AMERICAN JOURNAL OF HUMAN BIOLOGY 26:1–9 (2014)

Human Biology Toolkit

Development and Validation of Assay Protocols for use with Dried Blood Spot Samples THOMAS W. MCDADE* Department of Anthropology and Institute for Policy Research, Northwestern University, Evanston, Illinois, 60208

ABSTRACT: Dried blood spots (DBS)—drops of capillary whole blood collected from finger stick—represent a minimally invasive alternative to venipuncture that facilitates the collection of blood samples from research participants in naturalistic, field-based research settings. But the number of validated assays for quantifying biomarkers in DBS samples is relatively low in comparison with serum or plasma. The objective of this review is to discuss the advantages and disadvantages of DBS sampling, and to outline the steps involved in developing and validating an immunoassay for application to DBS samples. These steps include deciding on reagents, preparing calibration and quality control material, evaluating elution protocols, optimizing sample quantity, and assessing multiple aspects of assay performance, including intra- and interassay variation, lower limit of detection, accuracy, stability, and agreement between results from matched DBS and plasma samples. The broader goal of this “how-to” approach is to encourage investigators to validate, implement, and disseminate assay protocols for DBS samples in order to advance field-based research on human C 2013 Wiley Periodicals, Inc. V biology. Am. J. Hum. Biol. 26:1–9, 2014.

Biological anthropology has an established track record of innovation with respect to methods for quantifying hormones, immune factors, and other biomarkers in diverse field settings all over the world (Ellison, 1988; James, 1991; McDade et al., 2000b; O’Connor et al., 2003; Salvante et al., 2012; Worthman and Stallings, 1997). Methodological innovation has been essential in advancing our understanding of the causes and consequences of human biological variation, and in complementing—and at times challenging— prevailing clinic- and lab-based research paradigms that inform our understanding of human biology and health. In particular, the measurement of biomarkers in blood, saliva, and urine allows us to study the pathways through which social, cultural, and other physical ecological factors “get under the skin” to shape physiological function and health over the life course (Finch et al., 2001; Panter-Brick and Worthman, 1999; Weinstein et al., 2007). The collection of plasma or serum (the liquid fraction of blood that remains after whole blood is centrifuged to remove red and white blood cells) is the current standard for biomarker measurement, but the costs, participant burden, and logistics associated with venipuncture blood collection are major barriers to community-based research, particularly in remote field settings where access to a centrifuge, freezer, or even electricity, may be limited. Saliva and urine are useful alternatives for the subset of biomarkers that enter these fluids in a measurable form (e.g., cortisol, estradiol), but this is not an option for the majority of analytes that are accessible only in blood. Dried blood spots (DBS)—drops of whole blood collected on filter paper following a simple finger stick—represent a low cost, “field-friendly” alternative that allows investigators to collect blood from large numbers of participants in naturalistic settings, and to integrate physiological information with rich contextual data in ways not possible with clinic-based research designs (McDade et al., 2007; Mei et al., 2001). Biological anthropologists have been using DBS overseas for 20 years (Campbell 1994; Worthman and Stallings, 1994, 1997), and recently more than 35,000 DBS C 2013 Wiley Periodicals, Inc. V

samples have been collected for research purposes in the US, including applications in large surveys like the National Longitudinal Study of Adolescent Health and the Health and Retirement Study (McDade, 2011). Since the majority of biomarker assays are designed for use with serum or plasma, relatively few assays have been developed and validated for DBS samples (for a list of previously validated assays, see McDade et al., 2007). The dearth of validated methods constrains the scientific questions that can be addressed by studies collecting DBS samples. The objective of this review, therefore, is to outline the steps involved in validating an immunoassay for application to DBS samples so field-based research on human biology can keep pace with developments emerging from clinic and lab-based approaches. The focus here is on the enzyme immunoassay due to its established utility for quantifying proteins with high sensitivity and specificity, the relatively low costs of instrumentation, and the commercial availability of assay kits and reagents (Lequin, 2005). However, many of the issues and procedures outlined below are relevant to other forms of immunoassay (e.g., radioimmunoassay, fluorometric immunoassay), as well as other methods for evaluating proteins (e.g., mass spectrometry) or molecular markers (e.g., mRNA, DNA) in DBS samples. ADVANTAGES AND DISADVANTAGES OF DRIED BLOOD SPOTS The filter papers (Whatman #903, GE Healthcare, Piscataway, NJ) used for DBS sampling were originally developed in the early 1960s to facilitate the collection of heel prick blood samples from newborns (Guthrie and *Correspondence to: Thomas Mcdade, Department of Anthropology, Northwestern University, Institute for Policy Research, Evanston, IL 60208, USA. E-mail: [email protected] Received 30 July 2013; Revision received 30 August 2013; Accepted 4 September 2013 DOI: 10.1002/ajhb.22463 Published online 15 October 2013 in Wiley Online Library (wileyonlinelibrary.com).

