthesis. In this respect, blood levels of rapidly turning over transport proteins (retinol-binding protein, or RBP, transferring and prealbumin) are known to be sensitive to short-term changes in the available protein and energy supply (step 3. fig. 1), but do not necessarily reflect depletion in protein mass (step 2, fig. 1 ) or decreased functional level (step 4, fig. 1).
The slower turning over albumin level in blood best reflects longer term protein status (3). The transport proteins, for example, are most useful for evaluating the immediate responsiveness to an intervention that provides an energy or protein supplement, or that decreases the burden of infections that stress protein-energy requirements; albumin levels, on the other hand, better reflect true nutritional impact on protein status. Because some transport proteins, such as transferring and retinol-binding protein (RBP), are at least partially dependent upon the availability of the nutrient they carry, their use for assessing protein nutriture may be confounded by concurrent deficits of the dependent nutrient. Prealbumin, though a carrier of the RBP-retinol complex as well as the iodine-containing thyroxine, is not dependent on either of these nutrients for its hepatic synthesis or secretion, yet still has a short half-life (2-days) that makes it sensitive to the immediate availability of a balanced amino acid supply (3). Blood levels of prealbumin, like albumin and other transport proteins, are dependent upon adequate liver function, and therefore are depressed by liver disease independent of dietary adequacy.
To evaluate programme impact on protein status, a combined assessment of prealbumin and albumin provides information on both short and long-term dietary effects, respectively (4). This methodology is most appropriately applied for evaluation of intervention programmes targeted to vulnerable groups such as preschool children, pregnant and lactating mothers. Unless evidence exists for substantially inadequate protein or energy intake among other recipient populations, such as school children and adult male workers, prior to the intervention, laboratory assessment of protein nutriture is unlikely to reflect responsiveness to a dietary change achieved through food aid programmes.
Both prealbumin and albumin can be determined by relatively inexpensive, reliable laboratory technique requiring a routinely trained laboratory technician (see appendix). Radial immuno-diffusion (RID) is used to determine blood levels of prealbumin, and kits are available commercially for this purpose (5). The kits contain complete instructions and all the materials necessary for the assay, including the protein standards, with the exception of a microliter syringe and measuring ruler or caliper. The plates should be stored before use at refrigerator temperatures. The assay can be completed in as little as 18 hours.
There are several methods for determination of serum albumin, including standard electrophoresis, dyebinding, and salt fractionation. Specificity is highest for electrophoresis, followed by dye-binding and salt fractionation, and the relative cost for the analysis and sophistication of required equipment follow the same order. For evaluation purposes, the specificity and precision of the dye-binding procedure is adequate.
Vitamin A Nutriture
Interpretation of the biochemical measurement of vitamin A in blood short of deficient (< 10 µg/dl) or excess (> 70-80 µg/dl) levels is confounded by homeostatic controls, partially independent of diet, that modulate the release of liver reserve supplies (1). Hence, blood levels (step 3, fig. 1 ) do not necessarily reflect the level in the liver reserve (step 2, fig. 1 ) and, therefore, the relative level of vitamin A nutriture. Blood values that lie between about 15-30 µg/dl. particularly in young children and for some specific individuals, may reflect physiological conditions unrelated to vitamin A reserve stores. Under such circumstances, improved dietary intake of vitamin A through an intervention will not necessarily change the level in the blood. On the other hand, if the blood level for individuals is in a range that is difficult to interpret (15-30 µg/dl), for reasons of chronic inadequate intake and low liver reserves (step 2, fig. 1), blood levels will increase in response to an increased intake of vitamin A. Still, it cannot be assumed that adequate reserve stores have been established as a result of increased circulating levels, since blood levels must exceed a threshold before stores are replenished.
