Abstracts of Oak Ridge Posters - Clinical Chemistry

Abstracts of Oak Ridge Posters Factors Affecting Circulating mRNA for Nephrin, Emanuela Orlandi,1 Asif Butt,1 David Goldsmith,2 and R. Swaminathan1* (Departments of 1 Chemical Pathology and 2 Nephrology, Guy’s & St. Thomas’ Hospital, London, United Kingdom; * address correspondence to this author at: Department of Chemical Pathology, 5th Floor, North Wing, St. Thomas’ Hospital, London SE1 7EH, UK; fax 44-207-9284226, e-mail [email protected]) Kidney disease affects more than 20 million individuals in the United States alone. Although the causes of kidney failure are diverse, the glomerular filtration barrier is often the target of injury. Identification of the type of disease is important for targeting therapy. One area of particular interest is the ability to predict which patients with nephrotic syndrome might respond to steroids or cytotoxic therapies and which will not (1 ). Often this depends on renal biopsy, an invasive procedure with some risk of complications. A noninvasive test to determine the type of renal disease would be advantageous. During the past decade, nucleic acids have been detected in the circulation, and measurements of tissuespecific DNA and mRNA offer enormous potential for diagnosis and prognosis (2 ). Increased amounts of nucleic acids have been reported in disease, including lung cancer (3 ), thyroid cancer (4, 5 ), stroke (6 ), and trauma (7 ). We have recently shown that measurement of total DNA in plasma is a good prognostic marker of outcome in critically ill patients (8 ). In another study we observed that mRNA for rhodopsin is increased in diabetic retinopathy, and it may be a useful and inexpensive way to detect and monitor diabetic patients for retinopathy (9 ). Nephrin, a transmembrane protein with a large extracellular portion including 8 immunoglobulin-like domains, is expressed by visceral epithelial cells (podocytes) in the slit diaphragm of the glomerulus (10 ). A product of the NPHS1 gene located on chromosome 19, this protein is crucial for the integrity of the slit diaphragm, and abnormalities in this protein can lead to proteinuria and nephrotic syndrome. Measurement of nephrin mRNA in peripheral blood by real-time quantitative PCR (qPCR) may provide a clue to the etiology of the renal disease. In the present study, we developed a real-time qPCR assay for nephrin mRNA and used this assay to measure the concentrations in healthy individuals and in renal transplantation patients to determine the factors that may be associated with the observed nephrin mRNA concentrations in peripheral blood. Blood samples were collected from healthy volunteers (n ⫽ 14) with no known disease and from patients, after kidney transplantation (n ⫽ 53), from the transplant clinic at Guy’s Hospital, London. The protocol for the study was approved by the Guy’s Hospital Local Research Ethics Committee. All patients had stable renal function, and the period after transplantation ranged from months to ⬎10 years. Informed consent was obtained from each participant before blood collection. Peripheral vein blood (2.5 mL) was drawn directly into PAXgeneTM Blood RNA tubes specifically designed for the collection and stabili-

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Clinical Chemistry 51, No. 10, 2005

zation of RNA from whole blood (PreAnalytiX). Wholeblood RNA was extracted with the PAXgene Blood RNA Kit and treated with the supplied DNase to prevent genomic DNA contamination, strictly according to manufacturer’s instructions. Extracted RNA was stored at ⫺80 °C until required for cDNA synthesis. Reverse transcription was carried out with SuperScript IITM reverse transcriptase according to the manufacturer’s instructions (Invitrogen Life Sciences). The cDNA generated was stored at ⫺80 °C until required for quantification. Separately, samples were also subjected to the above procedure with the exception that SuperScript II was replaced with water (negative control). cDNA was amplified and PCR products were detected by use of sequence-specific oligonucleotide probes and intron-spanning specific primers on the ABI 7000 Sequence Detection System (PE Applied Biosystems). ␤-Actin cDNA was amplified by the manufacturer’s method (Pre-Developed Assay Reagents for the TaqMan® assay; PE Applied Biosystems). For the TaqMan nephrin assay, 300 nM each of the forward and reverse primers, 100 nM probe, 25 ␮L of 2⫻ TaqMan Universal Master Mix, and 10 ␮L of cDNA sample were used for each reaction. For both assays, calibrators and samples were analyzed in duplicate in a final reaction volume of 50 ␮L. Calibration curves were prepared from serial dilutions of cDNA (Ambion Corp.) obtained from healthy human kidney. A water blank was also incorporated in each run for the respective assays. Both assays were run simultaneously on 96-well optical reaction plates. PCR amplification included an initial phase of 2 min at 50 °C, followed by 10 min at 95 °C, 40 cycles of 15 s at 95 °C, and 1 min at 60 °C. Results for nephrin mRNA are reported as its ratio to total blood ␤-actin mRNA. Statistical analysis was performed with SPSS 11 (SPSS Inc.). Differences in nephrin mRNA between groups were analyzed by the Mann– Whitney U-test, and a P value ⬍0.05 was considered statistically significant. The slopes of 10 consecutive calibration curves (10-fold serial dilutions of kidney cDNA to obtain a 5-point curve) showed good reproducibility for both the target and housekeeping genes (CV for slopes, 4% for nephrin and 6% for ␤-actin). Nephrin mRNA was detected in peripheral blood from all healthy and transplant patients. In the healthy controls, the median nephrin mRNA concentration was 4.99 ⫻ 10⫺3 (n ⫽ 14), higher in females than in males (Table 1). In transplant patients, the median nephrin mRNA concentration was 1.29 ⫻ 10⫺3 (n ⫽ 53); in this group also, females had concentrations higher than males (Table 1). When healthy individuals and transplant patients of the same gender were compared, significant differences were observed (Table 1). Because there were few individuals in the healthy group, it was not possible examine the effect of age. In male transplant patients, an age effect was seen: Patients ⬍30 years of age had higher nephrin mRNA (2.41 ⫻ 10⫺3; n ⫽ 10) than patients ⬎50 years of age (1.07 ⫻ 10⫺3; n ⫽ 11; P ⫽ 0.02). In a multiple regression analysis of nephrin as the dependent variable and gender, age group, and patient group as independent

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Table 1. Blood nephrin mRNA concentrations in healthy persons and kidney transplant patients. Median (range)

Healthy persons Transplant patients a b

Females

Males

P, Mann–Whitney U-test

0.076 (0.003–0.180) 0.0284 (0.0023–1.383)a

0.0394 (0.0064–0.1339) 0.01065 (0.0013–0.4925)b

0.04 0.022

P ⫽ 0.05 compared with healthy males. P ⫽ 0.029 compared with healthy females.

variables, only gender (P ⫽ 0.019) and age group (0.045) were significant contributors to the variation in nephrin mRNA. To our knowledge, this is the first study to measure nephrin mRNA in blood by real-time qPCR. This method does not distinguish between the possible sources of RNA, i.e., renal cells, blood cells, or other cells. It is also possible that the nephrin mRNA detected by our assay may arise from the presence of “illegitimate transcription”, i.e., basal transcription of tissue-specific genes outside of the tissues where they are typically active. Although this phenomenon is known to occur for certain genes, the illegitimate transcripts are thought to be of very low abundance (11 ), and this may account, in part, for some of the nephrin mRNA detected in our study. Extensive searches of available genomic databases nevertheless seem to suggest that nephrin expression is overwhelmingly kidney specific. This study shows that nephrin mRNA in circulation is influenced by age and transplantation, both of which are associated with reduced renal mass: Renal function decreases with age, and transplant patients have lower renal mass. The mean serum creatinine concentration in the transplant group in this study was 178 mol/L. Serum creatinine and renal function are lower in females than in males, and nephrin mRNA would be expected to be lower in females. In our study, however, mRNA was higher in females, both in healthy individuals and in transplant patients. This finding suggests that the nephrin mRNA expression is influenced by estrogens or other female hormones. There were not enough female participants, particularly in the postmenopausal age group, to test this hypothesis, and further studies are necessary to confirm our findings. We conclude that nephrin mRNA in peripheral blood measured by real-time qPCR is higher in females and in individuals with higher renal mass. These factors must be taken into account in interpreting the results of nephrin mRNA.

We thank the staff of the Molecular Working Group at St. Thomas’ Hospital for the use of the facilities in the Molecular Diagnostic Laboratory and the staff in the renal transplant unit for helping with the study. References 1. Ly J, Alexander M, Quaggin SE. A podocentric view of nephrology. Curr Opin Nephrol Hypertens 2004;13:299 –305. 2. Chan AK, Chiu RW, Lo YM; Clinical Sciences Reviews Committee of the

3.

4.

5.

6.

7.

8.

9. 10.

11.

Association of Clinical Biochemists. Cell-free nucleic acids in plasma, serum and urine: a new tool in molecular diagnosis. Ann Clin Biochem 2003;40: 122–3. Ng EK, Tsui NB, Lam NY, Chiu RW, Yu SC, Wong SC, et al. Presence of filterable and nonfilterable mRNA in the plasma of cancer patients and healthy individuals. Clin Chem 2002;48:1210 –1. Bojunga J, Roddiger S, Stanisch M, Kusterer K, Kurek R, Renneberg H, et al. Molecular detection of thyroglobulin mRNA transcripts in peripheral blood of patients with thyroid disease by RT-PCR. Br J Cancer 2000;82:1650 –5. Li D, Butt A, Clarke S, Swaminathan R. Real-time quantitative PCR measurement of thyroglobulin mRNA in peripheral blood of thyroid cancer patients and healthy subjects. Ann N Y Acad Sci 2004;1022:147–51. Rainer TH, Wong LK, Lam W, Yuen E, Lam NY, Metreweli C, et al. Prognostic use of circulating plasma nucleic acid concentrations in patients with acute stroke. Clin Chem 2003;49:562–9. Lam NYL, Rainer TH, Chan LYS, Joynt GM, Lo YMD. Time course of early and late changes in plasma DNA in trauma patients. Clin Chem 2003;49:1286 – 91. Wijeratne S, Butt A, Burns S, Sherwood K, Boyd O, Swaminathan R. Cell-free plasma DNA as a prognostic marker in intensive treatment unit patients. Ann N Y Acad Sci 2004;1022:232– 8. Hamaoui K, Butt A, Powrie J, Swaminathan R. Concentration of circulating rhodopsin mRNA in diabetic retinopathy. Clin Chem 2004;50:2152–5. Tryggvason K. Unravelling the mechanism of glomerular ultrafiltration: nephrin a key component of the slit diaphragm. J Am Soc Nephrol 1999; 10:2440 –5. Chelly J, Concordet JP, Kaplan JC, Kahn A. Illegitimate transcription: transcription of any gene in any cell type. Proc Natl Acad Sci U S A 1989;86:2617–21. DOI: 10.1373/clinchem.2005.053124

