Parathyroid hormone (PTH) is an 84 amino acid peptide hormone which has important physiological roles in regulating bone metabolism. It stimulates renal reabsorption of calcium, bone resorption and activation of vitamin D, while also inhibiting renal phosphate reabsorption, bone formation and bone mineralisation. PTH measurement is integral to the diagnosis and management of hypoparathyroidism and hyperparathyroidism. Patients with chronic kidney disease (CKD), which is associated with progressive loss of renal mass and consequent reduction in the activation of vitamin D , may develop chronic kidney disease-mineral bone disorder (CKD-MBD). Current guidelines recommend that PTH should be maintained within two to nine times the upper limit of the reference interval in CKD-MBD patients .
Optimal implementation of such guidelines requires not only that PTH methods give comparable results, which at present they do not [3, 4], but also that pre-analytical conditions are clearly defined. Results from proficiency testing schemes demonstrate marked between-method differences in PTH results which parallel the more than three-times the differences that can be observed in the same sample from patients with CKD-MBD . Such variability represents significant clinical risk, which could result in opposite decisions, e.g., concerning whether or not a patient receives vitamin D treatment, cinacalcet or parathyroidectomy, depending upon the PTH method used.
Almost all clinical laboratories now use second or third generation PTH methods (Table 1). Factors likely to contribute to observed between-method variation include differences in method calibration, analytical specificity and assay design. The International Federation of Clinical Chemistry and Laboratory Medicine (IFCC) has established a Working Group , whose remit is to encourage manufacturers to calibrate their PTH assays in terms of the newly established recombinant International Standard for PTH (IS 95/646) (provided its commutability can be confirmed), to develop a reference measurement procedure for PTH, and to prepare a panel of reference plasma samples. This should ultimately improve between-method comparability of PTH methods.
As an important first step, and under the auspices of the IFCC PTH Working Group, the systematic review of published data reported here has enabled identification of optimal pre-analytical sampling and storage conditions for PTH, with the aim of producing good practice guidance for PTH measurement. The systematic review addresses the following three questions, developed according to the well-established population, intervention, comparator, outcome (PICO) approach :
How stable is PTH in ethylenediaminetetraacetic acid (EDTA) or lithium heparin anticoagulated human whole blood or plasma, as compared to clotted whole blood or separated serum at room temperature, 4°C, −20°C and −80°C?
Does the site of sampling affect measured PTH concentration?
Does the time of sampling affect measured PTH concentration?
Results of the systematic review are reported here together with some suggestions for additional issues requiring further investigation.
Materials and methods
Electronic searches of the Medline, Embase, Cochrane, Bandolier and Centre for Research and Dissemination databases were undertaken to identify relevant articles (up to 06 December 2012). Medical subject headings (MeSH) terms and free text as well as the full search strategy used are presented in Supplemental Data Table 1. Searches were further expanded by the inclusion of relevant papers that were either known to the authors, had been referenced in the original articles or had been retrieved from related articles. Only full papers and letters were included in the search. Titles and abstracts were read and relevant papers obtained and reviewed by two authors (EH, EJL). A manual search of abstracts from national meetings of the Association for Clinical Biochemistry (Focus 1987–2012) and the American Association of Clinical Chemistry (AACC, 1993–2012) was also performed, with abstracts including PTH, parathyroid hormone, parathormone or parathyrin in the title being selected for review. We then undertook forward searching of abstracts to identify whether these had subsequently been published as full papers.
The following inclusion criteria were used: human studies addressing the stability of PTH in blood samples ex vivo, the effect of the sampling site on PTH concentration in blood samples ex vivo, and the effect of seasonal, circadian and/or ultradian (cycles shorter than 24 h) rhythms on PTH concentration in blood samples ex vivo. Exclusion criteria included research in animals, PTH (1–34), parathyroid hormone-related peptide (PTHrP), PTH receptor, PTHrP gene, PTH mRNA, PTH-receptor antagonist and preproPTH. For the first of the PICO questions addressed, publications prior to 1987 were excluded as they referred to earlier less specific PTH assays [7, 8].
The authors agreed and trialled spreadsheet fields. Data were extracted from the selected papers by EH and checked by EJL. In some cases, confirmation of data was sought from authors through direct contact (e.g., e-mail). Data extracted included the number, nature and concentration range of samples, sample types studied and tube manufacturer, sample processing and storage conditions, assay method and manufacturer, comparator/reference used in each study, statistical analysis and a summary of findings including any information relating to circadian rhythm, pulsatile frequency and peak amplitude. The principal outcome measure was change in PTH concentration expressed in pmol/L or occasionally as a percentage. (N.B. 1 pmol/L is approx. equivalent to 9.43 ng/L).
Currently recommended methodological systems of assigning level of evidence and strength of recommendation could not easily be adopted to assess the evidence we identified . Such systems have primarily been designed to assess evidence from therapeutic trials  or diagnostic studies  as opposed to pre-analytical sampling practices. Furthermore, for most of the data we identified it was not possible to produce quantitative summaries across studies. We have used a transparent evidence-based approach to review systematically, appraise critically and then discuss the evidence, including highlighting the strengths and limitations of the evidence and identifying gaps in the knowledge base behind the recommendations. In assessing the literature, for each of the three PICO questions we considered study design, internal validity and consistency across studies and directness.
Most studies were prospective with clearly reported protocols, so the risk of bias was considered low. The risk of allocation bias was also low, particularly in studies addressing PICO question 1 where samples from all subjects within individual studies were compared under the same conditions (e.g., storage temperature, sample type and tube type). Strongest weight was placed on those studies which we considered to equate with “level 1” evidence as described by Hayes et al.  (evidence from a single, high-powered, prospective, controlled study that is specifically designed to test a marker) or with “high level” evidence as described in the GRADE approach . For PICO questions 2 and 3 all studies were observational cohort studies. When grading our recommendations we chose to use the approach described by Kidney Disease: Improving Global Outcomes (KDIGO) guideline group which acknowledges some of the restrictions of other systems when applied to data such as ours . Recommendations were classified as either “strong”, indicating a choice that most well-informed people would make, or “weak”, indicating a choice that a majority of well-informed people would make but a substantial minority would not: recommendations were consequently worded as “We recommend…” or “We suggest…” respectively .
