The use of liquid chromatography-tandem mass spectrometry (LC-MS/MS) to measure serum folate has been recommended for population monitoring as it allows a more accurate and reproducible measurement than the commonly used protein binding assays. The ability to differentiate between endogenous folate vitamers and folic acid makes LC-MS/MS an extremely valuable tool, not least due to the uncertainty surrounding the potential adverse effects of high concentrations of circulating folic acid.
by Sarah Meadows
Role of folates
Folate is the general term for a water-soluble B vitamin naturally found in foods such as leafy vegetables, legumes, egg yolks, liver and some citrus fruits. Folic acid itself does not occur naturally but it can be found in individuals who take vitamin supplements or eat fortified foods [1, 2]. There are many critical cellular pathways that depend on folate, including DNA, RNA and protein methylation, as well as DNA synthesis and maintenance [2, 3], and because of this there are many health consequences of folate deficiencies among all age groups. These include megaloblastic anemia, depression, cognitive impairment, low birth weight, risk of placental abruption, neural tube defects and other birth defects including orofacial clefts and heart defects [4, 5]. The most widely publicized public health issue surrounding folate is that of low folate during pregnancy causing birth defects associated with the nervous system. Folate is an essential micronutrient during fetal development because of its roles in transmethylation reactions and in synthesis of DNA in growing cells [3]. A significant portion of the 300 000 neural tube defects (NTDs) that occur yearly worldwide are preventable by the periconceptual consumption of folic acid and continue to be a great public health burden globally [2]. The demonstration that periconceptional supplementation with folic acid dramatically reduces the incidence of NTDs has generated considerable clinical and public health interest and has led to fortification with folic acid of the food supply in the United States and some other countries [6]. The recent World Health Organization (WHO) guidelines for ‘Optimal serum and red blood cell folate in women of reproductive age for prevention of neural tube defects’ states that in 2012 an estimated 270 358 deaths globally were attributable to congenital abnormalities during the first 28 days of life. NTDs were one of the most serious and most common abnormalities and increasing awareness of the significance of insufficient folate intake has emphasized the need for identification of accurate biomarkers for large scale assessment of folate status [7].
As well as neural tube closure, B vitamins, including folate, are required for essential brain metabolic pathways and are fundamental in all aspects of brain development and maintenance of brain health throughout the lifecycle. Observational and animal evidence appears to be supportive for a role of maternal folate status in later cognitive performance of the child and there are also studies linking low maternal folate status with a higher incidence of behavioural and emotional problems, inattention and hyperactivity in their offspring [8]. Recent studies have also shown links between low plasma folate and poor cognitive performances in children and adolescents, and similarly a positive association between higher dietary folate intake and academic achievement [8].
Cognitive dysfunction in the elderly (ranging from cognitive impairment to dementia) is also a matter of concern. Brain changes progress long before the diagnosis of dementia is made, and given the increase in life expectancy, the numbers of individuals suffering is set to double by 2025. Therefore it is important to find early biomarkers that would enable timely interventions to delay the onset or slow the progression of the disease. There is emerging evidence suggesting that suboptimal status of folate and metabolically related B vitamins may be linked with cognitive dysfunction and dementia; if this can be slowed or prevented by improving the B vitamin status in healthy older people it could offer a cost effective preventative public health strategy in ageing populations [8].
Evidence showing that supplementation with folic acid protects against NTDs has led to government recommendations, which are in place worldwide, advising all women planning a pregnancy to consume 400 µg/day folic acid from preconception until the end of the first trimester of pregnancy [2, 8–11]. Even with this knowledge, public health campaigns remain largely unsuccessful and limited [2, 9]. Mandatory fortification programmes have been implemented in many countries to improve folate status and reduce high costs associated with prevention programmes such as education campaigns and other interventions that require behavioural change [2]. The Scientific Advisory Committee on Nutrition (SACN) has called for mandatory fortification in the United Kingdom to replace voluntary fortification in a bid to increase the UK population’s folate status [12].
Despite the unequivocal success of folic acid in reducing NTD rates, several studies have questioned whether unmetabolized folic acid in blood may have adverse effects [3, 11]. Concerns have been raised that due to fortification the subsequent increase in folic acid intakes across the population may have harmful effects on health, such as the masking of pernicious anemia, colorectal cancer promotion in people with pre-existing lesions or adverse cognitive effects in the elderly with low vitamin B12 status [4, 9, 11, 13]. Measurement of unmetabolized folic acid has been suggested as a way of monitoring whether folic acid intake is in excess of body requirements [2, 5] and at a time when there are still questions regarding the effects of high levels of folic acid in the blood, the ability to differentiate between this and endogenous folates is valuable.
Serum folate is considered an indicator of recent folate intake whereas red blood cell folate concentrations indicate long term status [7, 10].
Measurement of folates
There are several methods currently in use to measure serum folates all with their own advantages and limitations (Table 1). Folate had traditionally been measured using a microbiological assay but, since the 1970s, commercial protein binding assays on automated clinical analysers have been widely used due to both the ease of use of these platforms and the increased throughput they offer. Microbiological assays have not been made obsolete by protein binding assays, as originally expected, due to these assays being well suited to low resource settings [6]. The microbiological assays are considered more accurate as they recover folate vitamers equally and are, therefore, considered the gold standard measurement [14], whereas the protein binding assays generally underestimate folate concentrations due to the different affinities of the folate vitamers for the binding protein used [1, 7].
In contrast to both the microbiological assay and protein binding assays, chromatography techniques are able to differentiate between individual folate species [7] and are now often coupled to mass spectrometers as this method has high sensitivity, specificity and selectivity compared to other detection methods such as fluorimetric or electrochemical detection [6, 7, 15].
The importance of measuring the different folate species is likely to become greater in the future as more information on genetic polymorphisms that affect nutritional status and folate distributions become available [1] and in order to determine the safety of free folic acid in the blood.
The differences seen in results produced by different assays have led the WHO to recommend the standardization of blood vitamin analysis [5, 7]. Initial steps to standardize folate methods began with the development of higher order reference methods that use isotope dilution/ liquid chromatography/tandem mass spectrometry and with recent advances in sample clean up procedures routine methods using LC/MS or LC-MS/MS are becoming more common [5].
Red cell folate is normally calculated using whole blood folate concentrations, serum folate concentrations and hematocrit. However, low concentrations of serum folate within an individual over the course of a month are also indicative of low folate or folate depletion [7, 16]. It has proven to be more technically challenging to measure whole blood folate by LC-MS/MS than it is to measure serum folate, in part because red blood cells first need to be hemolysed to release the polyglutamate folates, which then need to be deconjugated to monoglutamates without any folate loss before being analysed [6, 7]. This has led to whole blood assays only being carried out, commonly by microbiological assays, in specialist laboratories.
