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Laboratory for Human Biology Research, Department of Anthropology, Northwestern University, Evanston, IL 60208 and * Departments of Anthropology and International Health, University of Washington, Seattle, WA 98195
2To whom correspondence and reprint requests should be addressed. E-mail: t-mcdade{at}northwestern.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • iron deficiency • transferrin receptor • blood specimen collection • immunoassay
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). In part, this is the result of inadequate screening tools to overcome the logistical constraints imposed by impoverished and/or remote field settings in which iron deficiency is likely to be most prevalent. Here we present a method for measuring transferrin receptor (TfR) level, an index of iron deficiency anemia as well as preanemic iron-deficient erythropoiesis (2
–6
), in samples of whole blood dried on filter paper.
Dried blood spot methods are available for a growing number of analytes, and several community-based applications have shown this to be a convenient and reliable means to facilitate sample collection, storage, and transportation (7
–12
). A major advantage of finger-prick blood sampling is the elimination of venipuncture, a relatively invasive sampling procedure (particularly for infants and children) that requires readily accessible facilities where blood samples can be promptly processed and stored under controlled conditions.
Building on a previously developed serum assay for TfR (13
), Cook et al. (7
) recently validated a method for measuring TfR in dried spots of whole blood collected on filter paper. This effort has demonstrated the feasibility of measuring TfR in small quantities of dried blood spots, but three shortcomings may limit the application of the method in future field-based studies of iron deficiency:
) uses in-house assay standards and antibodies that have recently become commercially available in a kit manufactured by Ramco Laboratories, however the kit protocol differs substantially from the protocol evaluated by Cook et al. (7
), and has yet to be validated with respect to its application to whole blood spots.
) calculates concentrations of TfR for whole blood samples based on a standard curve generated from standards in a plasma-like matrix; our method uses whole blood standards that are stored, processed and assayed in a manner that is identical to whole blood unknowns to maximize comparability.
) requires a premeasured quantity of sample, i.e., whole blood is collected by venipuncture or in capillary tubes after finger sticks, and spotted with a pipette onto filter paper in 25-µL quantities. This procedure adds processing steps at the sampling site, increases the cost of blood collection and generates additional biohazardous waste. In our method, capillary blood is applied directly from the finger to filter paper, and a hole punch is used in the laboratory to obtain a uniform amount of sample that allows for the quantitative determination of TfR concentration without premeasurement at the collection site. |
MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Up to five drops of blood can be collected on each filter paper, but one drop (
50 µL) is sufficient because the assay can be performed in duplicate with two 3.2-mm discs of dried blood (one 50-µL drop yields up to seven 3.2-mm discs). Samples are dried overnight at room temperature, placed in air tight plastic bags or containers with desiccant and then frozen at -20° to -30°C until analysis. A set of in-house laboratory controls provided 37 matched plasma and whole blood samples to evaluate the correlation between blood spot and plasma concentrations of TfR. From each sample of venous blood (EDTA-anticoagulated), a portion of whole blood was removed immediately after collection and spotted onto filter paper; the remaining sample was centrifuged at 1,500 x g for 15 min and the plasma withdrawn. All procedures in this validation study comply with the Helsinki Declaration as revised in 1983.
Reagents and standards.
With the exception of blood spot standards and elution buffer, all necessary reagents are included in a commercially available enzyme immunoassay (EIA) kit (TF-94, Ramco Laboratories, Stafford, TX). Microwell strips are precoated with polyclonal anti-TfR capture antibody, and anti-TfR monoclonal antibody conjugated with horseradish peroxidase is included as the detection antibody. Tetramethylbenzidine substrate and 2.5 mol/L sulfuric acid stop solutions are also included. Following previous work by Cook et al. (7
), PBS (pH 7.2) plus 0.05% Tween-20 was used to elute dried blood spot samples, standards and controls.
To minimize matrix differences and maximize comparability between standards and unknowns, blood spot standards were made by diluting purified TfR protein stock (purchased separately from Ramco Laboratories) with washed erythrocytes, followed by application onto filter paper. Washed erythrocytes were obtained as follows: 1) whole blood was collected by venipuncture in 5 mL EDTA vacutainer tubes, and centrifuged at 1,500 x g for 15 min; 2) plasma and buffy coat were removed and discarded; 3) 3 mL normal saline (8.6 g NaCl/8.6 g/L dH2O) was added; 4) tubes were mixed gently for 5 min on a hematology rotor and centrifuged as before. Saline and any remaining buffy coat were removed, and steps 3 and 4 were repeated for a total of 3 washes.
