Printer Friendly

High-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders.

Mitochondria play a vital role in cellular energy metabolism. The enzymes of the mitochondrial energy-generating system (MEGS) [2] are localized in the mitochondrial matrix space [pyruvate dehydrogenase complex (PDHc) and tricarboxylic acid (TCA) or Krebs cycle] and in the inner mitochondrial membrane [oxidative phosphorylation (OXPHOS) complex] (1). Enzyme analysis of individual OXPHOS complexes in skeletal muscle biopsy remains the mainstay of the diagnostic process for patients suspected of mitochondrial cytopathy (2). In addition, measurement of the MEGS capacity is a powerful tool for the assessment of mitochondrial function. The MEGS capacity can be examined in detail by measurement of 14CO2 production rates from oxidation of [1-14C]pyruvate and carboxyl14C-labeled TCA cycle intermediates in combination with measurement of ATP production in intact mitochondria from a muscle biopsy (1), or by examining mitochondrial respiration via oxygen consumption measurements using polarography. These methods have been demonstrated to be very valuable in the examination of fresh muscle tissue. For fibroblasts, several methods have been described to measure ATP production or oxygen consumption (3,4). Analysis of fibroblasts is useful for several reasons (5), including confirmation of the defects observed in muscle, to exclude secondary mitochondrial dysfunction, e.g., due to poor feeding or disuse (6), and serving as a model system for follow-up and functional testing. One of the most informative ways of assessing mitochondrial function is by analysis of mitochondrial oxygen consumption by polarography using the Clark-type oxygen electrode (7). With this method, oxygen consumption can be measured in isolated mitochondria and in intact and detergent-permeabilized cells (8). Although this technique has proven very useful, a major limitation can be the relatively low sample throughput. An assay setup with a higher throughput has recently become available, but requires specialized equipment (9). Quenched-fluorescence oxygen sensing is an approach that may overcome these limitations. This measurement methodology allows the assessment of mitochondrial function using conventional instrumentation, thereby combining the high degree of information provided by oxygen consumption analysis with the simplicity, throughput, and scaling-up capabilities of standard microtiter plate assays (10). This approach (10) has previously been used to investigate mitochondrial toxicity of drugs in isolated mitochondria (7) and in cultured adherent cells (11).

The aim of our study was to adapt a protocol allowing quenched-fluorescence oxygen sensing to be used in the diagnostic evaluation of suspected mitochondrial disorders. We demonstrate that this assay provides a fast and nonlaborious high-content screening method for mitochondrial function in different digitonin-permeabilized cell types.

Materials and Methods

This study was carried out in the Netherlands in accordance with the applicable rules concerning research ethics committee review (Commissie Mensgebonden Onderzoek Regio Arnhem-Nijmegen) and informed consent.


We purchased pyruvate and oligomycin from Sigma, malate from Fluka, and ADP from Roche. All other chemicals were of the highest purity commercially available. Phosphorescent oxygen-sensitive probe, MitoXpress[TM] (A65N-1), was from Luxcel Biosciences. Black-body clear-bottom 96-well plates (Black Isoplate TC) and black backing tape were from PerkinElmer.


After receiving informed consent from patients and their caretakers, we obtained skin biopsy specimens for diagnostic purposes and cultured fibroblasts in E199 medium (Gibco; Invitrogen). We cultured transmitochondrial cybrids in high-glucose (4.5 g/L) Dulbecco's modified Eagle's medium (DMEM). Both media were supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (100 U/mL and 100 [micro]g/ mL, respectively). Before harvesting, fibroblasts were incubated for 48 h in galactose medium, consisting of DMEM without glucose, without pyruvate, with L-glutamine 2 mmol/L (Invitrogen) supplemented with 10% dialyzed FBS, 1 mmol/L uridine, 1% penicillin/streptomycin, and 5.5 mmol/L D-galactose. Cells were harvested and washed twice in ice-cold 1% FBS in PBS, and we resuspended the cell pellet at 30% (wt/vol) (4-6 x [10.sup.4] cells/[micro]L) in ice-cold 1% FBS in PBS.


