The development and establishment of the 1st WHO BKV International Standard for nucleic acid based techniques

Biologicals 60 (2019) 75–84

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The development and establishment of the 1st WHO BKV International Standard for nucleic acid based techniques

T

Sheila Govinda,∗, Jason Hockleyb, Clare Morrisa, Neil Almonda, the Collaborative Study Group1 a

Division of Infectious Disease Diagnostics, National Institute of Biological Standards and Control, Blanche Lane, South Mimms, Potters Bar, Hertfordshire, EN6 3QG, United Kingdom b Division of Analytical and Biological Sciences, National Institute of Biological Standards and Control, Blanche Lane, South Mimms, Potters Bar, Hertfordshire, EN6 3QG, United Kingdom

ARTICLE INFO

ABSTRACT

Keywords: BK virus WHO International standard NAT Standardisation

Immunocompromised patients are at significant risk from BKV reactivation, causing allograft dysfunction or loss. Patient management relies on viral DNA monitoring, using a viral load cut-off to reduce immunosuppression. However, consistency between viral load detection assays cannot be achieved without an effective means of standardisation. We have worked with the WHO's Expert Committee on Biological Standardisation to develop suitable reference materials and undertake an international collaborative study to establish the 1st WHO International Standard for BKV detection assays. We report on the evaluation of two lyophilised candidate cell culture derived, whole virus preparations, undertaken by 33 expert laboratories. By employing the principles of biological standardisation, we show improved agreement across laboratories, demonstrating the suitability of either candidate for use as a primary order calibrant. Candidate 14/212 was established by the WHO ECBS with an assigned potency of 7.2 log10 International Units/mL intended for the calibration of BKV secondary reference materials.

1. Introduction BK Virus is a member of the Polyomaviridae family of double stranded DNA viruses. Primary infection is acquired in early childhood and in most cases is asymptomatic. Consequently, seropositivity across the adult population is approximately 90% [1,2]. Following primary infection, the virus establishes latency in the kidneys and urinary tract with intermittent reactivation throughout adulthood, where viruria is detectable in < 5% of healthy individuals [3]. The clinical sequelae of opportunistic BKV reactivation is confined to the immunocompromised state, such as in renal transplant and haematopoietic stem cell transplantation (HSCT) patients. Under immunosuppression, reactivation of latent virus may result in BKV-associated nephropathy (BKVAN) characterised by interstitial nephritis and/or urinary tract stenosis, affecting up to 10–15% of patients. This can result in allograft dysfunction or loss for up to 60% of affected kidney transplant recipients [3,4]. In HSCT patients BKV reactivation can present with haemorrhagic cystitis which is associated with

significant morbidity and mortality [5]. In 2009 the international initiative Kidney Disease: Improving Global outcomes (KDIGO) published a set of guidelines for the management of kidney transplant recipients advising periodic screening of BK virus using nucleic acid amplification tests (NAT), initially at monthly intervals for up to 6 months post-transplantation and then for two further occasions within the first-year post transplantation. A persistent BK viral load in plasma of ≥10,000 copies/mL is presumed predictive for BKVAN, whereupon a reduction in immunosuppressive medication is recommended [6]. Therein the routine application of NAT assays for BK viral load assessment is integral to patient diagnosis and management. Viral monitoring for BKV reactivation is not limited to blood (whole blood or plasma) but may also include analysis of urine, since viral reactivation is detectable in the urine several weeks before viremia and may be beneficial as an early marker for the development of BKVAN when BK viruria exceeds 7.0 log10 copies/mL [7]. Whilst BKV NAT assays are in routine use across transplantation

Corresponding author. National Institute for Biological Standards and Control, Blanche Lane, South Mimms, Potters Bar, Hertfordshire, EN6 3QG, United Kingdom. E-mail address: [email protected] (S. Govind). 1 See Acknowledgements. ∗

https://doi.org/10.1016/j.biologicals.2019.04.004 Received 7 September 2018; Received in revised form 15 April 2019; Accepted 25 April 2019 Available online 17 May 2019 1045-1056/ © 2019 Published by Elsevier Ltd on behalf of International Alliance for Biological Standardization.