With respect to sample stability, the time points presented in the text and figures are the longest times for which PTH was reported to be stable in each study. Conclusions with respect to stability are conservative estimates taking all relevant studies into account. In addition to the literature search, information was acquired from manufacturers product inserts for several commonly used commercial PTH immunoassays.
The systematic review was written in accordance with preferred reporting items for systematic reviews and meta-analyses (PRISMA) guidelines  and a completed PRISMA checklist is presented in Supplemental Data Table 2.
Results and discussion
The electronic search identified 5478 papers (Figure 1). Thirty-three additional articles referenced in these papers were also included. Thirty-three meeting abstracts were also identified as relevant to the study of which five had subsequently been published. Of these five, four had already been identified by the search. One potentially relevant paper was excluded as it contained insufficient detail for data assimilation. A total of 96 papers were selected for full text review, of which 83 were finally included. Most studies identified by the search were performed using second generation assays. As described in the Methods section, for the first PICO question the search was limited to papers published after 1987 as data obtained using earlier assays [7, 8] were considered to be of limited relevance to contemporary clinical practice.
Variability in study design
Although PTH stability ex vivo has been extensively studied, authors have not always reached the same conclusions. Assimilating and collating data from different studies was complicated by the different approaches taken. Confounding factors included variability in the length of time and specific time points over which stability was studied, variability in the time delay between venepuncture and separation of serum/plasma from cellular components of blood, missing data for PTH stability at specific time points and for specific types of samples, and/or failure in some studies to extend the observation period for sufficient time to enable identification of differences between sample types and/or comparison with baseline results. The practice recommendations, described below and summarised in Table 2, focused on results from studies in which there were direct comparisons between sample types, and those studies that answered questions most relevant to clinical practice.
Stability of PTH in whole blood, serum and plasma
PICO Question 1: In human blood samples, how stable is PTH in EDTA or lithium heparin whole blood or plasma compared to clotted whole blood or separated serum at room temperature, 4°C, −20°C and −80°C?
Studies focusing on PTH stability in blood samples were performed with different types of blood collection tubes: plain tubes, tubes containing anticoagulant (potassium-EDTA, citrate or lithium heparin) and/or gel separating tubes (GST) containing a barrier gel to separate serum from cellular blood constituents (cells and protein clot) after centrifugation. One study used results from plasma or serum from fresh whole blood as comparator , but most used results from serum or plasma which had been separated and frozen prior to analysis. For the following analysis, data have been grouped according to sample type and assay characteristics (i.e., second or third generation).
Stability in anticoagulated and clotted whole blood at room temperature
Fifteen papers relating to PTH stability in whole blood were identified that had used a second generation assay (Figure 2, Supplemental Data Table 3) [15–29]. Collation of study results was difficult due to variation in experimental design (see above), but almost all studies suggested that PTH was more stable in whole blood samples containing EDTA than in anticoagulant-free samples (Supplemental Data Table 3). More definitive evidence to support this was provided by seven studies that assessed the stability of different sample types within the same study using second generation methods [15, 16, 18, 22, 25, 26, 29] and one study using a third generation method  (Table 3). These data suggested that PTH was stable in EDTA whole blood for at least 24 h at room temperature as compared to only 3 h in clotted whole blood.
Stability in separated serum and plasma at room temperature
Eighteen papers studying PTH stability in separated serum and plasma using second generation (Supplemental Data Table 4) [19, 20, 22–25, 27, 29, 31–35, 38–42] and four using third generation (Supplemental Data Table 5) [30, 36, 37, 43] assays were identified. PTH stability in EDTA plasma and serum was directly compared in seven studies using second generation assays [25, 29, 31–35]. In these, PTH was more stable in EDTA plasma than in serum (Table 3). In one study slight increases in measured PTH concentration in EDTA plasma after 24 and 48 h were observed, although it was concluded overall that PTH was stable for 72 h in EDTA plasma but for only 24 h in serum . Overall, these results suggested that PTH was stable in separated EDTA plasma for at least 48 h at room temperature but that in serum significant immunoreactivity may be lost after as little as 2 h.
Stability at other temperatures
Only one study directly compared the stability of PTH in clotted and EDTA whole blood at 4°C: PTH was reported stable for 72 h at 4°C in both sample types  (Figure 3). PTH was also reported to be stable in EDTA whole blood for at least 18 h  or 24 h  at 4°C and in clotted whole blood for 30 h  (Table 3, Supplemental Data Table 3). Nine studies addressed stability at 4°C in serum or plasma (Supplemental Data Table 4) [19, 22, 24, 25, 29, 33, 34, 38, 39]. Five of these directly compared the stability of PTH in serum and plasma at 4°C [25, 29, 33, 34, 38] using second generation assays generally confirming superior stability in EDTA (Table 3). In both sample types, PTH was more stable at 4°C than at room temperature.
Three studies addressed long-term frozen storage of PTH using second generation assays (Supplemental Data Table 4). When serum and EDTA plasma samples were kept at −20°C for 24 h, no difference in measured PTH was observed for the two sample types, while PTH degradation was significantly greater in EDTA plasma after 5 days at −20°C . At −80°C, Brinc et al.  demonstrated instability of PTH after 2 months (up to 16% loss compared to baseline) in serum and EDTA plasma. Using third generation methods, PTH was reportedly stable for 14 days in serum  and for 12 months in serum and plasma kept at −20°C and −80°C . Observed stability may be method dependent: Cavalier et al. [14, 16] reported that if stored at −20°C or −80°C PTH was more stable in serum than EDTA plasma when measured using the DiaSorin Liaison method (9 vs. 2 months, respectively). In contrast, PTH was stable in both frozen serum and plasma for at least 2 years when analysed using the Roche Elecsys method. It is possible that this might reflect conformational alterations during freezing but further confirmatory studies are required.
Using a variety of assays, PTH withstood four (serum and EDTA plasma ) to six (serum [23, 43]) freeze-thaw cycles without measured recovery being affected.