We obtained human skeletal muscle from healthy young adults after receiving informed consent. Muscle mitochondria were isolated as described (12). Briefly, we placed the muscle specimens in ice-cold 0.25 mol/L sucrose, 2 mmol/L EDTA, 10 mmol/L Tris-HCl, 5 x [10.sup.4] U/L heparin (SETH) medium. After chopping, homogenization was performed by hand with a Teflon-glass Potter-Elvehjem-type homogenizer. The resulting homogenate was centrifuged for 10 min at 600g. Mitochondria were isolated from the resulting supernatant by centrifugation for 10 min at 14 000g. The mitochondrial pellet was finally taken up in SETH-medium.


We dissolved MitoXpress (A65N-1) oxygen probe, supplied as dry reagent in a vial, in PBS to a concentration of 1 [micro]mol/L. Probe fluorescence is quenched by molecular oxygen via a nonchemical (collisional) mechanism, resulting in an increase in probe signal with decreasing concentration of dissolved oxygen due to mitochondrial respiration (10). We performed measurements immediately after harvesting in digitonin-permeabilized nonfractionated cells with the addition of substrates; we also tested isolated fresh human skeletal muscle mitochondria. The incubations were pyruvate + malate with or without ADP. The incubation volume in each well was 50 [micro]L, containing 5 [10.sup.4]L cell suspension (2-3 x [10.sup.5] cells), 100 nmol/L fluorescent probe, and a substrate mix consisting of 30 mmol/L potassium phosphate, 75 mmol/L potassium chloride, 8 mmol/L Tris, 1.6 mmol/L EDTA, 5 mmol/L [MgCl.sub.2], 0.2 mmol/L [p.sup.1],[p.sup.5]-di(adenosine-5') pentaphosphate (myo-adenylate kinase inhibitor), 32.6 [micro]mol/L digitonin for the permeabilization of the cell membranes, 1 mmol/L pyruvate, 1 mmol/L malate, and, where indicated, 2.0 mmol/L ADP, pH 7.4 (1). To regenerate ADP by creatine kinase in the cell preparations, we added 20 mmol/L creatine to all ADP-containing incubations. We added compound solutions to the wells to give the indicated final concentrations. Finally, we added 100 [micro]L mineral oil to each well to seal the samples from ambient oxygen, which would otherwise destroy oxygen gradients and compromise assay performance (10) and placed the plate in a time-resolved fluorescence plate reader (Viktor 2; PerkinElmer) equilibrated to a temperature of 37[degrees]C and monitored every 5 min over 55 min. Instrument settings were 340/ 642 nm excitation/emission filters, delay time of 30 [micro]s, measurement window of 100 [micro]s, and active temperature control of the microplate compartment at 37[degrees]C. After completion of fluorescence measurements, time profiles of fluorescence intensity in each well were analyzed using Viktor (PerkinElmer) and Excel (Microsoft) software. All measurements were performed in duplicate and the results averaged. We calculated relative oxygen consumption rate (ROCR), reflecting respiratory activity, from the maximal linearly increasing fluorescence intensity slope (Fig. 1) to compare respiration rates of different samples. The results were normalized for citrate synthase (CS) activity (13).


The different fibroblast groups were tested for goodness of fit to the normal distribution using the Shapiro-Wilk test. The Kruskal-Wallis test was applied to study differences in respiration rates between the investigated groups. Differences were considered statistically significant if the P value was lower than 0.05. When overall significance was found, we used the Mann-Whitney test with Bonferroni correction (P value <0.0125) as a posthoc test to compare each patient group to the control.


To evaluate the maximal mitochondrial oxygen consumption capacity of the samples, we did measurements in nonfractionated digitonin-permeabilized cells under optimal substrate concentrations and in the presence of an excess of ADP. Most experiments were performed in fibroblasts to be able to exclude secondary mitochondriopathies. Therefore, different media were tested (data not shown). We used galactose-based medium, which drives the cell toward the OXPHOS system for ATP production, thereby increasing the differences between control and patient fibroblasts (14, 15), for subsequent experiments in fibroblasts. Supplemental Table 1 (which accompanies the online version of this article at content/vol56/issue3) gives an overview of the cell lines used for oxygen consumption measurements in fibroblasts.