LC-MS/MS measurement of folates
The recommendation from an expert and stakeholder workshop for the use of an LC-MS/MS method to measure serum folate for UK population monitoring in 2009 [17] led to the establishment of a assay at the MRC Human Nutrition Unit, Cambridge for the measurement of serum folate in the UK National Diet and Nutrition Survey Rolling Programme (NDNS RP). This was developed from the published method used by the Centers for Disease Control and Prevention, Atlanta, GA for the US National Health and Nutrition Examination Survey (NHANES) [18]. The method described here is for a routine LC-MS/MS method allowing the determination and quantitation of six folate vitamers in serum: tetrahydrofolate (THF); 5-methyltetrahydrofolate (MTHF); 5-formyltetrahydrofolate (FTHF); free folic acid (polyglutamic acid/PGA); 5,10-methenyltetrahydrofolate (5,10 methenylTHF) and an oxidation product of MTHF, MeFox [19].
Samples undergo solid phase extraction, using phenyl columns, to isolate the folate forms in serum samples. Stable isotope-labelled internal standards are added during the extraction step and undergo processing identical to the analytes, thereby normalizing for sample preparation and instrument variability. Analytes are measured using isocratic reversed-phase UPLC prior to electrospray ionization tandem mass spectrometry with a run time of 3.5 minutes. The retention times for all the analytes are very similar and the internal standards are identical to their corresponding analytes, but due to their differing masses, there is clear distinction between them in the assay (Fig. 1). FTHF and MeFox have the same molecular weights and cannot be chromatographically separated, so transitions unique to each form have to be used. The ratio of analyte to internal standard signal is compared to that of a calibration curve to determine analyte concentration.
Recovery is 95.1% for MTHF and >78% for all the other analytes and within batch precision is <7% for all analytes. The calibration graphs are linear, R2 >0.99, for all analytes, from 1 to 100 nmol/L for MTHF and 0.5 to 20 nmol/L for all the other folate forms. Linearity extends above these ranges but these encompass the normal concentrations seen currently in the UK population. Total serum folate concentrations in the UK population lie mainly between 2 and 80 nmol/l. The main folate form in the serum is MTHF, free folic acid and THF are found at concentrations usually <2 nmol/L and MeFox is also found in the majority of samples at concentrations <10 nmol/L. FTHF and CH+THF are rarely found in the serum samples of the UK population.
Tandem mass spectrometry may require operation by experienced personnel but it can provide high throughput measurements in a routine environment. The big disadvantage of this method of analysis to most laboratories is the large financial outlay required to purchase the equipment, but the recent advances in mass spectrometry has led to cheaper and smaller instruments being available and this has led to many more being implemented into routine clinical laboratories.
The use of immunoassays for clinical purposes may still be the preferred option for many routine clinical labs but LC-MS/MS is a more accurate, precise and reliable tool for population studies and research purposes than other available methods.
This work was funded by the Medical Research Council MRC_MC_U105960384.
References
1. Shane B. Folate status assessment history: implications for measurement of biomarkers in NHANES. Am J Clin Nutr. 2011; 94(1): 337S–342S.
2. Crider KS, Bailey LB, Berry RJ. Folic acid food fortification-its history, effect, concerns, and future directions. Nutrients, 2011; 3(3): 370–384.
3. Obeid R, Kasoha M, Kirsch SH, Munz W, Herrmann W. Concentrations of unmetabolized folic acid and primary folate forms in pregnant women at delivery and in umbilical cord blood. Am J Clin Nutr. 2010; 92(6): 1416–1422.
4. Smith AD. Folic acid fortification: the good, the bad, and the puzzle of vitamin B-12. Am J Clin Nutr. 2007; 85(1): 3–5.
5. de Benoist B. Conclusions of a WHO Technical Consultation on folate and vitamin B12 deficiencies. Food Nutr Bull. 2008; 29(2 Suppl): S238–244.
6. Bailey LB. Folate in health and disease, 2nd ed. Taylor & Francis 2010.
7. WHO. In: Guideline: Optimal serum and red blood cell folate concentrations in women of reproductive age for prevention of neural tube defects. WHO 2015. (http://apps.who.int/iris/bitstream/10665/161988/1/9789241549042_eng.pdf)
8. McGarel C, Pentieva K, Strain JJ, McNulty H. Emerging roles for folate and related B-vitamins in brain health across the lifecycle. Proc Nutr Soc. 2015; 74(1): 46–55.
9. Hopkins SM, Gibney MJ, Nugent AP, McNulty H, Molloy AM, Scott JM, Flynn A, Strain JJ, Ward M, Walton J, McNulty BA. Impact of voluntary fortification and supplement use on dietary intakes and biomarker status of folate and vitamin B-12 in Irish adults. Am J Clin Nutr. 2015; 101(6): 1163–1172.
10. Clarke R, Bennett D. Folate and prevention of neural tube defects. BMJ 2014; 349: g4810.
11. European Food Safety Authority (EFSA). Folic acid: an update on scientific developments. EFSA 2010. (https://www.efsa.europa.eu/en/supporting/pub/2e)
12. Scientific Advisory Committee on Nutrition (sacn). Folate and disease prevention. The Stationery Office 2006.
13. Bailey RL, Mills JL, Yetley EA, Gahche JJ, Pfeiffer CM, Dwyer JT, Dodd KW, Sempos CT, Betz JM, Picciano MF. Serum unmetabolized folic acid in a nationally representative sample of adults >/=60 years in the United States, 2001–2002. Food Nutr Res, 2012; 56: DOI: 10.3402/fnr.v56i0.5616.
14. Pfeiffer CM, Hughes JP, Lacher DA, Bailey RL, Berry RJ, Zhang M, Yetley EA, Rader JI, Sempos CT, Johnson CL. Estimation of trends in serum and RBC folate in the U.S. population from pre- to postfortification using assay-adjusted data from the NHANES 1988-2010. J Nutr. 2012; 142(5): 886–893.
15. Wang X, Zhang T, Zhao X, Guan Z, Wang Z, Zhu Z, Xie Q, Wang J, Niu B. Quantification of folate metabolites in serum using ultraperformance liquid chromatography tandem mass spectrometry. J Chromatogr B Analyt Technol Biomed Life Sci. 2014; 962: 9–13.
16. McDowell MA, Lacher DA, Pfeiffer CM, Mulinare J, Picciano MF, Rader JI, Yetley EA, Kennedy-Stephenson J, Johnson CL. Blood folate levels: the latest NHANES results. NCHS Data Brief 2008; 6: 1–8.
17. Duthie SJ, Bird S, Mayer C, Macdonald H. FSA UK: Programme N08: Dietary surveys and nutrients in food: Informed systematic review and critical comparison of analytical methods for the quantification of blood folate status in the population. FSA Website April 2009.
18. Fazili Z, Whitehead RD Jr, Paladugula N, Pfeiffer CM. A high-throughput LC-MS/MS method suitable for population biomonitoring measures five serum folate vitamers and one oxidation product. Anal Bioanal Chem. 2013; 405(13): 4549–60.