Immunogenicity Testing by Electrochemiluminescent Detection for Antibodies Directed against Therapeutic Human Monoclonal Antibodies, Michael Moxness,* Suzanna Tatarewicz, Dohan Weeraratne, Nancy Murakami, Danika Wullner, Dan Mytych, Vibha Jawa, Eugen Koren, and Steven J. Swanson (Department of Clinical Immunology, Amgen, Inc., Thousand Oaks, CA; * address correspondence to this author at: Department of Clinical Immunology, Amgen, Inc., Thousand Oaks, CA 91320-1799; fax 805-480-1306, e-mail [email protected]) Monitoring the immune response against human therapeutic monoclonal antibodies is an important component of preclinical and clinical trials to assess drug exposure, efficacy, and safety (1, 2 ). Detection of antibodies (analyte) directed against therapeutic antibodies (drug) is difficult because of the similarity of structure between drug and analyte and the high concentrations of drug present in serum, particularly during preclinical toxicology studies (3, 4 ). The purpose of this study was to

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Abstracts of Oak Ridge Posters

Table 1. Assay characteristics.a Assay

Species

LOD, ␮g/L

Interassay CV at LOD, %

Acid pretreatment

Drug tolerated (␮g/L) at LOD

Drug tolerated (␮g/L) at 500 ␮g/L analyte

A B C D E

Mouse Monkey Human Rat Human

63 63 31 5 15

8 5 14 4 18

Yes Yes Yes No No

5000 1000 10 000 NDb ⬎5000

20 000 5000 ⬎100 000 ⬎50 000 ⬎50 000

a The analyte measured was a unique rabbit anti-drug–specific antibody generated for each assay that was diluted in pooled normal serum from each designated species. b ND, not determined.

develop a robust assay format applicable across species and drug compounds for rapid method development and validation. We developed 5 assays to detect antibodies against 5 distinct monoclonal antibody therapeutic drug products. Detection was dependent on bivalent antibodies binding both biotin- and ruthenium-conjugated drug compounds (for a schematic of the assay, see Fig. 1 of the Data Supplement that accompanies the online version of this abstract at http://www.clinchem.org/content/vol51/ issue10/). This “bridging” assay format eliminated the need for species–specific secondary antibodies and provided assay consistency in different serum matrices as a drug product moves from preclinical to clinical development. In addition, these assays theoretically detected all classes of immunoglobulins as long as they were functionally multivalent. We used 4 different serum matrices (mouse, monkey, rat, and human). Each drug was conjugated with a ruthenium complex (Meso Scale Discovery) that produces light [electrochemiluminescence (ECL)] on application of an electric potential. A separate sample of the drug was also conjugated with biotin (Pierce). A rabbit polyclonal affinity-purified antibody specific for each drug was generated and used as a surrogate analyte. Labeling ratios were optimized to retain immunochemical reactivity, which was determined by surface plasmon resonance measurements (Biacore 3000; Biacore) of the binding between conjugated drugs and the rabbit antidrug antibody. Relative binding was required to fall within 80%–120% of binding of the native drug compound. Typically, label-to-protein molar ratios were found to be optimal in the range of 2:1 to 5:1. Sera were diluted to 10%, and 25 ␮L was incubated overnight with 50 ␮L of biotin- and ruthenium-conjugated drug, each at a concentration of 500 ␮g/L. The mixture was then added to plates equipped with electrodes and coated with streptavidin (Meso Scale Discovery) to capture complexes formed between biotin-labeled drug, analyte, and ruthenium-labeled drug. After a 2-h incubation, the plates were washed, and the ECL signals were measured on an MSD Sector PR analyzer (Meso Scale Discovery). Optimization included maximizing conjugated drug concentrations and incubation times. Samples were pretreated with acid in some assays to disrupt immune complexes before analysis. Acid pretreatment enhanced the detection of analyte in the presence of high drug concentrations (5 ). Assay

precision, limits of detection (LOD), and tolerance to excess soluble drug were determined for each assay (Table 1). Analyte stability was also determined for each assay. As expected, antibodies withstood up to 10 freeze– thaw cycles and up to 24 h at room temperature without substantial loss of activity (data not shown). The wide dynamic range of the ECL technology allowed for a linear signal response between analyte concentrations of 10 and 10 000 ␮g/L. The upper limit of a 95% one-sided reference interval (threshold value) was estimated for each assay by measuring at least 60 individual serum samples from the designated species (Bioreclamation). All samples were from drug-naive individuals or animals. Each response was normalized against a pooled species-specific serum (Bioreclamation) run in every assay. The frequency distribution was analyzed for gaussian shape, and the data were transformed by log or squareroot function if necessary. The LOD was determined by adding a known amount of analyte (a unique rabbit anti-drug–specific antibody generated for each assay) to at least 30 donor samples (6 ). The normalized frequency distribution was examined, and the expected minimal response (mean ⫺ 2 SD) at the LOD was required to fall above the threshold value. The LOD varied between 5 and 63 ␮g/L; however, this variability probably depended primarily on the use of different drug-specific antibodies with differing affinities. Tolerance to the presence of excess drug was determined by mixing known amounts of drug and analyte in pooled species-specific serum and incubating for at least 1 h. Samples were analyzed and determined to be positive if the response was above the threshold value (Fig. 1). In summary, ECL bridging assays for the detection of antibodies directed against 5 different human therapeutic antibodies were developed and validated in sera of multiple species. The high sensitivity and reproducibility of this technique allowed for low LOD (5– 63 ␮g/L), as well as for good precision (CV ⬍20%), dynamic range (10 – 10 000 ␮g/L), and tolerance to excess drug present in serum (drug concentrations up to 100 000 ␮g/L tolerated at an analyte concentration of 500 ␮g/L). These assays can be used to detect antibodies against therapeutic antibody drugs from preclinical development through clinical trials and for postmarketing immunogenicity surveillance.


Fig. 1. Results for serum samples supplemented with 31 (E), 500 (Œ), 1000 (f), and 5000 (⽧) ␮g/L analyte, respectively. Each sample was also supplemented with increasing concentrations of drug and analyzed in assay C.

Amgen, Inc. supported all work described in this report. All authors are also employed by Amgen, Inc. References 1. Brekke OH, Sandlie I. Therapeutic antibodies for human diseases at the dawn of the twenty-first century. Nat Rev Drug Discov 2003;2:52– 62. 2. Chamberlain P. Immunogenicity of therapeutic proteins. Reg Rev 2002;5: 4 –9. 3. Wadhwa M, Bird C, Dilger P, Gaines-Das R, Thorpe R. Strategies for detection, measurement, and characterization of unwanted antibodies induced by therapeutic biologicals. J Immunol Methods 2003;278:1–17. 4. Mire-Sluis AR, Barrett YC, Devanarayan V, Koren E, Liu H, Maia M, et al. Recommendations for the design and optimization of immunoassays used in the detection of host antibodies against biotechnology products. J Immunol Methods 2004;289:1–16. 5. Moxness M, Foley J, Stene M, Finco-Kent D, Bedian V, Krasner A, et al. Development and validation of radioligand binding assays to measure total IgA, IgE, IgG, and IgM insulin antibodies in human serum. Ann N Y Acad Sci 2003;1005:265– 8. 6. Linnet K, Kondratovich M. Partly nonparametric approach for determining the limit of detection. Clin Chem 2004;50:732– 40.

time and making such systems suitable for point-of-care testing. However, decreased reaction volumes typically lead to other problems, such as imprecision of liquid metering, liquid evaporation, and problems related to the increased surface-to-volume ratio. To address these challenges, we have developed a new bioanalytical system composed of a disposable microfluidic compact disc (CD) device (Gyrolab BioaffyTM) that is intended for sandwich immunoassays and an instrument for automatic processing of CDs (GyrolabTM Workstation). The Gyrolab Bioaffy CD (diameter, 120 mm) contains 104 parallel microstructures conveniently fitting into the CD, each covering a surface area of ⬃2 ⫻ 15 mm. The CD is manufactured by injection molding using a cycloolefin polymer. The CD is covered with a lid in which appropriately placed holes enable liquid introduction by the robot into the microstructures; the lid protects liquids from evaporation once in the CD. Liquid movement and localization are achieved by a combination of capillary action, centrifugal force, and hydrophobic barriers within the microstructure. Each microstructure contains a 15-nL particle-based column coupled with streptavidin in which reactions take place (Fig. 1). The Gyrolab Bioaffy is designed to be an open system; i.e., it is compatible with various assays that use biotinylated reagents, which bind to the streptavidin column as the first layer of the sandwich. Thus the capture column in a microstructure can promptly be functionalized for different analytes by addition of selected biotinylated capture reagents in excess. Each microstructure contains integrated functions for several process steps, such as volume metering, and each microstructure can be ac-

DOI: 10.1373/clinchem.2005.053272

Integrated Microfluidic Compact Disc Device with Potential Use in Both Centralized and Point-of-Care Laboratory Settings, Mats Ingana¨s,* Helene De´rand, Ann Eckersten, Gunnar Ekstrand, Ann-Kristin Honerud, Gerald Jesson, Gunnar Thorse´n, Tobias So¨derman, and Per Andersson (Gyros AB, Uppsala Science Park, SE-751 83 Uppsala, Sweden; * author for correspondence: fax 46-18-566352, e-mail [email protected]) Reducing the size of reaction volumes in immunodiagnostic procedures can reduce not only reagent cost but also analysis time. With reduced sample and reagent volumes and use of a flow-through mode, reactions reach completion rapidly, potentially decreasing turnaround

1985

Fig. 1. Gyrolab Bioaffy CD for sandwich immunoassay.