Recommendations for sample handling
With respect to analyte stability ex vivo, most studies, with both second and third generation assays, indicate PTH to be more stable in EDTA whole blood than clotted whole blood [15, 16, 18, 25, 26, 29, 30], and in EDTA and lithium heparin plasma than in serum at room temperature [25, 31–35]. Whilst direct comparisons of lithium heparin whole blood with EDTA whole blood , and lithium heparin plasma with EDTA plasma [25, 32], suggested similar stability could be achieved in lithium heparin preserved tubes, there was limited evidence upon which to base a recommendation. Furthermore, one manufacturer (Diasorin) does not support the use of lithium heparin. The following statement therefore seems appropriate:
1. We recommend blood samples for PTH measurement should be taken into tubes containing EDTA and the plasma separated from the cells within 24 h of venepuncture [Strong recommendation].
This recommendation is consistent with guidance issued by the Clinical Laboratory Standards Institute  and the World Health Organization . It is also broadly in accord with advice provided in manufacturers’ kit inserts, most of which suggest measurement of PTH in either EDTA plasma or serum, but confirm that PTH is more stable in EDTA plasma than serum (Table 4, [35, 37, 47]). We accept that PTH is commonly measured in conjunction with calcium, and sometimes vitamin D, to permit interpretation and that this recommendation will necessitate an additional sample being taken since calcium cannot be measured in EDTA plasma. This is clearly a practical limitation, but it is not unique to assessment of bone mineral metabolism (e.g., assessment of the pituitary-adrenal axis requires different samples for ACTH and cortisol). Clearly under optimal sample handling conditions (<3 h between venepuncture and separation/measurement) clotted blood may also be a suitable sample , but such stringent requirements are not likely to be achievable in many clinical laboratories.
Once separated, PTH was more stable in EDTA plasma than in serum, but the stability of PTH in both sample types could be successfully extended by refrigeration.
While losses of PTH observed in clotted blood samples may be small within the time frame of a typical working day (e.g., 8%  or 10%  after 8 h; 10% after 12 h ), such differences could contribute to misdiagnosis or changes in management of patients. Given the many other factors that influence clinical interpretation of PTH results (e.g., between-method differences, specificity) , laboratory professionals should ensure samples are appropriately stored. Based on the evidence reviewed here, the following recommendation seems appropriate:
2. We recommend EDTA plasma samples for PTH measurement should be stored at 4ºC and analysed within 72 h of venepuncture [Strong recommendation].
Historically many laboratories have measured PTH in batch mode following freezing of samples at −20°C for variable periods of time. Many manufacturers’ recommendations will support this practice (Table 4). However, published evidence was inconsistent regarding the stability of PTH under frozen storage conditions. Should laboratories still need to freeze plasma prior to PTH measurement, we suggest that they establish the stability of PTH in frozen plasma as measured with their own assay.
Influence of sampling site
PICO Question 2: In human blood samples, does the sampling site affect PTH concentration?
In clinical practice, most samples are taken from the antecubital vein. However, in haemodialysis patients, samples are often taken through a central line. In haemodialysis patients, PTH concentrations were found to be 30% higher in central blood (superior vena cava, median 24.3 pmol/L, interquartile range 9.2–38.2 pmol/L) compared to peripheral blood (forearm vein, median 15.3 pmol/L, interquartile range 6.3–29.0 pmol/L)  (Supplemental Data Table 6). Similarly in patients with primary hyperparathyroidism undergoing parathyroidectomy with intra-operative PTH monitoring, central venous (internal jugular vein) PTH concentrations were higher compared to peripheral venous PTH concentration (17.5 vs. 10.8 pmol/L ; 1.3–20.0 pmol/L higher ).
In relation to the above two clinical situations, particular regard must be paid to the site of origin of the blood samples. Use of central venous catheters in haemodialysis is variable but significant. For example, in the UK 65% of haemodialysis patients commenced dialysis in 2005 using a central venous catheter although by 12 months this had fallen to 30% . In Canada between 2001 and 2004 the prevalence of catheter use in the haemodialysis population was 52% .
3. We recommend blood samples for PTH measurement should always be collected from the same sample site (central or peripheral) for comparison both within and between individuals. Clinical guidelines should explicitly state whether targets refer to peripheral or central venous concentrations [Strong recommendation].
Influence of time of sampling
PICO Question 3: In human blood samples, does the time of sampling affect PTH concentration?
PTH concentrations in blood have been reported to fluctuate according to the season [54–57] (Supplemental Data Table 7). With the exception of one small early study using a first generation assay , all studies identified reported a relative decrease in PTH concentration in summer and an increase in winter [54–57]. Studies undertaken with second generation assays suggested the difference between mean winter and summer concentrations was <1 pmol/L [57, 59]. All studies were undertaken in the northern hemisphere.
It seems likely that the observed pattern may mirror and reflect seasonal variation of vitamin D concentration [59, 60]. Given the clear inverse relationship between PTH and vitamin D it has been suggested that reference ranges for PTH should be established in vitamin D-replete individuals [60, 61]. However, this remains a controversial area. Whether the observed seasonal variation in vitamin D concentration is pathological and not normal physiology is difficult to assess. The definition of vitamin D sufficiency, often regarded as the concentration above which PTH cannot be suppressed further, varies widely (e.g., from 30 to 110 nmol/L ) and indeed, in the largest study to date, no threshold above which increasing vitamin D concentration failed to further suppress PTH could be identified . Furthermore, the relationship between PTH and vitamin D was highly dependent on age . The population in which PTH is most commonly measured is also known to have a high prevalence of vitamin D deficiency/insufficiency  and therefore use of a PTH reference range derived in vitamin D repleted individuals may be inappropriate.
4. We suggest season, latitude and vitamin D status should be considered and/or reported in all studies undertaking reference range determinations for PTH and when interpreting PTH results in individual patients [Weak recommendation].
