The 24,25-dihydroxyvitamin D [24,25(OH)₂D] is a catabolite of 25-hydroxyvitamin D [25(OH)D]. This transformation is performed by 1,25-hydroxyvitamin D 24-hydroxylase (or 24-hydroxylase, encoded by the CYP24A1 gene). Mutations in CYP24A1 can lead to severe diseases such as idiopathic infantile hypercalcemia (IIH). Explorations of hypercalcemia with suppressed parathyroid hormone levels and normal or high phosphatemia should now include 24,25(OH)2D determination to exclude CYP24A1 mutations. 24,25(OH)₂D and the vitamin D metabolite ratio (VMR) [i.e. 25(OH)D/24,25(OH)₂D] are now considered as new biomarkers for the assessment of functional vitamin D deficiency.

by L. Vranken, C. Fontaine, Prof. JC. Souberbielle and Prof. E. Cavalier

Vitamin D metabolism
Nowadays, there is an increased focus on the vitamin D and its benefits on health maintenance and disease prevention. Vitamin D is mainly produced following skin exposure to UVB rays. Additionally, it is found in several foods, such as oily fish, mushrooms and egg yolk. Vitamin D is considered as a pro-hormone owing to the fact that its production in the skin from 7-dehydrocholesterol could be sufficient when the sun exposure is adequate. Two forms of vitamin D coexist: vitamin D₂ produced by vegetables, and vitamin D₃ produced by animals and humans [2, 8]. After its synthesis in the skin or its intestinal absorption, this liposoluble vitamin is transported to the liver where it is hydroxylated by vitamin D 25-hydroxylase (or 25-hydroxylase, encoded by the CYP2R1 gene) to form 25-hydroxyvitamin D [25(OH)D]. This hydroxylation is very poorly regulated and, therefore, most of the circulating vitamin D will be metabolized into 25(OH)D. 25(OH)D is then transported to the kidney by a specific protein carrier [vitamin D binding protein (DBP)], and to a lesser extent by albumin, where it is hydroxylated by 25-hydroxyvitamin D-1 alpha hydroxylase (or 1α-hydroxylase, encoded by the CYP27B1 gene) on the carbon in position 1 to form the most active metabolite, 1,25-dihydroxy-vitamin D [1,25(OH)₂D]. This transformation is strictly regulated, notably by the parathyroid hormone (PTH), fibroblast growth factor 23 (FGF23) and 1,25(OH)₂D itself (Fig. 1). The major role of vitamin D is the maintenance of calcium homeostasis, by acting on the vitamin D receptor (VDR). Calcium regulation is very complex and not fully understood yet. When ionized calcium decreases, the calcium sensing receptors (CaSR) located on the surface of the parathyroid glands stimulate PTH secretion.

PTH then acts on different targets to increase serum calcium concentration: it stimulates the release of calcium (and phosphate) from bones by acting on osteoclasts through osteoblasts and the RANK/RANKL system. It also decreases calcium excretion by the kidney and stimulates 1α-hydroxylase to produce 1,25(OH)₂D which, in turn, acts on the VDR of intestinal cells to produce calbindin 9k, TRPV6 and the NCX1 Ca/Na exchanger increasing intestinal absorption of calcium. The resulting increase of calcium levels inhibits CaSR-stimulated PTH production, but 1,25(OH)₂D also acts as a feedback loop to stop PTH synthesis. 1,25(OH)₂D finally acts on the VDR of the FGF23 gene to stimulate FGF23 production. In turn, FGF23, which is the most potent phosphaturic hormone (it inhibits Npt2a and Npt2c sodium-dependent phosphate co-transporters in the proximal renal tubule), blocks the activity of 1α-hydroxylase and stimulates 24-hydroxylase which leads to 25(OH)D and 1,25(OH)₂D catabolism (Fig. 2).

24-Hydroxylase is a key enzyme that catalyses the inactivation of svitamin D. It is expressed in most vitamin D target cells and is also stimulated by 1,25(OH)₂D, which hence regulates its own metabolism, therefore protecting against hypercalcemia and limiting the levels of 1,25(OH)₂D in cells [1]. Production of 1,24,25(OH)₃D and 24,25(OH)₂D is the first step of a five-step pathway that transforms vitamin D in a more hydrophilic compound, calcitroic acid, and allows its excretion in urine and in bile [2–6, 8]. 24,25(OH)₂D has a half-life of approximately 7 days and a concentration in the range of 1 to 10 ng/mL in healthy individuals.