Therefore, to evaluate the impact of a food programme that seeks to improve vitamin A nutriture, it is important to look at changes in the lower end of the population distribution curve of blood levels rather than means or absolute values (6). When the lower end of the distribution curve shifts to the right following an intervention programme, this can be interpreted as programme impact even though there may be no significant change in mean or median values (7). Figures 4.2 (see
Retinol-binding protein has been suggested as a vitamin A adequacy indicator that can be simply assayed in the field by RID techniques. However, as noted above, RBP synthesis is influenced by acute protein deficiency (4) and liver function, as well as by the availability of vitamin A from the diet or reserve tissue stores (8). Furthermore, the RID assay determines total RBP, including that not bound to retinol. The unbound or apo-RBP is physiologically unimportant with respect to vitamin A status. It is important to note that total RBP may remain in the low normal range while available vitamin A (holo-RBP) has declined dangerously (1).
The analytic method of choice for determination of vitamin A status uses high-pressure liquid chromatography (HPLC), which is fast, determines retinol directly, and minimizes opportunity for oxidative losses, but is expensive.
Several other analytic methods are available, such as spectrophotometry, fluorometry, and colorimetry, and with care they can be used interchangeably, depending upon available laboratory resources. All methods require a carefully trained and standardized technician. These methods are described in detail elsewhere (1). When a spectrophotometer equivalent to a DU is available, the procedure of Bessey and Lowry, based on UV inactivation, is likely to be least expensive and most reliable. In the absence of a DU spectrophotometer, which involves a relatively high initial investment, the colorimetric assay using trifluoracetic acid, trichloracetic acid, or antimony bichloride can be satisfactory, provided a reliable vitamin A standard is available and proper precautions are exercised (1). Fluorometric procedures are more sensitive than the colorimetric ones. However. spurious high results are notable because of fluorescent contaminants difficult to avoir under most laboratory conditions in developing countries.
Serum levels of vitamin A obtained cross-sectionally and displayed in distribution curves can provide information on the percentage of individuals with low levels of vitamin A who may be subclinicaily malnourished. When evaluated against an appropriately matched comparison group, this information is useful in assessing differences in the magnitude of the "at risk" group among recipients of an intervention programme. The best way to determine the nutritional impact of a programme on vitamin A status is to have before and after treatment laboratory measurements. An alternative is to measure the response of a subsample of recipients with plasma values that are in the lower portion of the distribution to an additional short-term supplement (8).
Laboratory assessments of vitamin A status are appropriate for evaluation of nutrition intervention programmes in which the daily intake of vitamin A is increased (vitamin A-containing food fortification programmes), in single, massive-dose intervention programmes, and in programmes to correct severe forms of protein-energy malnutrition. Usefulness in the latter types of programme stems from the intimate interrelationship between protein and vitamin A status. Intervention programmes to correct serious protein deficiency should always provide sources of vitamin A concurrently, Since there is no practical way of knowing whether serum levels of vitamin A are lowered by depletion of tissue reserves or are secondary to protein deficiency and impairment of mobilization. Stimulation of growth by correcting protein deficiency elevates the need for vitamin A, and if the latter is not supplied, irreversible eye damage can be precipitated in a very short time.
Clinical examination and palpation of the thyroid gland are generally sufficient to evaluate the success of programmes to correct iodine deficiency and control endemic goiter. However, iodine nutriture can be assessed in the laboratory by measuring blood levels of protein-bound iodine (PBI), urinary excretion of iodine, and radioiodine uptake. Since all three of these parameters may be influenced by various physiological states and drugs, interpretation of iodine nutriture by biochemical methods must be done with caution. As for vitamin A, it is necessary to evaluate the iodine impact of food programmes by looking for shifts in the lower range of PBI and urinary excretion values rather than for absolute values for means or medians.
Iron Deficiency Anaemia
Nutritional anaemia is one of the most common and significant nutritional problems in the world today. It is likely, therefore, that anaemia will be prevalent in areas where nutrition intervention programmes take place. Iron deficiency, folate deficiency, protein-calorie malnutrition, acute infection, and chronic disease can all contribute to the occurrence of anaemia. Studies in several parts of the world have demonstrated, however, that iron deficiency is, in most situations, the main etiologic factor.