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Susi, 1963). These papers are often referred to as “Guthrie cards,” in recognition of Dr. Robert Guthrie’s pioneering efforts to develop convenient methods to screen for congenital metabolic disorders (Mei et al., 2001). Dried blood spots have played a central role in public health surveillance efforts in the US ever since, and as a result, the filter papers are certified to meet performance standards for sample absorption and lot-to-lot consistency. The CDC, which maintains an independent quality control monitoring program, notes that “The filter paper blood collection device has achieved the same level of precision and reproducibility that analytical scientists and clinicians have come to expect from standard methods of collecting blood, such as vacuum tubes and capillary pipettes” (Mei et al., 2001; p 1631). While this statement does not apply equally to all analytes in blood, it speaks to the confidence one can have in DBS-based results following successful assay validation. Advantages of DBS sampling for research applications include the following: 1. Sample collection is minimally-invasive and low cost. The participant’s finger is cleaned with isopropyl alcohol, and a sterile, single-use micro-lancet is used to deliver a controlled puncture (Table 1). Up to five drops of blood (50 ml per drop) are applied to filter paper, allowed to dry, and then stored at room temperature or refrigerated before shipment to the laboratory. Sample collection is relatively straightforward, but requires attention to important details that can affect DBS quantity and quality (Table 2). Supplies cost approximately $2/participant, and samples can be collected by non-medically trained interviewers in a participant’s home, or in some cases, by participants themselves. The ease of sample collection is a boon to field-based research, and is also advantageous in any setting for studies with infants, children, and the elderly, for whom venipuncture may be particularly problematic. The low burden of sampling also increases the feasibility of collecting multiple blood samples from the same individual over time (McDade et al., 2012a). 2. Requirements for sample processing are minimal. Unlike venipuncture sampling for serum or plasma, DBS samples do not need to be centrifuged, separated, or immediately frozen following collection. Whole blood is simply applied to filter paper, allowed to dry, and samples can be stacked and sealed in gas impermeable plastic bags for storage and shipment. A cold chain from the point of sample collection to receipt in the laboratory is not required since most analytes remain stable in DBS samples stored at room temperature for a week or more (McDade et al., 2007). However, it is

always advisable to refrigerate and freeze samples when possible to maximize future applications, and it is important to protect the samples from exposure to excessive heat. Since DBS samples are dried, requirements for shipping are less onerous in comparison with serum or plasma: Samples do not have to remain frozen in transit, dried blood poses a greatly reduced biohazard risk, and regulations associated with the shipment of DBS are greatly reduced. 3. A single finger prick can provide capillary whole blood for spots on filter paper, and for onsite assessments using “point-of-care” instruments. Relatively affordable, portable instruments for the analysis of hemoglobin (e.g., HemoCue), HbA1c (e.g., Bayer A1cNow, DCA Vantage), and lipid profiles (e.g., CardioChek, Cholestech) are currently available that provide an opportunity to collect physiological information in real time. Using the same finger prick sampling procedure detailed above, a drop of blood (or less) can be placed into one of these instruments, with subsequent drops applied to filter paper. By combining these procedures, biomarker results can be collected onsite and shared with participants, while DBS samples can be assayed in the lab for a broader range of analytes. In some cases this may provide a valuable health screening service, and act as an incentive for research participation. Advantages of DBS associated with sample collection in the field need to be weighed against potential disadvantages associated with quantification in the lab. 1. The volume of sample is very small. Whereas a typical venipuncture blood draw yields at least 5 ml of whole blood, and easily much more, a DBS sample with five large drops of blood contains approximately 250 ml—a volume of blood that is more than an order of magnitude smaller. Lancet selection (bigger is better) and adequate training/experience are key to collecting five large drops of blood (Table 2). Otherwise, only two or three drops will be obtained without a second finger stick. The constraint of small sample volume places a premium on the efficient use of DBS material, and may limit the number of analytes that can be quantified. Fortunately, improvements in assay sensitivity have reduced sample requirements for many analytes, and recent technological innovations are allowing for the simultaneous quantification of multiple biomarkers in a single volume of sample. For example, Luminex, Meso Scale Discovery, and Quansys Biosciences all offer multiplex immunoassay platforms (Chowdhury et al., 2009; Salvante et al., 2012; Skogstrand et al., 2005). Regardless, the number of biomarkers that can

TABLE 1. Protocol for collecting finger stick dried blood spot samples 1. Put on gloves and follow universal precautions for preventing transmission of bloodborne infections (http://www.cdc.gov/niosh/topics/bbp/ universal.html). 2. Clean the participant’s finger with isopropyl alcohol. Allow finger to dry. 3. Establish a firm grip on the participant’s hand and use your thumb and finger to create a taut surface on the skin of the middle or ring finger. 4. Use a sterile, single-use lancet to prick the finger just off the center of the tip of the finger. Immediately dispose of the lancet in a sharps container. 5. Wipe away the first drop of blood with a sterile gauze pad. Apply subsequent drops to filter paper (Whatman #903). Allow blood from the participant’s finger to be drawn onto the paper via capillary action. Do not blot the finger on the paper. 6. Collect at least two, and preferably five, drops of blood that fill the borders of the pre-printed circles on the filter paper (approximately 50 ml each). 7. After collection, place a bandage on the participant’s finger. 8. Allow filter papers to air dry for at least 4 h (or overnight). Do not stack the samples until they are dry. 9. Stack filter papers and place in an airtight container (e.g., sealable plastic bag or container) with desiccant.