19. Meadows S. Multiplex measurement of serum folate vitamers by UPLC-MS/MS. Methods in Molecular Biology 2016 (in press).
The author
Sarah Meadows MSc, CSci
MRC, Elsie Widdowson Laboratory, Cambridge, UK
E-mail: sarah.meadows@mrc-ewl.cam.ac.uk
DxN VERIS Molecular Diagnostics System
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, /in Featured Articles /by 3wmediaUp-to-Date Solution for Newcomers in Automated Testing
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, /in Featured Articles /by 3wmediaChromosomal rearrangement detection in lymphomas: digital FISH analysis
, /in Featured Articles /by 3wmediaDetection of chromosomal rearrangements using fluorescence
in situ hybridization (FISH) in lymphomas is an important diagnostic measure for treating this aggressive tumour type. This article aims to describe the recent advancements in the technology.
by Dr Michael Liew, Leslie Rowe and Prof. Mohamed E. Salama
Background
Lymphomas are cancers that affect cells of the lymphatic or immune system, affect a wide range of age groups and are not gender specific. Differentiation of different lymphomas types is important for prognosis and treatment regimens. Lymphomas can be differentiated according to the chromosomal rearrangements they have undergone. For example, Burkitt’s lymphoma is characterized by a rearrangement in the MYC gene (8q24). Diffuse large B-cell lymphoma (DLBCL) is a very aggressive tumour that is characterized by multiple gene rearrangements involving MYC, IGH, BCL2 or BCL6. Mantle cell lymphomas are characterized by the presence of the CCND1 gene rearrangement. All of these rearrangements are routinely detected by fluorescence in situ hybridization (FISH) [1].
Digital FISH analysis
Microscope slides prepared for FISH analysis are still currently viewed by eye using an epifluorescence microscope. An emerging technology is digital FISH analysis. The microscope slides are prepared in the same way, but the fluorescent signals are captured and analysed digitally. The entire field of microscopy has benefited from whole slide imaging (WSI). This enables the capture, analysis, storage and sharing of whole slide pathology images [2]. However, WSI is still in its infancy as a diagnostic tool because there is a lack of evidence that it can be used as a primary diagnostic tool compared to viewing the slide directly with a traditional microscope. Similarly FISH analysis of a tissue slide is starting to become digitized and laboratories are validating its use diagnostically.
There are three main components to digital FISH analysis. The first component, which is critical for its accuracy, is the need for z-stacking of multiple digital images for FISH applications. This is in contrast to WSI for bright field microscopy, which only needs to capture a single digital image. The second component is the identification, or segmentation, of individually stained nuclei that have been stained with a fluorescent dye, such as DAPI. With a suspension of cells, having software that identifies isolated nuclei is relatively straightforward. However, the digital analysis of formalin-fixed paraffin-embedded (FFPE) tissue sections, which can be more clinically informative, can be more challenging. The final component of digital FISH analysis is the signal count, or signal classification. Digital FISH analysis provides a powerful way of documenting and analysing, what can be complex signal patterns.
Validation of digital FISH assays
Our laboratory has validated digital FISH assays for the detection of MYC rearrangements in FFPE tissue (Fig. 1) [3]. We demonstrated a good correlation between traditional and digital FISH analysis for MYC rearrangements using both locus specific identifier (LSI) break-apart and IGH-MYC fusion probes. Our findings document improved diagnostic accuracy with the implementation of digital FISH analysis. Segmented and classified digital images allow for permanent storage of analysed specimens and easy accessibility for further review and/or educational purposes. The digital platform is also conducive to laboratory workflow as it allows timely segmentation and classification of nuclei, remote access for review of cases, elimination of manual slide transportation, and accurate identification/assessment of tumour from digital tissue matching.
We are not the first to adopt digital FISH analysis; however, we are among the first efforts to validate the system for clinical use, and results are in good agreement with previous studies that looked at automated analysis of FISH results from FFPE B-cell lymphoma tissue [4, 5]. The automated analysis yielded 100% agreement with conventional FISH in both studies. In contrast though, the amount of time it took to analyse a sample was approximately twice as long as the previous study. This difference in time was due to the manual editing required when using the digital system particularly in the initial phases of validation, whereas the other system was able to rely more on the automated analysis. We are developing the current workflow and system to reduce this step.
Implementation of digital FISH
The primary cost of switching over to a digital FISH system from a manual system is the new hardware including computer, scanning system and slide loaders. The digital FISH system increases the cost of a stand-alone epifluorescence microscope significantly. Depending upon whether whole slide images are captured, or just a few fields of view, additional computer servers may be needed for data storage. The promise is that if the digital FISH analysis is completely automated, and the results are 100% accurate, it is faster than manual FISH analysis. From our experience though, additional development is needed to reach the 100% accuracy and much development is needed to minimize the time needs to be spent manually editing the results. However, using the digital FISH analysis versus current direct visualization of the FISH slide with manual recording of a scoresheet, will come with several benefits including automatic digitized records and the possibility of remote connectivity for pathologists; to mention a few.
Future perspectives
There are several future considerations that arise from developing digital FISH imaging. WSI of a FISH slide is a possible application in digital FISH imaging. Similar to imaging an entire hematoxylin and eosin (H&E)-stained slide, it would mean that the entire section could be analysed, minimizing the possibility that a small area of tumour may be missed. However, magnification is an issue. FISH slides need to be analysed under higher magnification (at least 60×) to maintain the resolution between signals. Coupled with the acquisition of multiple z-stacked images, this would make the scanning time longer, and the image files extremely large, making data storage an issue. In addition, FDA concerns over digital pathology will affect future regulation of digital FISH analysis. This will mean that validated assays that are developed in the future will need to demonstrate that there is no loss of accuracy using a computer versus an epifluorescence microscope. Image formats, resolution and compression are important factors in accurate interpretation of digital FISH images. For the most accurate digital FISH data, images that do not lose data during compression (ie. TIF LZW or PNG) should be used. The drawback is that data storage becomes an issue, since files stay large, but is important to maintain image quality and accuracy.
Conclusion
Digital capture and analysis of FISH assays are a positive development for this important laboratory testing modality. The MYC FISH assays which we have converted to our digital imaging platform have provided numerous logistical and diagnostic advantages as indicated previously. In addition, individual signal patterns can be recorded and stored. These data alongside advances in computational power can potentially lead to correlation between signal pattern and unique tumour phenotypes, or overall tumour prognosis. There are still limitations to digital FISH analysis, in particular being able to reliably identify nuclei and hybridized signal. However, since the resolution of digital FISH images will only increase, and the algorithms used for detection of nuclei and signals will continually be refined, digital FISH analysis can only improve. As indicated, we feel that digital FISH analysis provides more efficient and accurate results and better patient care in comparison to traditional FISH methods. Efforts to convert other FFPE-based FISH assays to this digital platform are underway in our laboratory.