1986


cessed through an individual inlet or through a common channel connecting 8 microstructures. The workstation automatically handles the analytical process from addition of capture reagent to detection of reactions. It is equipped with a robotic station for liquid transfer from a microtiter plate to a CD, a CD spinner, and a detector for HeNe laser-induced fluorescence at 633 nm. Reagents and samples, kept in microtiter (MT) plates, are placed in a carousel in the instrument. The system is designed to handle maximally 5 CDs in a batch run. The instrument is also equipped with dedicated software to handle all process steps and evaluate the data. Fluorescence detection is carried out on column by rotating the CD while moving the detector in the radius direction. The fluorescent signal is integrated and represents the total response of the reaction. A few microliters of sample can be handled by the robotic system. For each data point, 200 nL of sample is analyzed. Wash liquids, reagents, and samples are sequentially introduced into the CD according to the method. All process steps, including the analyte capture step, are performed under flow conditions by spinning the CD at appropriate rotational speeds. More than 100 data points can be generated in 50 min. To avoid evaporation in MT plates, an aluminum foil that is penetrated by capillaries on aspiration of liquids is attached on top of MT plates. Commercially available antibodies specific for human analytes were selected for their assay performance in the system. Recombinant proteins were used to create calibration curves diluted in Gyros Standard Diluent (Gyros AB). Capture antibodies were labeled with biotin (cat. no. 21335; Pierce) and detection antibodies with ALEXA 647 (cat. no. A-20186; Molecular Probes) according to instructions from the manufacturers. Biotinylated capture reagents were used at a concentration of 100 mg/L, which is sufficient to saturate the streptavidin column, and detection reagents at 2–5 mg/L. Samples were prepared by mixing sample and equal volumes of Gyros Sample Diluent (Gyros AB). All quantitative assays are critically dependent on precise metering of sample volume. In the Gyrolab Bioaffy, volume definition is performed within the CD as part of the analysis procedure. When human EDTA plasma was subjected to volume definition within the CD microstructure, a 200-nL aliquoted sample volume was prepared with a CV of 0.75% (n ⫽ 560). The measuring range for a given analyte is dependent on the reagent properties and the binding capacity of the capture column. When working at low analyte concentrations, the analyte content of the 200-nL sample volume is enriched to a few nanoliters at the top of the capture column. When using high-quality immunoreagents, subpicomolar concentrations of analyte can be determined with a measuring range of 3– 4 orders of magnitude of analyte concentrations before the column becomes saturated (data not shown). To illustrate the intra-CD precision of the Gyrolab Bioaffy, recombinant human interleukin-2 (hIL-2) was

diluted to 50, 500, and 5000 ng/L. Each preparation was repeatedly analyzed for hIL-2 content in 3 separate CDs, each generating 104 data points per concentration. The CVs were 6.0%, 4.0%, and 4.2% for hIL-2 concentrations of 50, 500, and 5000 ng/L, respectively. To determine recovery in the Gyrolab Bioaffy, we added recombinant hIL-1␤ to 2 sets of human serum and plasma samples (citrate, EDTA, and heparin) at 2 concentrations (20 and 200 ng/L). The concentration of hIL-1␤ was measured with use of a set of immunoreagents specific for human hIL-1␤. Mean recovery of hIL-1␤ was 74%–117% in serum, 102%– 104% in citrate plasma, 95%–101% in EDTA plasma, and 88%–105% in heparin plasma. We developed a prototype CD containing microstructure functions for integrated separation of whole blood into plasma combined with an immunoassay for myoglobin as an illustrative example of how integration of functions for an intended process can be achieved with CD microfluidic technology. The process took place in 4 steps: First, 3 ␮L of fresh heparin blood was applied to a CD structure inlet that filled by capillary action. Plasma was then generated in the upper part of the microfluidic structure (not shown here) by a sedimentation spin at 1500 rpm for 180 s. Next, an overflow channel was activated and defined the plasma volume of 0.5 ␮L (CV ⬍1%). Finally, plasma was transferred to the top of the capture column through an outlet channel by increasing the rotational frequency to 5000 rpm. The subsequent assay procedure was similar to the Gyrolab Bioaffy method described above. To test the prototype functions, fresh heparin blood samples were collected from 3 donors. Plasma was prepared within the CD and analyzed for myoglobin. In parallel, plasma was generated outside the CD by conventional means, introduced into the CD through a downstream inlet, and analyzed for myoglobin. The fluorescence intensities of the reactions representing the amount of myoglobin were compared for each sample set (Table 1). In summary, the results obtained for hIL-2 and hIL-1␤ regarding precision, dynamic range, and recovery in different matrices illustrate the performance of the Gyrolab Bioaffy. The features of the CD microfluidic technology can be adapted to meet specific needs and requirements for in vitro diagnostic purposes, for example, by incorporating functions for blood separation in point-ofcare laboratory settings. Costs for reagents can be lowered 10- to 100-fold by

Table 1. Comparison of response from CD-based immunoassay of myoglobin in heparin plasma with separation of plasma from blood cells performed either inside or outside the CD. Mean (SD) fluorescence

Plasma prepared inside CD Plasma prepared outside CD

Sample 1

Sample 2

Sample 3

135 (36) 131 (34)

124 (23) 90 (34)

58 (19) 53 (2)


1987

reducing reagent consumption. Our experience to date suggests that existing assays developed for established in vitro diagnostic formats can be readily transferred into a CD microfluidic format.

Previously published online at DOI: 10.1373/clinchem.2005.053181

Development of a Novel Enzymatic Cycling Assay for Total Homocysteine, Chao Dou,† Dongyuan Xia,† Liqing Zhang, Xiaoru Chen, Patrick Flores, Abhijit Datta, and Chong Yuan* (Diazyme Laboratories Division, San Diego, CA; † these authors contributed equally to this work; * address correspondence to this author at: Diazyme Laboratories Division, General Atomics, 3550 General Atomics Court, San Diego, CA 92121; fax 858-455-4750, e-mail [email protected] or www.diazyme.com) Homocysteine (HCY) is a thiol-containing amino acid produced by the intracellular demethylation of the essential amino acid methionine. Intracellular HCY enters either the transsulfuration pathway or the remethylation cycle. When the enzymatic reactions involved in the 2 metabolic pathways of intracellular HCY are impaired, because of either genetic defects in enzymes for HCY metabolism or the nutritional deficiency of vitamins such as folate, B12, and B6, HCY accumulates in the cells and is exported to the circulation (1– 4 ). Approximately 80% of circulating HCY in the blood is protein bound by disulfide linkage (5 ). The remaining unbound HCY combines by oxidation, either with itself to form the dimer homocystine or with cysteine to form the mixed disulfide cysteineHCY. Only a small amount circulates as free HCY. Total HCY (tHCY) represents the sum of all forms of HCY, including forms of oxidized HCY, protein bound, and free. Highly increased concentrations of tHCY are found in persons with homocystinuria, a rare genetic disorder most commonly caused by a deficiency of cystathionine ␤-synthase. Typical clinical manifestations of patients with homocystinuria include mental retardation, early arteriosclerosis, and arterial and venous thromboembolism (4, 6 ). Other less severe genetic defects, including polymorphisms in genes encoding for enzymes that participate in HCY metabolism, such as methylenetetrahydrofolate reductase (MTHFR) and methionine synthase, are also found in patients who have moderately increased plasma concentrations of HCY (7, 8 ). Patients with chronic renal disease have high morbidity and mortality resulting from arteriosclerotic cardiovascular disease (CVD). Increased tHCY is a frequently observed finding in the blood of these patients. Initial investigation (9 ) suggested that the markedly increased plasma tHCY found in end-stage renal disease patients contributed independently to their very high incidence of fatal and nonfatal CVD outcomes. The increased concen-

Scheme 1.

trations of tHCY are the result mainly of impaired removal of HCY from the blood by the kidney and are independent of their vitamin concentrations. In most US clinical laboratories, 15 ␮mol/L is the cutoff value for a normal concentration of HCY for adults. Each laboratory is recommended to establish a range of normal values for the population in their region. Recently, we developed an enzymatic cycling assay for detection of HCY in human serum or plasma. The Diazyme enzymatic HCY assay is based on a cosubstrate conversion product cycling principle, as shown in Scheme 1. Oxidized HCY is first reduced to free HCY, which then reacts with a cosubstrate, S-adenosylmethionine (SAM), catalyzed by an HCY S-methyltransferase to form methionine (the HCY conversion product of HCY) and Sadenosylhomocysteine (SAH, the cosubstrate conversion product, which does not contain any element from the sample homocysteine molecule). SAH is hydrolyzed into adenosine and HCY by SAH hydrolase. The formed HCY that originated from the cosubstrate SAM is cycled back to the HCY conversion reaction by HCY S-methyltransferase, producing an enzymatic cycling system with substantial amplification of detection signals. The formed adenosine is hydrolyzed into inosine and ammonia, which reacts with glutamate dehydrogenase with concomitant conversions of NADH to NAD⫹. The concentration of HCY in the sample is proportional to the amount of NADH converted to NAD⫹ (⌬A340nm). The Diazyme HCY enzymatic assay has been formulated into a 3-reagent, homogeneous liquid system. For analyzers capable of handing 3 reagents, the assay is ready for use. For analyzers capable of handling only 2 reagents, premixing of Reagent 1 and Reagent 2 is needed. The assay has been implemented on various automated analyzers, and samples can be analyzed in as little as 10 min. We evaluated the precision of the Diazyme enzymatic HCY assay on a Cobas Mira analyzer, according to NCCLS guideline EP5-A. In the study, 3 specimens containing 7, 12, and 29.5 ␮mol/L homocysteine were tested with 2 runs per day and with duplicates over 20 working days. The results demonstrated that within-run impreci-

1988


sions (CVs) were 2.2% for 7 ␮mol/L, 3.0% for 12 ␮mol/L, and 1.8% for 29.5 ␮mol/L HCY, respectively; and total imprecisions (CVs) were 4.1% for 7 ␮mol/L, 5.9% for 12 ␮mol/L, and 4.0% for 29.5 ␮mol/L HCY, respectively. We determined the diagnostic accuracy of the assay by testing individual serum or plasma samples with our enzymatic cycling HCY assay and comparing that assay with both a commercial enzymatic HCY assay and a commercial HCY immunoassay. To ensure that the concentrations of HCY distributed across the reportable dynamic range, some HCY samples used for the study were enriched with a stock solution of HCY to targeted concentrations. Fig. 1 illustrates the correlation results with the Catch enzymatic HCY assay and the Bayer HCY immunoassay on varieties of clinical chemistry analyzers.

