Clinical Chemistry 50, No. 12, 2004
lated folates (quantified as the sum of 5,10CH⫽THF and 5- or 10CHOTHF) made up, on average, 30% (range, 0 – 69%) of the TF in erythrocytes of individuals with the T/T genotype. The preliminary results for individuals with the C/T genotype (n ⫽ 5) indicated that erythrocytes contained only 5CH3THF polyglutamates. No THF was detected in the erythrocytes of individuals with either genotype. The discrepancies in findings between our work and that of Bagley and Selhub (12 ), i.e., we found small amounts of formylated folates in individuals with the C/C genotype and significant amounts of THF in individuals with the T/T genotype, might be explained by the difference in folate extraction. Bagley and Selhub (12 ) extracted frozen erythrocytes at high pH to prevent enzymatic deconjugation of folate polyglutamates. We used WB lysates generated at low pH (4.0) to obtain folate monoglutamates by the action of plasma pteroylpoly-␥glutamate hydrolase. Although this is the generally recommended and clinically used procedure for WB folates, it is conceivable that folate metabolism takes place during the lysis process. As a result, some folate forms could be found in conventionally prepared WB lysates that might not be present in native erythrocytes. The THF we find is most likely the dissociation product of 5,10-methylenetetrahydrofolic acid losing the C-1 group. We analyzed ⬃100 paired serum and WB lysate samples to assess what proportion of folates found in WB originates from serum. In general, we found no FA in WB lysates with the exception of those from two individuals who had serum FA concentrations ⬎25 nmol/L, which spilled over into the WB lysate. Serum did not have quantifiable amounts of THF or 5,10CH⫽THF. On average, ⬃95% each of 5CH3THF and 5CHOTHF in WB lysates originated from erythrocytes, whereas the rest originated from serum. All of the THF and 5,10CH⫽THF found in WB lysates originated from erythrocytes. The presented method measures folate monoglutamates only. If deconjugation of polyglutamates is incomplete, the TF concentration will be underestimated. Our findings on folate extraction from WB and the comparability of results obtained with different methods will be presented in a subsequent report. This is the first reported method for measuring intact folate monoglutamates in serum and conventionally prepared WB lysates by LC/MS/MS using automated highthroughput SPE. The method displays excellent sensitivity and satisfactory precision, accuracy, and stability of folates during the 5-h SPE procedure. The specificity of this method is a clear advantage when it comes to quantifying individual folate forms. It can be a disadvantage when measurement of TF is of interest because the only folate forms measured are those that one is aware of. However, this highly specific method has a high potential of becoming a candidate reference method and helping to establish a standard reference material, which ultimately is needed to standardize clinical assays. The method’s automation and high sample throughput make it attractive for future use in clinical laboratories when further studies of the method have been completed.
2381
We thank Dr. Les McCoy for input and thoughtful discussions related to the mass spectrometry aspect of the method and Dr. Michael Rybak for coordinating the blood collection to evaluate the effect of sample type. References 1. MRC Vitamin Study Research Group. Prevention of neural tube defects: results of the Medical Research Council Vitamin Study. Lancet 1991;338: 131–7. 2. Mangoni AA, Jackson SHD. Homocysteine and cardiovascular disease: current evidence and future prospects. Am J Med 2002;112:556 – 65. 3. Choi SW, Mason JB. Folate and carcinogenesis: an integrated scheme. J Nutr 2000;130:129 –32. 4. Seshadri S, Beiser A, Selhub J, 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;346:476 – 83. 5. US Food and Drug Administration. Food labeling: health claims and label statements; folate and neural tube defects. Fed Regist 1993;58:53254 – 95. 6. Gunter EW, Bowman BA, Caudill SP, Twite DB, Adams MJ, Sampson EJ. Results of an international round robin for serum and whole-blood folate. Clin Chem 1996;42:1689 –94. 7. Pfeiffer CM, Gunter EW, Caudill SP. Comparison of serum and whole-blood folate measurements in 12 laboratories: an international study [Abstract]. Clin Chem 2001;47(Suppl 6):A62–3. 8. Hoppner K, Lampi B. Reversed phase high pressure liquid chromatography of folates in human whole blood. Nutr Rep Int 1983;27:911–7. 9. Leeming RJ, Pollock A, Barley C. A critical assessment of methods for measuring folate in human serum and red blood cells. In: Cortios H-C, Ghisla S, Blau U, eds. Chemistry and biology of pteridines. Berlin: Water de Gruyter & Co., 1990:188 –91. 