Definition of Anaemia
Anaemia is usually defined using criteria established by population studies. These studies have determined, for individuals of different sex, age, and physiological condition, levels of haemoglobin concentration under which anaemia is likely to be present. It must be borne in mind, however, that there is an overlap of haemoglobin concentration figures between normal and anaemic individuals. Therefore the use of fixed limits of normality will misdiagnose as anaemic a certain proportion of individuals with adequate haemoglobin concentration and include among the normal group some anaemic subjects (see
Stages in the Development of Iron Deficiency
A measureable decrease in haemoglobin concentration is a late effect of iron deficiency. The iron-replete individual not only has sufficient iron to synthesize haemoglobin and other essential iron containing compounds, but also has some iron reserves. The amount of iron stores in normal adult males have been estimated at 500 to 1,000 mg. Stores are lower in women of reproductive age and in infancy and childhood.
The first consequence of a negative iron balance is a decrease in the amount of storage iron, a condition known as iron depletion. Once stores are depleted there may not be a sufficient supply of iron for erythropoiesis, and haemoglobin synthesis is impaired. This state is known as iron-deficient erythropoiesis. After some time, this is reflected in a decrease in haemoglobin concentration and iron deficiency anaemia.
The most useful tests for the evaluation of iron nutritional status are the plasma ferritin, the per cent saturation of transferrin, the concentration of free erythrocyte protoporphyrin, and the haemoglobin concentration. There is a general correlation between the stages of iron deficiency and the changes in these laboratory tests.
Valuable information on the iron status of individuals can be obtained by the measurement of plasma or serum ferritin. It has been shown that the concentration of this compound (step 3, fig. 4.1.) reflects the amount of storage iron (step 2, fig. 4.1.), and that 1 µg/l of serum ferritin is roughly equivalent to 10 mg. of iron stores. With progressive depletion of iron stores there is a parallel drop in serum ferritin, with serum values below 12 µg/l representing absence of storage iron.
After iron stores are depleted there is a drop in the amount of iron being transported in the plasma (plasma iron). The amount of the transport protein transferrin that is saturated with iron (or total binding capacity) concomitantly increases, so that per cent saturation of transferrin with iron falls from values above 30 per cent to less than 15 per cent. At the same time, since not all the protoporphyrin synthesized by erythrocyte precursors in the bone marrow is formed into heme because of the insufficient iron supply, there is a rise in the amount of free erythrocyte protoporphyrin in red cells from normal values of about 30 µg/dl to above 100 µ/dl. Finally, there is a measurable drop in haemoglobin concentration.
The relationship of these measurements to iron stores and the values found in the different stages of iron deficiency are depicted in figure 4.5.(see
Monitoring Results of Intervention Programmes
As already mentioned, the two most useful laboratory tests for measuring the effect of interventions on nutritional anaemias are the haemoglobin concentration and the serum or plasma ferritin. In situations where there is a high prevalence of anaemia and supplementation strategies are used, effects will be most readily measured by changes in haemoglobin. With food fortification, on the other hand, especially in populations where there is little anaemia, one can expect relatively modest increases in the amount of daily absorbed iron, and results may be better monitored by measuring changes in iron stores as reflected in serum ferritin.
The effects of intervention programmes on iron deficiency anaemia can be better evaluated in vulnerable groups such as infants, preschool children, and pregnant women. Ideally, studies should be conducted in representative samples of the target populations and should include appropriate control groups.
1. Determination of Serum Albumin
Serum albumin levels can be determined by standard electrophoresis, salt fractionation, or dye-binding techniques. Although electrophoretic analyses can provide precise, specific quantitative data, for this the technique requires an electrophoresis apparatus and densitometer or colorimeter and a separate determination of total protein. In contrast, the salt fractionation procedure is rather non-specific and suffers from lack of precision. On the other hand, specific dye-binding methods are rapid, involve few manipulations and specialized equipment, and do not require a separate determination of total protein. A widely used method of this type, based on bromcresol green (BCG) (12), is described here.
The addition of albumin to a solution of bromcresol green in a 0.075 M succinate buffer, pH 4.20, results in an increase in absorbance of 628 nm. The absorbance-concentration relationship is linear for samples containing up to 6 g/dl albumin. Bilirubin, moderate lipaemia, and salicylate do not interfere with the analysis. The use of a non-ionic surfactant (Brij-35) reduces the absorbance of the blank, prevents turbidity, and provides linearity. The results with this method agree very well with those obtained by electrophoresis and salt fractionation. The method is simple, it has excellent precision, and the reagents are stable.