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centres, data from interlaboratory proficiency assessment initiatives reveal that BKV NAT assay measurements show considerable variability between testing laboratories [8,9]. Furthermore, a single transplant centre reported discordance between viral load estimates when comparing two assays, concluding the guideline cut-off of ≥4.0 log10/mL, underestimated the diagnosis of BKVAN with one of the assays compared to the other [10]. These studies highlight the heterogeneity between assays which encompass CE-marked and analyte specific reagent (ASR) commercial assays as well as numerous laboratory developed tests (LDT) for BKV NAT detection. Each routine method in turn differs in extraction technology and extraction platform, as well as differences in amplification assay design: such as target selection, primer design and probe chemistries. Consequently, there is an urgent need for standardisation of diagnostic assays to permit meaningful cross-laboratory comparison. The World Health Organisation's (WHO) Expert Committee on Biological Standardisation (ECBS) establishes reference standards for biological substances intended for use in the prevention, treatment or diagnosis of human disease. WHO International Standards (IS) are recognised as the highest order reference materials for biological substances. Their primary purpose is intended for use by diagnostic laboratories, reference laboratories and assay manufacturers to calibrate secondary and tertiary reference materials for use in routine clinical diagnostic assays, offering traceability to a single common reference preparation regardless of methodology. Since 1999, WHO International NAT standards for blood-borne viral pathogens have been established to harmonise viral load measurements [11] and their continued use has led to great improvement in the comparability of NAT assays essential for blood safety testing. Moreover, the first WHO IS's for clinical infections [12,13] are also improving the standardisation of CMV and EBV NAT assays, despite the added need and complexity of detecting viral DNA in multiple clinical matrices. Here we report on the preparation of two BKV candidate reference materials and their evaluation in a multi-centre international collaborative study which resulted in the establishment of the 1st WHO International Standard for BKV for NAT detection assays in October 2015.

concentration of 10% FCS which was then stored at −80 °C until required for the bulk preparation. Both preparations were formulated in a universal buffer: 10 mM Tris-Cl pH 7.4, 0.5% Human serum albumin (HSA), 0.1% D-(+)-Trehalose dehydrate to contain approximately 7.0 log10 BKV copies/mL in a final volume of 5.0 L. The HSA was obtained from a licensed product that was screened for anti-HIV-1, HBsAg, and HCV RNA (Zenalb 20, BPL).

2. Materials and methods

Filling and lyophilisation was undertaken at the Centre for Biological Reference materials (CBRM) at NIBSC. The candidate formulations B and D, were designated NIBSC codes 14/202 and 14/212 respectively and were filled sequentially on 20/10/14 and 10/11/14. The filling of the bulk material was performed in a Metall and Plastic GmbH (Radolfzell, Germany) negative pressure isolator that contains the entire filling line and is interfaced with the freeze dryer (CS150 12m2, Serail, Argenteuil, France). The bulk material was kept at 18 °C throughout the filling process and stirred constantly using a magnetic stirrer. The bulk was dispensed into 5 mL screw cap glass vials in 1 mL aliquots, using a Bausch & Strobel (Ilfshofen, Germany) filling machine FVF5060. Filled vials were partially stoppered and lyophilised over a three day cycle. Post-lyophilisation the candidate standards were examined for precision of the fill, appearance, residual moisture content and oxygen head space. Further details are reported in WHO ECBS report [16]. The sealed vials are stored at −20 °C at NIBSC under continuous temperature monitoring for the lifetime of the product.