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T.W. MCDADE TABLE 2. Tips for successful collection of finger stick dried blood spot samples

1. Heavy items should not be placed on top of the filter papers before they are used. This will compress the papers and prevent them from absorbing blood evenly. The papers should be transported in plastic containers or boxes for protection. After the samples are collected and the blood has dried, compression is not a concern and papers can be transported in sealable plastic bags. 2. Larger lancets with blades, rather than needles, produce better blood flow, particularly when individuals have calloused hands. The lancets typically used by diabetics to monitor blood glucose have small needles designed for frequent sampling, and will produce small volumes of blood. 3. If the participant’s hands are cold, rub the hand to warm it up and increase blood flow. Heating pads, or soaking in warm water, can also be used to warm hands. It may also be helpful to ask the participant to stand, to make a fist, and/or to swing the hand rapidly downward before the finger stick to try and move blood into the fingers. 4. After pricking the finger, frequently use the gauze pad to wipe away blood from the puncture site. If blood is allowed to stay at the puncture site it will begin to clot, and blood flow will stop. 5. If blood flow is not sufficient, use your fingers to squeeze the finger lightly or to try to move blood from the base of the finger to the tip. Avoid excessive squeezing or “milking” of the finger as this will dilute the sample. If blood flow is still not sufficient, the skin puncture may be repeated on another finger if the participant is willing. 6. It is absolutely essential that blood from the participant’s finger be drawn onto the paper via capillary action. The finger should never touch the paper. Blotting blood onto the paper will prevent the uniform diffusion of blood across the paper, which is essential to quantification. If samples are collected correctly, the spots should look identical on both sides of the filter paper once the spot has dried. If one side is larger than the other, or if borders are irregular, then the blood was blotted or smeared on the paper. 7. Do not place blood on top of blood. Once blood has been spotted on the card, another drop of blood should not be placed on top of it, even if the previous spot seems small. This will concentrate the sample and defeat the uniform diffusing properties of the paper. 8. If blood is not flowing freely, and five full drops cannot be collected, it is almost always better to collect one or two large drops of blood than four or five small drops. Uniform drops that fill the borders of the pre-printed circles on the #903 papers will provide more accurate results, and allow for the more efficient and flexible use of a limited quantity of sample.

be quantified in a given DBS sample is likely to be reduced in comparison with venipuncture-based approaches. 2. Some analytes cannot be measured in DBS. Although the default assumption should be that anything that can be measured in serum or plasma can also be quantified in DBS samples, there are potential obstacles that may prove to be insurmountable. For example, the presence of red and/or white blood cells may interfere with some assays. When whole blood dries on the filter paper, cellular fractions rupture and their contents are subsequently released into solution when DBS samples are reconstituted. Different assay systems and specific analytes will vary in their sensitivity to potential interference. This is not a common problem, but in some cases (e.g., ferritin; Ahluwalia, 1998) it may prevent accurate quantification. In addition, analytes in DBS have to come off the filter paper and enter solution in a form suitable for analysis. While whole blood on filter paper is relatively stable and easy to store and transport, the process of drying may alter the biochemical structure of a molecule and affect the efficiency with which analytes enter solution. Lastly, analytes have to be present in quantities sufficient for analysis. Even with improvements in immunoassay technology that reduce requirements for sample volume, there are limits. Serum- or plasmabased assays that require large volumes of undiluted sample (e.g., 50 or 100 ml) are not likely to translate easily to use with DBS samples. (Plasma differs from serum in containing clotting factors, but for most immunoassay applications both sample types provide similar results; for the sake of simplicity in the remainder of the article, I refer to plasma only). 3. DBS samples are not the clinical standard. With the exception of newborn screening, DBS sampling is rarely applied clinically and values from the analysis of venipuncture samples are considered the gold standard. Results from DBS samples represent the concentration of an analyte in whole blood, which by definition will differ from results determined in

plasma. But the correlations across matched plasma and DBS samples are typically very high, and correction factors can be applied to DBS values to generate plasma equivalents if desired. Such corrections will not be necessary for within-study comparisons, but will be important for any attempts to compare data to prior research based on results from plasma samples, or to make use of established clinical cut-points. These issues relate to the broader point that clinical chemistry is not oriented toward the analysis of DBS samples, and in some contexts this may pose additional challenges. Some of the disadvantages of DBS can be overcome by alternative methods for collecting and transporting capillary blood, which produce plasma for analysis in the lab. Whole blood from a finger stick can be collected in a capillary tube (a thin tube open on both ends) or a microtainer device (a small vial with a screw top). Like DBS, the burden of sample collection is low, but handling and transport are complicated by the fact that the samples remain in a liquid state: a cold chain is typically needed to maintain the integrity of the sample, the blood has to be centrifuged and separated before analysis or freezing, and the transportation of liquid blood poses greater biohazard risk. Serum separator cards are similar to DBS cards in terms of sample collection, but differ in using a diffusion gradient or membrane that isolates red and white blood cells while capturing and drying serum in the filter paper. The device therefore retains the advantages of traditional DBS sampling, with the added benefit of providing a serum sample that better approximates the clinical standard for most analytes. However, serum separator cards are more costly, and they pose challenges to precise quantification when concentration gradients are present across the cards. PRINCIPLES OF ENZYME IMMUNOASSAY The enzyme immunoassay can be applied in many forms, all of which involve a highly specific interaction between antibody and the target of interest (antigen), and American Journal of Human Biology