Acknowledgements
This work was funded by the Institute for Clinical and Experimental Pathology, ARUP Laboratories.
References
1. Martin-Subero JI, Gesk S, Harder L, Grote W, Siebert R. Interphase cytogenetics of hematological neoplasms under the perspective of the novel WHO classification. Anticancer Res. 2003; 23: 1139–1148.
2. Goacher E, Randell R, Williams B, Treanor D. The diagnostic concordance of whole slide imaging and light microscopy: a systematic review. Arch Pathol Lab Med. 2016; doi: http://dx.doi.org/10.5858/arpa.2016-0025-RA [Epub ahead of print].
3. Liew M, Rowe L, Clement PW, Miles RR, Salama ME. Validation of break-apart and fusion MYC probes using a digital fluorescence in situ hybridization capture and imaging system. J Pathol Inform. 2016; 7: 20.
4. Alpár D1, Hermesz J, Pótó L, László R, Kereskai L, Jáksó P, Pajor G, Pajor L, Kajtár B. Automated FISH analysis using dual-fusion and break-apart probes on paraffin-embedded tissue sections. Cytometry A. 2008; 73: 651–657.
5. Reichard KK, Hall BK, Corn A, Foucar MK, Hozier J. Automated analysis of fluorescence in situ hybridization on fixed, paraffin-embedded whole tissue sections in B-cell lymphoma. Mod Pathol. 2006; 19: 1027–1033.
The authors
Michael Liew1 PhD, Leslie Rowe1 MS, Mohamed E. Salama1, 2 MD
1ARUP Institute for Clinical and
Experimental Pathology, Salt Lake City, UT, USA
2Department of Pathology, University of Utah School of Medicine, Salt Lake City, UT, USA
*Corresponding author
E-mail: liewm@aruplab.com
Is fully integrated LC-MS/MS the future for the routine clinical lab?
, /in Featured Articles /by 3wmediaLiquid chromatography-mass spectrometry (LC-MS/MS) is an analytical chemistry technique that combines the physio-chemical separation capabilities of liquid chromatography (via conventional chromatography within a column) with the analytic power of mass spectrometry. It allows the user to properly ascertain the individual mass/charge ratio of analytes present in a chromatographic peak. The high throughput capabilities of this technique will bring value to the clinical lab, where time taken to analyse samples is paramount. Bringing LC-MS/MS testing into the clinical setting has been a slow process, however, the medical device industry is on the verge of a fundamental breakthrough that could help drive the adoption of this technique.
LC-MS/MS is used primarily for the identification and quantification of particular molecules within a substance, and its application in diagnostics is a promising venture due to its potential ability to increase throughputs and streamline the processes needed. As such, patient data can be analysed quickly and accurately in order to provide improved patient care. Broadly speaking, the methodology can be divided into three parts. Initially, sample preparation is undertaken; be it whole blood, plasma, saliva or urine – the sample must be prepared to ensure large proteins and salts that may dirty the instrumentation are removed. Conventionally, this phase has been undertaken manually, which can be time-consuming and prone to human error. As such there is a need for the automation of this step to improve efficiency and reliability before LC-MS/MS is adopted by the clinical laboratory. Once sample preparation is complete, the liquid chromatography and mass spectrometry steps can take place, in which the sample is separated and analysed respectively.
LC-MS/MS and the clinical laboratory
Although adoption of LC-MS/MS in the clinical laboratory has been slow but steady, this technique has demonstrated vast improvements in analytical specificity when compared to conventional immunoassays. Mass spectrometry’s strength lies in its ability to be extremely specific to the target analyte, due to the absence of cross reactivity; the likes of which can be common in antibody-based immunoassay (IA) methods. However, the uptake of this technique by clinical labs has not been as rapid as expected, with many choosing to continue using immunoassay-based methods instead.
There are a number of factors causing clinical labs to be cautious about the mainstream use of LC-MS/MS systems. There are numerous LC and MS systems available to choose from, something which in itself can seem overwhelming to a clinical scientist who is not an LC-MS/MS expert. In addition, there is a range of options for calibrators and controls available, along with the internal expertise required to develop and validate methods, and set-up and run the instruments. The final factor to impact the decision is often cost, since investment in such systems is commonly high, especially when taking into account the automated components required to help reduce labour needs for sample preparation. As such, finance options are often limited. When combined, these factors can make immunoassay analysers seem like the simpler option.
The emergence of connected components
Although used in many clinical labs, immunoassay techniques are not always accurate. For example small molecule biomarkers, such as steroid hormones, prove challenging due to the lack of specificity in the binding sites on small molecules, a fact that many clinical scientists are all too aware of. Recent improvements to LC-MS/MS systems have focused on advancing both ease of use and efficacy, essentially to make them a viable alternative to IA methods. Laboratory managers can find ample published documentation that shows just how beneficial LC-MS/MS systems are when used in place of IAs. For instance, a study by Nigel W. Brown and colleagues published in Clinical Chemistry in 2005 demonstrated that LC-MS/MS was far more precise than a microparticle enzyme immunoassay (MEIA), which was ‘significantly affected by patient cohort’ (Brown, N et al. Clinical Chemistry 2005; 51(3): 586-593).
Clinical laboratories are faced with increasing complexities in their daily workflows, and there are pressures to provide detailed analyses of patient samples using streamlined and well-coordinated practices. The need to provide efficient turnaround on samples is also on the increase. There is, therefore, a trend where system manufacturers are looking to provide laboratories with the ability to advance efficiency through the implementation of compatible technologies, such as the combination of stand-alone elements (automated sample handlers, LC-MS/MS reagent kits, and software), which are supplied together to better manage workflows. These connected component-based systems, by which the different components of the LC-MS/MS system (sample preparation, liquid chromatography, and mass spectrometry) are placed in tandem with each other, is a big step in the right direction to increase productivity and efficiency, while simplifying the number of decisions that the lab needs to make. However, there are still improvements that can be made. The issue lies in the fact that connected components are not the same as a fully-integrated, automated system with dedicated assays and diagnostic kits that are regulatory compliant. The development of properly synergized components can truly simplify the decisions faced by clinical scientists and enable LC-MS/MS to become an integral part of the clinical laboratory.
The needs of the lab
Clinical labs require a high level of automation with a number of its systems, owing to the high turnover rate demanded to meet the needs of patient care. In addition, easy to use technologies that include walk away operations are essential, and considered commonplace to clinical scientists, owing to the multitude of responsibilities placed on laboratory personnel. These busy labs require built-for-purpose, fully integrated analysers that are able to greatly reduce installation, validation, and training times, having the system ready to operate in a matter of weeks, rather than months. Streamlining the procedure without compromising the quality of the analysis via implementation of better integrated systems can be considered an essential next step in the medical devices industry. Furthermore, results obtained from these systems need not be in isolation: standardization between laboratories using the same system will be achievable owing to the inclusion of dedicated test kits that are fully validated and ready for use with the analyser. The ideal next-generation system for the clinical laboratory will encompass every step, including automated sample preparation, handling and LC-MS in one unit. Moreover, it will be labelled as a medical device, have dedicated assay kits, and be produced, serviced, and supported by a single manufacturer. Finally, such a device would ideally be able to connect bi-directionally with the laboratory information system (LIS) and furthermore to the laboratory automation system (LAS).