From these results, we can see that the Diazyme enzymatic HCY assay correlated very well with both assays. It appears that the Diazyme enzymatic cycling HCY assay correlated slightly better with the Catch enzymatic assay than with the Bayer HCY immunoassay. To demonstrate the linearity of the Diazyme HCY enzymatic assay, a serum sample containing 7 ␮mol/L HCY was enriched with l-HCY stock solution to 71.2 ␮mol/L HCY. Next, a serial dilution of the serum sample containing 71.2 ␮mol/L HCY was made with 7 ␮mol/L HCY serum to the indicated concentrations. The serum sample containing 7 ␮mol/L HCY was diluted with saline in series, and the final dilution contained 3.1 ␮mol/L HCY. The serum samples, prepared as described, were tested on Cobas Mira using the Diazyme enzymatic assay.

Fig. 1. Correlation with Catch enzymatic HCY assay and Bayer immunoassay. (A), correlation with Catch HCY assay on Cobas Mira; (B), correlation with Catch HCY assay on Hitachi 917; (C), correlation with Catch HCY assay on Synchron CX-7; (D), correlation with Bayer HCY immnoassay.


The result demonstrated that the assay has a linear range of at least 50 ␮mol/L HCY. In a separate study, we determined the limit of detection (LOD) of the Diazyme HCY enzymatic assay. A zero calibrator was tested with 12 replicates. The LOD, defined as the mean ⫹ 3 SD, was ⬍1.5 ␮mol/L HCY. We determined the extent of interference from the substances typically present in the serum by testing a 12.9 ␮mol/L HCY serum sample enriched with various concentrations of substances. We also determined the interference from common prescription and over-the-counter pharmaceuticals by testing 2 serum samples, containing 11.5 ␮mol/L and 42.7 ␮mol/L HCY, respectively, which were enriched with various concentrations of substances examined. The test results, with the respective concentration of each substance, are listed in Table 1. In conclusion, we have developed an innovative enzymatic cycling assay for the in vitro quantitative determination of total l-HCY in serum or plasma, which has been adapted on automated chemistry analyzers. The assay uses 3 reagents and has been implemented on the Hitachi 917, Synchron CX-7, and Cobas Mira, testing human serum and

Table 1. Results of interference study. Interfering substances

Endogenous substancesa NH4⫹ NaF Inorganic phosphorus Triglycerides Ascorbic acid Bilirubin Hemoglobin Glutathione L-Cysteine SAM Adenosine Cystathionine Pharmaceuticalsb Acetylsalicylic acid Ampicillin Carbamazepine Cefoxitin Cyclosporin Ibuprofen Lidocaine Methotrexate Methyldopa Penicillamine Rifampicin Salicylic acid Theophylline Valproic acid

Concentration

500 ␮mol/L 1 mmol/L 1 mmol/L 25 g/L 10 mmol/L 200 mg/L 12 g/L 0.5 mmol/L 1 mmol/L 20 ␮mol/L 100 ␮mol/L 20 ␮mol/L 1 g/L 1 g/L 100 mg/L 2.5 g/L 5 mg/L 500 mg/L 100 mg/L 100 mg/L 20 mg/L 100 mg/L 60 mg/L 1 g/L 100 mg/L 100 mg/L

a The listed endogenous substances typically present in serum produced ⬍10% deviation when tested at the concentrations given. b The listed prescription and over-the-counter pharmaceuticals typically present in serum produced ⬍15% deviation when tested at the concentrations given.

1989

plasma. Within-run imprecision (CV) was ⬍3%, and total imprecision (CV) was ⬍6% for the 3 concentrations of HCY controls tested. The study testing human serum and heparin-plasma samples demonstrated excellent correlation with another enzymatic HCY assay on various automated chemistry analyzers. The assay also correlated well with an HCY immunoassay. Limit of detection in serum is ⬍1.5 ␮mol/L HCY in serum, and the assay is linear up to 50 ␮mol/L HCY. Interference is ⬍10% with 1 mmol/L l-cysteine, 0.5 mmol/L glutathione, 100 ␮mol/L adenosine, 20 ␮mol/L cystathionine, and 500 ␮mol/L NH4Cl. A panel of commonly prescribed drugs did not contribute interference to the assay at the concentrations indicated. References 1. Eikelboom JW, Lonn E, Genest J Jr, Hankey G, Yusuf S. Homocyst(e)ine and cardiovascular disease: a critical review of the epidemiologic evidence. Ann Intern Med 1999;131:363–75. 2. Scott J, Weir D. Homocysteine and cardiovascular disease. QJM 1996;89: 561–3. 3. Nygard O, Nordrehaug JE, Refsum H, Ueland PM, Vollset SE. Plasma homocysteine levels and mortality in patients with coronary artery disease. N Engl J Med 1997;337:230 – 6. 4. Seshadri S, Beiser A, Selhub S, Jacques PF, Rosenberg IH. D’Agostino RB, et al. Plasma homocysteine as a risk factor for dementia and Alzheimer’s disease. N Engl J Med 2002;346:477– 83. 5. McLean R, Jacques PF, Selhub J, Tucker KL, Samelson EJ, Broe KE, et al. Homocysteine as a predictive factor for hip fracture in older persons. N Engl J Med 2004;350:2042–9. 6. Refsum H. Total homocysteine guidelines for determination in the clinical laboratory. Clin Lab News 2002;5:12– 4. 7. Guttormsen AB, Schneede J, Fiskerstrand T, Ueland PM, Refsum HM. Plasma concentrations of homocysteine and other aminothiol compounds are related to food intake in healthy human subjects. J Nutr 1994;124:1934 – 41. 8. Vilaseca MA, Moyano D, Ferrer I, Artuch R. Total homocysteine in pediatric patients. Clin Chem 1997;43:690 –2. 9. Faure-Delanef L, Quere I, Chasse JF, Guerassimenko O, Lesaulnier M, Bellet H, et al. Methylenetetrahydrofolate reductase thermolabile variant and human longevity. Am J Hum Genet 1997;60:999 –1001.

DOI: 10.1373/clinchem.2005.053421

Automated Enzymatic Assay for Measurement of Lithium Ions in Human Serum, Chao Dou, Olga Aleshin, Abhijit Datta, and Chong Yuan* (Diazyme Laboratories Division, General Atomics, 3550 General Atomics Ct., San Diego, CA 92121; * author for correspondence: fax 858455-4754, e-mail www.diazyme.com) For decades, lithium carbonate has been one of the most effective agents for the treatment of bipolar disorder (manic depressive psychosis). Lithium acts by altering intraneuronal metabolism of catecholamines, inhibiting noradrenaline-sensitive adenylate cyclase, reducing synaptic transmission, and increasing neuronal excitability with modification of the central nervous system (CNS) amine concentrations. Recent studies have shown that lithium holds promise against Alzheimer disease. Lithium has many side effects, however. Overdosage of lithium can cause acute Li⫹ intoxication, which occurs quite often because of lithium’s narrow therapeutic index. For example, serum Li⫹ concentrations ⬎1.5 mmol/L 12 h after a dose usually indicate a significant risk of intoxication. Therefore, the timely and accurate monitoring of serum

1990


concentrations of lithium after a therapeutic dose is critical (1– 4 ). The most commonly used methods to detect serum lithium are ion-selective electrodes (ISEs) and flame emission photometry. ISE analyses rely on ion-specific electrodes. Ideally, each electrode possesses a unique ionselective property that allows it to respond to the desired ion. In practice, however, interference from other ions in the sample compromises the specificity of the detecting electrode, rendering the electrodes susceptible to false readings. The instrumentation for ISE analysis is relatively expensive, requires routine maintenance that is sometimes cumbersome and time-consuming, and demands that the operating technician have considerable skill and knowledge for accurate and consistent readings. Flame emission photometry relies on the principle that certain atoms, when energized by heat, become excited and emit a light of characteristic wavelength. Radiant energy produced by atoms in the flame is directly proportional to the number of atoms excited in the flames, which is directly proportional to the concentration of the substance of interest in the sample. Like ISE analysis, the instrumentation required for this method is complex and expensive. Moreover, flame emission photometry requires the use of combustible gas, necessitating expensive hazard prevention measures. More recently, a dye-based lithium assay has been developed and used in certain automated chemistry analyzers (5 ). We recently developed an enzymatic lithium assay, which has been adapted to most automated clinical chemistry analyzers. In this assay, lithium is determined through a kinetic coupling system involving a lithiumsensitive enzyme, 3⬘,5⬘-bisphosphate nucleotidase, from yeast. This enzyme is sensitive to lithium inhibition, with an IC50 of 0.1 mmol/L. The enzyme is also sensitive to sodium inhibition, with a much higher IC50 (⬎20 mmol/ L). The inhibitive effect of serum sodium ions is effectively masked by the sodium-specific binding reagent Kryptofix 221. Through enzymatic coupling, substrate adenosine 3⬘,5⬘-bisphosphate (PAP) is converted to hypoxanthine by enzymatic reactions to generate uric acid and hydrogen peroxide (H2O2). The generated H2O2 then reacts with N-ethyl-N-(2-hydroxy-3-sulfopropyl)-3-mtoluidine (EHSPT) and 4-aminoantipyrine (4-AA) to form a quinone dye with an absorbance maximum at 556 nm. The rate of dye formation is inversely proportional to the lithium concentration in serum: 3⬘,5⬘-Bisphosphate nucleotidase