2.2. Determination of viral stock potency and Sanger sequence analysis Nucleic acid extractions were performed using the QIAamp Viral RNA mini Kit according to the manufacturer's instructions (QIAGEN, Germany) using the QIAcube (QIAGEN, Germany) with the inclusion of an internal control to act as a positive control for DNA extraction from non-cellular biological samples (ELITechGroup, Torino, Italy). 20 μl of purified nucleic acid sample was amplified by qPCR using probe-based quantification according to manufacturer instructions on the ABI 7900HT Real-time PCR instrument (Applied Biosystems, California USA). An estimate of viral quantification was achieved by the inclusion of four plasmid-based quantification standards in the amplification reaction, to generate a standard curve with a dynamic range of 102-105 quantifiable copies/mL (ELITechGroup, Torino, Italy). The complete viral genomes of both candidates were sequenced at NIBSC using Sanger sequencing and a base pair comparison across the VP1 region was used to verify the genotype. Briefly, PCR fragments were amplified on the Veriti 96 well thermal cycler (Applied Biosystems) using Platinum® Taq DNA Polymerase (Life Technologies) and sequencing reactions were performed using purified PCR amplicons on the 3130 XL Genetic Analyzer (Applied Biosystems). Candidate B was identified to be BKV1b-1 subtype, using base pair comparison within the VP1 region, showing a 98.9% sequence identity with GenBank Accession AB301095 and Candidate D showed 98.4% sequence identity to GenBank Accession AB301093 BKV1b-2 subtype. The subtype classification places 1b-1 to be most prevalent in SouthEast Asian populations and 1b-2 to show greatest geographical prevalence in Europe and USA [15]. 2.3. Filling and lyophilisation process

2.1. Preparation of candidate standards Two candidate formulations were prepared; candidate one (also referred to herein as Sample B) was donated by Dr JL Murk (UMC Utrecht, Netherlands) as a high titre ∼9.0 log10 copies/mL cell culture supernatant propagated in diploid human lung fibroblast MRC-5 cells. The viral isolate used for propagation was obtained from a urine sample of a bone marrow transplant recipient diagnosed with BK-cystitis. Candidate two (also referred to herein as Sample D) was propagated inhouse from an isolate donated to NIBSC by Wendy Knowles (Personal communication Pam Pipkin), from the Gardner laboratory where the original BKV Gardner isolate was discovered in 1971 [14]. Briefly, for candidate two dilution of a high titre cell culture supernatant was used to inoculate MRC-5 cells suspended in 1 mL of cell culture medium; cells were left for 1 h for infection, then briefly centrifuged at low speed to pellet the cells and remove the residual inoculum. Cells were maintained at 37 °C, 5% CO2 in MEM (Sigma Cat# G8644), 10% FCS, 2% 200 mM L-Glutamine (Sigma Cat# G7513), and 1.5% 1M HEPES (Sigma Cat# H0887). Cell culture supernatant was periodically tested for viral propagation using quantitative PCR to track increase in BK viral load using a CE-marked in-vitro diagnostic (IVD) assay BKV ELITE MGB kit (ELITech, Torino, Italy). The cell culture supernatant was harvested when maximal CPE was visible, approximately after 4 weeks and cleared from cellular debris by centrifugation for 15 min at 150 g. FCS was added to the cleared BKV viral supernatant to obtain a final

2.4. Stability assessment: accelerated thermal degradation To determine the stability of the candidate standards post lyophilisation, 30 vials of each candidate were taken at random and placed in monitored storage temperatures of −70 °C, +4 °C, +20 °C, +37 °C and +45 °C to perform periodic accelerated degradation assessments for up to 10 years or for the life time of the product. Three vials are removed from each temperature and tested for viral potency using the qPCR method described, to provide an indication of stability relative to the 76

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Table 1a Production data of lyophilised BKV preparations. NIBSC code

BKV WHO IS 14/212 (Sample D)

Candidate IS 14/202 (Sample B)

Product content Mean fill weight (g) CV of fill weight (%) Mean residual moisture (%) CV of residual moisture (%) Mean of Oxygen content (%) CV of Oxygen content (%) Viral potency Log10 copies/mL CV of potency (%) SD

BK Virus genotype 1b-2 1.0075 0.27 (n = 142) 0.91 (n = 12) 8.6 0.75 (n = 12) 14.33 6.50 (n = 12) (Range 6.49–6.56) 1.6 0.105

BK Virus genotype 1b-1 1.0074 0.23 (n = 142) 0.66 (n = 12) 12.3 0.82 (n = 12) 8.65 6.53 (n = 12) (Range 6.43–6.82) 4.4 0.029

storage temperature of −20°C. Three independent tests were performed in the first year and annually thereafter.