10. Wigertz K, Jaegerstad M. Comparison of a HPLC and radioprotein-binding assay for the determination of folates in milk and blood samples. Food Chem 1995;54:429 –36. 11. Santhosh-Kumar CR, Kolhouse JF. Molar quantitation of folate by gas chromatography-mass spectrometry. Methods Enzymol 1997;261:26 –34. 12. Bagley PJ, Selhub J. A common mutation in the methylenetetrahydrofolate reductase gene is associated with an accumulation of formylated tetrahydrofolates in red blood cells. Proc Natl Acad Sci U S A 1998;95:13217–20. 13. Dueker SR, Lin Y, Jones D, Mercer R, Fabbro E, Miller JW. Determination of blood folate using acid extraction and internally standardized gas chromatography-mass spectrometry detection. Anal Biochem 2000;283:266 –75. 14. Lin Y, Dueker SR, Jones AD, Clifford AJ. A parallel processing solid phase extraction protocol for the determination of whole blood folate. Anal Biochem 2002;301:14 –20. 15. Lin Y, Dueker SR, Clifford AJ. Human whole blood folate analysis using a selected ion monitoring gas chromatography with mass selective detection protocol. Anal Biochem 2003;312:255–7. 16. Pfeiffer CM, Fazili Z, McCoy L, Zhang M, Gunter EW. Determination of folate vitamers in human serum by stable-isotope-dilution tandem mass spectrometry and comparison with radioassay and microbiologic assay. Clin Chem 2004;50:423–32. 17. Friso S, Choi S-W, Girelli D, Mason JB, Dolnikowski GG, Bagley PJ, et al. A common mutation in the 5,10-mehylenetetrahydrofolate reductase gene affects genomic DNA methylation through an interaction with folate status. Proc Natl Acad Sci U S A 2002;99:5606 –11. Previously published online at DOI: 10.1373/clinchem.2004.036541
Nonparametric Determination of Reference Intervals for Plasma Metanephrine and Normetanephrine, Emily C. Heider,1 Bret G. Davis,2 and Elizabeth L. Frank3* (1 ARUP Institute for Clinical and Experimental Pathology, Salt Lake City, UT; 2 ARUP Laboratories, Inc., Salt Lake City, UT; 3 Department of Pathology, University of Utah Health Sciences Center, Salt Lake City, UT; * address correspondence to this author at: Department of Pathology, University of Utah, c/o ARUP Laboratories, 500 Chipeta Way, Salt Lake City, UT 84108; fax 801-584-5207, e-mail e.frank@ aruplab.com)
2382
Technical Briefs
Measurement of plasma metanephrine and normetanephrine concentrations has recently been acknowledged as one of the leading tests for the diagnosis of the neuroendocrine tumor, pheochromocytoma (1 ). The occurrence of the tumor is rare, and it is often difficult to diagnose. The wide spectrum of signs and symptoms includes hypertension, headaches, and diaphoresis. Located primarily in the adrenal gland, these tumors produce excess catecholamines and their metabolites. Assays have been developed to measure catecholamines and metanephrines in both plasma and urine. Although tests for all of these compounds can be diagnostically useful, measurement of metanephrine and normetanephrine in plasma has been reported as a particularly sensitive and specific indicator of the disease. The utility of the test is limited by uncertainty of the predictive value of measured concentrations. This is attributable in part to ambiguity in published reference intervals for these analytes. Lenders et al. (1 ) reported upper reference limits of 0.3 nmol/L for plasma metanephrine and 0.6 nmol/L for normetanephrine, whereas a reference laboratory (2 ) reported values ⬍0.5 nmol/L for metanephrine and ⬍0.9 nmol/L for normetanephrine as within the reference interval. In this study, we established a reference interval by measuring metanephrine and normetanephrine concentrations in plasma collected from 120 volunteers after obtaining informed consent. Reference individuals (53 females and 67 males; age range, 20 –73 years) fasted for 12 h before sampling. Interviews were conducted to excuse individuals taking medications, particularly for hypertension. All participants were sitting upright as their blood was collected by needlestick. The whole blood, in EDTA, was stored at 2– 8 °C until it was centrifuged within a few hours after collection. Plasma was separated and stored at ⫺70 °C until analysis. The conditions under which the volunteers were prepared and blood was collected were selected based on evaluation of studies in the literature. Volunteers were fasting to eliminate the interference of dietary constituents in the HPLC analysis, as observed by Lenders et al. (1 ). For this reason, medical practitioners are advised to instruct patients to fast for 12 h before blood collection for plasma metanephrine analysis. Additionally, as