¡ AMP ⫹ Pi PAP ⫹ H2O O

5⬘-Nucleotidase/ adenosine deaminase/ PNP

¡ Hypoxanthine ⫹ Pi AMP O ⫹ NH3 ⫹ R-1-Pi

where PNP is purine nucleoside phosphorylase. Xanthine oxidase

¡ Uric Hypoxanthine ⫹ 2 H2O ⫹ 2 O2 O acid ⫹ 2 H2O2 Peroxidase

H2O2 ⫹ 4-AA ⫹ EHSPT O ¡ 4 H 2O ⫹ dye (550 nm) The assay is formulated into a lyophilized 2-reagent system with MES buffer (pH 6.0). The assay analyzes nonhemolyzed serum samples on a variety of automated analyzers in as little as 10 min. The precision of the Diazyme enzymatic lithium assay was evaluated on a Cobas Mira analyzer according to Clinical and Laboratory Standards Institute (formerly NCCLS) guideline EP5-A. In the study, 2 specimens containing 1 and 2.3 mmol/L lithium, respectively, were tested in 2 runs per day with duplicates for a period of 20 working days. The results showed that within-run imprecision (CVs) was 4.7% for 1.0 mmol/L Li⫹ and 3.3% for 2.3 mmol/L Li⫹, respectively; and total imprecision (CVs) was 6.9% for 1 mmol/L and 5.5% for 2.3 mmol/L Li⫹, respectively. To demonstrate accuracy, the Diazyme lithium enzymatic assay was tested with individual serum samples and compared with both ISE and Thermo Trace colorimetric methods. Three concentrations of serum calibrators containing 0 mmol/L (low), 1 mmol/L (medium), and 3 mmol/L lithium (high) were prepared by adding 1 mol/L lithium acetate stock solution to the pooled lithium-free serum. After verification with a Trace lithium assay, the calibrators were sent out to a certified laboratory for confirmation. The pooled serum and the individual patient serum samples used for this study were from a certified commercial source and were accompanied by an Institutional Review Board certificate of approval for all protocols, including informed consent, used to collect samples. To ensure that the lithium concentrations were distributed across the reportable dynamic range, some lithium serum samples (especially at lithium concentrations ⬎2.5 mmol/L) were supplemented to the targeted concentrations with a stock solution of 1.0 mol/L lithium. The Diazyme enzymatic lithium assay and ISE were compared on Cobas Mira [n ⫽ 56; lithium concentration, 0 –3 mmol/L; y ⫽ 1.008x ⫹ 0.10 mmol/L (r2 ⫽ 0.953; Sy兩x ⫽ 0.042 mmol/L)], Hitachi 717 [n ⫽ 67; lithium concentration, 0 –3 mmol/L; y ⫽ 0.987x ⫹ 0.072 mmol/L (r2 ⫽ 0.962; Sy兩x ⫽ 0.19 mmol/L)], and Synchron CX-7 [n ⫽ 38; lithium concentration, 0 –3 mmol/L; y ⫽ 0.980x ⫹ 0.028 mmol/L (r2 ⫽ 0.989; Sy兩x ⫽ 0.019 mmol/L)] analyzers. The Diazyme lithium enzymatic assay and Trace lithium reagent were also compared [n ⫽ 61; lithium concentration, 0 –3 mmol/L; y ⫽ 1.083x ⫺ 0.071 mmol/L (r2 ⫽ 0.950; Sy兩x ⫽ 0.20 mmol/L)].


We also determined the linearity of the Diazyme lithium enzymatic assay. A serum sample containing 0 mmol/L Li⫹ was supplemented with lithium acetate stock solution to a concentration of 3.0 mmol/L Li⫹. The serum sample containing 3.0 mmol/L was then serially diluted with a serum sample containing 0 mmol/L lithium to obtain final lithium concentrations of 0, 0.5, 1.0, 1.5, 2.0, 2.5, and 3.0 mmol/L. The serum samples prepared as described were tested in triplicate with the Diazyme enzymatic lithium assay on the Cobas Mira. The result demonstrated that the assay was linear at least up to 3.0 mmol/L lithium (y ⫽ 0.9857x ⫹ 0.0548 mmol/L; r2 ⫽ 0.9863). To determine the extent of interference from other cations and substances typically present in serum, we supplemented 1 mL of serum containing 1.0 mmol/L lithium with various concentrations of substances, and then assayed the sera (6 replicates) with the Diazyme enzymatic lithium assay on the Hitachi 917. The results are shown in Table 1. In summary, we have developed an enzymatic coupling assay for quantitative measurement of lithium in nonhemolyzed human sera and have developed applications for commonly used automated chemistry analyzers. The assay uses 2 reagents, and applications have been developed for testing human serum specimens on the Cobas Mira, Synchron CX-7, and Hitachi 717. The withinrun CV was ⬍4.7%, and the total CV was ⬍6.9%. The study testing human sera with lithium concentrations ranging from 0 to 3 mmol/L demonstrated good correlation with both a commercially available ISE method and a colorimetric method on various automated analyzers. The assay was linear up to 3.0 mmol/L. No interference was detected from the following substances at the indicated concentrations: sodium, 200 mmol/L; ammonium, 0.5 mmol/L; calcium, 4.0 mmol/L; magnesium, 2.0 mmol/L; ascorbic acid, 5.0 mmol/L; zinc, 0.25 mmol/L; iron, 0.25

Table 1. Interference by cations and substances typically found in serum. Interferent

Concentration

Deviation, %

NH4⫹ Pi Ca2⫹ Mg2⫹ Na⫹ K⫹ Cu2⫹ Fe3⫹ Zn2⫹ Triglycerides Ascorbic acid Bilirubin

0.5 mmol/L 1.5 mmol/L 5.0 mmol/L 2.0 mmol/L 200 mmol/L 10 mmol/L 0.25 mmol/L 0.25 mmol/L 0.25 mmol/L 2.82 mmol/L (250 mg/dL) 5.0 mmol/L 770 ␮mol/L (45 mg/dL)

⫹0.8 ⫺1.6 ⫹4.7 ⫹6.2 ⫹3.9 ⫺0.8 ⫺0.8 ⫹0.8 ⫹0.8 ⫺1.2 ⫹0.8 ⫹6.5

a The interference study was carried out by adding various concentrations of the substances examined to individual patient serum samples and then testing with the Diazyme enzymatic lithium assay (6 replicates) on a Hitachi 917 analyzer. The cations and substances listed produced a ⬍10% deviation from the result for 1.0 mmol/L lithium at the concentrations given.

1991

mmol/L; copper, 0.25 mmol/L; potassium, 10 mmol/L; triglycerides, 2.82 mmol/L (250 mg/dL); and bilirubin, 770 ␮mol/L (45 mg/dL). References 1. Murguia JR, Belles JM, Serrano R. A salt-sensitive 3⬘(2⬘),5⬘-bisphosphate nucleotidase involved in sulfate activation. Science 1995;267:232– 4. 2. Moyer TP, Pippenger CE. Therapeutic drug monitoring. In: Burtis CA, Ashwood ER, eds. Tietz textbook of clinical chemistry, 2nd ed. Philadelphia: WB Saunders, 1994:1094 –154. 3. Amdisen A. Serum lithium determinations for clinical use. Scand J Clin Lab Invest 1967;20:104 – 8. 4. Wachtel MS, Paulson R, Plese C. Creation and verification of reference intervals. Lab Med 1995;26:593–7. 5. Rumbelow B, Peake M. Performance of a novel spectrophotometric lithium assay on a routine biochemistry analyzer. Ann Clin Biochem 2001;38: 684 – 6. DOI: 10.1373/clinchem.2005.053439

Development of an Automated Enzymatic Assay for the Determination of Glycated Serum Protein in Human Serum, Yuping Wang, Chao Dou, Chong Yuan, and Abhijit Datta* (Diazyme Laboratories Division, General Atomics, 3550 General Atomics Ct., San Diego, CA 92121; * author for correspondence to this author: fax 858-455-4750, e-mail [email protected] or www.diazyme.com) Fructosamine, or glycated serum protein (GSP), is formed via a slow Maillard reaction between glucose and amino acid residues of proteins. In patients with diabetes, increased blood glucose correlates with increased formation of fructosamine. The fructosamine value is representative of the mean blood glucose concentration over the preceding 2–3 weeks (medium-term indicator), whereas measurement of glycohemoglobin (hemoglobin A1c) offers the mean glucose concentration over a 6 – 8 week period (long-term indicator) and measurement of blood glucose gives only the current glucose concentration (transient or short-term indicator). Studies have shown that measurements of both glycohemoglobin and GSP are the most useful for monitoring long-term control of diabetes (1, 2 ). The reported reference GSP value for nondiabetic adults is ⬍285 mmol/L (3 ); however, it is generally recommended that each clinical testing laboratory establish its own reference intervals to reflect the ages, sexes, diet, and geographic location of the testing population. The Diazyme enzymatic assay for GSP uses a mixture of proteases, including proteinase K, to digest GSP into low–molecular-weight glycated protein fragments and uses Diazyme’s specific fructosaminaseTM, a microorganism-originated amadoriase to catalyze the oxidative degradation of Amadori product glycated protein fragments to yield protein fragments or amino acids, glucosone, and H2O2. The H2O2 released is measured by a colorimetric Trinder end-point reaction, and the absorbance generated is proportional to the concentration of GSP present in the sample (Fig. 1A). The assay has been formulated into a 2-reagent– based endpoint assay system, and applications

1992


Fig. 1. Schematic of Diazyme GSP enzymatic assay (A), and correlation between GSP results obtained for samples tested with the Diazyme GSP assay and the Randox Fructosamine assay (B). (A), TOOS, N-ethyl-N-sulphohydroxypropyl-m-toluide; 4-AAP, 4-aminoantipyrine. (B), the values obtained were plotted and are shown as a correlation graph. The slope is 0.998, the y-intercept is ⫺4.98 ␮mol/L, and the correlation coefficient (R2) between the methods is 0.98.