Blood spot TfR standards were made as follows. Stock TfR was serially diluted in dilution buffer (PBS containing bovine serum albumin and normal rabbit serum; provided by Ramco) to generate concentrations across the desired assay range. In this case, TfR stock was diluted to 35 mg/L, and then serially diluted to 17.5, 8.75, 4.38 and 2.19 mg/L. Each diluted standard was added to an equal volume of washed erythrocytes (1:2 dilution). The calibrator dilution buffer was also added to an equal volume of washed erythrocytes for a zero standard. All solutions were mixed thoroughly. Standards were then applied to labeled filter paper cards in 50-µL drops using a manual pipette, dried overnight at room temperature and stored at -20°C in zip-lock plastic bags with desiccant. Final standard concentrations were 17.50, 8.75, 4.38, 2.19 and 1.10 mg/L.
Blood spot procedure
Elution protocol.
The day before an assay was to be performed, blood spot standards, unknowns and controls were removed from the freezer, and two discs of each were punched out using a standard 3.2-mm (1/8 inch) hole punch (available from office supply stores) and placed in disposable glass tubes. Elution buffer (250 µL) was added and the tubes were covered with parafilm. Because each disc contains 1.5 µL of serum, this corresponds to a dilution factor of 1:83.3. Samples were incubated overnight at 4°C. The day of the assay, samples were removed from refrigeration and rotated at 350 rpm at room temperature for 2 h.
Assay protocol. The blood spot assay was performed as follows: 1) sample eluate (100 µL) was added (in duplicate) to microtiter wells; 2) horseradish peroxidase–conjugated detection antibody (100 µL) was added to all wells; 3) plates were sealed with adhesive film, rotated for 10 min at 200 rpm, then incubated at room temperature for 2 h; 4) plates were washed 4 times with wash buffer (included with Ramco kit); 5) substrate solution (200 µL) was added to each well; 6) plates were incubated in the dark at room temperature for 1 h; 7) stop solution (50 µL) was added; 8) absorbances were read at 450 nm on a microplate reader (BioTek Instruments Elx808, Winooski, VT); 9) Unknown concentrations were calculated from the best-fit 4 parameter logistic standard curve (KCJunior, BioTek).
Analysis of assay performance. The performance of the blood spot assay was investigated by evaluating the difference between blood spot and plasma TfR concentrations in paired samples, assay linearity, precision and reliability, and sensitivity. In addition, we investigated potential variation in TfR concentration in discs punched from different locations within a single blood spot, and the stability of blood spot TfR under a range of storage conditions. Plasma samples were assayed using the standard Ramco TfR EIA protocol, and blood spot samples were assayed according to the modified procedure detailed above. Statistical analyses were performed using STATA (version 6.0; STATA, College Station, TX).
Linearity was assessed by eluting and then serially diluting (1:2, 1:4) three blood spot samples at the high end of the assay range. Assay precision and reliability were evaluated by calculating within-assay and between-assay CV (SD/mean) from multiple determinations of two in-house laboratory controls at the low and high end of the assay range, respectively. Precision was evaluated with 10 determinations of each control in a single assay, and reliability was evaluated with duplicate measurements of each control across seven assays performed within a 2-wk period.
Even though elevated concentrations of TfR are of primary interest due to their association with iron deficiency, assay sensitivity (minimal detectable dose) was evaluated on the basis of 10 determinations of the zero standard (dilution buffer and washed erythrocytes) measured in the same assay. The mean absorbance of the zero standard was calculated, and the point 2 SD above zero was plotted on the assay standard curve to determine the lowest TfR concentration that could be differentiated from zero.
It has been reported previously that TfR concentrations may vary in discs of whole blood punched out at varying distances from the center of the same blood spot (7
). A typical 50-µL spot of dried blood will yield seven 3.2-mm discs: one in the center of the spot and 6 around the periphery. We investigated the possibility of uneven distribution of TfR by comparing the concentration of TfR in discs punched from the center and from the periphery of ten 50-µL blood spot samples.
The stability of TfR in blood spots was determined over a 4-wk period in which blood spots for three control samples were exposed to one of three temperature conditions [37°C, room temperature (21–23°C), 4°C], and one oscillating condition (12 h at 32°C and 12 h at 21°C to represent ambient conditions in tropical environments), in the presence or absence of 2 desiccant packs (Humidity Sponge, VWR # 61161–319, Chicago, IL). Samples were considered to be stable if values remained within a 10% CV range of the initial baseline values (from the mean of 10 determinations of a baseline sample).