Spectrophotometer or colorimeter equipped with 630 nm interference filter.
A sample of 25 µl of serum or working standard solutions are added to 5.0 ml of working dye solution. The solution is vigorously mixed and allowed to stand 10 minutes at 25° C. The absorbance is measured at 628 nm after adjusting the instrument to zero absorbance with the working dye solution. If a serum sample is extremely lipaemic, a serum blank is prepared by adding 25 µl of the sample to 5.0 ml of 0.075 M succinate buffer. Its absorbance with water as a reference is subtracted from the absorbance of the unknown. The albumin concentration of the serum is obtained from an absorbance-concentration plot, or, if the response of the instruments is linear, only a single standard solution (2.0 g/dl) is required and the serum albumin concentration is calculated in the usual manner. A typical standard curve is seen in figure 4.A.1 (see
2. Determination of Serum Prealbumin by Radial Immunodiffusion (RID)
Procedure as Described by Manufacturer:
Although the antibody for human prealbumin can be developed by standard techniques, commercial kits are available from Calbiochem-Behring Corp. (10399 North Torrey Pines Rd., La Jolla, CA 92037, USA) and may be more economical if a limited number of assays are anticipated. Since prealbumin is synthesized in the liver, acute and chronic liver disease will reduce serum concentrations independent of protein nutriture. Total fasting for more than 48 hours also will reduce serum levels by limiting the substrate available for hepatic protein synthesis. Hence, in the absence of liver disease and prolonged absolute fasting, prealbumin is a sensitive parameter for improvement in protein nutritional status.
A protein (antigen) solution is applied to a cylindrical well cut in a gel matrix containing a uniform concentration of monospecific antibodies. Antigen placed in the well diffuses radially, producing a precipitin ring. Precipitin rings can be read any time after overnight incubation, or endpoint. Results are quantitated by comparing the diameter of the precipitin ring produced by the sample to the precipitin rings produced by standards of known concentrations.
Plate preparation: Carefully remove the plate from the container. Open the plate by pressing the thumbs firmly on the centre of the lid while holding the lid at edges. Allow the plate to stand open at room temperature to permit evaporation of any moisture that may have condensed in the wells.
The user can choose, according to workload requirements, to make determinations based either on an overnight readout from a reference curve or on an endpoint readout (after 48 hours). The initial steps of the procedure are the same for either method.
The assay range of M-Partigen(tm) prealbumin radial immunodiffusion plates is indicated on the label. If the sample concentration exceeds the upper limit of the assay range, the test should be repeated with appropriate dilutions of the specimen.
If the reference curve is not linear, or does not intercept the ordinate at 11 + 3.5 mm2 when the endpoint curve is plotted, the procedure should be repeated; errors in technique or product instability should be considered.
1. Spectrophotometric Method Based on UV Inactivation (1)
Vitamin A (retinol) is destroyed when exposed to ultraviolet light. After saponification with alcoholic KOH, retinol (and carotenoids*) is extracted by solvent partition using a mixture of xyiene-kerosene. The optical absorbance of the sample extract is read at 460 nm for the determination of total carotenoids and at 328 nm for the determination of retinol. The sample extract is then irradiated with ultraviolet light and its absorbance read again at 328 nm. The difference in optical absorbance at 328 nm before and after irradiation of the sample corresponds to the amount of retinol present. The concentration of carotenoids and retinol are calculated. based on their respective extinction coefficients in the solvent mixtures.
Since cleanliness is a critical factor in this type of analysis, it is recommended that all the glassware be treated as follows: After regular washing, glassware should be rinsed with a 50 per cent solution of nitric acid and rinsed again with sufficient distilled water to remove all traces of nitric acid. The microcells are first washed with a 1: 1 mixture of 3N HCI-ethanol, rinsed again with ethanol, and finally with the xylenekerosene mixture.
In this method, the introduction of a standard serves only to check the assay conditions, and its absorbance is not used in the calculation of the retinol concentration.