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an enzymatic label that changes in color in proportion to the quantity of antigen in the sample (Crowther, 2009). The enzyme-linked immunosorbent assay (ELISA) is a specific application that uses a 96-well microtiter plate as the solid phase, where each well is coated with a capture antibody specific to the target. Immunoassay principles are comparable regardless of sample type. But for the purposes of clarity, and to define terms that will be used below in the discussion of assay development and validation, it may be helpful to present a brief example. To quantify C-reactive protein (CRP) in DBS (Brindle et al., 2010; McDade et al., 2004), eluted sample is added to identified wells in the assay plate, where CRP binds to the anti-CRP antibody that has already been coated to the bottom of the well. The CRP in the sample is therefore “captured” by the plate, and remains bound while the rest of the sample is removed through a series of wash steps. In a sandwich ELISA, a second antibody is then added which binds to available epitopes on the CRP molecule, the plate is washed to remove excess antibody, and a CRP “sandwich” remains bound to the bottom of each well. The second antibody—or in this case, the detection antibody— has been previously conjugated to an enzyme that catalyzes a color change following the addition of the enzyme’s substrate (in some cases the second antibody may be conjugated to biotin, which in turn binds to streptavidin which has been conjugated to the enzyme). The intensity of color change is directly proportional to the concentration of CRP in each well, and it can be precisely measured using a microplate absorbance reader. The amount of color change specific to bound CRP is also referred to as “signal,” whereas nonspecific color change is considered “background” or “noise.” The concentration of CRP in each sample is determined by comparing the amount of color change in samples (unknowns) with the amount of color change generated by material with known CRP concentration (calibrators, or standards). ISSUES TO CONSIDER BEFORE DBS ASSAY DEVELOPMENT In most cases, the process of developing an immunoassay for use with DBS samples is relatively straightforward under the following conditions: (1) prior work has demonstrated that the analyte of interest can be quantified by immunoassay in serum or plasma; (2) the presence of lysed red and/or white blood cells does not interfere with quantification; and (3) personnel involved with assay development have access to the necessary laboratory equipment as well as prior experience with immunoassay implementation and troubleshooting. The immunoassay is a truly remarkable tool for gaining insight into key aspects of human biology, but there are technological limits and tradeoffs in assay design. The following issues may be useful to consider as one evaluates different assay approaches for specific applications. ASSAY PERFORMANCE AND RANGE OF MEASUREMENT In order to convince ourselves, and our colleagues, that a DBS protocol produces valid results we need to demonstrate a reasonable level of accuracy, precision, and reliability in our laboratory measurements. Operational definitions of these performance parameters are provided below, but at this point it is sufficient to say that measurement error is a reality, and the level of error will American Journal of Human Biology

vary across analytes and across measurement protocols. It is therefore essential to identity sources of error, to minimize their impact to the extent possible, and to consider acceptable margins of error for particular applications. In addition, it is important to consider the range of values that can be quantified, and the level of accuracy, precision, and reliability across this range. In other words, how low can the assay go? How high? The immunoassay’s enzyme signal has a limited dynamic range, and attempts to optimize quantification at very low analyte concentrations will inevitably constrain quantification at higher concentrations. Determining whether the assay range corresponds to the range of expected values in a given study is therefore an important aspect of assay design. LOWER LIMIT OF DETECTION LIMIT VERSUS SAMPLE QUANTITY When the relevant range of measurement includes values that approach zero, the use of DBS samples poses additional challenges. Using a larger quantity of sample in an assay is usually an effective way to improve the lower limit of detection (LLD) and enhance precision and reliability at the low end of the assay range, but DBS samples do not provide much material, and we often want to conserve sample for other analyses. A typical drop of blood will contain approximately 50 ml of whole blood, and will yield seven 3.2 mm (1/8 in) discs of blood for laboratory analyses. A full card of five blood spots will therefore contain enough sample for 35 analyses requiring one 3.2 mm disc, 17 analyses requiring two discs, 11 analyses requiring three discs, etc. However, in practice, five perfect blood spots are rarely obtained, and sufficient sample for 10 to 20 3.2 mm discs is a more reasonable expectation for a single finger prick. Given the tradeoff between assay performance and sample volume, DBS assays typically strive to use the smallest amount of material possible, but need to evaluate the extent to which using additional material improves the LLD and other aspects of assay performance. ASSAY SUPPLIES: OFF-THE-SHELF OR DO-IT-YOURSELF? Commercially available immunoassay kits—typically designed for serum or plasma—can often be adapted for use with DBS samples. Advantages of these kits are as follows: (1) all necessary supplies (e.g., coated plates, detection antibody, calibrators, and sample/wash buffers) are typically included; (2) kit components are known to work well together (e.g., capture and detection antibodies pair effectively, enzyme and substrate are complementary, buffers do not interfere with signal development); and (3) commercial availability increases the likelihood of protocol implementation in other labs, and comparability of results across labs. Disadvantages include the relatively high cost of immunoassay kits ($150 to $1,000 per kit, each of which can analyze 40 samples in duplicate), the possibility that production of a particular kit may end without notice, and the lack of flexibility in modifying important aspects of the protocol. For example, kits typically provide an immunoassay plate that is precoated with capture antibody. The convenience of a precoated plate—almost certainly optimized for use with plasma— precludes increasing the concentration of coating