CYP24A1 mutations
Loss-of-function mutations of the CYP24A1 gene have been identified in children presenting with idiopathic infantile hypercalcemia (IIH). These CYP24A1 gene product (24-hydroxylase) defects can be inherited as an autosomal recessive biallelic mutation. Infants present with severe hypercalcemia, suppressed PTH levels, hypercalciuria and medullary nephrocalcinosis owing to hypersensitivity to
vitamin D [4]. Indeed, there is no transformation of 25(OH)D and 1,25(OH)₂D to 24,25(OH)₂D and 1,24,25(OH)₃D leading to a prolonged and excessive elevation of 25(OH)D and 1,25(OH)₂D concentrations and an incapacity to clear them from plasma. By feedback, there will be a decrease of PTH and an increase in FGF23 concentrations (Fig. 2). These symptoms are similar to those met in vitamin D intoxication and it is important to make the distinction between these two diseases. In IIH, the vitamin D metabolite ratio (VMR), the ratio between 25(OH)D and 24,25(OH)₂D, allows the differential diagnosis of 24-hydroxylase defects from vitamin D intoxication. In IIH, the VMR will be high (>50–80); that is to say high 25(OH)D with low 24,25(OH)₂D, and is indicative of idiopathic hypercalcemia due to CYP24A1 gene mutations. In vitamin D intoxication, the VMR is normal because both 25(OH)D and 24,25(OH)₂D are increased. Moreover, the VMR may be more accurate for revealing this mutation than 24,25(OH)₂D alone because the ratio takes into consideration the circulating 25(OH)D and provides a clear distinction from a vitamin D deficiency, in which both 25(OH)D and 24,25(OH)₂D are low. Indeed, if the substrate decreases, in this case 25(OH)D, the activity of 24-hydroxylase is reduced, thus the production of 24,25(OH)₂D is low [4]. These genetic mutations indicate that vitamin D supplementation in children could be potentially deleterious. In these children, vitamin D supplementation must be eliminated. Indeed, they may have failure to thrive, vomiting, dehydratation, spikes of fever and nephrocalcinosis. Supplementation of mothers with 24-hydroxylase defects during pregnancy could lead to hypercalcemia associated with prematurity and intra-uterine growth retardation. Treatment of IIH encompasses the avoidance of sun and calcium- and vitamin D-rich foods. However, recently, it has been shown that isoniazid could induce the cytochrome P450 3A4, which is another vitamin D degradation pathway [9].

Thereafter, Molin et al. found that CYP24A1 gene mutations are frequently associated with renal complications including renal failure, nephrolithiasis and nephrocalcinosis. Also, they suggest that this loss-of-function of 24-hydroxylase is the most recently elucidated cause of hypercalcemia after parathyroid hypercalcemia, vitamin D intoxication and poorly regulated 1α-hydroxylation [3]. They have described patients with CYP24A1 heterozygous mutations, mostly asymptomatic, implying a hypothesis of an autosomal-dominant trait from which clinical consequences would vary throughout life and where hypercalcemia would appear only when vitamin D intakes are excessive.

Less severe mutations have been observed in patients with moderate hypercalcemia and inappropriately low PTH (<20 pg/mL). Those patients are likely to develop nephrolithiasis. 24,25(OH)₂D evaluation should be done on subjects with hypercalcemia and low PTH, especially as they suffer from nephrolithiasis. Not all the mutations have been discovered yet and further genetic studies are required. Moreover Ginsberg et al. found that lower 24,25(OH)₂D concentrations and lower VMR are associated with increased hip-fracture risk in community-living older men and women. They also noticed that higher 24,25(OH)₂D concentrations were associated with higher bone mineral density (BMD), whereas VMR was not. Additionally, 1,25(OH)₂D concentrations were not associated with BMD, consistent with previous studies in older adults [1]. In addition to catabolism, many studies tend to demonstrate that the 24,25(OH)₂D may have its own biological activity in vitro in calcium regulation [5, 6]. Finally, recent studies suggest that the assessment of 24,25(OH)₂D or the assessment of the VMR could better reflect the activity of the VDR and could be used as an index of vitamin D clearance [1, 3, 4]. The VMR may have the advantage of being uninfluenced by DBP concentrations, which affects both the numerator and denominator of the ratio.

Vitamin D metabolite evaluation
Quantitative evaluation of 24,25(OH)₂D is complicated by its presence at low concentrations. LC-MS/MS is currently the only alternative to evaluate 24,25(OH)₂D levels and has the great advantage to distinguish simultaneously the different metabolites and 25(OH)D in serum [6, 10]. The NIST (National Institute of Standards and Technology) has recently issued a new serum-matrix standard reference material [11] and Tai et al. published a reference measurement procedure for the determination of 24,25(OH)₂D in human serum using isotope-dilution LC-MS/MS [10].

Conclusion
In conclusion, the assessment of 25(OH)D alone is not always enough. 24,25(OH)₂D and VMR are other available tools to help for the diagnosis and the monitoring of abnormalities in  phosphocalcic metabolism. The drawback is that it requires the determination of vitamin D metabolites by LC-MS/MS, and very few laboratories perform this determination [only 10 labs participate in the 24,25(OH)2D proficiency testing provided by the Vitamin D External Quality Assessment Scheme (DEQAS)]. Collaboration with a reference lab may be a good compromise. It is important to be aware of hypercalcemia caused by CYP24A1 mutants and their consequences on health. Further studies will be needed to explore the others mutations of CYP24A1 and the potential biological activity of 24,25(OH)₂D in vivo.