In the end, technologies that are able to advance the state of play for laboratory sample analysis are required in order to ensure laboratory personnel can be confident in the analyses they are making. Beyond connected components, the introduction of integrated LC-MS/MS systems into the laboratory could lead to a paradigm shift with regards to specificity in small molecule analysis that is expected by clinical scientists. Systems that can lead to better quality of care for patients and improved analysis for physicians will essentially help healthcare systems operate more efficiently.
The author
Sarah Robinson, Ph.D,
Market Development Specialist,
Thermo Fisher Scientific
& Expert Consultant to the EFLM
Working Group on Test Evaluation
Measurement of serum folate vitamers by LC-MS/MS
, /in Featured Articles /by 3wmediaThe use of liquid chromatography-tandem mass spectrometry (LC-MS/MS) to measure serum folate has been recommended for population monitoring as it allows a more accurate and reproducible measurement than the commonly used protein binding assays. The ability to differentiate between endogenous folate vitamers and folic acid makes LC-MS/MS an extremely valuable tool, not least due to the uncertainty surrounding the potential adverse effects of high concentrations of circulating folic acid.
by Sarah Meadows
Role of folates
Folate is the general term for a water-soluble B vitamin naturally found in foods such as leafy vegetables, legumes, egg yolks, liver and some citrus fruits. Folic acid itself does not occur naturally but it can be found in individuals who take vitamin supplements or eat fortified foods [1, 2]. There are many critical cellular pathways that depend on folate, including DNA, RNA and protein methylation, as well as DNA synthesis and maintenance [2, 3], and because of this there are many health consequences of folate deficiencies among all age groups. These include megaloblastic anemia, depression, cognitive impairment, low birth weight, risk of placental abruption, neural tube defects and other birth defects including orofacial clefts and heart defects [4, 5]. The most widely publicized public health issue surrounding folate is that of low folate during pregnancy causing birth defects associated with the nervous system. Folate is an essential micronutrient during fetal development because of its roles in transmethylation reactions and in synthesis of DNA in growing cells [3]. A significant portion of the 300 000 neural tube defects (NTDs) that occur yearly worldwide are preventable by the periconceptual consumption of folic acid and continue to be a great public health burden globally [2]. The demonstration that periconceptional supplementation with folic acid dramatically reduces the incidence of NTDs has generated considerable clinical and public health interest and has led to fortification with folic acid of the food supply in the United States and some other countries [6]. The recent World Health Organization (WHO) guidelines for ‘Optimal serum and red blood cell folate in women of reproductive age for prevention of neural tube defects’ states that in 2012 an estimated 270 358 deaths globally were attributable to congenital abnormalities during the first 28 days of life. NTDs were one of the most serious and most common abnormalities and increasing awareness of the significance of insufficient folate intake has emphasized the need for identification of accurate biomarkers for large scale assessment of folate status [7].
As well as neural tube closure, B vitamins, including folate, are required for essential brain metabolic pathways and are fundamental in all aspects of brain development and maintenance of brain health throughout the lifecycle. Observational and animal evidence appears to be supportive for a role of maternal folate status in later cognitive performance of the child and there are also studies linking low maternal folate status with a higher incidence of behavioural and emotional problems, inattention and hyperactivity in their offspring [8]. Recent studies have also shown links between low plasma folate and poor cognitive performances in children and adolescents, and similarly a positive association between higher dietary folate intake and academic achievement [8].
Cognitive dysfunction in the elderly (ranging from cognitive impairment to dementia) is also a matter of concern. Brain changes progress long before the diagnosis of dementia is made, and given the increase in life expectancy, the numbers of individuals suffering is set to double by 2025. Therefore it is important to find early biomarkers that would enable timely interventions to delay the onset or slow the progression of the disease. There is emerging evidence suggesting that suboptimal status of folate and metabolically related B vitamins may be linked with cognitive dysfunction and dementia; if this can be slowed or prevented by improving the B vitamin status in healthy older people it could offer a cost effective preventative public health strategy in ageing populations [8].
Evidence showing that supplementation with folic acid protects against NTDs has led to government recommendations, which are in place worldwide, advising all women planning a pregnancy to consume 400 µg/day folic acid from preconception until the end of the first trimester of pregnancy [2, 8–11]. Even with this knowledge, public health campaigns remain largely unsuccessful and limited [2, 9]. Mandatory fortification programmes have been implemented in many countries to improve folate status and reduce high costs associated with prevention programmes such as education campaigns and other interventions that require behavioural change [2]. The Scientific Advisory Committee on Nutrition (SACN) has called for mandatory fortification in the United Kingdom to replace voluntary fortification in a bid to increase the UK population’s folate status [12].
Despite the unequivocal success of folic acid in reducing NTD rates, several studies have questioned whether unmetabolized folic acid in blood may have adverse effects [3, 11]. Concerns have been raised that due to fortification the subsequent increase in folic acid intakes across the population may have harmful effects on health, such as the masking of pernicious anemia, colorectal cancer promotion in people with pre-existing lesions or adverse cognitive effects in the elderly with low vitamin B12 status [4, 9, 11, 13]. Measurement of unmetabolized folic acid has been suggested as a way of monitoring whether folic acid intake is in excess of body requirements [2, 5] and at a time when there are still questions regarding the effects of high levels of folic acid in the blood, the ability to differentiate between this and endogenous folates is valuable.
Serum folate is considered an indicator of recent folate intake whereas red blood cell folate concentrations indicate long term status [7, 10].
Measurement of folates
There are several methods currently in use to measure serum folates all with their own advantages and limitations (Table 1). Folate had traditionally been measured using a microbiological assay but, since the 1970s, commercial protein binding assays on automated clinical analysers have been widely used due to both the ease of use of these platforms and the increased throughput they offer. Microbiological assays have not been made obsolete by protein binding assays, as originally expected, due to these assays being well suited to low resource settings [6]. The microbiological assays are considered more accurate as they recover folate vitamers equally and are, therefore, considered the gold standard measurement [14], whereas the protein binding assays generally underestimate folate concentrations due to the different affinities of the folate vitamers for the binding protein used [1, 7].
In contrast to both the microbiological assay and protein binding assays, chromatography techniques are able to differentiate between individual folate species [7] and are now often coupled to mass spectrometers as this method has high sensitivity, specificity and selectivity compared to other detection methods such as fluorimetric or electrochemical detection [6, 7, 15].