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antibody, which can be an effective way to increase signal when small quantities of sample are used, as with DBS. An alternative do-it-yourself approach involves sourcing antibodies and calibration material, immunoassay plates, and chemicals for sample/wash buffers directly from various suppliers. The cost savings of designing an assay from scratch are substantial in terms of materials, but these savings should be weighed against the time required to assemble the necessary components (e.g., make buffers, coat plates with antibody), and the extensive time required up front to evaluate components during assay development. In addition, a higher level of technical expertise and immunoassay experience is required to develop protocols from scratch. INSTRUMENTATION AND EQUIPMENT There are no special instrumentation requirements for DBS assays, and any lab that is equipped to implement immunoassay analysis of serum, plasma, saliva, or urine should have the capacity to implement DBS assays. The only exception is the requirement for a hole punch to remove sample from the DBS card, which can be purchased from office supply stores. Two highly recommended pieces of equipment, which are often but not always available in the immunoassay lab, include an orbital plate shaker and a microplate washer. Most immunoassays include extended periods of incubation, and increasing incubation times and/or rotating assay plates during incubation are often effective ways to increase performance of DBS assays. In addition, microplate washers automate the process of washing plate wells between assay steps (e.g., after sample incubation and before addition of detection antibody), which is essential for proper color development and to reduce background noise. Manual washing is possible, but it is timeconsuming and generally introduces more variation across wells that can reduce assay precision and reliability. Regardless of sample type, enzyme immunoassays require an absorbance microplate reader to quantify the amount of color change that is proportional to analyte concentration. Basic models that provide accurate endpoint readings are sufficient for the vast majority of applications (e.g., BioTek ELx808, Molecular Devices EMax), but more advanced (and more expensive) multimode models (e.g., BioTek Synergy, Molecular Devices SpetraMax) provide the flexibility to use fluorescence and luminescence, in addition to absorbance, as modes of detection. If prior work has shown that the analyte of interest can be quantified in plasma at better resolution using modes of detection other than absorption, then it may be worth applying the same mode to protocols designed for DBS samples. SIX STEPS TO DEVELOPING AN IMMUNOASSAY FOR DBS Recommended procedures for assay development and validation are described here in six steps, although it should be emphasized that the process is iterative. Results from step 5, for example, might lead to a reconsideration of procedures evaluated in steps 3 or 4. Flexibility, creativity, and patience early in the process will pay dividends later by preventing one from getting locked into a protocol that could have been better.

Step 1. Decide on reagents The range of options for commercially available immunoassay kits will vary widely by biomarker, and the first key decision involves whether to consider these kits or to pursue a do-it-yourself protocol. The following factors are useful to keep in mind when evaluating potential kit options: cost, required volume of sample for the serum/ plasma protocol, and key aspects of assay performance as evaluated by the manufacturer, if available (e.g., lower limit of detection, antibody cross-reactivity, dynamic range). If a clear choice does not emerge, it is probably worth evaluating two or more kits since it is not always easy to predict which kits will perform best when applied to DBS samples (customer service representatives can often be convinced to provide a free or discounted kit for evaluation). A major determinant of final assay performance is the affinity and specificity of the antibodies selected, and it cannot be assumed that antibodies that work well with plasma will function similarly in the more complex sample matrix of DBS. Antibodies will differ across kits, and this will be a major determinant of kit performance. When pursuing a do-it-yourself approach, the selection of effective capture and detection antibodies is critical, and it is worth making the initial investment to evaluate multiple potential combinations. Step 2. Prepare calibration and quality control material In order to minimize matrix differences and maximize comparability between calibrators and unknowns, DBS calibration material should be manufactured. This material is comprised of a known concentration of the analyte of interest in a plasma-like matrix (for examples see McDade et al., 2004, b; McDade and Shell-Duncan, 2002; Miller and McDade, 2012; Tanner and McDade, 2007), added to an equal volume of washed red blood cells to approximate whole blood with a hematocrit of 50%. Washed erythrocytes are obtained as follows: (1) collect whole blood by venipuncture in EDTA vacutainer tubes, and centrifuge at 1,500g for 15 min; (2) remove plasma and buffy coat with a Pasteur or other pipet; (3) add an approximately equal volume of normal saline (0.86 g NaCl/100 ml deionized H2O) to the packed red blood cells; (4) mix gently with inversion for 5 min; and (5) centrifuge as before. Remove saline and any remaining buffy coat, and repeat steps 3 to 5 for a total of three washes. The supernatant should be clear after each centrifugation; a pinkish tone indicates hemolysis of red blood cells, in which case the centrifugal force should be reduced. Remove saline following the final wash so only red blood cells remain. Calibrator dilutions should be prepared during the final wash steps. Calibration material is typically supplied in a concentrated form, but occasionally immunoassay kits provide prediluted calibrators representing concentrations across the range of measurement. When using concentrated material, calibrators should first be diluted in a plasma-like matrix to known concentrations across the desired range of measurement. Each concentration of calibrator is then added to an equal volume of washed erythrocytes (1:2 dilution). Mix gently with inversion for 5 min, then apply each calibrator to labeled filter paper cards in 50 ml drops using a manual pipette. Dry overnight at room temperature. Calibrators should then be stored at 230 C American Journal of Human Biology