For the purpose specified, the USP reference standard for vitamin A from USPC Inc., Rockville, MD, USA, can be used. This is a solution of retinol acetate in cottonseed oil. Several procedures to prepare adequate standard solutions using this substance have given excellent results. One is outlined below. Weigh out approximately 30 mg of the oily UPS reference standard solution of vitamin A, which contains approximately 34.4 mg retinyl acetate/g, and dissolve in 100 ml ethanol. The solubility of the oil in ethanol does not permit more concentrated solutions. This solution can be used as a stock if kept in a dark bottle and refrigerated. If a calibration curve is desired, prepare suitable dilutions of the stock standard in ethanol. Carefully determine the absorbance of these solutions at 328 nm before (A0) and after (A1) irradiation with ultraviolet light.
Determine the time required for A, to reach a plateau under the assay conditions. Calculate net absorbency values (A0 - A1) of retinyl acetate solutions for each dilution used under conditions of optimal bleaching. Under proper conditions of irradiation, A, should be 3 per cent or less of A0. Plot the values with A0 - A1 on the y axis and relative vitamin A concentration on the x axis curve. Determine the method-specific absorbency factor as a function of the retinyl acetate concentration. For the latter, an E 1%(1cm) value at 328 nm of 1565 for retinyl acetate in ethanol is used. To obtain the concentration of the retinyl acetate solution in terms of retinol, an E 1%(1cm) value of 1795 in ethanol is used. This value corrects for the 3 per cent decrease in the E 1%(1cm) value of retinyl esters in ethanol.
The calculations for retinol and carotene are based on their respective extinction coefficients (E 1%(1cm). The factor 637 is used to calculate the retinol concentration and it corresponds to a E 1%(1cm) of 1,570 adjusted to yield directly g of retinol/dl. The factor 480 used for carotenes corresponds to a E 1%(1cm) of 2080 of this substance adjusted to yield directly g of carotenes/dl. The respective calculations are as follows:
Retinol (g/dl) = A (328) - A' x 637
Carotenes (g/dl) = A (460) x 480 where A = initial optical absorbance reading
A' = optical absorbance after ultraviolet irradiation
Effect of Sample Storage
If care is taken to minimize the air/serum interface (storage under nitrogen is strongly recommended) and exposure to light, samples may be stored at -20 C for several months without affecting the vitamin A levels. Retinol is more stable on storage than carotenes are.
Variations and Modifications
The size of the sample can be changed as desired from 50-200 l to larger volumes up to 1.5 or 2.0 ml, depending on sample volume availability. A large sample volume may facilitate the analytical procedure by avoiding the use of specialized glassware, micro-cell adaptors in the spectrophotometer, and the meticulous and sometimes difficult handling of small amounts of sample extracts.
Although the ratio of sample volume to the volume of alcoholic KOH added for saponification must always be 1:1, the ration of sample size to the volume of solvent added for the extraction procedure can be modified. Instead of using a 1:1 ratio, larger volumes of solvent (xylene-kerosene) can be used. This modification may facilitate the extraction procedure and will provide a larger volume of sample extracts that would be easier to handle. Keep in mind, however, that the more solvent added the more diluted the vitamin A content of the sample, and the lower the optical absorbance readings. This may be a critical factor to consider when dealing with samples with low vitamin A levels.
If the proportions of sample size to the volume of solvent added is changed from a 1:1 ratio, make sure that proper correction is made for the dilution of the sample when doing the calculations.
Because of its high boiling point, the xylene-kerosene mixture is the preferred solvent. Only kerosene with an initial O.D. below 2 is suitable. However, laboratory workers can use other organic solvents, such as purified jet fuel (Turbo-fuel A-1), which is available at most major airports. About 300 ml of the solvent are distilled in 15 ml fractions. Fractions with an O.D. < 0.5 at 328 nm are pooled for use. Appropriate E 1%(1cm) for retinol and carotenes must be selected for the specific solvent used. The volatility of cyclohexane, otherwise an excellent alternative, reduces its utility. This constraint is critical, especially when using small amounts of sample.