have been developed for a variety of automated analyzers. The simple test procedure is as follows: In a suitable cuvette, 20 ␮L of serum is mixed with 200 ␮L of Reagent 1, which includes proteases to digest the serum proteins, and the reaction mixture is incubated at 37 °C for 5 min. Reagent 2 (50 ␮L), which contains the Diazyme fructosaminase, is then added, and the absorbance at 550 nm is measured after an additional 5 min of incubation at 37 °C. On an automated analyzer, samples can be tested in 10 min. The Diazyme GSP enzymatic assay uses a 340 ␮mol/L GSP calibrator. The lyophilized calibrator is made with serum proteins, and the values are adjusted to the targeted concentrations by addition of appropriate amounts of glycated amino acid. We verified the calibrator values by using it as a sample in the Randox Fructosamine assay; the values were also confirmed by independent testing at Quest Diagnostics, a certified testing laboratory in the United States. To examine the performance of the Diazyme GSP enzymatic assay, we obtained a representative number of patient samples and used them for accuracy studies and method– comparison studies comparing the novel Diazyme GSP enzymatic assay with a commercially available fructosamine assay (Randox Laboratories). The pooled serum and the individual patient serum samples used for the accuracy study were obtained from a certified commercial source (ProMedDx, LLC) and were obtained with Institutional Review Board (IRB) certification on the handling and informed consent protocols. To ensure that the GSP concentrations were distributed across the reportable dynamic range, some serum samples used for the

study were prepared by addition of human GSP purchased from Sigma-Aldrich. We evaluated the assay precision on the Hitachi 917 general chemistry analyzer according to the guidelines in Clinical and Laboratory Standards Institute (formerly NCCLS) document EP5-A. We tested 2 specimens containing 280 and 586 ␮mol/L GSP with 2 runs per day, with duplicate runs over 10 working days. The results demonstrated that within-run imprecision (CVs) was 1.7% for 280 ␮mol/L and 1.8% for 586 ␮mol/L GSP, respectively, and total imprecision (CVs) was 2.9% for both 280 and 586 ␮mol/L GSP. We also completed a method– comparison study as follows: Individual serum samples were tested with the Diazyme GSP enzymatic assay and compared data obtained for the same samples with the Randox Fructosamine assay (Randox Laboratories). A total of 65 serum samples were tested with both the Diazyme GSP and Randox Fructosamine assays. The correlation between the results is shown in Fig. 1B. We determined the linearity of the Diazyme GSP enzymatic assay as follows. Human GSP was purchased from Sigma-Aldrich and reconstituted to a stock concentration of 1354 ␮mol/L. Serial dilutions of this stock then prepared to give GSP concentrations of 1083, 1015, 677, 338, 169, 84.6, 42.3, and 21.2 ␮mol/L. Each sample was tested in duplicate; the results demonstrated that the assay had a linear range from 21 to 1354 ␮mol/L GSP. We also examined the assay performance with regard to interference from substances commonly present in serum. In this interference study, we added the interferents listed below to a serum sample containing 160 ␮mol/L GSP and tested the sample with the Diazyme GSP enzymatic assay, in 3 replicates. We found that the following substances produced ⬍10% deviation when tested at the following target concentrations: uric acid, 2 mmol/L; hemoglobin, 2000 mg/L; glucose, 133 mmol/L; triglycerides, 20 g/L; ascorbic acid, 0.23 mmol/L; and bilirubin, 200 mg/L. In conclusion, we have developed an enzymatic assay for the in vitro quantitative determination of GSP in serum and have developed applications for commonly used automated chemistry analyzers. The Diazyme GSP enzymatic assay is linear up to 1354 ␮mol/L. The within-run and total CVs are ⬍2% and ⬍3%, respectively. The assay shows good correlation with the Randox Fructosamine assay (R2 ⫽ 0.98) and is not affected by ascorbic acid, bilirubin, hemoglobin, glucose, triglycerides, or uric acid at concentrations commonly found in patient samples. References 1. Armbuster DA. Fructosamine: structure, analysis and clinical usefulness. Clin Chem 1987;33:2153– 63. 2. Kouzuma T, Usami T, Yamakoshi M, Takahashi M, Imamura S. An enzymatic method for the measurement of glycated albumin in biological samples. Clin Chim Acta 2002;324:61–71. 3. Schleicher ED, Olgemoller B, Wiedenmann E, Gerbitz KD. Specific glycation of albumin depends on its half-life. Clin Chem 1993;39:625– 8. DOI: 10.1373/clinchem.2005.053447


Toward Multianalyte Immunoassays: A Flow-Assisted, Solid-Phase Format with Chemiluminescence Detection, Aldo Roda,1* Mara Mirasoli,1 Dora Melucci,2 and Pierluigi Reschiglian2 (1 Department of Pharmaceutical Sciences and 2 Department of Chemistry “G. Ciamician”, University of Bologna, Bologna, Italy; * address correspondence to this author at: Department of Pharmaceutical Sciences, University of Bologna, Via Belmeloro 6, 40126 Bologna, Italy; fax 39-051-343398, e-mail aldo.roda@ unibo.it and http://www.anchem.unibo.it) Chemiluminescence immunoassays are widely used in clinical chemistry. A multianalyte format is of particular interest to increase throughput and diagnostic utility. Field-flow fractionation (FFF) can separate dispersed particles in the micrometer-sized range (1 ). Separation is achieved within an empty capillary channel by the combined action of a transporting laminar flow and a field applied perpendicularly to the flow. According to their characteristics (e.g., dimension, density, and surface properties), particles are distributed at different positions within the parabolic flow profile, are thus transported along the channel at different velocities, and elute at different times. Methods combining FFF with chemiluminescence (CL) detection (FFF-CL) have recently been developed to efficiently separate horseradish peroxidase (HRP) in solution from HRP immobilized on micrometersized spheres and to quantify each fraction by means of the CL-HRP substrate based on H2O2–luminol– enhancer (2, 3 ). This was the basis for the development of FFF-CL– based solid-phase competitive immunoassays, in which micrometer-sized beads coated with the capture antibody are used as a solid phase and an analyte–HRP conjugate is used as a tracer. Once the competitive immunologic reaction takes place within the injection loop of the system, the antibody-bound tracer (bound fraction) is separated from tracer in solution (free fraction) in a few minutes by means of FFF. FFF-based immunoassays offer several advantages. For example, the kinetics of the immunologic reaction on micrometer-sized beads are faster than in the case of conventional microtiter plate assays. In addition, multianalyte FFF-based immunoassays can be developed by use of beads of different sizes (1–50 ␮m), each coated with the specific antibody for 1 analyte. The beads are fractionated by FFF before CL detection using different enzymes, such as HRP or alkaline phosphatase, detected by CL. Here we report the development of a CL-FFF– based competitive immunoassay for chloramphenicol (CAF), chosen as a model analyte. Flow FFF, in which the applied field structuring the separation is a flow perpendicular to the transporting flow, was used. The analytical conditions suitable for the monoanalyte case were optimized as a premise for the development of multianalyte applications. Polyclonal antibodies against CAF were produced in rabbits by use of a CAF– bovine serum albumin derivative as described previously (4 ). CAF– bovine serum albumin and CAF-HRP were synthesized by a previously reported

1993

method (5 ), and the purified derivatives were characterized by matrix-assisted laser desorption/ionization mass spectrometry. Antibody-coated microspheres were prepared from carboxylated polystyrene beads (Polysciences) with a mean (SD) diameter of 6.10 (0.57) ␮m; they were first coated with protein A and then incubated with the antibody diluted in incubation buffer [IB; 50 mmol/L phosphate buffer (pH 7.4) containing 1 g/L bovine serum albumin]. CAF calibrators (0.03–30 ␮mol/L, corresponding to 0.01–10 ppm) were prepared by serially diluting a CAF stock solution in IB. All conventional chemicals were purchased (Sigma). We used a Model F-1000 Fractionator (FFFractionation LLC; 29.4 ⫻ 2.0 ⫻ 0.0254 cm). No membrane was used at the accumulation wall (6 ). The longitudinal flow was set at 4.0 mL/min and the cross-flow at 0.5 mL/min. Mobile phase was 10 mmol/L Tris-HCl (pH 7.4), containing 0.5 g/L Tween 20. The injection port was a Rheodyne valve with a calibrated loop (17.83 ␮L; SD, 0.06 ␮L). Samples were injected in stop-flow mode (1 ) with a 6-min stopflow time. The flow FFF channel outlet was fed to an HPLC ultraviolet (UV)/visible detector (Dynamax Model UV-1; Varian) operating at 600 nm and then to a photomultiplier-based flow luminometer (FB12; Berthold Detection Systems) (7 ). The CL signal is given in relative light unit(s)/s. To perform CL detection, the CL enzymatic substrate, ECL® (Amersham Pharmacia Biotech) was flow-injected postcolumn, downstream of the UV/ visible detector and upstream of the luminometer, at a flow rate of 0.4 mL/min (7 ). To perform the FFF-CL– based CAF immunoassay, we mixed 100 ␮g of antibody-coated microphages with 100 ␮L of CAF calibrator and 100 ␮L of CAF-HRP diluted in IB; this mixture was then injected into the flow FFF system. After injection, the transporting flow was stopped for 6 min to allow for sphere relaxation. Free and bound tracer fractions were then separated during the flow FFF run and measured by online CL detection. The resulting fractogram contained a void peak at 20 s, corresponding to the free fraction, and a retained peak at 1.5 min, corresponding to the bound fraction. Total analysis time was ⬃10 min. After each run, a 3-min washing step in back-flushing mode with mobile phase was performed. A dose–response curve for CAF was obtained by plotting the peak area of the retained peak in the CL fractogram against CAF concentration (Fig. 1). Preliminary optimization of experimental conditions was necessary to achieve 3 main requirements: (A) Preservation of the immunologic binding during the FFF run. Effective FFF separation with high recovery usually requires high pH, up to 1 g/L surfactant, and 10 –100 mL of organic solvent (e.g., ethanol) per liter. Unfortunately, these conditions are likely to perturb the immunologic binding; thus, the mobile phase composition must be optimized to obtain efficient FFF separation while preserving antibody–antigen binding.

1994


Fig. 1. Principle of the multianalyte CL-FFF-based immunoassay (top), and results obtained with CL-FFF-based immunoassay for CAF (bottom). (Top left; a), injection and relaxation; (b), separation of the free tracer fraction and the various microsphere populations, with on-line CL measurement of enzyme activity. (Top right), representative CL fractogram in which the void peak corresponds to the unbound tracer fractions, whereas each of the retained peaks corresponds to tracer bound to one microsphere population. (Bottom left), optimized dose–response curve for the CAF assay. Data are shown as the bound-enzyme activity ratio (B/B0) plotted against the log of the CAF concentration. The bound-enzyme activity at each CAF concentration (B) and the bound-enzyme activity in the absence of CAF (B0) were calculated as the area of the retained peak in CL fractograms. Each data point represents the mean (SD; error bars) of 6 determinations. (Bottom right), representative CL fractograms. Shown are retained peaks at CAF concentrations of 0.03 (c), 0.3 (d), and 9.0 (e) ␮mol/L.