Studies addressing circadian variation of PTH are summarised in Supplemental Data Table 8 [64–90]. Most studies reported a circadian bimodal rhythm with a nocturnal acrophase, a mid-morning nadir and a smaller afternoon peak (Table 5). Peak times varied between studies and were affected by gender [75, 77]. There was also interindividual variability in the return to baseline (between 06:00 and 10:00) [81, 91]. Most studies reported a circadian amplitude amongst healthy individuals of between 0.3 and 0.8 pmol/L [64, 66, 68, 69, 72, 73, 75, 77, 79, 81, 82, 90] although higher amplitudes of 1.2  and 1.9  pmol/L, respectively, were also reported. Circadian variation was absent in patients with thalassemia  and primary hyperparathyroidism [80, 81].
Logue et al.  recommend that blood samples should be collected between 10:00 and 16:00 and results interpreted against a reference range based on this sampling time. Since the data were derived using first or second generation assays, it is possible that the reported diurnal variation of PTH could reflect differential clearance of PTH fragments over a 24 h period, e.g., with reduced renal clearance of (7-84) PTH at night reflecting decreased glomerular filtration rate at night .
5. We suggest blood samples for PTH measurement should be collected between 10:00 and 16:00 and results interpreted against a reference interval derived for this sampling time [Weak recommendation].
We found no studies that addressed the relative diagnostic accuracy of PTH measurement at different time of the day. Generally studies were undertaken in small cohorts and no studies specifically addressing this question in CKD patients were identified. Partly because of this there are concerns about the validity of the data identified and we have only made a weak recommendation. Our recommendation is based upon practical considerations since it avoids the times of day over which PTH peaks are observed.
Studies addressing pulsatile variation of PTH are summarised in Supplemental Data Table 9 [68, 76, 79, 93–102]. As for many peptide hormones, a pulsatile secretory pattern was superimposed on the circadian rhythm of PTH. Thirteen studies addressed this issue. In all but one , PTH secretion was found to be pulsatile with one to seven secretory pulses per hour [68, 79, 94–96, 98, 99, 101, 102]. Reported pulse amplitudes were 0.5 , 0.8  and 1.8  pmol/L.
The ability to detect secretory pulses depends on the frequency of blood sampling, which varied greatly between studies. Not all studies covered an entire 24 h period and no studies were undertaken using a third generation assay. Most data suggested that pulse amplitude was similar in magnitude to the effects of circadian variation noted above, approximately 1 pmol/L. In clinical practice it is not possible to mitigate against the effects of pulse amplitude in terms of measuring PTH and this probably contributes to the high biological variation observed for PTH [103, 104]. The effects of pulsatile secretion should be considered when interpreting PTH results (e.g., by recommending confirmatory sampling when appropriate). In the future this may suggest a role for other bone markers (e.g., bone alkaline phosphatase), which might provide a more time-averaged measure of PTH effect, analogous to the use of insulin-like growth factor 1 (IGF1) measurement to integrate the pulsatility of growth hormone secretion .
Other pre-analytical factors influencing PTH measurement
We have focused on pre-analytical sampling and storage conditions for PTH. However, a number of studies describing other pre-analytical influences on PTH concentration were identified. Food ingestion may  or may not  affect PTH concentration. In many of the studies reported above [64, 66, 67, 69, 71, 74, 75, 77, 79, 83–85, 87, 89, 90] related to circadian variation, subjects were provided with meals at specified times of day: no clear relationship between feeding and PTH concentration was reported. Fasts of 33 h and 96 h led to the loss of PTH circadian rhythm [107, 108]. A vegetarian diet led to a higher PTH concentration compared to a meat diet (5.9 vs. 4.9 pmol/L ). The reported effect of strenuous exercise was inconsistent [110–113], and sleep did not seem to affect the circadian and/or pulsatile release of PTH . Higher mean PTH concentrations were reported in men compared to women (4.4 vs. 2.8 pmol/L) , in African-American women compared to white women (2.0 vs. 3.3 pmol/L)  and in postmenopausal females compared to males and premenopausal females (5.8 pmol/L vs. 5.4 and 4.7 pmol/L, respectively) .
There were many limitations to the studies we included. Most used as comparator a sample which had been frozen at baseline, as opposed to a freshly analysed sample, used by one study only , so interpretation of test results could have been confounded by changes in PTH concentration following freezing. Different approaches were taken to define significance of change, e.g., some studies used a prespecified percentage change compared to baseline [16, 17, 25, 29, 33, 38, 44], whilst others used paired statistical analyses [15, 18, 19, 21–24, 26, 28, 30–32, 34, 35, 37, 39, 40, 42, 43]. PTH as measured by most clinical assays is not a single entity with assays recognising different molecular forms to different extents and the prevalence of these forms varying between individuals and disease states. Most studies were performed on samples from patients with kidney disease or hyperparathyroidism, and we cannot be confident therefore that our conclusions are necessarily generalisable to other patient groups or healthy individuals. Nevertheless in the majority of studies, PTH was more stable in EDTA whole blood or plasma than clotted whole blood or serum, irrespective of experimental design.
We found no direct published comparisons of clotted and EDTA whole blood stability at 4°C. Data with respect to stability of serum and plasma at −20°C and −80°C were extremely limited and we are unable to draw conclusions in this regard. We were somewhat surprised by this given the relative consistency of manufacturer’s recommendations in this respect (Table 4) and the fact that, until relatively recently, many laboratories analysed PTH in batch mode with frozen storage prior to analysis. We may have missed such evaluation data by limiting our searches to the era of second and third generation PTH assays. However, we consider that data derived using earlier PTH assays is not reliable in this respect. Clinical laboratories and researchers undertaking studies where samples are to be stored frozen should establish analyte stability for their own sample types and assays.
Second and third generation PTH assays measure different mixtures of peptides which may have differing stability [114, 115]. We found no direct published comparisons of observed analyte stability with second versus third generation PTH assays but there were some data to suggest that PTH may be more stable when measured by third rather than second generation assays . This could be explained on the basis that the peptide fragments detected by the second generation assays may be less stable than the intact molecule detected by third generation assays. If confirmed, this should provide further impetus to support the use of third generation assays.