The importance of measuring the different folate species is likely to become greater in the future as more information on genetic polymorphisms that affect nutritional status and folate distributions become available [1] and in order to determine the safety of free folic acid in the blood.
The differences seen in results produced by different assays have led the WHO to recommend the standardization of blood vitamin analysis [5, 7]. Initial steps to standardize folate methods began with the development of higher order reference methods that use isotope dilution/ liquid chromatography/tandem mass spectrometry and with recent advances in sample clean up procedures routine methods using LC/MS or LC-MS/MS are becoming more common [5].
Red cell folate is normally calculated using whole blood folate concentrations, serum folate concentrations and hematocrit. However, low concentrations of serum folate within an individual over the course of a month are also indicative of low folate or folate depletion [7, 16]. It has proven to be more technically challenging to measure whole blood folate by LC-MS/MS than it is to measure serum folate, in part because red blood cells first need to be hemolysed to release the polyglutamate folates, which then need to be deconjugated to monoglutamates without any folate loss before being analysed [6, 7]. This has led to whole blood assays only being carried out, commonly by microbiological assays, in specialist laboratories.
LC-MS/MS measurement of folates
The recommendation from an expert and stakeholder workshop for the use of an LC-MS/MS method to measure serum folate for UK population monitoring in 2009 [17] led to the establishment of a assay at the MRC Human Nutrition Unit, Cambridge for the measurement of serum folate in the UK National Diet and Nutrition Survey Rolling Programme (NDNS RP). This was developed from the published method used by the Centers for Disease Control and Prevention, Atlanta, GA for the US National Health and Nutrition Examination Survey (NHANES) [18]. The method described here is for a routine LC-MS/MS method allowing the determination and quantitation of six folate vitamers in serum: tetrahydrofolate (THF); 5-methyltetrahydrofolate (MTHF); 5-formyltetrahydrofolate (FTHF); free folic acid (polyglutamic acid/PGA); 5,10-methenyltetrahydrofolate (5,10 methenylTHF) and an oxidation product of MTHF, MeFox [19].
Samples undergo solid phase extraction, using phenyl columns, to isolate the folate forms in serum samples. Stable isotope-labelled internal standards are added during the extraction step and undergo processing identical to the analytes, thereby normalizing for sample preparation and instrument variability. Analytes are measured using isocratic reversed-phase UPLC prior to electrospray ionization tandem mass spectrometry with a run time of 3.5 minutes. The retention times for all the analytes are very similar and the internal standards are identical to their corresponding analytes, but due to their differing masses, there is clear distinction between them in the assay (Fig. 1). FTHF and MeFox have the same molecular weights and cannot be chromatographically separated, so transitions unique to each form have to be used. The ratio of analyte to internal standard signal is compared to that of a calibration curve to determine analyte concentration.
Recovery is 95.1% for MTHF and >78% for all the other analytes and within batch precision is <7% for all analytes. The calibration graphs are linear, R2 >0.99, for all analytes, from 1 to 100 nmol/L for MTHF and 0.5 to 20 nmol/L for all the other folate forms. Linearity extends above these ranges but these encompass the normal concentrations seen currently in the UK population. Total serum folate concentrations in the UK population lie mainly between 2 and 80 nmol/l. The main folate form in the serum is MTHF, free folic acid and THF are found at concentrations usually <2 nmol/L and MeFox is also found in the majority of samples at concentrations <10 nmol/L. FTHF and CH+THF are rarely found in the serum samples of the UK population. Tandem mass spectrometry may require operation by experienced personnel but it can provide high throughput measurements in a routine environment. The big disadvantage of this method of analysis to most laboratories is the large financial outlay required to purchase the equipment, but the recent advances in mass spectrometry has led to cheaper and smaller instruments being available and this has led to many more being implemented into routine clinical laboratories.
The use of immunoassays for clinical purposes may still be the preferred option for many routine clinical labs but LC-MS/MS is a more accurate, precise and reliable tool for population studies and research purposes than other available methods.
This work was funded by the Medical Research Council MRC_MC_U105960384.
References
1. Shane B. Folate status assessment history: implications for measurement of biomarkers in NHANES. Am J Clin Nutr. 2011; 94(1): 337S–342S.
2. Crider KS, Bailey LB, Berry RJ. Folic acid food fortification-its history, effect, concerns, and future directions. Nutrients, 2011; 3(3): 370–384.
3. Obeid R, Kasoha M, Kirsch SH, Munz W, Herrmann W. Concentrations of unmetabolized folic acid and primary folate forms in pregnant women at delivery and in umbilical cord blood. Am J Clin Nutr. 2010; 92(6): 1416–1422.
4. Smith AD. Folic acid fortification: the good, the bad, and the puzzle of vitamin B-12. Am J Clin Nutr. 2007; 85(1): 3–5.
5. de Benoist B. Conclusions of a WHO Technical Consultation on folate and vitamin B12 deficiencies. Food Nutr Bull. 2008; 29(2 Suppl): S238–244.
6. Bailey LB. Folate in health and disease, 2nd ed. Taylor & Francis 2010.
7. WHO. In: Guideline: Optimal serum and red blood cell folate concentrations in women of reproductive age for prevention of neural tube defects. WHO 2015. (http://apps.who.int/iris/bitstream/10665/161988/1/9789241549042_eng.pdf)
8. McGarel C, Pentieva K, Strain JJ, McNulty H. Emerging roles for folate and related B-vitamins in brain health across the lifecycle. Proc Nutr Soc. 2015; 74(1): 46–55.
9. Hopkins SM, Gibney MJ, Nugent AP, McNulty H, Molloy AM, Scott JM, Flynn A, Strain JJ, Ward M, Walton J, McNulty BA. Impact of voluntary fortification and supplement use on dietary intakes and biomarker status of folate and vitamin B-12 in Irish adults. Am J Clin Nutr. 2015; 101(6): 1163–1172.
10. Clarke R, Bennett D. Folate and prevention of neural tube defects. BMJ 2014; 349: g4810.
11. European Food Safety Authority (EFSA). Folic acid: an update on scientific developments. EFSA 2010. (https://www.efsa.europa.eu/en/supporting/pub/2e)
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13. Bailey RL, Mills JL, Yetley EA, Gahche JJ, Pfeiffer CM, Dwyer JT, Dodd KW, Sempos CT, Betz JM, Picciano MF. Serum unmetabolized folic acid in a nationally representative sample of adults >/=60 years in the United States, 2001–2002. Food Nutr Res, 2012; 56: DOI: 10.3402/fnr.v56i0.5616.
14. Pfeiffer CM, Hughes JP, Lacher DA, Bailey RL, Berry RJ, Zhang M, Yetley EA, Rader JI, Sempos CT, Johnson CL. Estimation of trends in serum and RBC folate in the U.S. population from pre- to postfortification using assay-adjusted data from the NHANES 1988-2010. J Nutr. 2012; 142(5): 886–893.