sample dispatch, to ascertain the types of clinical samples and quantity of sample routinely assayed in their laboratory. Sample sets were thereby customised for each participant based on the responses received. All participants received the candidate BKV IS materials in both the lyophilised and liquid state (Samples B-E) and a plasmid construct (Sample A), encoding BKV 1a sequence (Genbank ID EU681733). The construct was prepared using viral DNA extracted and amplified (missing a small proportion of the non-coding regulatory region) from a clinical urine specimen which was then cloned into a pACY477 backbone (donated by Dr P Vallone at National Institute of Standards and Technology, USA). Depending on the routine test practices of each laboratory, samples obtained from a Bone marrow transplant (BMT) patient; Sample F urine and Sample G plasma, were also included in the laboratory test panels (donated by Dr CB Christiansen of Rigshospitalet Department Clinical Microbiology, Copenhagen, Denmark). Each anonymised patient sample was diluted into negative urine and plasma respectively, to ensure sufficient volume for participants to perform duplicate tests across 3 independent experimental runs using extraction and amplification methods commonly performed in each laboratory. Upon receipt participants were directed to store samples C, E, F and G at −70 °C and samples A, B and D at −20 °C. Participants were directed to reconstitute the lyophilised sample (B and D) in 1 mL of nuclease-free molecular grade water for a minimum of 20 min with occasional agitation before use. Participants performing quantitative analyses were directed to test samples B and D undiluted and then at a minimum of 3–4 serial ten-fold dilutions in a single sample matrix commonly used in their laboratory, such that at least 1 of these dilutions should fall into the linear range of their quantitative assay. For subsequent runs participants were requested to test a minimum of two serial dilutions of samples B and D that fell within the linear range of their assay. All participants reported their quantitative estimates in copies/mL. Participants performing qualitative analysis were requested for the first assay, to test an undiluted and then an additional minimum of several serial 10-fold dilutions of Samples B and D in a single sample matrix commonly used in their laboratory, in order to determine end point detection. Participants were requested to ensure their data included at least 2 dilution points at which an amplification product was no longer detectable. For the remaining qualitative assays, participants were requested to re-test the dilutions around the assay end point as determined in the first assay and requested to include a minimum of two half-log serial dilutions either side of the determined end point dilution. Participants performing qualitative analysis reported their findings as either negative or positive, which were used to calculate sample potencies that are herein referred to as NAT detectable units. Of the 36 recruited participants, 33 laboratory datasets were returned and used for analysis. Where laboratories submitted more than one dataset they were referenced with their numerical code and the addition of the letter a and b to signify multiple datasets. There was considerable variability in the methods employed by the participants; full details are provided in Supplementary Table 4. Except for one

2.5. Collaborative study design Thirty six collaborative study participants were recruited. Participants from research and clinical laboratories were selected based on peer reviewed publications on BKV NAT detection assays. Manufacturers of BKV NAT IVD kits were also included as well as reference and External quality assurance (EQA) laboratories. Each laboratory was randomly assigned a numerical code by which to reference their data, assuring laboratory anonymity. A total of 7 study samples coded A-G (Table 2) were prepared for evaluation. Participating laboratories were sent a questionnaire prior to Table 1b Thermal stability data of lyophilised BKV preparations (14/212) and (14/202) at different storage temperatures. Temperature

BKV WHO IS 14/212 mean log10 copies/mL (Difference in Mean log10 copies/mL from -20⁰C baseline)

Minus 70 °C Minus 20⁰C Plus 4 °C Plus 20 °C Plus 37 °C Plus 45 °C

3 months

6 months

12 months

24 months

6.63 −0.04 6.67 6.67 0.00 6.65 −0.02 6.80 0.13 6.81 0.14

6.52 −0.03 6.49 6.49 0.00 6.50 0.01 6.70 0.21 6.69 0.20

6.61 0.06 6.55 6.65 0.10 6.65 0.10 6.78 0.23 6.71 0.16

6.55 0.03 6.52 6.55 0.03 6.57 0.05 6.60 0.08 6.55 0.03

Candidate B 14/202 mean log10 copies/mL (Difference in log10 copies/mL from -20⁰C baseline) Temperature