References
1. Ginsberg C, Katz R, de Boer IH, Kestenbaum BR, Chonchol M, Shlipak MG, Sarnak MJ, Hoofnagle AN, Rifkin DE, et al. The 24,25 to 25-hydroxyvitamin D ratio and fracture risk in older adults: the cardiovascular health study. Bone 2018; 107: 124–130.
2. Vranken L, Emonts P, Bruyère O, Cavalier E. Prévalence de l’hypovitaminose D chez la femme enceinte: quelle est la situation en région liégeoise? Revue Médicale de Liège 2018; 73 (1): 10–16 [in French].
3. Molin A, Baudoin R, Kaufmann M, Souberbielle JC, Ryckewaert A, Vantyghem MC, Eckart P, Bacchetta J, Deschenes G, et al. CYP24A1 mutations in a cohort of hypercalcemic patients: evidence for a recessive trait. J Clin Endocrinol Metab 2015; 100(10): E1343–E1352.
4. Schlingmann KP, Kaufmann M, Weber S, Irwin A, Goos C, John U, Misselwitz J, Klaus G, Kuwertz-Bröking E, et al. Mutations in CYP24A1 and idiopathic infantile hypercalcemia. N Engl J Med 2011; 365(5): 410–421.
5. Van Leeuwen JPTM, an den Bemd GJCM, van Driel M, Buurman CJ, Pols HAP. 24,25-Dihydroxyvitamin D3 and bone metabolism. Steroids 2011; 66: 375–380.
6. Wagner D, Hanwell HE, Schnabl K, Yazdanpanah M, Kimball S, Fu L, Sidhom G, Rousseau D, Cole DEC, Vieth R. The ratio of serum 24,25-dihydroxyvitamin D3 to 25-hydroxyvitamin D3 is predictive of 25-hydroxyvitamin D3 response to vitamin D3  supplementation. J Steroid Biochem Mol Biol 2011; 126: 72–77.
7. Lu X, Chen Z, Mylarapu N, Watsky MA. Effects of 1,25 and 24,25 vitamin D on corneal epithelial proliferation, migration and vitamin D metabolizing and catabolizing enzymes. Sci Rep 2017; 16951: 1–12.
8. Bikle DD. Vitamin D and bone. Curr Osteoporos Rep 2012; 10(2): 151–159.
9. An inducible cytochrome P450 3A4-dependent vitamin D catabolic pathway. Wang Z, Lin YS, Zheng XE, Senn T, Hashizume T, Scian M, Dickmann LJ, Nelson SD, Baillie TA, et al. Mol Pharmacol 2012; 81(4): 498–509.
10. Tai SSC, Nelson MA. Candidate reference measurement procedure for the determination of (24R),25-dihydroxyvitamin D3 in human serum using isotope-dilution liquid chromatography-tandem mass spectrometry. Anal Chem 2015; 87: 7964–7970.
11. Tai SS, Nelson MA, Bedner M, Lang BE, Phinney KW, Sander LC, Yen JH, Betz JM, Sempos CT, Wise SA. Development of standard reference material (SRM) 2973 vitamin D metabolites in frozen human serum (high level). J AOAC Int 2017; 100(5): 1294–1303.

The authors
Laura Vranken¹, Corentin Fontaine¹, Jean-Claude Souberbielle² PhD, Etienne Cavalier¹ PhD
1Clinical Chemistry, University of Liège, CHU Sart-Tilman, Belgium
²Service des Explorations Fonctionnelles, Hôpital Necker-Enfants Malades, Paris, France

*Corresponding author
E-mail: Laura.vranken@chuliege.be

Dermatomycoses are extremely widespread, and are characteristically long-lasting, recurring and very difficult to cure. Early and accurate identification of the causative agent is essential for targeted therapy. A new DNA microarray provides direct detection and differentiation of the most important dermatomycosis pathogens in one reaction. The assay simultaneously detects up to 50 dermatophyte species, and provides species identification for 23 of these, as well as 6 yeasts and moulds. The microarray analysis aids differential diagnosis of dermatomycoses from other dermatoses (e.g. psoriasis), and specifically identifies mixed infections with yeasts and moulds. The dermatomycosis microarray is part of the established EUROArray platform, which also includes microarrays for multiplex identification of sexually transmitted infections (STI) and complete detection and typing of human papillomaviruses (HPV).

by Dr Jacqueline Gosink

Dermatomycosis
Dermatomycoses are infections of the skin, hair and nails which are typically caused by dermatophytes and in rarer cases by yeasts and moulds. Fungal infections of the skin are the most frequently occurring infectious diseases globally with high and growing relapse rates. Elderly people and immunocompromised patients are especially at risk. Worldwide, around 20 to 25% of the population is affected by fungal skin diseases.