Conclusions and suggestions for further study
Pre-analytical sampling and storage conditions affecting PTH concentration have been systematically reviewed and recommendations for good practice developed (Table 2). Several areas of uncertainty remain which should be the subject of further research. These include: 1) how stability of PTH in EDTA whole blood as measured with a third generation assay compares with that as measured with a second generation assay; 2) the circadian and seasonal variation of PTH as measured with a third generation assay; 3) how CKD and vitamin D status affect these rhythms; and 4) further data are also required regarding the stability of PTH at −20°C, −30°C (i.e., below the eutectic point) and −80°C with both second and third generation assays. There is also an urgent requirement for sound reference range data derived under clearly defined sampling conditions. For the present useful progress can be made by adopting and implementing the recommendations in Table 2. This should significantly improve the comparability and consistency of PTH data in all clinical settings.
This systematic review was initiated at the suggestion of a working group chaired by C. Sturgeon which convened at the Royal College of Pathologists in London in September 2010 to discuss aspects of standardisation of PTH measurement . We are grateful to M. Kerr from the Library at Kent and Canterbury Hospital for his help and advice. We are extremely grateful to the following members and supporters of the IFCC Scientific Division Working Group on PTH who advised and commented on an earlier draft of this manuscript, although their acknowledgement here should not necessarily be taken as an endorsement of this review: A. Algeciras-Schimnich, J. Barth, G. Beastall, G. Bobba, C. Burns, E. Cavalier, A. Dawnay, J. Middle, B. Schodin, P. Sibley, J.C. Souberbielle, S. Sprague, H. Vesper and I. Young.
Conflict of interest statement
Authors’ conflict of interest disclosure: There are no conflicts of interest regarding the publication of this article.
Research funding: None declared.
Employment or leadership: None declared.
Honorarium: None declared.
Kidney Disease: Improving Global Outcomes (KDIGO) CKD–MBD Work Group. KDIGO clinical practice guideline for the diagnosis, evaluation, prevention, and treatment of chronic kidney disease-mineral and bone disorder (CKD-MBD). Kidney Int 2009;76(Suppl 113): S1–130.Google Scholar
Souberbielle JC, Boutten A, Carlier MC, Chevenne D, Coumaros G, Lawson-Body E, et al. Inter-method variability in PTH measurement: implication for the care of CKD patients. Kidney Int 2006;70:345–50.PubMedCrossrefGoogle Scholar
Sturgeon CM, Sprague SM, Metcalfe W. Variation in parathyroid hormone immunoassay results – a critical governance issue in the management of chronic kidney disease. Nephrol Dial Transplant 2011;26:3440–5.CrossrefGoogle Scholar
Almond A, Ellis AR, Walker SW. Current parathyroid hormone immunoassays do not adequately meet the needs of patients with chronic kidney disease. Ann Clin Biochem 2012;49:63–7.PubMedCrossrefGoogle Scholar
Nussbaum SR, Zahradnik RJ, Lavigne JR, Brennan GL, Nozawa-Ung K, Kim LY, et al. Highly sensitive two-site immunoradiometric assay of parathyrin, and its clinical utility in evaluating patients with hypercalcemia. Clin Chem 1987;33:1364–7.PubMedGoogle Scholar
Brown RC, Aston JP, Weeks I, Woodhead JS. Circulating intact parathyroid hormone measured by a two-site immunochemiluminometric assay. J Clin Endocrinol Metab 1987;65:407–14.CrossrefPubMedGoogle Scholar
Atkins D, Best D, Briss PA, Eccles M, Falck-Ytter Y, Flottorp S, et al. Grading quality of evidence and strength of recommendations. Br Med J 2004;328:1490–4.Google Scholar
Hayes DF, Bast RC, Desch CE, Fritsche H, Jr., Kemeny NE, Jessup JM, et al. Tumor marker utility grading system: a framework to evaluate clinical utility of tumor markers. J Natl Cancer Inst 1996;88:1456–66.PubMedCrossrefGoogle Scholar
Uhlig K, Macleod A, Craig J, Lau J, Levey AS, Levin A, et al. Grading evidence and recommendations for clinical practice guidelines in nephrology. A position statement from Kidney Disease: Improving Global Outcomes (KDIGO). Kidney Int 2006;70:2058–65.PubMedGoogle Scholar
Liberati A, Altman DG, Tetzlaff J, Mulrow C, Gotzsche PC, Ioannidis JP, et al. The PRISMA statement for reporting systematic reviews and meta-analyses of studies that evaluate health care interventions: explanation and elaboration. J Clin Epidemiol 2009;62:e1–34.CrossrefGoogle Scholar
Cavalier E, Delanaye P, Hubert P, Krzesinski JM, Chapelle JP, Rozet E. Estimation of the stability of parathyroid hormone when stored at -80 degrees C for a long period. Clin J Am Soc Nephrol 2009;4:1988–92.CrossrefGoogle Scholar
Morales Garcia AI, Gorriz Teruel JL, Plancha Mansanet MC, Escudero Quesada V, Pallardo Mateu LM. Analysis of variability in determining intact parathyroid hormone (iPTH) according to the method used to process the sample. Nefrologia 2009;29:331–5.PubMedGoogle Scholar
Omar H, Chamberlin A, Walker V, Wood PJ. Immulite 2000 parathyroid hormone assay: stability of parathyroid hormone in EDTA blood kept at room temperature for 48 h. Ann Clin Biochem 2001;38:561–3.Google Scholar