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or lower in gas impermeable plastic bags with desiccant. DBS-based control samples—to be used for assay validation or to monitor between-assay variation—can also be manufactured using these procedures. Step 3. Evaluate elution protocols Since the blood contained in a DBS sample has been dried on filter paper, analytes must be brought into solution before analysis. This is a key step in the use of DBS samples. A standard hole punch is typically used to cut out discs of whole blood of uniform size, and one or more discs are placed into an elution buffer for a fixed amount of time. In effect, the dried blood spot is reconstituted as hemolyzed liquid whole blood, and then transferred to the assay plate where the protocol proceeds just as it would with plasma. Three variables should be considered in designing an elution protocol: (1) type of elution buffer; (2) duration and temperature of elution; and (3) whether to mix during elution. Phosphate-buffered saline is a good place to start for an elution buffer, and in some applications results may be improved with the addition of protein (e.g., bovine serum albumin), polysorbate 20 (i.e., Tween 20, a surfactant), or protease inhibitor. The addition of protein or Tween 20 may improve assay signal by stabilizing proteins as they come into solution, and by blocking nonspecific binding sites in the assay plate. When using a commercial kit it is a good idea to evaluate buffers provided with the kit (e.g., sample dilution buffer) as potential elution buffers. This approach has the advantage that the buffer is already provided, and it is known to perform well with other components of the kit. Its effectiveness as an elution buffer, however, is uncertain and should be evaluated against potential alternatives. Examples of elution buffers and protocols can be found in previously validated methods (Dowd et al., 2011; McDade et al., 2000a, 2004, 2012b; McDade and Shell-Duncan, 2002; Miller and McDade, 2012; Shirtcliff et al., 2001; Tanner and McDade, 2007). In all of the assays I have developed I apply overnight incubations during elution. Shorter elution periods are possible (e.g., 2 h, 4 h), but I have found that overnight incubations maximize elution efficiency and improve work flow in the lab. For example, punching out DBS samples in the morning, eluting for 2 or 4 h, and then implementing an immunoassay protocol (which typically takes 3 or 4 h) makes for a very long day. With overnight elution, samples punched out the previous day can be transferred to the plate first thing in the morning to start the assay, and DBS samples for the next day’s assay can be punched out during incubations later in the day. A potential problem associated with longer elution periods is sample degradation. A great advantage of DBS is that the paper matrix stabilizes the blood sample, but when analytes are brought into solution they are more susceptible to degradation. The extent to which degradation occurs during elution will vary across analytes. Incubating samples at 4 C during elution may prevent or attenuate degradation. In general, incubating at room temperature or higher (e.g., 37 C), and adding mixing (end-over-end, or orbital) during elution will reduce elution times, but may increase rates of sample degradation. In evaluating different elution protocols the efficiency of elution can be formally calculated by dividing the conAmerican Journal of Human Biology

centration of analyte recovered from whole blood eluted from filter paper by the concentration of analyte obtained from the same quantity of hemolyzed whole blood in liquid form. Alternatively, results from different elution protocols can be compared directly to determine which method provides the best signal:noise ratio in the assay. Step 4. Optimize the quantity of sample Hole punches for cutting out uniform discs of dried blood come in various sizes, and a larger hole punch is an obvious way to introduce more sample into an assay. But I have used a relatively small 3.2 mm hole punch in prior work because there is more flexibility in removing sample from smaller or irregularly shaped drops of whole blood on the card. Assays developed so far have required 1 to 8 discs of whole blood for duplicate measures, and increasing the amount of material is typically an effective way to improve the lower limit of detection, as well as precision and reliability at the low end of the assay range. The goal of improved assay performance, therefore, is often traded off against limited quantity of sample and the desire to preserve sample for other analyses. The impact of sample quantity on assay performance can be evaluated simply by comparing results when using, for example, 1 versus 2 discs in a fixed amount of elution buffer. More discs should yield higher signal, but there is often a clear point of diminishing returns, both in terms of assay signal as well as use of sample. Step 5. Optimize the assay protocol Standard immunoassay protocols involve a series of time-standardized incubations that facilitate antigenantibody interactions in samples and calibrators. Again, duration of incubations (e.g., increasing from 1 to 2 h or overnight), temperature during incubation (4 C, RT, or 37 C), and mixing (e.g., incubation on plateshaker vs. benchtop) are all variables that can be manipulated to optimize assay performance. Extending incubation times after DBS eluates are added to the assay plate, and using an orbital plateshaker to rotate the assay plate during incubation, can be effective ways to increase signal. When using a do-it-yourself protocol, determining the optimal concentration of capture antibody and detection antibody is an important early step in assay development. Higher antibody concentrations typically increase assay signal, but at a cost to higher non-specific background noise. Checkerboard titrations (Crowther, 2009) can be particularly useful at this point because they allow one to optimize two or three assay components simultaneously. For example, a 96-well plate can be prepared to compare different concentrations of capture antibody (e.g., 2 mg/ml, 10 mg/ml, 20 mg/ml, each set up in different columns on the assay plate) and different concentrations of detection antibody (e.g., 1 ng/ml, 5 ng/ml, 10 ng/ml, set up in different rows on the assay plate). The “checkerboard” allows one to evaluate all potential combinations of capture and detection antibody concentrations. By setting up the checkerboard twice, once on each half of the 96-well plate, one can determine the optimal signal:noise ratio that is obtained with different quantities of DBS sample across antibody concentrations (e.g., eluate from 1 disc on the left side of the plate, eluate from 2 discs on the right side of the plate).