Infections which are caused exclusively by dermatophytes are referred to as dermatophytoses or tinea. Tinea pedis, which occurs on the soles of the feet and between the toes, is one of the most frequent forms worldwide, followed by tinea unguium, which affects the nails, and tinea corporis, which affects the neck, back or trunk. Further forms, for example, on the face, legs, beard area, arms and hands, are rarer. Nail infections caused by dermatophytes and/or yeasts/moulds are called onychomycoses. They are typically accompanied by deformation of the nail.

Pathogens of dermatomycosis
Dermatophytes encompass fungi of the genera Trichophyton, Epidermophyton, Nannizzia, Paraphyton, Lophophyton, Microsporum and Arthroderma. Individual species are classified as anthropophilic, zoophilic or geophilic according to their main occurrence. Human pathogenic yeasts and moulds include Candida spp., Scopulariopsis brevicaulis, Fusarium spp. and Aspergillus fumigatus.

Around 70% of human dermatophyte infections are caused by anthropophilic species. Trichophyton rubrum, in particular, is the most frequent cause of fungal skin infections worldwide. Infections can spread easily to other areas of the body or to other persons, for example, via showers, bathtubs or floors. Zoophilic dermatophytes are transmitted to humans by close contact with animals, especially pets, which are often asymptomatic. They can cause severe inflammatory reactions in humans. Geophilic dermatophytes cause disease less frequently in humans. Infections typically occur in gardeners and farm workers or children who play outside. Moulds and yeasts often cause opportunistic infections, benefitting from damage to the skin or nail caused by an existing dermatophyte infection. In immunocompromised individuals, local fungal infections may develop into systemic mycosis.

Clinical picture
Dermatomycoses are clinically heterogeneous and cannot always be differentiated from other dermatoses, such as eczema, psoriasis, erysipelas, or autoimmune diseases such as Lichen ruber planus. Furthermore, 5 to 15% of onychomycosis cases comprise mixed infections of dermatophytes with yeasts and moulds. Simultaneous bacterial infection of the damaged skin, pretreatment with corticosteroid-containing preparations, or secondary contact allergy can also hinder diagnosis.

Dermatomycoses must always be treated. This is generally undertaken using various topical antifungal drugs, with severe cases sometimes requiring oral medication. Each drug has a limited activity spectrum. Positive pathogen identification prior to treatment enables targeted selection of the most suitable drug and optimal planning of the oftentimes lengthy therapy. In multiple infections, a change of the primary pathogenic agent may occur during the therapy and it may seem like the therapy is failing. This must be taken into account in the treatment of fungal infections of the nails, which may only yield first success after months. Identification of the causative pathogen also helps to determine the source of the infection. In the case of zoophilic pathogens, for example, this is usually a pet.

Laboratory diagnostics
Laboratory diagnostic methods for identifying dermatomycosis pathogens include microscopic detection and an attempt at culturing from clinical material. Successful culture in most cases enables species assignment based on micro- and macromorphological presentation of the fungus. Culturing is, however, time-consuming and is not possible for all dermatophytes. In mixed infections, false diagnoses may occur since slowly growing species may be overgrown or overlooked. Furthermore, antifungal therapy started before the sampling can hinder the culture.

Direct detection of pathogen genomic material by DNA microarray enables secure and accurate identification of the causative agent. Microarray analysis offers a significant time advantage over detection by culturing, and is especially useful for detecting dermatophytes that are difficult to cultivate. It provides higher sensitivity and specificity, even in patients already undergoing treatment. The EUROArray Dermatomycosis analysis includes a universal dermatophyte detection encompassing 50 species of the genera Trichophyton, Epidermophyton, Microsporum, Nannizzia, Arthoderma, Lophophyton, as well as species identification for the 23 dermatophyte and 6 yeast and mould species listed in Table 1.

EUROArray procedure
The EUROArray procedure (Figure 1) is performed on DNA samples isolated from skin scales, nail shavings or hair stubs. Defined gene sections of the pathogens are first amplified by multiplex polymerase chain reaction (PCR). The fluorescently labelled PCR products are then incubated with biochip microarray slides containing immobilized complementary probes. Specific binding (hybridization) of the PCR products to their corresponding oligonucleotide probes is detected using a special microarray scanner. The signals are evaluated and interpreted automatically by the EUROArrayScan software (Figure 2). A detailed result report is produced for each patient and all data are documented and archived. Meticulously designed primers and probes, ready-to-use PCR components and integrated controls all contribute to the reliability of the analysis. The entire EUROArray procedure from sample arrival to report release is IVD-validated and CE-registered, supporting quality management in diagnostic laboratories.

Specifications and evaluation
The lower detection limit of the test system depends on the pathogen and lies between 50 and 600 DNA copies per reaction, in individual cases also higher. Evaluation studies verified that template DNA in concentrations ranging from the lower detection limit to 50 ng can be used in the PCR without generating any false positive results. Furthermore, potential cross reactivity with 37 microorganisms of the resident and transient skin flora was excluded experimentally.