3 months

6 months

12 months

24 months

Minus 70 °C

6.56 −0.01 6.57 6.64 0.07 6.59 0.02 6.64 0.07 6.63 0.06

6.39 0.01 6.40 6.49 0.09 6.49 0.09 6.46 0.06 6.46 0.06

6.49 −0.04 6.53 6.54 0.01 6.59 0.06 6.59 0.06 6.52 −0.01

6.32 0.00 6.32 6.31 −0.01 6.32 0.00 6.35 0.03 6.28 −0.05

Minus 20⁰C Plus 4 °C Plus 20 °C Plus 37 °C Plus 45 °C

Mean results presented from single extractions from 3 independent vials (combinations of neat, 1 in 10 and 1 in 100 dilution) at each time-point and temperature, each amplified in duplicate. 77

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Table 2 Collaborative study overall mean estimates and inter-laboratory variation (log10 copies/mL for quantitative or log10 NAT-detectable units/mL for qualitative assays). Sample

ID

Assay

n

Mean

SD

GCV

Min

Max

Log10 difference

A B

NIST BKV1a Plasmid Lyophilised Candidate IS 14/202

C D

Liquid bulk 14/202 Lyophilised Candidate IS 14/212

E F G

Liquid bulk 14/212 Urine BMT patient Plasma BMT patient

Quantitative Qualitative Quantitative Combined Quantitative Qualitative Quantitative Combined Quantitative Quantitative Quantitative

34 3 35 38 34 3 35 38 35 28 28

8.97 4.56 6.62 6.46 6.71 4.67 7.17 6.97 7.21 2.67 2.99

0.85 0.4 0.65 0.84 0.77 1.33 0.61 0.95 0.71 0.88 0.72

607% 152% 344% 596% 494% 2034% 306% 787% 415% 660% 427%

7.01 4.12 4.38 4.12 3.62 3.6 5.69 3.60 4.92 1 1.24

11.24 4.9 7.53 7.53 8.05 6.19 8.33 8.33 8.35 4.82 4.6

4.23 0.78 3.15 3.59 4.43 2.59 2.64 4.73 3.43 3.82 3.36

Excluding laboratory 13a.

laboratory, all participants used commercially available nucleic acid extraction kits, where the majority of the extraction protocols were performed on automated extraction platforms 28/39 (72%). 15 datasets (39%) were obtained using laboratory developed amplification assays, compared with 24 datasets obtained using commercially available amplification kits (61%) or analyte-specific reagents (ASR).

moisture (0.66% and 0.91%), and the mean residual oxygen content (0.82% and 0.75%) for 14/202 and 14/212 respectively. The CVs of the fill mass and mean residual moisture were within acceptable limits for WHO International Standard preparations [17] and the residual oxygen content was well within the NIBSC working limit of 1.1% (Table 1a). Further stability evaluation of the candidate materials is still under continual assessment. Currently four accelerated degradation time points have been assessed at 3, 6, 12 and 24 months post production. Following reconstitution of the lyophilised sample further dilutions were made in universal buffer to ensure at least one of the estimates fell within the linear range (2.0–5.0 log10 copies) of the quantitative assay. The mean estimated log10 copies/mL for three vials of BKV WHO IS stored at each temperature for each time point were calculated. Estimates from vials stored at a temperature other than the storage temperature of −20 °C were then subtracted from this baseline estimate to assess any loss in potency. The differences (log10 ‘copies/mL’) from the −20 °C baseline sample are presented in Table 1b. To date there is no consistent detectable loss in potency at each elevated storage temperature compared with the −20 °C baseline samples.