2. Colorimetric Method Based on Carr-Price Reactions Using TFA (1)
The proteins of plasma or serum are precipitated with ethanol and the vitamin A and carotene are extracted into hexane (or petroleum ether). The carotene concentration is determined by measuring the absorption of the extract at 450 nm (A450). Following evaporation of the solvent, vitamin A dissolved in chloroform is determined by reading, at two time points the intensity of blue colour developed after addition of trifluoracetic acid-chloroform reagent. A correction is made for the concentration of carotene, since carotene contributes to the intensity of blue colour when present in high amounts. When serum levels of vitamin A are low in the presence of carotene in high amounts. falsely low vitamin A values can be avoided by first removing carotenoids by chromatography on alumina columns.
All procedures should be carried out in dim light and caution exercised to avoid excessive exposure to oxidation.
Duplicate 2 ml aliquots of serum or plasma are pipetted into glass-stoppered test tubes. An equal volume (2 ml) of ethanol is added dropwise with mixing to give a 50 per cent solution (v/v). At this concentration the proteinretinol blond is disrupted and the free retinol and retinyl esters are extracted by addition of 3 ml hexane (or petroleum ether). The tubes are stoppered and contents are mixed vigorously by mechanical mixer for 2 minutes, then centrifuged 5-10 minutes at 600-1000 xg to obtain a clean separation of phases. Then 2 ml of the upper hexane (or petroleum ether) extract is pipetted into cuvettes and the cuvettes capped. Absorbance at 450 nm due to carotenoids is read against a hexane (or petroleum ether) blank (A450).
After determining A450, the cuvettes are removed and the hexane (or petroleum ether) evaporated just to dryness under a stream of nitrogen in a 40-60 C. water bath in dim light. If evaporation cannot be carried out in the cuvettes, transfer to another tube with rinsing and proceed. Just at the point of dryness, the residue is immediately redissolved and dehydrated in 0.1 ml of a mixture of chloroform acetic ahnydride (1 :1 v/v). The cuvettes or tubes should be capped to minimize evaporation and protected from light.
The spectrophotometer at 620 nm is set at zero absorbance with a blank consisting of 0.1 ml chloroform-acetic anhydride mixture and 1.0 ml TFA-chloroform chromagen reagent.
The cuvette containing the sample is placed in the spectrophotometer and 1.0 ml TFA chromagen reagent added to the cuvette from a rapid delivery pipette. Alternatively, the reagent can be added to the extract in a separate tube and rapidly transferred to the cuvette. These steps must be carried out rapidly and with care since the blue colour fades quickly and the chromagen reagent is highly corrosive. Record the absorbance reading (A620) at exactly 15 seconds (t15) and 30 seconds (t30) after addition of the reagent (A620).
The microprocedure is essentially the same as the macroprocedure except adapted to a smaller scale. Accurate results are dependent upon great care in pipetting and transfering small volumes and minimizing possible losses from evaporation, oxidation, and light exposure.
A minimum of 50 Ill and preferably 100-200 l serum or plasma are pipetted into 6 x 50 mm glass stoppered test tubes. An equal volume of ethanol is added with mixing followed by 1.5 volumes hexane or petroleum ether (40-60 BP) (1 :1: 1.5 v/v/v ratio serum-alcohol-solvent, respectively).
The tubes are immediately stoppered and vortexed for 2 minutes, then centrifuged 5-10 minutes at 600-1000 xg to achieve a clean phase separation. 100 l hexane (petroleum ether) extract is transferred to a microcuvette by means of a micropipette and the absorbance due to carotenoids at 450 nm read against a hexane (or petroleum) blank.
The spectrocolorimeter, or preferably a spectrophotometer of quality equivalent to a Beckman DU, is set at 620 nm and then zeroed against a reagent blank containing 10 l chloroform-acetic anhydride reagent and 100 l freshly prepared TFA-chloroform chromagen reagent.
The sample is then transferred from the microcuvette to a clean 6 x 50 mm test tube, cuvette rinsed once with 50 l hexane (petroleum ether), and the rinsing added to the sample in test tube. The extract is then evaporated just to dryness under a stream of nitrogen in a water bath in dim light.
The residue is redissolved in 10 l chloroform-acetic anhydride (1:1, v/v) reagent.