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Step 6. Evaluate assay performance Once the protocol is set and the assay optimized, it is time to evaluate the assay for precision, reliability, accuracy/recovery, lower limit of detection, and agreement with plasma results (Nexo et al., 2000; Vikelsoe et al., 1974). Quantitative assessment of these aspects of assay performance is important for identifying sources and magnitudes of measurement error that may inform study design, the interpretation of assay results, and comparability with prior research. INTRA-ASSAY VARIATION The level of imprecision of an assay can be estimated by calculating the coefficient of variation (%CV; 100 3 standard deviation/mean) of multiple determinations (usually 10 or more) of a single sample, all measured in a single assay. This is typically done with two or three samples across the range of measurable values since imprecision may vary, and is often higher at lower analyte concentrations. Repeating this analysis across multiple runs will provide a better estimate of intra-assay variation. INTERASSAY VARIATION The day-to-day variation, or reliability, of a method can be estimated by calculating the CV for multiple determinations of a single sample measured on different days. As with precision, reliability should be assessed using two or three samples with values that span the assay range. Acceptable levels of intra- and inter-assay variation will vary across assays and particular applications, but %CV 0.95) between DBS and gold standard, venous blood results. But in some cases investigators may have to settle for less, and the advantages associated with DBS in the field will have to be weighed against lower correlation with the gold standard. Interference from cellular material in DBS samples, inconsistent or improper application of blood to the filter paper, the process of drying and reconstituting the sample, the application of different antibodies across DBS and plasma protocols, and metabolic/circulatory dynamics can all contribute to differences in results based on the analysis of finger stick DBS samples versus venous blood. Analysis of matched DBS and plasma samples can also be used to generate a conversion formula to derive plasma-equivalent values from results based on DBS samples (Worthman and Stallings, 1997). Converted values may be particularly useful for comparisons with prior research, and for the application of clinically-relevant cutoff values (it should also be noted that converted values can be used in the Bland-Altman analyses described above, in which case the difference across methods, rather than the ratio, should be plotted on the y-axis). However, caution should be used in the application of converted values, since the relationship between DBS and plasma American Journal of Human Biology

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DEVELOPMENT AND VALIDATION OF ASSAY PROTOCOLS

values will vary across analytic methods, and may also vary across populations (e.g., Shirtcliff et al., 2001). If conversion to plasma equivalent values is a priority for a particular study, it is worth dedicating time and resources to develop a study-specific conversion formula based on DBS samples that are collected, handled, and analyzed in exactly the same way as participant samples (Buxton et al., under review).

benchtop at room temperature for 1 h, and then returned to the freezer. This protocol should be repeated over 5 different days. Baseline and freeze/thaw samples should then be analyzed together to inspect for evidence of degradation relative to samples that have never been thawed. The paper matrix of DBS seems to provide considerable protection against freeze/thaw degradation, but it is worth evaluating this possibility nonetheless.

STABILITY

CONCLUSIONS

A major advantage of DBS sampling is the ability to collect blood in field settings, and to store and transport samples without refrigeration or freezing. There are, however, limits to the stability of analytes in filter paper stored at ambient temperatures. These limits should be determined empirically for each analyte before sample collection, based on the range of conditions DBS samples may be exposed to during sample collection and transport (McDade et al., 2004; Worthman and Stallings, 1997). In prior work I have used the following protocol to analyze stability across a range of potential conditions. First, collect at least three sets of DBS samples, representing low, mid, and high levels of analyte concentration. At this point it is important to calculate the quantity of sample needed for the evaluation, which will vary depending on the number of stability parameters being considered, and the quantity of sample required for the assay. If large volumes of sample are required it may not be feasible to collect finger stick DBS samples; instead, collect a larger volume of whole blood via venipuncture, and then use a pipette to transfer whole blood to the filter paper. Second, after samples are allowed to dry, place one set of each into the freezer (230 C or lower). These samples will be analyzed to represent “baseline” concentrations of the analyte of interest. Third, expose samples to one of three temperature conditions (4 C, room temperature, 37 C) and one oscillating condition (12 h at 32 C and 12 h at 22 C to represent ambient conditions in tropical environments), in the presence of desiccant in a gas impermeable bag, for varying lengths of time up to 4 weeks (humidity is also a parameter that may affect analyte stability, but it can largely be eliminated as a factor in the field by storing cards in sealed bags before their use, and by placing DBS samples in sealed bags with desiccant after they have dried). Samples should be stored in the same freezer as the baseline samples after their exposure time is completed. Baseline and exposed samples should then be analyzed together, on the same assay plate if possible, to maximize comparability. Samples can be considered stable so long as values remain within a 10% CV range of the initial baseline values. In most cases there will be a clear linear trend toward degradation after a certain amount of time, particularly at elevated storage temperatures. It is also useful to evaluate the stability of analytes in DBS following repeated cycles of freezing and thawing. Since the same set of samples is often used for multiple analyses, it is possible that removing samples from the freezer during assay set up may lead to degradation that adversely affects subsequent analyses. As recommended above, three or more samples across the assay range should be used, with baseline samples stored in the freezer. Freeze/thaw samples should be removed from the freezer, taken out of their plastic bags, placed on the