2.6. Statistical analysis Qualitative and quantitative assay results were evaluated separately. In the case of qualitative assays, for each laboratory and assay method, data from all assays were pooled to give a number of positive outcomes from the total number tested at each dilution step. A single ‘end-point’ for each dilution series was calculated, to give an estimate of NAT detectable units/mL, as described previously [11–13]. For quantitative assays, for each assay run, a single estimate of log10 copies/mL was obtained for each sample, by taking the mean of the log10 estimates of copies/mL across replicates, after correcting for any dilution factor. A single estimate for the laboratory and assay method was then calculated as the mean of the log10 estimates of copies/mL across assay runs. Overall analysis was based on the log10 estimates of copies/mL or log10 NAT detectable units/mL. Overall mean estimates were calculated from the means of all individual laboratories. Variation between laboratories (inter-laboratory) was expressed as the SD of the log10 estimates and percentage geometric coefficient of variation (%GCV) of the actual estimates. The ability of a candidate standard to reduce the inter-laboratory variability in BKV viral load measurements was assessed by calculating the potency of the candidate relative to the study sample. The relative potency of, for example, sample E relative to sample D was calculated for each individual assay by adding the difference in estimated log10 ‘units per mL’ (test sample – candidate standard) to the assigned log10 IU/mL of the WHO BKV IS (7.2 log10 IU/mL). The relative potency calculation using either sample B or A was calculated as above by using the overall quantitative mean estimate for either sample (6.62 and 8.97 respectively) as a nominal IU value.

3.2. Collaborative study data A coded seven-member panel was assembled for analysis in the multi-centre international collaborative study which included two lyophilised candidate IS preparations (sample B and D) and their liquid equivalents (samples C and E respectively), a plasmid construct (A) and two clinical samples (F and G) to provide a preliminary assessment of commutability (Table 2). Evaluation of the panel was completed by 33 participants, from 16 different countries. For the candidate materials sample B and D, 35 quantitative datasets and 3 qualitative datasets were analysed. For all other samples, only quantitative estimates were evaluated (Table 2). In general, participants performed their experimental runs using one assay method, with the use of one matrix type for the dilution of Sample B and D performing duplicate extractions of each sample over three independent runs. The results returned by the participants for each sample of the panel are depicted in Fig. 1, illustrating the spread of data. This variation of reported quantitative estimates across laboratories ranged between 2.64 and 4.43 log10 copies/mL irrespective of whether plasmid or whole viral template, supplied as lyophilised or frozen liquid, were measured (Table 2). Interestingly the mean estimates obtained by limiting dilution using qualitative assays were markedly lower than those obtained by quantitative assays, with an Δ log10 detectable units/ mL of 2.06 and 2.5 for Sample B and D respectively (Table 2). Paradoxically assay reproducibility within laboratories was overall very robust. Individual sample mean estimates per participant and associated intra-laboratory SD values are provided in Supplementary Tables 1 and 2

3. Results 3.1. Homogeneity and stability of lyophilised candidate materials Twelve vials of each candidate preparation were evaluated for BKV DNA potency using a commercial NAT detection assay to confirm the homogeneity of each fill. The mean viral potency was estimated at NIBSC as 6.53 and 6.50 log10 copies/mL for candidate 14/202 and 14/ 212 respectively (Table 1a). A total of 140 ampoules were weighed for 14/202 and 142 for 14/212, approximately 3% of the total vials filled per candidate, to determine the mean fill weight which were 1.0074 g and 1.0075 g for 14/202 and 14/212 respectively (Table 1a). A further 12 vials of each candidate were used to confirm the mean residual 78

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Fig. 1. Quantitative and qualitative laboratory mean estimates of collaborative study samples. Column scatter plot of mean BKV viral load estimates returned by each laboratory for samples A-G, presented with 95% CI for the mean. Quantitative assays (open circles) are shown as log10 units of copies/mL and qualitative assays (half-shaded circles) are shown as log10 units of NAT detectable units/mL for Samples B and D.