In control fibroblasts, pyruvate-malate--driven respiration yielded substantially higher respiratory activity upon ADP stimulation (Fig. 1A), indicating proper functional coupling of the mitochondria. This was also clearly seen in isolated mitochondria obtained from control fresh muscle biopsies. Muscle showed both a higher respiratory activity rate and a higher coupling efficiency than fibroblasts (Fig. 1B). During measurement, the initial reduction in raw signal reflects plate temperature equilibration (11). This is followed by a linear phase of oxygen consumption that can be used to analyze mitochondrial respiration. The linear phase of oxygen consumption ends owing to oxygen exhaustion (10), based on results of pyruvate oxidation rate measurements following the methods outlined in Janssen et al. (1). In cultured control fibroblasts, the pyruvate oxidation rate is between 0.1 and 0.15 nmol/min for 5 [micro]L cells. At this rate, 40 min of pyruvate oxidation produces 16-24 nmol of NADH and reduces 8 -12 nmol [O.sub.2], whereas the reaction volume of 50 [micro]L at 37[degrees]C and 1 bar (100 kilopascals) pressure contains only 10.7 nmol [O.sub.2]. In control muscle mitochondria, the pyruvate oxidation rate is 0.55-0.65 nmol/min for 5 [micro]L mitochondrial suspension, producing 28-32 nmol NADH and reducing 14-16 nmol [O.sub.2] in 12 min. The addition of a layer of mineral oil to each sample limited back-diffusion of oxygen (10). The presence of low-rate back-diffusion through the mineral oil seal and polystyrene plate (oxygen diffuses through polystyrene at considerable rates) determined the lower limit of assay sensitivity, i.e., mitochondrial oxygen consumption below this rate cannot be analyzed (10). This is in line with previous observations (7, 11). The signal stabilizes when equilibrium is reached between the rates of oxygen consumption and back-diffusion, so successful analysis in the microplate assay requires an oxygen consumption rate sufficient to produce a measurable signal change (10). Because of the back-diffusion, the rates of change of dissolved oxygen measured using this assay are lower than the actual mitochondrial oxygen consumption rate. Therefore, they were expressed as ROCR in relative fluorescence units (RFU) per unit CS. The correlation between initial rates of increase of probe signal and the corresponding mitochondrial protein concentration has been demonstrated to be close to linear (7), which made it possible to compare respiration rates of different samples by analyzing the ROCR of the linearly increasing respiratory activity slope (Fig. 1). Control fibroblasts typically exhibit an ROCR >3 RFU/U CS [mean (SD) 5.0 (2.0)]. We saw that the oxygen consumption rate can decrease when reaching a higher cell culture passage number (Fig. 2 and online Supplemental Table 1).


To study the sensitivity of the assay, several classical inhibitors of the mitochondrial energy-generating system were added to control fibroblasts: rotenone (for complex I), antimycin A (complex III), azide (complex IV), oligomycin (complex V), atractyloside [adenine nucleotide transporter (ANT)], [alpha]-hydroxycyanocinnamate (pyruvate carrier), and arsenite (PDHc and [alpha]-ketoglutarate dehydrogenase). Fig. 3A shows that all inhibitors significantly reduced oxygen consumption. Similar results were obtained with isolated mitochondria from a control fresh muscle biopsy (Fig. 3B). This indicates that oxygen consumption measured in this assay depends on a fully functional MEGS system. Moreover, it should be feasible to pick up deficiencies in any of its components.



For diagnostic purposes, it is important that deficiencies in the MEGS system can be diagnosed or confirmed in fibroblasts. Therefore, we measured patient fibroblasts with various known OXPHOS defects (Fig. 4). Patient cell lines with high passage number and low respiratory activity were not included in this study to prevent possible bias by aging (see Fig. 2). There was an overall difference in respiration rates between the groups (Kruskal-Wallis test, P = 0.002). All complex I- and complex IV-deficient fibroblasts showed an ADP-stimulated respiratory activity rate below the lowest control value. There was a significant difference between the control group and the complex I- (P = 0.001) and complex IV- (P = 0.008) deficient fibroblasts, and no significant difference when comparing the controls with the complex V-deficient fibroblasts (P = 0.427) (Fig. 4). A characteristic feature of the complex V-deficient cells was the clear stimulation of oxygen consumption by addition of an uncoupler [2 [micro]mol/L carbonyl cyanide 3-chlorophenyl hydrazone (CCCP)] (Fig. 5A). This phenomenon was not seen in control cells (Fig. 5B) or cells with other OXPHOS enzyme deficiencies (data not shown). These data indicate that the fluorescence-based oxygen sensing probe assay is a sensitive tool to measure mitochondrial function in fibroblasts.