In an evaluation study with 409 clinical samples, the EUROArray Dermatomycosis yielded a good agreement with the precharacterization. In many cases additional pathogens that were not included in the precharacterization were detected. The additional findings were confirmed by further independent tests or sequencing. Thus, the microarray provides reliable results and broad detection capabilities.

STI detection
The EUROArray STI is based on the same technology and provides parallel direct detection of the pathogenic agents of eleven sexually transmitted infections (STIs) in one reaction, namely Chlamydia trachomatis, Neisseria gonorrhoea, Mycoplasma genitalium, Mycoplasma hominis, Ureaplasma urealyticum, Ureaplasma parvum, Haemophilus ducreyi, Treponema pallidum, Trichomonas vaginalis and herpes simplex viruses 1 and 2.
STIs are often asymptomatic, but can nevertheless lead to serious sequelae, for example infertility, fetal damage during pregnancy, and severe postnatal infections in newborns. The PCR-based detection allows identification of both manifest and silent infections, and is thus suitable for diagnosis of symptomatic patients as well as for general screening. It offers a huge time advantage over cultivation and is especially useful for detecting sexually transmitted pathogens that are difficult or impossible to cultivate, e.g. C. trachomatis, Mycoplasma, Ureaplasma, T. pallidum. Due to amplification of the pathogen DNA, infections with a reduced pathogen number can also be reliably detected. A broad screening for STI pathogens is particularly important in asymptomatic or clinically ambiguous cases and for detecting multiple infections, which are often missed during single-parameter testing.

HPV detection and typing
The EUROArray HPV provides detection and typing of all 30 genitally relevant HPV subtypes in one test. HPV are involved in the development of cervical carcinoma, and HPV testing plays a central role in risk assessment and early diagnosis of this cancer. In contrast to Pap examinations, HPV detection is not dependent on subjective evaluation and it offers very high sensitivity even in the early stages of infection.
The EUROArray HPV is based on detection of the viral oncogenes E6 and E7, which provides the highest possible sensitivity. Using an extensive panel of specific primers and probes, the EUROArray detects and distinguishes between 18 high-risk subtypes that may trigger cancer (16, 18, 26, 31, 33, 35, 39, 45, 51, 52, 53, 56, 58, 59, 66, 68, 73, 82) and 12 low-risk subtypes that cause benign genital warts (6, 11, 40, 42, 43, 44, 54, 61, 70, 72, 81, 89). Multiple infections are reliably identified, and primary and persistent infections can be differentiated. A positive result for a high-risk subtype indicates an increased risk for cervical carcinoma, which can then be minimised by more frequent follow-up examinations to detect morphological cell changes at an early stage. Based on the recommendations of the respective professional societies, HPV-negative women can forgo subsequent HPV tests and Pap smears for a longer time interval.

Conclusions
Molecular diagnostic tests such as the EUROArray are an important tool for identifying the precise pathogenic agent in various infectious diseases, supporting decision-making on specific treatment. The new EUROArray Dermatomycosis provides the most comprehensive direct detection of dermatomycosis pathogens currently available commercially, and complements existing assays for STI and HPV. The EUROArray procedure is easy to perform and does not require extensive expertise in molecular biology. Moreover, the fully automated evaluation ensures objective, accurate and reproducible results. Further infectious disease microarrays based on the same technology are in development.

The author
Jacqueline Gosink, PhD
EUROIMMUN AG, Seekamp 31,
23560 Luebeck, Germany

www.euroimmun.com

The activated partial thromboplastin time coagulation assay is one of the most frequently performed tests in hematology, and has a variety of uses in clinical practice. Accurate interpretation of the test depends on both clinical context (i.e. why the test was ordered) as well as an understanding of each laboratory’s normal reference range and assay sensitivity regarding detection of factor deficiencies, (unfractionated) heparin therapy and lupus anticoagulant.

by Dr Julianne Falconer and Dr Emmanuel J. Favaloro

Introduction
The activated partial thromboplastin time (APTT) assay is a commonly requested coagulation test, perhaps second only to the prothrombin time (PT)/international normalized ratio (INR), as used to monitor vitamin K antagonist (VKA) therapy such as warfarin. The APTT test assesses the intrinsic pathway of coagulation and has a variety of clinical uses; however, it is primarily used to screen for hemostasis issues, factor deficiencies, lupus anticoagulant (LA) or to monitor unfractionated heparin (UFH) therapy dosing. The test is sensitive to, but not specific for, detection of these abnormalities or influences. APTT prolongation may also be seen in liver disease, disseminated intravascular coagulation (DIC) and in the presence of factor inhibitors. Interpretation of an APTT result, be it normal or prolonged, is dependent on both the clinical context and the characteristics of the reagents and the assay as performed on particular instruments. The establishment of normal reference intervals (NRIs) and assessment of the assay in terms of its sensitivity to heparin, LA and clotting factors are important to provide accurate information for clinical interpretation [1].