published elsewhere, plasma catecholamine concentrations are posture-dependent, but plasma metanephrine concentrations are not significantly altered if a patient is sitting or lying supine (3 ). Metanephrines were assayed by a procedure similar to the method developed by Lenders et al. (4 ). Contaminants were removed by solid-phase extraction with BondElut® AccuCAT columns (Varian, Inc.). Plasma (1 mL) was applied to the columns with an internal standard, 4-hydroxy-3-methoxybenzylamine. The analytes were eluted with 10% ammonium hydroxide in methanol (1:3 by volume), and the eluate was evaporated to dryness under nitrogen at 37 °C. The residue was reconstituted in 115 L of 0.2 mol/L acetic acid, and 80 L was injected for separation by reversed-phase HPLC. The mobile phase consisted of acetonitrile (70 mL/L) in sodium phosphate
buffer, pH 3.25. The mobile phase flow rate was 1.2 mL/min. The analytical column was a LUNA C18 reversed-phase column [250 ⫻ 4.6 mm (i.d.); 5-m bead size; Phenomenex]. The ESA 580 pump, 540 autosampler, and ESA 316 CoulArray Detector were from ESA, Inc. The analytes were detected coulometrically with a conditioning cell (5021; ESA) electrode at 410 mV and an analytical cell (5011; ESA) at ⫺310 mV. All of the compounds were separated and detected within 20 min. The retention times for metanephrine, normetanephrine, and the internal standard were 9.5, 7.1, and 11.7 min, respectively. The total run-time was extended to 31 min because of lateeluting compounds. A five-point calibration curve with the internal standard was used to calibrate the method. Analytes were quantified by comparison of the peak height and internal standard ratio with the calibration curve. A range of concentrations of metanephrine and normetanephrine was injected to determine the linearity of the method. The recovery of this method was established by analyzing solutions of known concentrations and plotting the known samples against the values acquired by the analysis. Over the linear range of the method, recovery for metanephrine was 100.3% of the expected amount and recovery for normetanephrine was 102.5%. Imprecision was determined by assaying each of three quality-control solutions in duplicate on 15 different days. This method was compared with a similar HPLC method (with cation-exchange, solid-phase extraction, and electrochemical detection) at a reference laboratory. Thirty samples, with concentrations spanning the linear range of the method, were analyzed by both methods, and data were evaluated by plotting the points for comparison. For metanephrine, the standard error of the estimate (Sy兩x) was 0.20 nmol/L, unfortunately high because of a difference in detection limits for the two methods. Results ⬎0.15 nmol/L were calculated by this method, but the reference laboratory did not quantify results ⬍0.2 nmol/L. The standard error of the estimate was 0.28 nmol/L for normetanephrine. The performance characteristics data are summarized in Table 1. Acetaminophen, caffeic acid, and ephedrine did not interfere with this assay at physiologic concentrations. The reference interval data were analyzed according to the NCCLS guideline for determining reference values (5 ). Data points were placed in decreasing order and then evaluated to eliminate outliers. The distribution of results was nongaussian for both analytes (Fig. 1). The central 95% of the data were taken as the reference interval for each analyte. Results ranged from ⬍0.15 to 0.37 nmol/L for metanephrine and from ⬍0.16 to 0.94 nmol/L for normetanephrine. Fifty-three percent of the metanephrine data fell below 0.15 nmol/L, the limit of quantification for this assay. The mean and median values were 0.16 and 0.15 nmol/L, respectively, for metanephrine and 0.40 and 0.36 nmol/L for normetanephrine. With 90% confidence, the upper reference limits were determined to be 0.29 nmol/L for metanephrine and 0.77 nmol/L for normetanephrine.
Clinical Chemistry 50, No. 12, 2004
Table 1. Validation data for the plasma metanephrine assay.a Metanephrineb
Imprecision Low control Mid control High control Method comparison n Slope Intercept, nmol/L R Sy兩x, nmol/L Reference interval, nmol/L
Normetanephrinec
Mean, nmol/L
CV, %
Mean, nmol/L
CV, %
0.21 0.43 0.84
22 16 8.2
0.35 0.80 2.70
15 10 8.6
30 1.226 0.022 0.977 0.20 0.00–0.29
30 0.931 0.041 0.968 0.28 0.00–0.77
a
Statistics were obtained using EP Evaluator. Linear range, 0.15–5.07 nmol/L. c Linear range, 0.16 –5.50 nmol/L. b
The reference intervals calculated in this study were comparable to those reported previously in the literature and fall between those of the two cited reports (1, 2 ). Reference interval values are included in Table 1.