The stability of TfR to repeated cycles of freezing and thawing was also evaluated to consider the potential effects of removing samples from the freezer during assay set up. Two blood spot controls were removed from their plastic bags and placed on the benchtop at room temperature for 1 h, and then returned to the freezer. The procedure was repeated for 4 days.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). The Pearson correlation is reported for comparability with previous publications, but an examination of the mean difference between the two methods is a more appropriate measure of agreement (14
). On average, plasma TfR concentrations were 0.741 mg/L higher than blood spot TfR (SD = 0.991). This result was expected because analyte concentrations are typically higher in plasma due to the diluting effect of erythrocytes in whole blood spots. All paired differences were within a 2 SD range around the mean, indicating an acceptable level of agreement (14
).
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Analysis of TfR stability in blood spots indicated that samples can be stored at room temperature or 4°C for at least 4 wk, but decline rapidly at 37°C, consistent with the pattern of stability reported by Cook et al. (7
). After 3 d, TfR concentrations in samples subjected to the oscillating condition declined on average to 85.0% of baseline in the presence of desiccant, and to 81.9% of baseline in the absence of desiccant. After 7 d, concentrations dropped to 76.7 and 75.1% in the presence and absence of desiccant, respectively. These results indicate that an effort should me made to keep samples at a stable room temperature (21–23°C) or lower after sample collection to maintain the integrity of TfR in filter paper. Blood spot samples showed no consistent pattern of decline associated with freezing and thawing.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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–6
), but prior attempts to quantify TfR have been hindered by requirements for sample collection and processing. We developed and validated a "field-friendly" method for assessing TfR that minimizes these requirements by using samples of whole blood applied directly to filter paper after finger stick. The blood spot TfR assay is precise and reliable, sensitive, and agrees well with plasma TfR. In addition, the assay uses commercially available materials, and can be performed at any facility with a microplate reader and basic laboratory equipment.
The collection of capillary blood on filter paper has several advantages over venipuncture that make it ideal for field-based research. Sample collection is relatively painless and noninvasive; samples do not have to be centrifuged, separated or immediately frozen, and sample storage and transportation are facilitated by the fact that filter papers can be stacked and stored in air-tight containers and kept at ambient temperatures. Finger prick blood sampling eliminates the need for a trained phlebotomist to collect and process blood samples, and reduces the burden imposed upon participants. Samples can be collected virtually anywhere; infants and children can provide blood without great difficulty, and repeat sampling becomes more feasible. In addition, capillary blood from the same finger prick can be used to screen for anemia using a portable photometer (15
), and assays for C-reactive protein (10
), retinol (12
) and other markers of health or nutritional status can be performed using remaining spots of whole blood (11
).
Previous analyses have shown that investigators can attain the same degree of precision and reliability from spots of whole blood applied directly to filter paper that they expect from more standard methods of sample collection (11
). However, important sources of potential error should be considered. First, proper placement of whole blood on the filter paper is essential. The uniform absorbing properties of the filter paper will be defeated if blood is blotted or smeared onto the paper, or if a drop of blood is placed on top of a previously collected drop. In addition, the volume of whole blood applied to filter paper as a blood spot can influence the volume of serum contained within a single disc punched out of that spot (16
). For this reason, an attempt should be made to ensure that all blood spots used for analyses are of a comparable size. Variation in blood spot size can be minimized by collecting samples on filter papers with preprinted circles as guides (available from Schleicher & Schuell) to standardize the volume of whole blood collected from each individual. When filled to its border, each circle will contain 50 µL of whole blood.
Chromatographic effects, i.e., the uneven spread of blood across a filter paper spot, are an additional potential source of error. Although previous analyses have documented < 2% variation in the concentration of absorbed blood taken from various locations within a single blood spot (16
), specific analytes such as retinol (12
) and TfR (7
) have been shown to vary across a single spot. In our assay, we found TfR concentrations to be 12% higher in discs taken from the periphery of a 50-µL spot. Thus, we recommend that discs be punched out from the periphery of dried blood spots exclusively, and that center discs be avoided. At the very least, to avoid biased assay results, it is imperative that discs be removed in a consistent manner for all samples, standards, and controls within an assay to maximize comparability.
Finally, a recent evaluation of the reliability of hemoglobin assessment in capillary blood reported significant within-individual variation over four consecutive days of sampling, and when blood was drawn from the left compared with the right hand (17
). A similar evaluation of these potential sources of variation with respect to the measurement of TfR remains to be performed.
There are potential disadvantages to our blood spot TfR method that should be considered. First, current TfR reference values apply to plasma and serum, and whole blood references will differ substantially due to the diluting effects of erythrocytes. This problem can be partially overcome by comparing plasma and whole blood spot TfR concentrations for a large set of matched samples to derive a conversion formula that produces plasma equivalent values for blood spot TfR results. Our plasma-blood spot comparison indicates a high degree of correlation in TfR concentrations, but a much larger, and more diverse sample will be required to establish a reliable quantitative conversion factor. A better approach would be to develop TfR reference values specific to the whole blood spot assay, as has been done for current plasma assays (3
,18
).