Uses of the APTT assay
The APTT test is a global assay that measures the time to fibrin clot formation via the contact factor (‘intrinsic’) pathway (Fig. 1). The APTT test is usually performed on fully automated platforms, and involves activation of coagulation within the test (plasma) sample by the addition of specific reagents (containing phospholipids, contact factor activator and calcium chloride). The type of contact factor activator, and the type and concentration of phospholipid, used in the APTT reagent affects the sensitivity of the assay to, and thus its prolongation by, factor deficiencies, as well as to the presence of UFH and LA [1, 2].

The APTT is commonly used to monitor anticoagulation therapy using UFH (Table 1). It may also be prolonged, however, in the presence of VKAs including warfarin, as well as direct oral anticoagulants (DOACs) such as dabigatran (direct thrombin inhibitor) and rivaroxaban (anti-FXa inhibitor). The APTT is generally less sensitive to, but may still be slightly prolonged, by anticoagulation with low molecular weight heparin (LMWH) and with apixaban, another DOAC (anti-FXa inhibitor).

In the absence of anticoagulation therapy, an ‘isolated’ prolonged APTT may be used to determine a clinically important factor deficiency, for example as a screen for hemophilia A (FVIII deficiency), hemophilia B (FIX deficiency), or hemophilia C (FXI deficiency), or even von Willebrand Disease (VWD; which may be associated with loss of FVIII) [1]. An ‘isolated’ prolonged APTT, however, could instead be indicative of a clinically unimportant factor deficiency such as FXII or other contact factor deficiency. Other alternatives for an ‘isolated’ prolonged APTT include a factor inhibitor or LA. Despite causing prolongation of APTT in vitro, LA may be associated clinically with increased risk of thrombosis rather than bleeding. A prolonged APTT may be accompanied by a prolonged PT in the context of liver disease, DIC or fibrinogen (or other ‘common factor pathway’ deficiency/ies). Clinical context, therefore, must form the basis for accurate interpretation of APTT, be it either normal or prolonged, and together with other routine coagulation studies is essential to guide further investigations (Fig. 2).

A large number of commercial APTT reagents are now available, with wide variation in the type of contact factor activator and phospholipid source and concentration used. This will result in variation in sensitivity to all typical influences; thus also causing substantial variation in NRIs between APTT reagents, and requiring the establishment and verification of NRIs based on both the reagent and instrument in use. Unawareness of variation in APTT reagent sensitivity in context of clinical picture will lead to flawed clinical interpretation of results.

Establishing and verification of NRIs
A minimum of 20 normal individuals may be sufficient to establish a NRI for PT and APTT, according to guidance documents provided by the Clinical and Laboratory Standards Institute (CLSI) [3, 4]. However, a larger number of normal individuals is recommended to establish an initial NRI, following which a smaller sample of normal individuals may be used for future verification purposes [1].

As an example, Figure 3 shows an initial (historical) NRI estimation for APTT testing using a dataset of nearly 80 normal individuals. This included one outlier sample result (Fig. 3a), which was removed to produce the cleaner dataset used to produce the subsequent NRI. A statistical normality test was performed and showed the distribution to be near Gaussian, allowing parametric statistical assessment. For APTT testing, the NRI would aim to evaluate the 95 % confidence interval, approximating a mean
± 2 standard deviation (SD) assessment (Fig. 3b). Logarithmic transformation can instead be used to normalize test data when it is non-parametric and fits a log distribution (e.g. Fig. 3c).

If a NRI has been previously established by the laboratory or by the manufacturer of the APTT reagent using a specific reagent/instrument combination, the laboratory could use a process of transference to verify the ‘established’ NRI as fit for purpose. This may be done by establishing that a majority of samples in a small set of normal donors give values within the established NRI (e.g. >18 out of a set of 20 normal samples). Samples obtained from normal individuals or a dataset of normal patient test results may be used to assess a new lot of reagent to establish whether an existing NRI can be maintained when changing reagent lots.

Factor (deficiency) sensitivity
Factor sensitivity of an APTT assay (representing a specific reagent/instrument combination) can be assessed in a number of ways. One method involves serial dilution of either in-house or commercially derived normal plasma, into single-factor deficient plasma, in order to generate a series of aliquots with reducing factor levels. These samples are then tested by APTT and for factor level. The APTT reagent is regarded to be sensitive to the level of factor that correlates with the upper limit of the NRI.

A more accurate process, though particularly difficult to perform outside of a hemophilia centre, is to establish APTT values from true patients with various known factor levels [1, 2] (e.g. Fig. 4).

As a general guide, if the APTT is used for screening factor deficiencies, then the patient APTT value should be above the NRI when their factor level is below around 30–40 U/dL for FVIII, FIX, and FXI.