Parent X, Alenabi F, Brignon P, Souberbielle JC. Delayed measurement of PTH in patients with CKD: storage of the primary tube in the dialysis unit, which temperature? Which kind of tube? Nephrol Ther 2009;5:34–40.PubMedCrossrefGoogle Scholar
Stokes FJ, Ivanov P, Bailey LM, Fraser WD. The effects of sampling procedures and storage conditions on short-term stability of blood-based biochemical markers of bone metabolism. Clin Chem 2011;57:138–40.CrossrefPubMedGoogle Scholar
Zwart SR, Wolf M, Rogers A, Rodgers S, Gillman PL, Hitchcox K, et al. Stability of analytes related to clinical chemistry and bone metabolism in blood specimens after delayed processing. Clin Biochem 2009;42:907–10.CrossrefPubMedGoogle Scholar
Cavalier E, Carlisi A, Bekaert AC, Rousselle O, Chapelle JP, Delanaye P. New insights on the stability of the parathyroid hormone as assayed by an automated 3rd generation PTH assay. Clin Chim Acta 2012;413:353–4.CrossrefGoogle Scholar
Hermsen D, Franzson L, Hoffmann JP, Isaksson A, Kaufman JM, Leary E, et al. Multicenter evaluation of a new immunoassay for intact PTH measurement on the Elecsys System 2010 and 1010. Clin Lab 2002;48:131–41.Google Scholar
Holmes DT, Levin A, Forer B, Rosenberg F. Preanalytical influences on DPC IMMULITE 2000 intact PTH assays of plasma and serum from dialysis patients. Clin Chem 2005;51:915–7.Google Scholar
Gao P, Scheibel S, D’Amour P, John MR, Rao SD, Schmidt-Gayk H, et al. Development of a novel immunoradiometric assay exclusively for biologically active whole parathyroid hormone 1-84: implications for improvement of accurate assessment of parathyroid function. J Bone Miner Res 2001;16:605–14.CrossrefGoogle Scholar
Parent X, Alenabi F, Etienne E, Brignon P, Chantrel F, Meynaud-Kraemer L. Preanalytical variability of parathyroid hormone measurement in dialysis patients; application to Elecsys 2010 (Roche) automate. Ann Biol Clin (Paris) 2008;66:53–8.Google Scholar
Joly D, Drueke TB, Alberti C, Houillier P, Lawson-Body E, Martin KJ, et al. Variation in serum and plasma PTH levels in second-generation assays in hemodialysis patients: a cross-sectional study. Am J Kidney Dis 2008;51:987–95.CrossrefPubMedGoogle Scholar
Inaba M, Nakatsuka K, Imanishi Y, Watanabe M, Mamiya Y, Ishimura E, et al. Technical and clinical characterization of the Bio-PTH (1-84) immunochemiluminometric assay and comparison with a second-generation assay for parathyroid hormone. Clin Chem 2004;50:385–90.CrossrefGoogle Scholar
Brinc D, Chan MK, Venner AA, Pasic MD, Colantonio D, Kyriakopolou L, et al. Long-term stability of biochemical markers in pediatric serum specimens stored at -80 degrees C: a CALIPER substudy. Clin Biochem 2012;45:816–26.CrossrefGoogle Scholar
Clinical and Laboratory Standards Institute. Procedures for the handling and processing of blood specimens for common laboratory tests; approved guideline, 4th ed. Document H18-A4. Wayne, PA: Clinical and Laboratory Standards Institute, 2010:1–57.Google Scholar
World Health Organization. Use of anticoagulants in diagnostic laboratory investigations and stability of blood, plasma and serum samples. WHO/DIL/LAB/99.1 Rev.2. Geneva: WHO, 2002:1–64.Google Scholar
Clinical and Laboratory Standards Institute (formerly NCCLS). Procedures for the handling and processing of blood specimens; approved guideline, 3rd ed. NCCLS Document H18-A3. Wayne, PA: Clinical and Laboratory Standards Institute, 2004.Google Scholar
Vulpio C, Bossola M, Speranza D, Zuppi C, Luciani G, Di Stasio E. Influence of blood sampling site on intact parathyroid hormone concentrations in hemodialysis patients. Clin Chem 2010;56:489–90.PubMedCrossrefGoogle Scholar
Broome JT, Schrager JJ, Bilheimer D, Chambers EP, Jacobs JK, Phay J. Central venous sampling for intraoperative parathyroid hormone monitoring: are peripheral guidelines applicable? Am Surg 2007;73:712–6.PubMedGoogle Scholar
Beyer TD, Chen E, Ata A, DeCresce R, Prinz RA, Solorzano CC. A prospective evaluation of the effect of sample collection site on intraoperative parathormone monitoring during parathyroidectomy. Surgery 2008;144:504–9; discussion 9–10.CrossrefGoogle Scholar
Ansell D, Feest T, Hodsman A, Rao R, Tomson C, Udayaraj U, et al. The Renal Association UK Renal Registry Ninth Annual Report. 2006;1–288.Google Scholar
Lips P, Hackeng WH, Jongen MJ, van Ginkel FC, Netelenbos JC. Seasonal variation in serum concentrations of parathyroid hormone in elderly people. J Clin Endocrinol Metab 1983;57:204–6.CrossrefPubMedGoogle Scholar
Meller Y, Kestenbaum RS, Galinsky D, Shany S. Seasonal variation in serum levels of vitamin D metabolites and parathormone in geriatric patients with fractures in Southern Israel. Isr J Med Sci 1986;22:8–11.PubMedGoogle Scholar
Woitge HW, Knothe A, Witte K, Schmidt-Gayk H, Ziegler R, Lemmer B, et al. Circaannual rhythms and interactions of vitamin D metabolites, parathyroid hormone, and biochemical markers of skeletal homeostasis: a prospective study. J Bone Miner Res 2000;15:2443–50.PubMedCrossrefGoogle Scholar
Guagnano MT, Del Ponte A, Menduni P, Nuzzo A, Palummeri E, Angelucci E, et al. Time structure of endocrine secretion. II. Circannual variations in the free fractions of tri- and tetraiodothyronine, cortisol, human growth hormone and plasma insulin in healthy subjects. Boll Soc Ital Biol Sper 1983;59:1243–7.Google Scholar