(B) Effective separation of bound and free tracer. In an FFF-assisted immunoassay, this is achieved when no free tracer is retained and no particles elute at the void time. Because a soluble molecule cannot be as retained as a micrometer-sized particle in a flow FFF system, it is then possible to detect particles at the void time when the relaxation procedure is not optimized.

(C) Run-to-run carryover effects must be avoided. This aspect is strictly related to the analytical recovery because low microsphere recovery might arise from accumulation of particles in the channel, and these might be eluted during the following run. High reproducible recovery, therefore, must be obtained.


These aspects were investigated as follows: 100 ␮g of antibody-coated microspheres was incubated with 100 ␮L of CAF-HRP diluted in IB, washed with IB through centrifugation (5000g for 5 min at room temperature) to eliminate free tracer, and then injected in the flow FFF-CL system. The stability of the immunologic binding was demonstrated when the CL and UV fractograms gave the same ratio of void-peak area to retained-peak area. The absence of unretained particles in the void peak was demonstrated by the absence of a void peak in the UV fractogram. Recovery was calculated as the ratio of the area of the retained peak (as detected by the UV/visible detector) to the peak area obtained by directly injecting spheres in the UV/visible detector cell at the same flow rate (8 ). Various mobile phase compositions were tested. In all cases, a longitudinal flow rate as high as possible was used to maximize the lift effect and thus increase analytical recovery (9 ). The maximum flow rate was limited by the postcolumn flow-injection setup: the backpressure generated by a high flow rate would hamper injection of the CL substrate at a flow rate sufficient for obtaining high tracer detectability. The experimental conditions that allowed relatively high mean (SD) recovery [79 (2)%; n ⫽ 15], nonsignificant presence of unretained microspheres, and negligible perturbing of the immunologic binding were as follows: 10 mmol/L Tris-HCl (pH 7.4) containing 0.5 g/L Tween 20 as mobile phase; longitudinal flow rate, 4.0 mL/min; cross-flow rate, 0.5 mL/min; and CL substrate injection flow rate, 400 ␮L/min. With these conditions, a sigmoidal dose–response curve was obtained for CAF (Fig. 1). The curve ranged from 0.03 to 15 ␮mol/L (0.01–5 ppm), with a limit of detection of 0.3 ␮mol/L (0.1 ppm), which was determined as the mean signal for the blank ⫺ 3 SD. This is the first example of an FFF-assisted CL immunoassay in which the area of a fractogram peak is used to obtain a dose–response curve. It should be noted that among liquid chromatographic techniques, FFF is particularly suited for this application because it ensures highly efficient bound/free separation in nondenaturing conditions. The coupling of FFF with CL detection allows fast, sensitive tracer detection. Finally, the use of a flowinjection setup allows high system flexibility because the composition of the CL substrate can be easily varied according to the type of enzymatic tracer. The results demonstrate the feasibility of an CL-FFF– based competitive immunoassay, thus opening the way for multianalyte immunoassays. In particular, various microsphere populations, characterized by different diameters, can be coated with an antibody specific for 1 analyte, mixed with a sample and with a combination of analyte–HRP tracers, and then separated by the FFF system. On-line detection of the CL signal would then allow independent quantification of each analyte based on the area of the corresponding retained peak, as shown in Fig. 1. In addition, several different enzymes suitable for CL detection can be conjugated to different analytes and then independently quantified with appropriate CL

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substrates. This approach would further increase the number of analytes that can be quantified in each sample. Finally, a multirun FFF system could be developed, in which multiple parallel channels (a channel array) simultaneously analyze different samples in 1 run. A combination of these approaches would allow the development of multiplexed, competitive immunoassays. This work was partially funded by MIUR (Italian Ministry for University and Research). References 1. Shimpf M, Caldwell K, Giddings JC, eds. Field-flow fractionation handbook. New York: John Wiley & Sons, 2000:616pp. 2. Melucci D, Guardigli M, Roda B, Zattoni A, Reschiglian P, Roda A. A new method for immunoassays using field-flow fractionation with on-line, continuous chemiluminescence detection. Talanta 2003;60:303–12. 3. Reschiglian P, Zattoni A, Melucci D, Roda B, Guardigli M, Roda A. Flow field-flow fractionation with chemiluminescence detection for flow-assisted, multianalyte assays in heterogeneous phase. J Sep Sci 2003;26:1417–21. 4. Roda A, Manetta AC, Portanti O, Mirasoli M, Guardigli M, Pasini P, et al. A rapid and sensitive 384-well microtitre format chemiluminescent enzyme immunoassay for 19-nortestosterone. Luminescence 2003;18:72– 8. 5. Degand G, Bernes-Duyckaerts A, Delahaut P, Maghuin-Rogister G. Determination of ␤-agonists in urine by an enzyme immunoassay based on the use of an anti-salbutamol antiserum. Anal Chim Acta 1993;275:241–7. 6. Reschiglian P, Melucci D, Zattoni A, Mallo´ L, Hansen M, Kummerow A, et al. Working without accumulation membrane in flow field-flow fractionation. Anal Chem 2000;72:5945–54. 7. Melucci D, Roda B, Zattoni A, Casolari S, Reschiglian P, Roda. A field-flow fractionation of cells with chemiluminescence detection. J Chromatogr A 2004;1056:229 –36. 8. Melucci D, Zattoni A, Casolari S, Reggiani M, Sanz R, Reschiglian P. Working without accumulation membrane in flow FFF. Effect of sample loading on recovery. J Liq Chromatogr Relat Technol 2002;25:2211–24. 9. Williams PS, Moon MH, Xu Y, Giddings JC. Effect of viscosity on retention time and hydrodynamic lift forces in sedimentation/steric field-flow fractionation. Chem Eng Sci 1996;51:4477– 88. DOI: 10.1373/clinchem.2005.053108

Recombinant Cell-Based Bioluminescence Assay for Androgen Bioactivity Determination in Clinical Samples, Elisa Michelini,1,2 Maria Magliulo,1 Piia Leskinen,3 Marko Virta,3 Matti Karp,3,4 and Aldo Roda1* [1 Department of Pharmaceutical Sciences, University of Bologna, Bologna, Italy; 2 Center for Applied Biomedical Research (CRBA), S. Orsola-Malpighi Hospital, Bologna, Italy; 3 Department of Biochemistry, University of Turku, Turku, Finland; 4 Tampere University of Technology, Institute of Environmental Engineering and Biotechnology, Tampere, Finland; * address correspondence to this author at: Department of Pharmaceutical Sciences, University of Bologna, Via Belmeloro 6, 40126 Bologna, Italy; fax 39-051343398, e-mail [email protected]] Androgens regulate several physiologic processes, such as normal prostate development and maintenance of male sexual function in adult life, and they are also involved in pathologic conditions, including prostate cancer, prepubertal gynecomastia, and premature pubarche. Androgens enter the target cell and bind to androgen receptor (AR), a ligand-dependent transcription factor in the nu-

1996


CH3

OH

CH3

(A), testosterone enters the cells, binds to hAR, activates the ARE sequences, and drives the expression of luciferase. (B), F, testosterone; f, DHT; Œ, androstenedione. The recombinant yeast cells were incubated with increasing concentrations of various androgens to evaluate the transactivation capacity of each steroid. Each point represents the mean (SD; error bars) of at least 3 replicates. The dose–response curves were obtained from 5 independent experiments.

Testosterone

O

CH 3

hAR

25

OH

CH 3

CH 3

OH

CH 3 O

hAR hAR

O

hAR hAR CH3

OH

CH3

CH3

Luc-skl

OH

CH3 O

O

B

Luc-skl

ARE

Corrected light signal

A Fig. 1. Schematic view of the recombinant yeast bioassay (A), and corrected dose–response curves for testosterone reflecting the transactivation capacity of different androgens (B).

20 15 10 5 0 10 -2

10 -1

10 0

10 1

10 2

10 3

10 4

10 5

Analyte (nmol/L) Bioluminescent signal

clear receptor superfamily comprising receptors for vitamin D3 and thyroid and steroid hormones (1, 2 ). After binding of the hormone, AR enters the nucleus and binds to the regulatory region of the target gene as a homodimer. Recent findings suggest that some environmental chemicals disrupt the endocrine system and cause adverse effects such as cancers and sexual abnormalities in humans and wildlife (3 ). These endocrine-disrupting compounds include several xenoestrogens, androgens, and antiandrogens, which present a variety of chemical structures and mechanisms of action (4, 5 ). Conventional detection methods for androgenic and antiandrogenic compounds cannot evaluate hormonal bioactivity, which is important in a wide range of clinical conditions, and no alternative direct and simple methods for measurement of plasma androgen bioactivity are available. In addition, the increased use of androgenic substances as therapeutic drugs and their abuse to enhance athletic performance require sensitive and rapid screening tests. The very low amounts of single analytes in anabolic cocktails and nutritional supplements are difficult to detect with standard techniques such as gas chromatography–mass spectrometry. Thus, bioassays that reveal the overall androgen bioactivity of complex mixtures would be more appropriate tools with great implications in antidoping analysis (6 – 8 ). We developed and validated a yeast-based bioassay to measure androgen bioactivity in clinical samples, including human serum, for the detection of androgen-like and antiandrogenic compounds (9 ). The bioassay is based on recombinant Saccharomyces cerevisiae BMA64-1A strain genetically engineered with the introduction of 2 plasmids: an expression plasmid (pG1AR), in which the expression of human androgen receptor (hAR) is regulated by a constitutive yeast promoter; and a reporter plasmid (YipLuc-ARE), in which the ARE sequences drive the expression of a truncated form of Photinus pyralis luciferase, used as reporter gene. The latter plasmid was integrated into the yeast genome to provide greater stability. In the presence of androgenic compounds, hAR moves into the nucleus and binds ARE sequences, leading to luciferase expression directly proportional to the an-