Distribution of results for metanephrine
Frequency
A 70 60 50 40 30 20 10 0 0.15
0.19
0.23
0.27
0.31
Metanephrine (nmol/L)
Distribution of results for normetanephrine
Frequency
B 30 25 20 15 10 5 0
2383
We analyzed patient results over a 6-month period with the newly determined reference limits. Of 1657 patient samples, 20.0% were increased with our upper reference limit, 0.29 nmol/L; 17.6% of the metanephrine values were increased by the criterion of 0.5 nmol/L suggested by Sawka et al. (2 ). For normetanephrine, 25.1% were increased based on our upper reference limit of 0.77 nmol/L, and 4.6% of the values were increased according to the upper reference limit of 0.9 nmol/L reported by Sawka et al. (2 ). As reported by Kudva et al. (6 ), cutoff concentrations for some analytes used to diagnose pheochromocytoma are adjusted to approximately twice the upper limit found in a healthy population. This technique has been found to provide optimal specificity while maintaining acceptable sensitivity. Although the authors do not state that this method was used to establish reference limits for plasma metanephrine and normetanephrine concentrations, it is interesting to note that this would account for the differences between reference limits used by Sawka et al. (2 ) and those determined in our study of a healthy population as well as those used by Lenders et al. (1 ). Definition of a positive result as a multiple of the upper limit of the 95% population reference interval may be of value in minimizing false-positive results and is consistent with the accepted medical use of biochemical testing to aid clinical judgment in the evaluation of pheochromocytoma (7 ). Adoption of lower reference limits for metanephrine and normetanephrine, such as those determined in our study, may lead to an increase in reported positive results. In this case, additional biochemical testing and radiologic imaging will be required for the diagnostic confirmation of pheochromocytoma. Although false-positive tests may lead to increased costs to the patient, false-negative results are far more destructive and are potentially fatal. This study established concise numerical limits for metanephrine and normetanephrine concentrations in the plasma of a population of healthy individuals with no indication of disease. The results of the study may be of value in the interpretation of plasma metanephrine concentrations used to support a clinical diagnosis of pheochromocytoma.
Research for this work was supported by the ARUP Institute for Clinical and Experimental Pathology. References
0.16
0.31
0.48
0.66
0.83
Normetanephrine (nmol/L) Fig. 1. Distribution of results for plasma metanephrine (A) and normetanephrine (B). n ⫽ 120 for both sets of results.
1. Lenders JW, Pacak K, Walther MM, Linehan WM, Manneli M, Friberg P, et al. Biochemical diagnosis of pheochromocytoma: which test is best? JAMA 2002;287:1427–34. 2. Sawka AM, Jaeschke R, Singh RJ, Young WF Jr. A comparison of biochemical tests for pheochromocytoma: measurement of fractionated plasma metanephrines compared with the combination of 24-hour urinary metanephrines and catecholamines. J Clin Endocrinol Metab 2003;88:553– 8. 3. Raber W, Raffesberg W, Bischof M, Scheuba C, Niederle B, Gasic S, et al. Diagnostic efficacy of unconjugated plasma metanephrines for the detection of pheochromocytoma. Arch Intern Med 2000;160:2957– 63. 4. Lenders JW, Eisenhofer G, Armando I, Keiser HR, Goldstein DS, Kopin IJ. Determination of metanephrines in plasma by liquid chromatography with electrochemical detection. Clin Chem 1993;39:97–103. 5. NCCLS—Second edition. NCCLS document C28-A2. How to define and
2384
Technical Briefs
determine reference intervals in the clinical laboratory; approved guideline. Wayne, PA: National Committee for Clinical Laboratory Standards, 2000. 6. Kudva YC, Sawka AM, Young WF. Clinical review 164: the laboratory diagnosis of adrenal pheochromocytoma: the Mayo Clinic experience [Review]. J Clin Endocrinol Metab 2003;88:4533–9. 7. Bravo EL, Tagle R. Pheochromocytoma: state-of-the-art and future prospects [Review]. Endocr Rev 2003;24:539 –53. DOI: 10.1373/clinchem.2004.035089
Gene Expression Profiles in Formalin-Fixed, ParaffinEmbedded Tissues Obtained with a Novel Assay for Microarray Analysis, Marina Bibikova,1 Joanne M. Yeakley,1 Eugene Chudin,1 Jing Chen,1 Eliza Wickham,1 Jessica WangRodriguez,2 and Jian-Bing Fan1* (1 Illumina, Inc., San Diego, CA; 2 Veterans Affairs Hospital, University of CaliforniaSan Diego, San Diego, CA 92161; * address correspondence to this author at: Illumina, Inc., 9885 Towne Centre Dr., San Diego, CA 92121-1975; fax 858-202-4680, e-mail