Sensitivity of APTT to UFH
Despite the changing landscape of anticoagulation therapy with the addition of direct anti-Xa inhibitors (rivaroxaban and apixaban) and a direct thrombin inhibitor (dabigatran) [5, 6], both LMWH and UFH continue to be frequently used in clinical practice. In turn, the APTT continues to be a generally preferred method of UFH monitoring over anti-FXa, given the wide availability and relative low cost of the assay. However, unlike the calibrated anti-FXa assay, APTT results are subject to variation between different instruments, be they be based on optical or mechanical clot detection methods [7], different APTT reagents (including variation between different lots of the same reagent type) and algorithms used on instruments for raw data processing. This poses a substantial problem with regards to historical recommendations to maintain patients on UFH between 1.5 and 2.5 times the ‘normal reference value’ (as based on limited evidence [8]). Therapeutic ranges should therefore be defined with specific reference to the instrument/reagent combination used locally [9].

One ‘spiking method’ involves testing samples containing known quantities of UFH diluted into normal pool plasma, as then tested by APTT and anti-FXa methods, allowing an estimation of the APTT therapeutic interval [1]. However, variations in certain components of patient plasma, as well as the non-physiologically processed nature of the UFH used, can impact on the interpretation of data obtained using this method. A better method involves ex vivo assessment of plasma obtained from patients on UFH therapy, with these tested for both APTT and anti-FXa, and then to establish a UFH therapeutic range for APTT that matches the therapeutic range for anti-FXa (e.g. 0.3–0.7 U/mL). It is important to recognize that individual response to UFH according to APTT is affected by many influences, including (but not limited to): antithrombin level; high or low levels of coagulation factors and proteins such as von Willebrand factor or proteins released from endothelial cells or platelets, competing with antithrombin for heparin binding; or increased FVIII levels in acute phase response; or reduction in FXII; or presence of LA (etc).

To obtain a cleaner data set to establish UFH therapeutic ranges, the following steps can be undertaken during sample collection and processing [1].
• Ensure baseline PT, APTT and INR testing prior to commencement of UFH are within their NRIs.
• Exclude underfilled samples, samples with visible hemolysis or likely platelet activation and release of heparin neutralizer platelet factor 4 (PF4).
• Exclude samples containing LMWH or other anticoagulants (e.g. VKAs, DOACs).
• Adhere to manufacturer guidelines with regards to the window from time of blood collection to testing.
• Double centrifuge samples when freezing them for batch testing (to remove residual platelets, which release PF4 and phospholipids on thawing).
• Accumulate data over a suitable time period to account for day-to-day test result variability.
• Aim for 30 or more data points.
• Appropriately dilute samples with anti-Xa activity above the test’s linearity limit.
• Remove data points reflecting ‘gross’ outliers.

LA sensitivity
The LA sensitivity of a particular APTT reagent can be assessed by comparing APTT tests of samples containing LA, for example by comparison of mean clotting times for each reagent.
Given that the APTT is a phospholipid-dependent assay, the test may be susceptible to prolongation in the presence of LA. However, differences in the phospholipid type and concentration between APTT reagents account for wide variation seen in the degree of prolongation of APTT, including due to LA. The LA sensitivity of the APTT reagent also has bearing on the use of APTT to monitor UFH and must inform the establishment of an algorithm to further investigate unexpectedly prolonged APTTs.
In one empirical method, initial testing using an LA sensitive method (e.g. dilute Russell viper venom time; dRVVT) is initially used to formulate a set of LA-positive samples of various ‘strengths’. Different APTT reagents can then be used to test the samples and the data for each sample can be plotted again the upper reference limit of the APTT for each reagent [1]. The ratio of clotting time of each LA-positive sample (of varying strengths) to the mean normal APTT derived from normal plasma samples is calculated. The median of these ratios allows different reagents to be ranked according to LA sensitivity. It can then become clear which APTT reagents are most (versus least) sensitive to LA. These can then be differentially selected according to the laboratory desire. For example, a laboratory may prefer to select an APTT reagent that is relatively LA ‘insensitive’, as combined with good factor VIII/IX/XI and UFH sensitivity if there is a desire to use a general purpose APTT screening reagent (i.e. hospital laboratory monitoring UFH, but wishing to avoid LA detection in asymptomatic patients). Alternatively, a laboratory may select an LA sensitive and an insensitive APTT reagent pair if they wish to assess for LA in symptomatic (thrombosis and/or pregnancy morbidity) patients.

Conclusion
Interpretation of a normal or a prolonged APTT must take into account both clinical context, including presence of anticoagulant therapy, as well as the methods and reagents used by the laboratory. The sensitivity of a particular APTT reagent to detect UFH therapy, LA and factor deficiencies has significant bearing on diagnostic assessment and therapy monitoring, and thus reflects essential knowledge for laboratory and clinical staff alike.