Krall EA, Sahyoun N, Tannenbaum S, Dallal GE, Dawson-Hughes B. Effect of vitamin D intake on seasonal variations in parathyroid hormone secretion in postmenopausal women. N Engl J Med 1989;321:1777–83.PubMedCrossrefGoogle Scholar
Eastell R, Arnold A, Brandi ML, Brown EM, D’Amour P, Hanley DA, et al. Diagnosis of asymptomatic primary hyperparathyroidism: proceedings of the third international workshop. J Clin Endocrinol Metab 2009;94:340–50.CrossrefPubMedGoogle Scholar
Souberbielle JC, Cormier C, Kindermans C, Gao P, Cantor T, Forette F, et al. Vitamin D status and redefining serum parathyroid hormone reference range in the elderly. J Clin Endocrinol Metab 2001;86:3086–90.PubMedCrossrefGoogle Scholar
Levin A, Bakris GL, Molitch M, Smulders M, Tian J, Williams LA, et al. Prevalence of abnormal serum vitamin D, PTH, calcium, and phosphorus in patients with chronic kidney disease: results of the study to evaluate early kidney disease. Kidney Int 2007;71:31–8.CrossrefGoogle Scholar
Ahmad AM, Hopkins MT, Fraser WD, Ooi CG, Durham BH, Vora JP. Parathyroid hormone secretory pattern, circulating activity, and effect on bone turnover in adult growth hormone deficiency. Bone 2003;32:170–9.CrossrefGoogle Scholar
Bell NH, Williamson BT, Hollis BW, Riggs BL. Effects of race on diurnal patterns of renal conservation of calcium and bone resorption in premenopausal women. Osteoporos Int 2001;12:43–8.PubMedCrossrefGoogle Scholar
Calvo MS, Eastell R, Offord KP, Bergstralh EJ, Burritt MF. Circadian variation in ionized calcium and intact parathyroid hormone: evidence for sex differences in calcium homeostasis. J Clin Endocrinol Metab 1991;72:69–76.PubMedCrossrefGoogle Scholar
Chapotot F, Gronfier C, Spiegel K, Luthringer R, Brandenberger G. Relationships between intact parathyroid hormone 24-hour profiles, sleep-wake cycle, and sleep electroencephalographic activity in man. J Clin Endocrinol Metab 1996;81:3759–65.CrossrefGoogle Scholar
El-Hajj Fuleihan G, Klerman EB, Brown EN, Choe Y, Brown EM, Czeisler CA. The parathyroid hormone circadian rhythm is truly endogenous – a general clinical research center study. J Clin Endocrinol Metab 1997;82:281–6.PubMedGoogle Scholar
Even L, Bader T, Hochberg Z. Nocturnal calcium, phosphorus and parathyroid hormone in the diagnosis of concealed and subclinical hypoparathyroidism. Eur J Endocrinol 2007;156:113–6.CrossrefPubMedGoogle Scholar
Fraser WD, Logue FC, Christie JP, Gallacher SJ, Cameron D, O’Reilly DS, et al. Alteration of the circadian rhythm of intact parathyroid hormone and serum phosphate in women with established postmenopausal osteoporosis. Osteoporos Int 1998;8:121–6.PubMedCrossrefGoogle Scholar
Generali D, Berruti A, Tampellini M, Dovio A, Tedoldi S, Bonardi S, et al. The circadian rhythm of biochemical markers of bone resorption is normally synchronized in breast cancer patients with bone lytic metastases independently of tumor load. Bone 2007;40:182–8.CrossrefPubMedGoogle Scholar
Goodman WG, Misra S, Veldhuis JD, Portale AA, Wang HJ, Ament ME, et al. Altered diurnal regulation of blood ionized calcium and serum parathyroid hormone concentrations during parenteral nutrition. Am J Clin Nutr 2000;71:560–8.PubMedGoogle Scholar
Greenspan SL, Holland S, Maitland-Ramsey L, Poku M, Freeman A, Yuan W, et al. Alendronate stimulation of nocturnal parathyroid hormone secretion: a mechanism to explain the continued improvement in bone mineral density accompanying alendronate therapy. Proc Assoc Am Physicians 1996;108:230–8.PubMedGoogle Scholar
Joseph F, Chan BY, Durham BH, Ahmad AM, Vinjamuri S, Gallagher JA, et al. The circadian rhythm of osteoprotegerin and its association with parathyroid hormone secretion. J Clin Endocrinol Metab 2007;92:3230–8.PubMedCrossrefGoogle Scholar
Jubiz W, Canterbury JM, Reiss E, Tyler FH. Circadian rhythm in serum parathyroid hormone concentration in human subjects: correlation with serum calcium, phosphate, albumin, and growth hormone levels. J Clin Invest 1972;51:2040–6.CrossrefPubMedGoogle Scholar
Kitamura N, Shigeno C, Shiomi K, Lee K, Ohta S, Sone T, et al. Episodic fluctuation in serum intact parathyroid hormone concentration in men. J Clin Endocrinol Metab 1990;70: 252–63.CrossrefPubMedGoogle Scholar
Lobaugh B, Neelon FA, Oyama H, Buckley N, Smith S, Christy M, et al. Circadian rhythms for calcium, inorganic phosphorus, and parathyroid hormone in primary hyperparathyroidism: functional and practical considerations. Surgery 1989;106:1009–16; discussion 16–7.Google Scholar
Logue FC, Fraser WD, Gallacher SJ, Cameron DA, O’Reilly DS, Beastall GH, et al. The loss of circadian rhythm for intact parathyroid hormone and nephrogenous cyclic AMP in patients with primary hyperparathyroidism. Clin Endocrinol 1990;32:475–83.CrossrefGoogle Scholar
Logue FC, Fraser WD, O’Reilly DS, Beastall GH. The circadian rhythm of intact parathyroid hormone (1-84) and nephrogenous cyclic adenosine monophosphate in normal men. J Endocrinol 1989;121:R1–3.CrossrefGoogle Scholar
Markowitz ME, Arnaud S, Rosen JF, Thorpy M, Laximinarayan S. Temporal interrelationships between the circadian rhythms of serum parathyroid hormone and calcium concentrations. J Clin Endocrinol Metab 1988;67:1068–73.PubMedCrossrefGoogle Scholar