For patients with non-OXPHOS-related decreases in MEGS capacity in fresh muscle biopsies, the oxygen consumption rate would be an important tool to confirm the deficiency, since traditional OXPHOS enzyme assays are not discriminative. We included several fibroblast cell lines from this patient category. The level of oxygen consumption upon ADP stimulation ranged from control respiratory activity rates to, predominantly, low to very low rates (Fig. 4). As a group, they were not statistically significant (P = 0.089), but nevertheless several of the non-OXPHOS-related MEGS deficiencies could be confirmed in cultured fibroblasts, suggesting that these are most likely caused by 1 or more primary defects.



Because transmitochondrial cybrid cells are often used to verify the pathogenicity of mitochondrial DNA (mtDNA) mutations, we included both control and complex I-deficient (ND1 mutation) cybrid cells. The clear difference in respiratory activity between control and complex I-deficient cybrids (Fig. 6; ROCR after ADP stimulation, 10.1 and 1.1 RFU/U CS, respectively) shows that the assay can also be used to rapidly screen cybrid clones for mitochondrial dysfunction due to mtDNA mutations.



Despite the availability of many tools to diagnose mitochondrial disease (16, 17), current methods are still insufficient for the evaluation of a large subgroup of patients suspected to have a mitochondriopathy. Here, we show that quenched-fluorescence oxygen sensing provides a fast and efficient additional diagnostic tool that can be used to examine the mitochondrial energy-generating system in cultured patient-derived cells.

The assay showed a clear difference between control and patient cell lines: control cells showed a substantial increase in oxygen consumption rate upon ADP stimulation, in contrast to the patient cell lines. This was illustrated in particular by the cybrid cells. Complex I- and complex IV-deficient fibroblasts showed a significantly decreased oxygen consumption rate. Complex V-deficient fibroblasts could be recognized because of a substantial increase after uncoupling. Addition of an uncoupler leads to a decrease in the mitochondrial membrane potential, which subsequently gives rise to a maximal stimulation of the respiratory rate (18). Uncoupling makes the oxygen consumption rate independent of complex V, and this explains the initial increase in complex V- deficient cells. It has also been shown that CCCP stimulates apoptosis via loss in mitochondrial transmembrane potential and reactive oxygen species (ROS) production (19). Cell death, combined with back-diffusion of oxygen, could explain the subsequent decrease of the respiratory activity slope that was sometimes observed. Inhibitors of the entire pathway from pyruvate import to ATP export, and several steps in between (the MEGS), reduced oxygen consumption in fibroblasts and isolated muscle mitochondria, confirming that the integrity of the entire OXPHOS machinery can be tested successfully in both permeabilized cells and isolated mitochondria. A peculiar finding was a slight loss of inhibition of oligomycin in the fibroblasts after 35 min; this was not seen in isolated muscle mitochondria. Further research is needed to understand this finding.



Some cautionary notes must be taken into account when measuring the oxygen consumption rate in fibroblasts. We saw that the rate can decrease when reaching a higher cell culture passage number (Fig. 2 and online Supplemental Table 1). To get reliable results, it is very important to measure cells at an early passage stage and to measure samples in at least 2 independent experiments. Usually, the passage number increases by 1-2 between measurements. In case of poor reproducibility of multiple measurements with increasing passage number, a possible negative effect of the passage stage should be considered. Results should then be distrusted and cells from an earlier passage should be tested, if available.