Research into the causes and consequences of global human biological variation encourages investigators to collect data in the field, while studies of biological mechanisms tether them to the lab. “Field-friendly” methods like DBS sampling provide tools that bridge this gap, and encourage an epistemological shift that reframes the study of human biology as a holistic, integrative endeavor (Stinson et al., 2012). By bringing our methods to research participants in the community, rather than relying on select individuals willing to come to the clinic or lab, we are in a much stronger position to obtain generalizable results, to link data across levels of analysis, and to foreground contextual factors that are important determinants of human physiological function and health across the life course. Finger stick DBS sampling has become an important part of the human biology toolkit, with applications in a growing number of studies in the U.S. and internationally (McDade et al., 2007). The same advantages that fostered the development of DBS sampling for large-scale neonatal screening programs—low costs and burdens of blood collection, stability of analytes on filter paper, simplified logistics associated with sample handling and transport— greatly facilitate blood collection in field-based settings, with participants of all ages. Convenience in the field, however, is traded off against challenges associated with quantification in the lab. In most cases investigators can expect results that are comparable to those obtained with gold standard, venous blood-based methods, but only after investing considerable effort in assay development and validation. My hope is that this review will encourage more investigators to go down this path, and to develop and disseminate DBS protocols that advance research into human biological variation around the world.

American Journal of Human Biology

USEFUL RESOURCES SUPPLIES FOR COLLECTING DBS SAMPLES  #903 filter papers are manufactured by Whatman, and produced in multiple formats (http://www.whatman. com/903ProteinSaverCards.aspx). The “Protein Saver Card” (#10534612) is compact, and has a flap to cover the sample once it is dry.  Lancets come in a variety of styles and sizes. The BD Microtainer contact-activated lancet contains a blade (rather than a needle) and produces good blood flow (#366594).  Desiccant comes in many forms. Self-contained pouches with color-changing indicator are useful for determining when the desiccant is no longer effective (e.g., VWR Humidity Sponge, Indicating, #61161-319)

T.W. MCDADE

ADDITIONAL INFORMATION ON DBS SAMPLING  For more information on the use of DBS in newborn screening programs: http://www.cdc.gov/labstandards/ nsqap.html  For guidance on regulations associated with shipping DBS samples: http://www.cdc.gov/labstandards/pdf/nsqap/ Bloodspot_Transportation_Guidelines.pdf  For information from Whatman on assessing blood spot quality: http://www.whatman.com/References/Simple%20 Spot%20Check%20Literature%20Piece%20LR%20FINAL %2011.02.09.pdf  For useful information and videos associated with the application of DBS sampling, see the website developed by the Biomarker Network, sponsored by the National Institute of Aging: http://gero.usc.edu/CBPH/ network/index.shtml LITERATURE CITED Ahluwalia N. 1998. Spot ferritin assay for serum samples dried on filter paper. Am J Clin Nutr 67:88–92. Bland JM, Altman DG. 1986. Statistical methods for assessing agreement between two methods of clinical measurement. Lancet i:307–310. Bland JM, Altman DG. 1999. Measuring agreement in method comparison studies. Stat Methods Med Res 8:135–160. Brindle E, Fujita M, Shofer J, O’Connor KA. 2010. Serum, plasma, and dried blood spot high-sensitivity C-reactive protein enzyme immunoassay for population research. J Immunol Methods 362:112–120. Campbell KL. 1994. Blood, urine, saliva and dip-sticks: Experiences in Africa, New Guinea, and Boston. Ann NY Acad Sci 709:312–330. Chowdhury F, Williams A, Johnson P. 2009. Validation and comparison of two multiplex technologies, Luminex and Mesoscale Discovery, for human cytokine profiling. J Immunol Methods 340:55–64. Crowther JR. 2009. The ELISA handbook. New York: Humana Press. Dowd JB, Aiello AE, Chyu L, Huang YY, McDade TW. 2011. Cytomegalovirus antibodies in dried blood spots: a minimally invasive method for assessing stress, immune function, and aging. Immun Ageing 8:3. Ellison P. 1988. Human salivary steroids: Methodological considerations and applications in physical anthropology. Yearb Phys Anthropol 31: 115–142. Finch CE, Vaupel JW, Kinsella KG, editors. 2001. Cells and surveys : Should biological measures be included in social science research? Washington, D.C.: National Academy Press. p 374. Guthrie R, Susi A. 1963. A simple phenylalanine method for detecting phenylketonuria in large populations of newborn infants. Pediatrics 32: 338–43. James GD. 1991. Blood pressure response to the daily stressors of urban environments: Methodology, basic concepts, and significance. Yearb Phys Anthropol 34:189–210. Lequin RM. 2005. Enzyme immunoassay (EIA)/enzyme-linked immunosorbent assay (ELISA). Clin Chem 51:2415–2418. McDade T, Stallings J, Angold A, Costello E, Burleson M, Cacioppo J, Glaser R, Worthman C. 2000a. Epstein-Barr virus antibodies in whole blood spots: A minimally-invasive method for assessing an aspect of cellmediated immunity. Psychosom Med 62:560–567. McDade TW. 2011. The state and future of blood-based biomarkers in the Health and Retirement Study. Forum Health Econ Policy 14:5.

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