Nielsen HK, Brixen K, Kassem M, Christensen SE, Mosekilde L. Diurnal rhythm in serum osteocalcin: relation with sleep, growth hormone, and PTH(1-84). Calcif Tissue Int 1991;49:373–7.CrossrefGoogle Scholar
Nielsen HK, Laurberg P, Brixen K, Mosekilde L. Relations between diurnal variations in serum osteocalcin, cortisol, parathyroid hormone, and ionized calcium in normal individuals. Acta Endocrinol (Copenh) 1991;124:391–8.PubMedGoogle Scholar
Rejnmark L, Lauridsen AL, Vestergaard P, Heickendorff L, Andreasen F, Mosekilde L. Diurnal rhythm of plasma 1,25-dihydroxyvitamin D and vitamin D-binding protein in postmenopausal women: relationship to plasma parathyroid hormone and calcium and phosphate metabolism. Eur J Endocrinol 2002;146:635–42.CrossrefGoogle Scholar
Robinson MF, Body JJ, Offord KP, Heath H, 3rd. Variation of plasma immunoreactive parathyroid hormone and calcitonin in normal and hyperparathyroid man during daylight hours. J Clin Endocrinol Metab 1982;55:538–44.CrossrefGoogle Scholar
Sinha TK, Miller S, Feming J, Khairi R, Edmondson J, Johnston CC, Jr., et al. Demonstration of a diurnal variation in serum parathyroid hormone in primary and secondary hyperparathyroidism. J Clin Endocrinol Metab 1975;41:1009–13.CrossrefGoogle Scholar
White HD, Ahmad AM, Durham BH, Chandran S, Patwala A, Fraser WD, et al. Effect of active acromegaly and its treatment on parathyroid circadian rhythmicity and parathyroid target-organ sensitivity. J Clin Endocrinol Metab 2006;91:913–9.PubMedGoogle Scholar
Koopman MG, Koomen GC, Krediet RT, de Moor EA, Hoek FJ, Arisz L. Circadian rhythm of glomerular filtration rate in normal individuals. Clin Sci (Lond) 1989;77:105–11.Google Scholar
De Francisco AL, Amado JA, Cotorruelo JG, Gonzalez M, De Castro SS, Canga E, et al. Pulsatile secretion of parathyroid hormone in patients with chronic renal failure. Clin Nephrol 1993;39:224–8.PubMedGoogle Scholar
Harms HM, Schlinke E, Neubauer O, Kayser C, Wustermann PR, Horn R, et al. Pulse amplitude and frequency modulation of parathyroid hormone in primary hyperparathyroidism. J Clin Endocrinol Metab 1994;78:53–7.CrossrefPubMedGoogle Scholar
Harms HM, Neubauer O, Kayser C, Wustermann PR, Horn R, Brosa U, et al. Pulse amplitude and frequency modulation of parathyroid hormone in early postmenopausal women before and on hormone replacement therapy. J Clin Endocrinol Metab 1994;78:48–52.PubMedCrossrefGoogle Scholar
Heidbreder E, Naujoks H, Brosa U, Schramm L. The calcium/parathyroid hormone regulation in chronic renal failure investigation of its dynamic secretion pattern. Horm Metab Res 1997;29:70–5.CrossrefGoogle Scholar
Samuels MH, Veldhuis J, Cawley C, Urban RJ, Luther M, Bauer R, et al. Pulsatile secretion of parathyroid hormone in normal young subjects: assessment by deconvolution analysis. J Clin Endocrinol Metab 1993;77:399–403.CrossrefPubMedGoogle Scholar
Samuels MH, Veldhuis JD, Kramer P, Urban RJ, Bauer R, Mundy GR. Episodic secretion of parathyroid hormone in postmenopausal women: assessment by deconvolution analysis and approximate entropy. J Bone Miner Res 1997;12:616–23.CrossrefPubMedGoogle Scholar
Schmitt CP, Huber D, Mehls O, Maiwald J, Stein G, Veldhuis JD, et al. Altered instantaneous and calcium-modulated oscillatory PTH secretion patterns in patients with secondary hyperparathyroidism. J Clin Endocrinol Metab 1998;9:1832–44.Google Scholar
Schmitt CP, Locken S, Mehls O, Veldhuis JD, Lehnert T, Ritz E, et al. PTH pulsatility but not calcium sensitivity is restored after total parathyroidectomy with heterotopic autotransplantation. J Am Soc Nephrol 2003;14:407–14.PubMedCrossrefGoogle Scholar
Schmitt CP, Schaefer F, Bruch A, Veldhuis JD, Schmidt-Gayk H, Stein G, et al. Control of pulsatile and tonic parathyroid hormone secretion by ionized calcium. J Clin Endocrinol Metab 1996;81:4236–43.PubMedCrossrefGoogle Scholar
Gardham C, Stevens PE, Delaney MP, LeRoux M, Coleman A, Lamb EJ. Variability of parathyroid hormone and other markers of bone mineral metabolism in patients receiving hemodialysis. Clin J Am Soc Nephrol 2010;5:1261–7.CrossrefPubMedGoogle Scholar
Ankrah-Tetteh T, Wijeratne S, Swaminathan R. Intraindividual variation in serum thyroid hormones, parathyroid hormone and insulin-like growth factor-1. Ann Clin Biochem 2008;45:167–9.CrossrefGoogle Scholar
Sethi R, Kukreja SC, Bowser EN, Hargis GK, Henderson WJ, Williams GA. Effect of meal on serum parathyroid hormone and calcitonin: possible role of secretin. J Clin Endocrinol Metab 1983;56:549–52.PubMedCrossrefGoogle Scholar
Fraser WD, Logue FC, Christie JP, Cameron DA, O’Reilly DS, Beastall GH. Alteration of the circadian rhythm of intact parathyroid hormone following a 96-hour fast. Clin Endocrinol 1994;40:523–8.CrossrefGoogle Scholar
Moe SM, Zidehsarai MP, Chambers MA, Jackman LA, Radcliffe JS, Trevino LL, et al. Vegetarian compared with meat dietary protein source and phosphorus homeostasis in chronic kidney disease. Clin J Am Soc Nephrol 2011;6:257–64.CrossrefPubMedGoogle Scholar
Ljunghall S, Joborn H, Roxin LE, Skarfors ET, Wide LE, Lithell HO. Increase in serum parathyroid hormone levels after prolonged physical exercise. Med Sci Sports Exerc 1988;20:122–5.PubMedCrossrefGoogle Scholar
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Published Online: 2013-09-27
Published in Print: 2013-10-01