Figure 1. The activated partial thromboplastin time (APTT) assay measures the clot time to formation of fibrin via the contact factor pathway and is dependent on contact factors (FXII and above), and then FXI, FIX, FVIII, FX, FV, and FII. The APTT is also affected by vitamin K antagonists (VKAs; ‘W’), but more importantly is used to monitor unfractionated heparin (UFH; ‘H’) therapy and also to assess for potential hemophilia (FVIII, FIX or FXI deficiency). The APTT is also sensitive to the presence of other anticoagulants, including direct oral anticoagulants (DOACs) such as dabigatran (‘D’) and rivaroxaban (‘R’), and potentially also apixaban (‘A’) for some reagents. The APTT may also be utilized as part of a panel of tests to help assess for lupus anticoagulant (LA). (Modified from Favaloro EJ, et al. How to optimize activated partial thromboplastin time (APTT) testing: solutions to establishing and verifying normal reference intervals and assessing APTT reagents for sensitivity to heparin, lupus anticoagulant, and clotting factors. Semin Thromb Hemost 2019; 45: 22–35 [1].)
Figure 2. An algorithm that provides one recommended approach for the follow-up of an abnormal APTT. Always exclude an anticoagulant effect first – there is no point investigating a prolonged APTT associated with anticoagulant use. Then consider the patient’s history, or the clinical reason for the test order, both of which assist in terms of follow-up approach. APTT, activated partial thromboplastin time; FBC/CBC, full blood count (UK/Australia)/complete blood count (USA); DIC, disseminated intravascular coagulation; DOAC, direct oral anticoagulant; EDTA, ethylenediaminetetraacetic acid; F, factor; LA, lupus anticoagulant; PT, prothrombin time. (Modified from Favaloro EJ, et al. How to optimize activated partial thromboplastin time (APTT) testing: solutions to establishing and verifying normal reference intervals and assessing APTT reagents for sensitivity to heparin, lupus anticoagulant, and clotting factors. Semin Thromb Hemost 2019; 45: 22–35 [1].)
Table 1. The APTT test. A multipurpose and sensitive assay, but not specific for any individual parameter. List is not meant to be all inclusive.
DOACs, direct oral anticoagulants; VWD, von Willebrand disease.
*PT should also be prolonged if APTT is prolonged in the indicated setting.
(Modified from Favaloro EJ, et al. How to optimize activated partial thromboplastin time (APTT) testing: solutions to establishing and verifying normal reference intervals and assessing APTT reagents for sensitivity to heparin, lupus anticoagulant, and clotting factors. Semin Thromb Hemost 2019; 45: 22–35 [1].)
Figure 3. Historical data from our laboratory to illustrate the process of deriving a normal reference interval (NRI) for the APTT, and using nearly 80 normal individual plasma samples. (a) APTT of all samples tested shown as a dot plot; one clear outlier shown as a red asterisk. (b) Data cleaned of outliers [i.e. in this case the single red asterisk sample in (a)]. (c) NRR estimate as mean ± 2 standard deviations (SDs) to provide approximate 95 % coverage. Bar graphs of parametric data processing and log transformed data processing shown. The NRI for this data set approximates 27–38 sec. (Modified from Favaloro EJ, et al. How to optimize activated partial thromboplastin time (APTT) testing: solutions to establishing and verifying normal reference intervals and assessing APTT reagents for sensitivity to heparin, lupus anticoagulant, and clotting factors. Semin Thromb Hemost 2019; 45: 22–35 [1].)
Figure 4. Ex vivo heparin versus APTT evaluation. (a) Samples from all patients identified to be on heparin (as identified by our laboratory information system) and for which an APTT was performed at the time of evaluation are also tested for anti-FXa level. The APTT therapeutic range is that corresponding to a heparin level of 0.3–0.7 U/mL by anti-Xa. However, many data points in this figure do not reflect UFH alone. Some points may instead reflect low molecular weight heparin (e.g. likely to be the sample yielding an anti-Xa value close to 0.7 U/mL but with normal APTT) or alternatively UFH co-incident to FXII deficiency or LA, or else patients potentially transitioning from UFH to VKAs. These data points can be removed to yield a ‘cleaner’ data set, as shown in (b). (Modified from Favaloro EJ, et al. How to optimize activated partial thromboplastin time (APTT) testing: solutions to establishing and verifying normal reference intervals and assessing APTT reagents for sensitivity to heparin, lupus anticoagulant, and clotting factors. Semin Thromb Hemost 2019; 45: 22–35 [1].)

Disclaimer: The views expressed in this paper are those of the authors, and are not necessarily those of NSW Health Pathology.

References
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The authors
Julianne Falconer1 MBBS and Emmanuel J. Favaloro*1,2 PhD, FFSc (RCPA)
1Haematology, Institute of Clinical Pathology and Medical Research (ICPMR), NSW Health Pathology, Westmead Hospital, NSW, Australia.
2Sydney Centres for Thrombosis and Hemostasis, Westmead Hospital

*Corresponding author
E-mail: Emmanuel.Favaloro@health.nsw.gov.au