Then 100 l TFA-chloroform chromagen reagent is rapidly added with vigorous mixing and the solution rapidly transferred to the microcuvette by means of a microtransfer pipette.
A reading at A620 is obtained against a TFA reagent blank at exactly 15 sec. (t15) and 30 sec. (t30) after addition of the chromagen. Careful timing is essential since the colour fades rapidly.
Preparation of Standard Curves
carotene-blue colour at A450
Weigh exactly 50 mg freshly opened all bans carotene standard and dissolve in a few millilitres of chloroform. Bring to exactly 100 ml in a volumetric flask with hexane (or petroleum ether). Prepare just prior to use in establishing the calibration curve since the solution deteriorates on storage. This is the stock carotene solution containing 0.5 mg/ml. Protect from light.
An intermediate standard containing 5 g/ml is prepared by diluting 1 ml stock carotene solution to 100 ml in a volumetric flask with hexane (or petroleum ether). This solution is stable only for a few hours and should be made just prior to use.
Working standards are prepared from the intermediate standard by diluting with hexane (or petroleum ether) in each of four 10 ml volumetric flasks 1, 2, 4, and 8 ml of intermediate standard solution. This results in solutions containing 0.5, 1.0, 2.0, and 4.0 g/ml of ,13-carotene, respectively.
Fill the cuvette with the carotene working standards and read A450 against a hexane (or petroleum ether) blank. Plot a standard curve and from it determine the factor (F)
g carotene/ml. for carotene (C) where FC450 = (g carotene/ml.) / A450
carotene-blue colour at A620
Carotenoids react with the TFA-chloroform to add to the blue colour at A620. Therefore, it is necessary to run a chromagen-carotene standard curve in order to calculate a correction factor in obtaining vitamin A values. This correction is not necessary if serum (plasma) contains carotene in concentration under 50 g/dl as the contribution to the blue color in this concentration range is negligible.
Carotene standards in chloroform are prepared to contain 4.0, 8.0, and 10.0 g/ml. Aliquots of 0.1 ml are pipetted into cuvettes. To this are added rapidly 1.0 ml TFA-chloroform chromagen with vigorous mixing. Absorbance at 620 mm is read at 15 sec. (t15) and 30 (t30) sec. exactly as described previously. The t15 and t30 values are plotted on rectangular coordinate graph paper with the ordinate containing A620 values and the abscissa time after addition of chromagen. Extrapolate to to with a ruler to determine the A620 at to. The to absorbance value, thus, can be determined by the formula At0 = At5 + (At5 - At30). Determine a factor (FC620) where: g carotene/ml FC620= A
A620 The carotene correction factor for vitamin A at A620 is FC5620 C in which the factor 2 derives from the difference in the dilution of the carotenoids and vitamin A in their respective assays.
Retinyl acetate or retinol can be used as standards for preparation of reference curves, since both have identical blue color characteristics in the analytic procedure after making appropriate molecular weight adjustments to convert retinyl acetate (MW = 328) to retinol (MW = 286) equivalents (i.e., when the acetate is used 286/328 = 0.872). The USP reference capsule of retinyl acetate in oil is suitable. It is said to contain 34.4 mg all bans retinyl acetate/gin solution, but it is necessary to check the concentration of dilutions by spectrophotometry. Standards should be kept refrigerated and protected from light. They should not be used beyond storage of two days without redetermining the concentration spectrophotometrically.
A stock vitamin A standard containing approximately 50/60 g/ml is prepared by carefully weighing an appropriate amount of the vitamin A standard and diluting with hexane in a volumetric flask. The exact concentration in g retinol/ml is obtained by determining the absorbance at 325 nm and using the retinol extinction coefficient, E 1%(1cm) = 1,850.
Working standards are prepared in 10 ml volumetric flasks by diluting the appropriate volume of the stock solution with hexane (or petroleum ether) to obtain concentrations in the range of 6, 12, 24, 36, and 60 g/ml.
Then 0.1 ml of each working standard is pipetted into cuvettes for reaction with 1.0 ml TFA-chloroform chromagen exactly as previously described, reading A620 at 15 sec (t15) and 30 sec (t30).