The assay can also be used to support the search for new defects in mitochondrial energy metabolism. In a group of patients with decreased ATP production in fresh muscle, a subgroup could be identified with clearly reduced oxygen consumption rate, despite normal OXPHOS enzymes and PDHc activities. Such data indicate that another still-unidentified defect in the mitochondrial energy metabolism may be responsible for the reduced oxygen consumption rates. This will encourage further studies to uncover new genetic defects causing mitochondrial disease. Another subgroup of patients showed normal or near-normal oxygen consumption rates in fibroblasts that may be a reflection of a tissue-specific mitochondriopathy, which is not uncommon, e.g., mtDNA mutations with uneven tissue distribution or POLG [polymerase (DNA directed), [gamma], formerly POLG1] gene mutations. A normal oxygen consumption rate in fibroblasts can also be expected when the MEGS dysfunction in muscle tissue is secondary (poor feeding, disuse) (6). Differentiating between primary and secondary mitochondrial abnormalities is of great importance with respect to counseling. Thus, this assay provides a valuable tool in the diagnostics of mitochondrial disorders. Advantages of the assay compared to substrate oxidation measurements include its simplicity, the requirement of less cell material, and the elimination of the need of an isotope laboratory. When comparing oxygen-sensitive fluorescence-based methods to classical polarography, it has been shown that, due to the low rate of back-diffusion, the rates of change of dissolved oxygen measured using the microplate approach are lower than the consumption rates measured using a sealed polarographic system (7).

In conclusion, quenched-fluorescence oxygen sensing provides a simple, high-throughput, high-content screening method for mitochondrial function in different cell types. It requires a relatively low sample volume and allows for simultaneous testing of multiple assay conditions. In addition, this approach is also suitable for the analysis of cell lines with relatively low mitochondrial activity (e.g., fibroblasts). Patients with a primary mitochondriopathy that does not affect the complexes of the respiratory chain can be identified. In combination with functional studies and other diagnostic assays (for instance, homozygosity mapping), the availability of this approach will facilitate the search for hitherto-unknown genetic defects leading to mitochondrial disease.

Author Contributions: All authors confirmed they have contributed to the intellectual content of this paper and have met the following 3 requirements: (a) significant contributions to the conception and design, acquisition of data, or analysis and interpretation of data; (b) drafting or revising the article for intellectual content; and (c) final approval of the published article.

Authors' Disclosures of Potential Conflicts of Interest: Upon manuscript submission, all authors completed the Disclosures of Potential Conflict of Interest form. Potential conflicts of interest:

Employment or Leadership: None declared.

Consultant or Advisory Role: None declared.

Stock Ownership: None declared.

Honoraria: None declared.

Research Funding: A.I. Jonckheere, grant from Prinses Beatrix Fonds; J.A.M. Smeitink, grant from Prinses Beatrix Fonds; M. Huigsloot, grant IGE05003 from Senternovem.

Expert Testimony: None declared.

Role of Sponsor: The funding organizations played no role in the design of study, choice of enrolled patients, review and interpretation of data, or preparation or approval of manuscript.


(1.) Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, et al. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin Chem 2006;52: 860-71.

(2.) Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem 2003;40:3-8.

(3.) Wanders RJ, Ruiter JP, Wijburg FA. Studies on mitochondrial oxidative phosphorylation in permeabilized human skin fibroblasts: application to mitochondrial encephalomyopathies. Biochim Biophys Acta 1993;1181:219-22.

(4.) Rizza T, Vazquez-Memije ME, Meschini MC, Bianchi M, Tozzi G, Nesti C, et al. Assaying ATP synthesis in cultured cells: a valuable tool for the diagnosis of patients with mitochondrial disorders. Biochem Biophys Res Commun 2009;383: 58-62.

(5.) van den Heuvel LP, Smeitink JA, Rodenburg RJ. Biochemical examination of fibroblasts in the diagnosis and research of oxidative phosphorylation (OXPHOS) defects. Mitochondrion 2004;4: 395-401.

(6.) Morava E, Rodenburg R, van Essen HZ, De Vries M, Smeitink J. Dietary intervention and oxidative phosphorylation capacity. J Inherit Metab Dis 2006;29:589.

(7.) Hynes J, Marroquin LD, Ogurtsov VI, Christiansen KN, Stevens GJ, Papkovsky DB, Will Y. Investigation of drug-induced mitochondrial toxicity using fluorescence-based oxygen-sensitive probes. Toxicol Sci 2006;92:186-200.

(8.) Rustin P, Chretien D, Bourgeron T, Gerard B, Rotig A, Saudubray JM, Munnich A. Biochemical and molecular investigations in respiratory chain deficiencies. Clin Chim Acta 1994;228:35-51.

(9.) Ferrick DA, Neilson A, Beeson C. Advances in measuring cellular bioenergetics using extracellular flux. Drug Discov Today 2008;13:268 -74.

(10.) Will Y, Hynes J, Ogurtsov VI, Papkovsky DB. Analysis of mitochondrial function using phosphorescent oxygen-sensitive probes. Nat Protoc 2006;1:2563-72.