Second, the blood spot TfR method is a relatively expensive measure of iron status at $10–12/sample for reagents and supplies. However, the minimal requirements for sample handling, storage and transportation make the blood spot assay less expensive than serum or plasma assays. In addition, the cost of the TfR assay is likely to decline in the future as its use becomes more widespread. We hope that this blood spot TfR method will provide investigators with additional options for future field-based studies of iron deficiency.
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FOOTNOTES |
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Manuscript received 1 August 2002. Initial review completed 18 August 2002. Revision accepted 16 September 2002.
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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1. Stoltzfus, R. (2001) Defining iron-deficiency anemia in public health terms: a time for reflection. J. Nutr. 131:565S-567S.
2. Skikne, B. S., Flowers, C. H. & Cook, J. D. (1990) Serum transferrin receptor: a quantitative measure of tissue iron deficiency. Blood 75:1870-1876.
3. Cook, J. D., Skikne, B. S. & Baynes, R. D. (1993) Serum transferrin receptor. Annu. Rev. Med. 44:63-74.[Medline]
4. Punnonen, K., Irjala, K. & Rajamaki, A. (1997) Serum transferrin receptor and its ratio to serum ferritin in the diagnosis of iron deficiency. Blood 89:1052-1057.
5. Cook, J. D. (1999) The measurement of serum transferrin receptor. Am. J. Med. Sci. 318:269-276.[Medline]
6. Verhoef, H., West, C. E., Ndeto, P., Burema, J., Beguin, Y. & Kok, F. J. (2001) Serum transferrin receptor concentration indicates increased erythropoiesis in Kenyan children with asymptomatic malaria. Am. J. Clin. Nutr. 74:767-775.
7. Cook, J. D., Flowers, C. H. & Skikne, B. S. (1998) An assessment of dried blood-spot technology for identifying iron deficiency. Blood 92:1807-1813.
8. Worthman, C. M. & Stallings, J. F. (1997) Hormone measures in finger-prick blood spot samples: new field methods for reproductive endocrinology. Am. J. Phys. Anthropol. 104:1-22.[Medline]
9. McDade, T., Stallings, J., Angold, A., Costello, E., Burleson, M., Cacioppo, J., Glaser, R. & Worthman, C. (2000) Epstein-Barr virus antibodies in whole blood spots: a minimally-invasive method for assessing an aspect of cell-mediated immunity. Psychosom. Med. 62:560-567.
10. McDade, T. W., Stallings, J. F. & Worthman, C. W. (2000) Culture change and stress in Western Samoan youth: methodological issues in the cross-cultural study of stress and immune function. Am. J. Hum. Biol. 12:792-802.[Medline]
11. Mei, J. V., Alexander, J. R., Adam, B. W. & Hannon, W. H. (2001) Use of filter paper for the collection and analysis of human whole blood specimens. J. Nutr. 131:1631S-1636S.
12. Erhardt, J. G., Craft, N. E., Heinrich, F. & Bielaski, H. K. (2002) Rapid and simple measurement of retinol in dried whole blood spots. J. Nutr. 132:318-321.
13. Flowers, C. H., Skikne, B. S., Covell, A. M. & Cook, J. D. (1989) The clinical measurement of serum transferrin receptor. J. Lab. Clin. Med. 114:368-377.[Medline]
14. Bland, J. M. & Altman, D. G. (1986) Statistical methods for assessing agreement between two methods of clinical measurement. Lancet i:307-310.
15. von Schenck, H., Falkensson, M. & Lundberg, B. (1986) Evaluation of "HemoCue," a new device for determining hemoglobin. Clin. Chem. 32:526-529.
16. Adam, B. W., Alexander, J. R., Smith, S. J., Chace, D. H., Loeber, J. G., Elvers, L. H. & Hannon, W. H. (2000) Recoveries of phenylalanine from two sets of dried-blood-spot reference materials: prediction from hematocrit, spot volume, and paper matrix. Clin. Chem. 46:126-128.
17. Morris, S. S., Ruel, M. T., Cohen, R. J., Dewey, K. G., de la Briere, B. & Hassan, M. N. (1999) Precision, accuracy, and reliability of hemoglobin assessment with use of capillary blood. Am. J. Clin. Nutr. 69:1243-1248.
18. Allen, J., Backstrom, K. R., Cooper, J. A., Cooper, M. C., Detwiler, T. C., Essex, D. W., Fritz, R. P., Means, R. T., Meier, P. B., Pearlman, S. R., Roitman-Johnson, B. & Seligman, P. A. (1998) Measurement of soluble transferrin receptor in serum of healthy adults. Clin. Chem. 44:35-39.
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