drogenic activity of the sample (Fig. 1A). The use of a luciferase without the peroxisomal targeting sequence (luc-skl) avoided luciferase importation into peroxisomes and produced higher light emission without interfering with normal host physiology (10 ). A recombinant yeast strain constitutively expressing the same luciferase in the plasmid PRS316-Luc was used as control to correct the bioluminescent signal according to cell vitality and nonspecific matrix effects (11 ). The 2 strains were routinely grown in selective synthetic complete (sc) medium containing 6.7 g/L yeast nitrogen base without amino acids, 1.4 g/L yeast synthetic drop-out media supplement, and 40 mL/L of a 500 g/L d-glucose solution. The control strain was grown in sc medium without uracil, whereas the androgen-responsive strain was grown in sc medium without uracil and tryptophan. Glycerol stocks of the 2 recombinant strains were prepared and kept at ⫺80 °C for long-term storage. Stock solutions of the compounds to be tested, all purchased from Sigma, were prepared in ethanol at a concentration of 10 mmol/L and stored at ⫺20 °C. The bioassay procedure was as follows: a 2-mL overnight culture was used to inoculate 18 mL of fresh selective sc medium. The A600 nm was monitored until it reached 0.4, at which point 90-␮L volumes of the cell suspension were transferred to wells in a sterile white 96-well microtiter plate with 10 ␮L of sample or calibrator solution. Plates were sealed to prevent evaporation and shaken for 1 min before incubation. After 2.5 h of incubation at 30 °C, luminescence was measured with a Luminoskan microplate reader as follows: 100 ␮L of 1 mmol/L dluciferin (supplied by Synchem) in 0.1 mol/L sodium citrate buffer (pH 5.0) was injected with a built-in reagent dispenser. After brief shaking, luminescence was measured with 5-s integration. Light emissions were expressed as relative light units. Dose–response curves for testosterone were produced in each plate for testosterone concentrations ranging from 0.01 to 104 nmol/L. The analytical signal was plotted as either light emission against log[testosterone] (uncorrected curves) or as the ratio of biosensor light emission


over control light emission against log[testosterone] (corrected curves). The corrected light signal was proportional to the testosterone concentration in the concentration range from 0.1 to 100 nmol/L. The detection limit of the bioassay, defined as the testosterone concentration that produced a luciferase activity 2 SD above the mean of 10 samples of charcoal-stripped serum (used to reduce the contaminating steroids from the serum), was 0.05 nmol/L. Increasing sample volumes did not yield an improvement of the assay sensitivity. The intraassay (estimated by assaying the same serum pools 6 times in a single assay) and the interassay (estimated by assaying duplicate samples from a serum pool on 6 separate days) variations were evaluated at 3 testosterone concentrations: low (1 nmol/L), middle (100 nmol/L), and high (1 ⫻ 104 nmol/L). The intra- and interassay CVs were ⬍13% and ⬍22%, respectively. In particular, for the lowest concentration (1 nmol/L), the intraassay and interassay CVs were 11% and 21%, respectively. We compared the results obtained by the yeast bioassay with results obtained by commercial androgen enzyme immunoassays. Serum androgen activity correlated well with serum testosterone concentration measured with standard enzyme immunoassays (r ⫽ 0.91; P ⬍0.0001; n ⫽ 10; see Fig. 1S in the Data Supplement that accompanies the online version of this poster abstract at http://www. clinchem.org/content/vol51/issue10/). Other hormones, such as 4-androstene-3␤,17␤-dione (or androstenedione) and dihydrotestosterone (DHT), were tested to verify the biosensor specificity. Different amounts of testosterone and other androgenic compounds were added to charcoal-stripped fetal calf serum, and the corresponding dose–response curves were obtained and compared with that of testosterone (Fig. 1B). DHT was the most active androgen: fetal calf serum containing 0.1 nmol/L DHT induced a light signal equal to ⬃1 nmol/L testosterone. 4-Androstene-3␤,17␤-dione was slightly less active than testosterone. These results agree with reported data (7 ). For analysis of samples with unknown composition, androgenic bioactivity was expressed in terms of testosterone equivalents. We investigated the effect of antiandrogens on the hAR by performing competitive assays. Testosterone was first added to charcoal-stripped FCS at a subsaturating concentration (10 nmol/L), after which increasing amounts of the tested compounds (bisphenol A, a plasticizer used in the production of epoxy resins and polycarbonate plastics, and vinclozolin, an antiandrogenic fungicide) were added; the resulting solution was subjected to the bioassay. The median inhibitory concentrations (IC50s) were 5 ␮mol/L for bisphenol A and 10 ␮mol/L for vinclozolin (data not shown). In addition, hydroxyflutamide and diethylstilbestrol (DES) were also shown to block androgen stimulation in a dose-dependent way. Hydroxyflutamide also displayed some agonistic activity at high concentrations, which is in agreement with previous findings (see Fig. 2S in the online Data Supplement) (12 ). Hydroxyflutamide had an IC50 of 0.5 nmol/L in the

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presence of 10 nmol/L testosterone. The difference between the IC50 values reported here and those reported earlier in other cell lines can be attributed to the presence of coactivators in different cell lines and to interference through other signal transduction pathways (13 ). The antiandrogenic activity of DES, together with other xenoestrogens, could explain the occurrence of several pathologies, such as hypospadias and cryptorchidism, associated with maternal exposure to DES and pesticides (14 ). To test a clinical application of the bioassay, we performed a preliminary screening of human serum samples, which were assayed without any sample pretreatment. The bioassay allowed differentiation between samples from males, females, and prepubertal boys, and all of the tested samples were above the assay detection limit. For clinical applications, this bioassay appears to be superior to conventional assays for steroid hormones based on immunologic detection. This assay allows evaluation of the overall androgenic effect instead of measuring single androgenic compounds; thus, it can be used to estimate circulating androgen bioactivity. Another advantage of this yeast-based bioassay is the absence of expression of endogenous receptors, such as glucocorticoid or progesterone receptor, which might interfere with the screening. Such interference is present in cell-based AR assays that use the Chinese hamster ovary (CHO) cell line and breast cancer cell lines (14, 15 ). Hence, the developed bioassay is a more robust model for screening both androgenic and antiandrogenic compounds. Because of the short incubation time (2.5 h), the assay should be less affected by toxic effects from the matrix and thus can be used for different sample matrices without sample pretreatment. Because preanalytical procedures required in bioassays using mammalian cells or other reporter genes are avoided, less time is required (16 ). The use of modified luciferase simplifies the assay procedure, allowing 1-step measurement and increasing the bioassay sensitivity with respect to previously reported bioassays. In view of the potentially extensive application of this assay, lyophilized strains could also be used to avoid the need for routine cultivation of the recombinant strains. This feature, associated with the 96-well plate format, makes the test even more suitable for different laboratories and for high-throughput screening. Further applications of this bioassay include measurement of serum and urine androgen bioactivity in children and adults to investigate the relationship between androgenic status and endocrine diseases as well as the detection of illegally used synthetic androgens for human and animal doping.

The work was partly supported by the Italian Ministry for University and Research (MIUR) and by Fondazione Cassa di Risparmio di Bologna. References 1. Kemppainen JA, Lane MV, Sar M, Wilson EM. Androgen receptor phosphorylation, turnover, nuclear transport, and transcriptional activation. Specificity for steroids and antihormones. J Biol Chem 1992;267:968 –74.

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2. Lin B, White JT, Lu W, Xie T, Utleg AG, Yan X, et al. Evidence for the presence of disease-perturbed networks in prostate cancer cells by genomic and proteomic analyses: a systems approach to disease. Cancer Res 2005;65: 3081–91. 3. Gaido KW, Leonard LS, Lovell S, Gould JC, Babai D, Portier CJ, et al. Evaluation of chemicals with endocrine modulating activity in a yeast-based steroid hormone receptor gene transcription assay. Toxicol Appl Pharmacol 1997;143:205–12. 4. Lathers CM. Endocrine disruptors: a new scientific role for clinical pharmacologists? Impact on human health, wildlife, and the environment [Review]. J Clin Pharmacol 2002;42:7–23. 5. Hutchinson TH, Brown R, Brugger KE, Campbell PM, Holt M, Lange R, et al. Ecological risk assessment of endocrine disruptors [Review]. Environ Health Perspect 2000;108:1007–14. 6. Lee HJ, Lee YS, Kwon HB, Lee K. Novel yeast bioassay for detection of androgenic and antiandrogenic compounds. Toxicol In Vitro 2003;17:237– 44. 7. Raivio T, Palvimo JJ, Dunkel L, Wickman S, Janne OA. Novel assay for determination of androgen bioactivity in human serum. J Clin Endocrinol Metab 2001;86:1539 – 44. 8. Bricout V, Wright F. Update on nandrolone and norsteroids: how endogenous or xenobiotic are these substances? [Review]. Eur J Appl Physiol 2004;92: 1–12. 9. Michelini E, Leskinen P, Virta M, Karp M, Roda A. A new recombinant cell-based bioluminescent assay for sensitive androgen-like compound detection. Biosens Bioelectron 2005;20:2261–7.

10. Leskinen P, Virta M, Karp M. One-step measurement of firefly luciferase activity in yeast. Yeast 2003;20:1109 –13. 11. Leskinen P, Michelini E, Picard D, Karp M, Virta M. Bioluminescent yeast assays for detecting estrogenic and androgenic activity in different matrices. Chemosphere;in press (dx.doi.org/10.1016/j.chemosphere. 2005.01.080). 12. Yeh S, Miyamoto H, Chang C. Hydroxyflutamide may not always be a pure antiandrogen. Lancet 1997;349:852–3. 13. Lemaire G, Terouanne B, Mauvais P, Michel S, Rahmani R. Effect of organochlorine pesticides on human androgen receptor activation in vitro. Toxicol Appl Pharmacol 2004;196:235– 46. 14. Roy P, Salminen H, Koskimies P, Simola J, Smeds A, Saukko P, et al. Screening of some anti-androgenic endocrine disruptors using a recombinant cell-based in vitro bioassay. J Steroid Biochem Mol Biol 2004;88:157– 66. 15. Paris F, Servant N, Terouanne B, Sultan C. Evaluation of androgenic bioactivity in human serum by recombinant cell line: preliminary results. Mol Cell Endocrinol 2002;198:123–9. 16. Blankvoort BM, de Groene EM, van Meeteren-Kreikamp AP, Witkamp RF, Rodenburg RJ, Aarts JM. Development of an androgen reporter gene assay (AR-LUX) utilizing a human cell line with an endogenously regulated androgen receptor. Anal Biochem 2001;298:93–102. DOI: 10.1373/clinchem.2005.053017