(11.) Hynes J, Hill R, Papkovsky DB. The use of a fluorescence-based oxygen uptake assay in the analysis of cytotoxicity. Toxicol In Vitro 2006;20: 785-92.

(12.) Bookelman H, Trijbels JM, Sengers RC, Janssen AJ, Veerkamp JH, Stadhouders AM. Pyruvate oxidation in rat and human skeletal muscle mitochondria. Biochem Med 1978;20:395-403.

(13.) Srere PA. Citrate synthase. Methods Enzymol 1969;13:3-11.

(14.) van der Westhuizen FH, van den Heuvel LP, Smeets R, Veltman JA, Pfundt R, van Kessel AG, et al. Human mitochondrial complex I deficiency: investigating transcriptional responses by microarray. Neuropediatrics 2003;34:14-22.

(15.) Rossignol R, Gilkerson R, Aggeler R, Yamagata K, Remington SJ, Capaldi RA. Energy substrate modulates mitochondrial structure and oxidative capacity in cancer cells. Cancer Res 2004; 64:985-93.

(16.) Wolf NI, Smeitink JA. Mitochondrial disorders: a proposal for consensus diagnostic criteria in infants and children. Neurology 2002;59:1402-5.

(17.) Taylor RW, Schaefer AM, Barron MJ, McFarland R, Turnbull DM. The diagnosis of mitochondrial muscle disease. Neuromuscul Disord 2004;14: 237-45.

(18.) Mracek T, Pecina P, Vojtiskova A, Kalous M, Sebesta O, Houstek J. Two components in pathogenic mechanism of mitochondrial ATPase deficiency: energy deprivation and ROS production. Exp Gerontol 2006;41:683-7.

(19.) Chaudhari AA, Seol JW, Kim SJ, Lee YJ, Kang HS, Kim IS, et al. Reactive oxygen species regulate Bax translocation and mitochondrial transmembrane potential, a possible mechanism for enhanced TRAIL-induced apoptosis by CCCP. Oncol Rep 2007;18:71-6.

An I. Jonckheere, [1] Merei Huigsloot, [1] Antoon J.M. Janssen, [1] Antonia J.H. Kappen, [1] Jan A.M. Smeitink, [1] and Richard J.T. Rodenburg [1] *

[1] Laboratory of Pediatrics and Neurology, Department of Pediatrics, Nijmegen Center for Mitochondrial Disorders, Radboud University Nijmegen Medical Center, Nijmegen, the Netherlands.

* Address correspondence to this author at: Radboud University Nijmegen Medical Center, Nijmegen Center for Mitochondrial Disorders, Laboratory of Pediatrics and Neurology, PO Box 9101, 6500 GA Nijmegen, the Netherlands. Fax +31-24-361-8900; e-mail

Received June 8, 2009; accepted December 14, 2009.

Previously published online at DOI: 10.1373/clinchem.2009.131441

[2] Nonstandard abbreviations: MEGS, mitochondrial energy-generating system; PDHc, pyruvate dehydrogenase complex; TCA, tricarboxylic acid; OXPHOS, oxidative phosphorylation; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; SETH, sucrose, EDTA, Tris-HCl, and heparin; ROCR, relative oxygen consumption rate; CS, citrate synthase; RFU, relative fluorescence units; ANT, adenine nucleotide transporter; CCCP, carbonyl cyanide 3-chlorophenyl hydrazone; mtDNA, mitochondrial DNA; ROS, reactive oxygen species.
COPYRIGHT 2010 American Association for Clinical Chemistry, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2010 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Title Annotation:Endocrinology and Metabolism
Author:Jonckheere, An I.; Huigsloot, Merei; Janssen, Antoon J.M.; Kappen, Antonia J.H.; Smeitink, Jan A.M.;
Publication:Clinical Chemistry
Date:Mar 1, 2010
Previous Article:Implementation of a closed-loop reporting system for critical values and clinical communication in compliance with goals of the joint commission.
Next Article:Assessment of vitamin [B.sub.12] absorption based on the accumulation of orally administered cyanocobalamin on transcobalamin.

Terms of use | Privacy policy | Copyright © 2018 Farlex, Inc. | Feedback | For webmasters