15. Wang X, Zhang T, Zhao X, Guan Z, Wang Z, Zhu Z, Xie Q, Wang J, Niu B. Quantification of folate metabolites in serum using ultraperformance liquid chromatography tandem mass spectrometry. J Chromatogr B Analyt Technol Biomed Life Sci. 2014; 962: 9–13.
16. McDowell MA, Lacher DA, Pfeiffer CM, Mulinare J, Picciano MF, Rader JI, Yetley EA, Kennedy-Stephenson J, Johnson CL. Blood folate levels: the latest NHANES results. NCHS Data Brief 2008; 6: 1–8.
17. Duthie SJ, Bird S, Mayer C, Macdonald H. FSA UK: Programme N08: Dietary surveys and nutrients in food: Informed systematic review and critical comparison of analytical methods for the quantification of blood folate status in the population. FSA Website April 2009.
18. Fazili Z, Whitehead RD Jr, Paladugula N, Pfeiffer CM. A high-throughput LC-MS/MS method suitable for population biomonitoring measures five serum folate vitamers and one oxidation product. Anal Bioanal Chem. 2013; 405(13): 4549–60.
19. Meadows S. Multiplex measurement of serum folate vitamers by UPLC-MS/MS. Methods in Molecular Biology 2016 (in press).
The author
Sarah Meadows MSc, CSci
MRC, Elsie Widdowson Laboratory, Cambridge, UK
E-mail: sarah.meadows@mrc-ewl.cam.ac.uk
Inductively coupled plasma mass spectrometry: the future of sweat analysis?
, /in Featured Articles /by 3wmediaSweat chloride is the gold standard diagnostic test for cystic fibrosis (CF) offering direct measurement of cystic fibrosis transmembrane conductance regulator (CFTR) protein function. Current methods are labour-intensive, complex, time-consuming and require relatively large sample volumes. Inductively coupled plasma mass spectrometry (ICP-MS) is an emerging technology capable of providing rapid and accurate sweat chloride concentrations from small sample volumes.
by Dr Anna Robson, Dr Alexander Lawson and Dr Stephen George
Background
Cystic fibrosis (CF) is a life limiting inherited disorder with an incidence of 1 in 2500–3500 live births in the UK and USA [1]. Mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene translate to dysfunction of the CFTR protein, responsible for controlling transepithelial chloride transport. CF is a multisystem disease, due to the ubiquitous location of the CFTR, however, it is most commonly associated with pulmonary and pancreatic pathologies. Data from the Cystic Fibrosis Foundation Patient Registry (CFFPR) annual report showed that approximately 80% of CF patients are on pancreatic enzyme replacement therapy (PERT) and over 70% of CF mortality is a consequence of respiratory or cardiorespiratory related causes [2].
Implementation of newborn screening pathways for CF has enabled early diagnosis, with advancements in treatment strategies showing significant benefits to patients over the last 25 years. CFFPR data shows that patients now have a median predicted survival of 39.3 years and a higher proportion of adults than children, with CF, was reported in the USA for the first time in 2014 [2]. Patients’ quality of life is also improving with better lung function at the age of 18 years, more patients graduating from higher education and an increased number of viable pregnancies in female patients [2].
There are currently over 2000 known genetic mutations of the CFTR gene associated with CF, many of which can be categorized into five classes [3]. Classes I–III give rise to the most severe phenotypes due to absence or non-function of CFTR, whereas residual CFTR function is observed in patients with class IV–V mutations [2, 3]. Molecular genetic testing has become an increasingly important aid to the diagnostic pathway for CF, especially for patients with rarer mutations and milder forms of the disease. However, molecular analysis is insufficiently sensitive and specific enough to be used as a first line test as rare variants can be missed. Use of next generation sequencing is currently prohibited by cost but may become a more viable option in the future. The UK newborn screening program for CF utilizes a combination of immunoreactive trypsinogen (IRT) and genetic testing for the most common mutations. Although IRT is a good screening tool in neonates, it is not diagnostic and is unsuitable for use in adults. All babies that screen positive for CF are referred for diagnostic confirmation by sweat testing.
Diagnosis of CF
More than six decades since its inception in 1959 [4], quantification of chloride ions in sweat remains the gold standard diagnostic test for diagnosis of CF. In normal functioning sweat glands, isotonic sweat is secreted into the secretory coil. Sodium and chloride are then reabsorbed in the water-impermeable reabsorptive duct via the epithelial sodium channel (ENaC) and the CFTR respectively. Reabsorption of chloride is reduced in CF patients with defective or absent CFTR, thus resulting in sweat electrolyte loss. Sweat concentrations, therefore, provide a direct measurement of electrolyte secretion. Elevated sweat sodium levels are also observed in CF due to the dependence of ENaC activation on CFTR function.
Sweat chloride measurements demonstrate 98% diagnostic specificity for CF [5]. Research has also shown correlations between the type of genetic mutation and chloride concentration [2, 6]. Methods employed in sweat analysis include osmolality, conductivity and electrolyte concentration, however current guidelines developed by the UK Royal College of Paediatrics and Child Health (RCPCH) and the Association of Clinical Biochemistry (ACB) recommend sweat chloride as the analyte of choice for CF diagnosis [7]. Sweat conductivity measurements are accepted for screening purposes in patients over 6 months of age provided that all positive and borderline results are followed up with a chloride measurement [7]. Sweat sodium is no longer recommended for CF diagnosis as it is less reliable than chloride as an indicator of CFTR function [7]. Most laboratories providing sweat analysis measure a combination of analytes, typically chloride with either conductivity or sodium, using the latter two for quality control purposes only [5].
There are currently two accepted methods for collecting sweat following pilocarpine stimulation, the Gibson and Cooke technique (GCT) and the Wescor Macroduct® collection system (WMCS). Using the GCT, sweat is collected onto pre-weighed chloride-free filter paper and eluted in the laboratory for analysis. Sweat is collected into a plastic capillary in the WMCS closed system, thus reducing analytical errors associated with weighing, dilution and evaporation. A minimum sweat secretion rate of 1 g/m2/min is recommended to obtain an accurate chloride concentration. This equates to approximately 75 mg or 15 μL of sweat, depending on the collection method, in 30 minutes [1]. Low sweat rates indicate either suboptimal sweat secretion by the patient or sample evaporation, both of which can affect the accuracy of electrolyte measurements [5]. RCPCH/ACB guidelines therefore recommend duplicate analysis, on each sweat sample collected, to minimize analytical imprecision due to the manual nature of sweat testing [7]. Studies report conflicting data in relation to which collection system yields a higher insufficient rate for sweat analysis [8, 9]. However, the primary limitation of WMCS compared to GCT is reduced sample volume for analysis when a sufficient sweat rate has been achieved. A recent UK audit at Heart of England NHS Foundation Trust (HEFT) showed that more than 30% of samples with a sufficient sweat rate (>15 μL) were insufficient for duplicate ion selective electrode (ISE) analysis when using the WMCS.