The A620 values for t15 and t30 are plotted on a graph where the ordinate is the A620 values and the abscissa the time after addition of chromagen. Using a ruler, extrapolate to to to obtain A620 for each working standard. This is the theoretical time of maximum colour intensity obtainable, since the decay in blue colour is linear at least up to 30 seconds after chromagen additions.
Plot a standard curve from the A620 values at to on ordinary rectangular coordinate paper where the ordinate is the A620 value and the abscissa the number of micrograms of vitamin A per tube. From the curve calculate a factor (FA620) where:
FA620 = (g vit A/tube) / A620
Based on the procedure outlined using 2 ml serum (plasma) extracted into 3 MP solvent, serum values are calculated by the following formula:
Total carotenoids (g/dl) =A450 x FC450 x 150
Where FC450 is the constant determined in each laboratory and 150 accounts for dilution factors.
Vitamin A (g/dl) = (A620 - 2A450 x FC620) x FA62o x 75 FC620
If the microprocedure is used, appropriate modifications in the calculations must be made for the volume of serum actually used.
Following collection of blood, serum or plasma should be separated within 24 hours. Preferably, the analysis will be carried out immediately. If this is not possible, serum or plasma should be frozen at -20 C until analysed. The samples should be placed in tubes that allow a minimum air head space, tightly stoppered and protected from light. Stability is increased by flushing the tubes with nitrogen or other inert gas prior to a tight seal. Storage without thawing results in little or no loss up to about one month, with approximately a 10% loss over a four-month storage period. Storage in a freezer has been reported not to affect values when TFA chromagen is used. Freezing may induce spurious high values when antimony bichloride is used. Each laboratory should determine the stability of standardized samples under its own storage conditions.
Haemoglobin concentration can be measured in venous or capillary blood by colorimetric determination of derivatives of hemoglobin such as cyanmethemoglobin, oxyhemoglobin, or acid hematin. Various automated methods exist that are based on some of these principles. The preferred method, as recommended by the International Committee of Standardization in Hematology (14), involves the conversion of ail hemoglobin derivatives except sulfhemoglobin to cyanmethemoglobin by dilution of blood in a solution containing potassium cyanide and potassium ferricyanide. Absorbance is then measured in a photoelectric colorimeter or spectrophotometer at a wavelength of 540 nm. Reference standards of cyanmethemoglobin that conform to the specifications of the ICSH are commercially available.
Comparable information to that obtained from hemoglobin determinations can be obtained by the measurement of the packed red-cell volume, a simple technique that only requires a microcentrifuge and capillary tubes. However, due to changes in the mean corpuscular hemoglobin concentration that occur in iron deficiency anaemia, changes in hemoglobin concentration are more marked than those in the packed cell volume.
Choice of methods for serum iron and total iron-binding capacity (TIBC) will depend on equipment and amount of specimen available. Most current methods consist of the colorimetric measurement of an ironchromogen complex and require a spectrophotometer. The TIBC is determined by adding excess iron to the serum sample and removing unbound iron by some absorbent such as magnesium carbonate. Percent saturation of the TIBC (transferrin) is then calculated. Careful handling of specimens with iron-free material is essential. The Iron Panel of the ICSH has recommended reference methods employing bathophenathroline sulfonate as the colour reagent (15-16). These methods have recently been reviewed in detail (17).
Recommended methods for the determination of FEP (18-19) require a fluorometer and relatively elaborate extraction stages. An alternate method determines Zn protoporphyrin directly from a blood smear using a special apparatus called a hematofluorometer (20). This method is simple, inexpensive, and well adapted for field work, but needs further validation. According to some authors (19), the instrument does not have the precision or accuracy to detect small changes in fluorescence that occur with mild iron deficiency.
Ferritin can be determined in a small amount of serum or plasma by radio-immunoassay (21), and there are several commercial kits that give satisfactory results. The need for radio-isotope facilities has been recently eliminated by the introduction of enzyme immunoassays that only require a spectrophotometer. EIA methods seem to give results comparable to those obtained by RIA. The relatively large variability of results that currently exists among different laboratories may be reduced in the future by the use of ferritin standards that are becoming available from the ICSH.