[email protected]) Gene expression profiling using microarrays has revolutionized the analysis of biological samples. In clinical applications, microarray data have been used to successfully distinguish among patients exhibiting similar symptoms (1, 2 ). Early demonstrations of this power were in the diagnosis of subtypes of acute leukemia (3 ) and diffuse large B-cell lymphomas (4 ), and such analyses are gradually gaining acceptance for diagnostic and prognostic applications (5–7 ). Investigators are currently accumulating microarray data for a broad assortment of such studies but are limited by the requirement of fresh/frozen tissues for sample preparation and labeling (8 ). This limitation requires the accumulation of fresh samples throughout the course of the disease, which may involve years of monitoring. However, formalin-fixed, paraffinembedded (FFPE) tissues are widely available and have the advantage of a known patient outcome and drug response history. RNAs derived from these samples are commonly badly degraded and have not been useful for conventional microarray studies (9, 10 ). We applied a novel expression assay to simultaneously monitor 502 cancer-related genes in RNAs derived from FFPE samples, using microarrays assembled on fiber optic bundles. Our results suggest that this approach can be used for extending microarray analyses to RNAs derived from archival tissue samples. We have recently developed a gene expression method called the DASLTM assay (cDNA-mediated annealing, selection, extension, and ligation) (11 ). This assay targets gene-specific sequences, using pools of chimeric query oligonucleotides. The oligonucleotides all share common primer landing sites so that once the upstream oligonucleotide is extended and ligated to the downstream oligonucleotide, an amplifiable product is generated. One PCR primer pair is used to amplify all of the amplifiable templates and generate amplicons of similar size (⬃100 bp). This uniformity minimizes potential bias during
amplification of many different targets. Currently, the DASL assay can be multiplexed to monitor hundreds of genes (11 ). To allow the use of universal microarrays, the downstream query oligonucleotides also contain a unique address sequence that is associated with each gene. This address sequence allows the amplified product, which is labeled during PCR with a fluorescent primer, to hybridize to a microarray bearing the complementary address sequences. This feature provides ready flexibility: to change the genes being monitored, the address sequences can be reassigned and the query oligonucleotide pool resynthesized, using the same arrays. Finally, the cDNA synthesis step is performed with both oligo(dT) and random priming, which frees the assay from dependence on an intact polyA tail, unlike the usual T7 promoter-oligo(dT) priming method for microarray sample preparation (12 ). The use of random hexamers or nonamers in the cDNA synthesis allows representation of targeted cDNA sequences despite RNA degradation. Less than 50% of target probes in FFPE samples were detected when only oligo(dT) primer was used for cDNA synthesis, compared with the use of both oligo(dT) and random primers. To monitor gene expression in FFPE samples, we prepared RNA from 5-m human tissue sections mounted on microscope slides, obtained from BioChain Institute according to an Institutional Review Board-approved protocol. Briefly, the slides were deparaffinized in xylenes, and tissue samples were then scraped off with razor blades and held in xylenes until subsequent processing. RNA was extracted by use of the High Pure RNA Paraffin Kit (Roche Applied Science) and quantified by use of RiboGreen (Molecular Probes). When measured on Bioanalyzer (Agilent) RNA Pico Chips, the size of the RNA fragments ranged from 100 to ⬎500 nucleotides, with a peak maximum at ⬃130 nucleotides. On average, 1–2 g of total RNA was isolated from five 5-m tissue sections. For each sample, 200 ng of total RNA was converted to cDNA and processed in the DASL assay as described previously (11, 13 ). Oligonucleotides targeting 502 cancer-related genes were used in these experiments, at a density of three nonoverlapping probes per gene, giving a 1506-plex measurement for each sample. Our previous study showed that three probes per gene lend the assay sufficient sensitivity and reproducibility for quantitative detection of differential expression in FFPE tissues (13 ). Mean signal values were computed for each gene by determining the mean signal for the three representative probes (11 ). Because DASL uses random priming in the cDNA synthesis, the probes can be designed to target any unique regions of the gene without the need to limit the selection of optimal probes to the 3⬘ end of transcripts. We profiled 16 FFPE samples of four tissue types— prostate, colon, breast, and lung—with each tissue represented by one nondiseased and several cancer samples. As shown in Fig. 1, the DASL assay gave highly reproducible intensity measurements for FFPE samples preserved for 1.5–3 years. The correlations (r2) between the