Current methods of analysis
Currently accepted methods for sweat chloride quantification include coulometry, colourimetry, and ISE analysis, of which coulometry is the most commonly used (112/161 UK laboratories enrolled in the UKNEQAS external quality assurance (EQA) scheme). Sweat-Chek equipment is recommended for conductivity measurements and accepted methods for sodium include flame photometry, ISE and atomic absorption spectroscopy [6, 7]. All of these methods require manual measurement of each sample, thus occupying the time of a specially trained member of staff for the analysis duration. Dedicated instrumentation is often used for each analyte and sample volume requirements are relatively large compared to the minimum accepted collection using the WMCS. Sweat analysis is therefore complex and time-consuming. The need for dedicated instrumentation, specifically trained staff and laboratory time also carries a cost burden for NHS laboratories with increasing budget restrictions.
Inductively coupled plasma mass spectrometry
At HEFT, we have developed a method for the analysis of sweat sodium and chloride using inductively coupled plasma mass spectrometry (ICP-MS). ICP-MS is an emerging technology in the clinical laboratory, primarily used for determining the elemental composition of samples and recently applied for sweat analysis [10, 11]. Benefits of ICP-MS include rapid and batched measurements, reproducibility, increased sensitivity and specificity compared to traditional methods, simultaneous quantification of multiple elements and ‘walk-away’ analysis. The ICP-MS method for sweat sodium and chloride uses a simple dilute and shoot approach, requiring just 2 μL of sample. The method was found to be both accurate and precise. Comparison studies using EQA samples showed a 3.6 % bias compared to target values [UKNEQAS EQA scheme all laboratory trimmed mean (ALTM)] (Fig. 1a); however, this was not statistically significant at clinical decision limits (30–60 mmol/L) and results were comparable to the coulometry method currently in use. Recommended acceptable precision (<5% CV), as defined by RCPCH/ACB guidelines, was obtained for all clinically relevant concentrations for quality control (QC) samples (Fig. 1b) [1, 7]. ICP-MS has numerous advantages, compared to coulometry, for sweat chloride analysis. Firstly, the low sample volume requirement allows for duplicate measurements on minimum viable samples (15 μL). Analysis run time is approximately 30–60 minutes depending on the number of samples, and ICP-MS is a ‘walk-away’ method. Staff are, therefore, available to carry out other work once the samples have been prepared and placed on the auto-sampler which is advantageous compared to current methods that can occupy a dedicated member of staff for up to half a day (Fig. 2). Improvements in laboratory efficiency are gained by moving away from dedicated chloride and conductivity meters to analysis using equipment already in use for the trace metal service. The main limitation of chloride analysis by ICP-MS at present is contamination due to the instrument tuning solution containing hydrochloric acid. Hence, care must be taken to ensure that the lines of the inlet system have been rinsed for long enough to remove any residual chloride before analysis. Clearly a tuning solution containing a different acid (e.g. nitric acid) would be beneficial and work is underway to source such a reagent. Overall, ICP-MS provides a much more efficient and cost-effective process for sweat analysis as illustrated in Figure 2. Summary
Sweat chloride analysis is the gold standard test for diagnosis of CF; however, current methods are time-consuming, costly and require large sample volumes relative to the minimum acceptable collection. ICP-MS is a relatively new analysis platform in the clinical environment and is therefore not yet included in any guidelines. However, this technique presents an attractive alternative to current methods for rapid and accurate sweat analysis using small sample volumes. ICP-MS has the potential to benefit sweat testing, improving efficiency and reducing costs in the clinical laboratory.
Acknowledgements
The authors would like to acknowledge Dr Chris Chaloner PhD FRCPath and Lesley Tetlow FRCPath, Central Manchester University Hospitals NHS Foundation Trust, for their assistance in proofreading the manuscript.
References
1. Farrell PM, Rosenstein BJ, White TB, Accurso FJ, Castellani C, Cutting GR, Durie PR, Legrys VA, Massie J, et al. Guidelines for diagnosis of cystic fibrosis in newborns through older adults: Cystic Fibrosis Foundation consensus report. J Pediatr. 2008; 153:S4–S14.
2. Cystic Fibrosis Foundation Patient Registry Annual Data Report 2014. (https://www.cff.org/2014-Annual-Data-Report.pdf)
3. Veit G, Avramescu RG, Chiang AN, Houck SA, Cai Z, Peters KW, Hong JS, Pollard HB, Guggino WB, et al. From CFTR biology toward combinatorial pharmacotherapy: expanded classification of cystic fibrosis mutations. Mol Biol Cell. 2016; 27(3): 424–433.
4. Gibson LE, Cooke RE. A test for concentration of electrolytes in sweat in cystic fibrosis of the pancreas utilizing pilocarpine by iontophoresis. Paediatrics 1959; 23(3): 545–549.
5. LeGrys VA. Sweat testing for the diagnosis of cystic fibrosis: Practical considerations. J Pediatr. 1996; 129: 892–897.
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7. Royal College of Paediatrics and Child Health. Guidelines for the performance of the sweat test for the investigation of cystic fibrosis in the UK, 2nd version. March 2014. (http://www.rcpch.ac.uk/system/files/protected/page/Sweat%20Guideline%20v3%20reformat_2.pdf)
8. Laguna TA, Lin N, Wang Q, Holme B, McNamara J, Regelmann WE. Comparison of quantitative sweat chloride methods after positive newborn screen for cystic fibrosis. Pediatr Pulmonol. 2012; 47: 736–742.
9. Hammond KB, Turcios NL, Gibson LE. Clinical evaluation of the macroduct sweat collection system and conductivity analyzer in the diagnosis of cystic fibrosis. J Paediatr. 1994; 124(2): 255–260.
10. Pullan NJ, Thurston V, Barber S. Evaluation of an inductively coupled plasma mass spectrometry method for the analysis of sweat chloride and sodium for use in the diagnosis of cystic fibrosis. Ann Clin Biochem. 2013; 50(Pt 3): 267–270.
11. Collie JT, Massie RJ, Jones OA, Morrison PD, Greaves RF. A candidate reference method using ICP-MS for sweat chloride quantification. Clin Chem Lab Med. 2016; 54(4): 561–567.
The authors
Anna Robson*1 PhD; Alexander Lawson2 PhD, FRCPath; Stephen George2 PhD, FRCPath
1Department of Clinical Biochemistry, Central Manchester University Hospitals NHS Foundation Trust, Newborn Screening Laboratory, Genetic Medicine, St. Mary’s Hospital, Oxford Rd, Manchester, UK
2Department of of Clinical Chemistry and Immunology, Birmingham Heartlands Hospital, Bordesley Green East, Birmingham, UK
*Corresponding author
E-mail: anna.robson@cmft.nhs.uk