3.3. Relative potency assessment

were compared between commercial assays and LDTs P value < 0.01 (Fig. 3b). To assess whether the reported potency estimates for sample D and B varied depending on the amplification target region within the BK viral genome, a one-way analysis of variance between the groups with more than 3 data points (VP1, LTAg and stAg) was performed. For sample D the difference between the mean of the estimates was not statistically significant, irrespective of the target region amplified (Fig. 4a). Similarly, for sample B no significant difference was observed between the mean of the estimates reported for the different target regions across the BK genome (Fig. 4b). However, a significant difference in the standard deviation of the estimates was observed (p = 0.03). A significant difference in the standard deviation of estimates was also detected with data obtained for Sample A, F and G (data not shown) where greater SD values for VP1 estimates were observed compared with either LTAg or stAg targeted assays.

To determine whether the variability in quantitative estimates across laboratories reported for each sample could be improved through calibration to a common reference material, we calculated the relative potency using either of the BKV candidates (B and D) to assess their suitability for calibration purposes (Fig. 2a–d). For sample E, when expressed as a relative potency to Sample D, the range in viral load estimates was reduced to 5.35–7.87 IU/mL (Table 3) compared with 4.92–8.35 (Table 2) when expressed in copies/mL. Using Sample B as a calibrator we also observed a similar reduction in variability (Table 3) (Fig. 2a). When expressing Sample C estimates as a relative potency to Sample D the range in viral load estimates was reduced to 5.13–7.73 IU/mL compared with 3.62–8.05 copies/mL, a reduction in SD to 0.54 from 0.77. Furthermore, using Sample B as a calibrator, demonstrated a greater reduction in variability (5.08–7.33 nominal IU/mL) (Table 3) (Fig. 2b). Since the intended purpose of the IS material is to harmonise viral load data from clinical samples, a similar assessment was performed on Sample F and Sample G. The variability in estimates of relative potency for each sample were reduced by 0.25 and 0.26 log10 respectively using Sample D as the calibrator. Similarly using Sample B as the calibrator, the variability in estimates of relative potency for sample F was reduced by 0.25 and for Sample G by 0.33 log10 (Table 3) (Fig. 2c–d). By contrast when the same calibration was performed using sample A (plasmid construct) as the reference for either sample E or C, no reduction in variance was observed for E and only a marginal reduction was observed for Sample C (0.1 log10) (Fig. 2a–b). Likewise, for Sample F and G only a modest reduction in variance of the potency estimates was observed (0.15 and 0.1 log10 respectively) compared with either sample D or B (Table 3) (Fig. 2c–d).

3.5. Assessment of diluent effects on reported estimates Since the proposed IS preparation is intended for use with multiple diluents, we assessed whether the quantitative estimates were skewed in a diluent dependent manner. Because the majority of participants used either plasma (n = 24) or urine (n = 14) as diluents, we plotted the quantitative laboratory mean estimates obtained for samples D and B and a t-test was performed to compare the mean estimates obtained. For sample D a significant difference between the mean log10 copies/ mL estimate obtained using plasma as a diluent compared with urine was observed (p = 0.024) (Fig. 5a) suggesting a possible matrix effect. However, no significant difference between the mean estimates was obtained for sample B when diluted in either plasma or urine (Fig. 5b). To investigate this further we compared data from a single laboratory that used three diluents in their evaluations. A comparison of the estimates reported using each matrix revealed a consistent and significant skewing of the data, where higher mean estimates were reported using plasma, compared with either whole blood or urine. For sample D the mean estimate of log10 copies/mL in plasma was 7.15, compared with 6.52 in whole blood and 6.24 in urine (Fig. 6a). For sample B the mean estimate in plasma was 6.85, compared with 6.31 in whole blood and 5.99 in urine (Fig. 6b). A one-way ANOVA analysis returned a significance value of p < 0.001 for both sample D and B. Two other laboratories also performed their analysis using multiple diluents for Sample B and D, however with fewer data points the significance of any differences could not be determined.

3.4. Assessment of methodological differences on reported estimates of the WHO IS Since differences in methodological practice influence viral load measurements, we next investigated whether differences in extraction or amplification methods influenced the consistency of data between laboratories. There was a greater consistency in results generated by automated extraction and/or using commercial amplification assays (Fig. 3a). However these different approaches did not alter the estimation of the mean viral load significantly, although a significant difference in variance (F-test value) was observed when performances 79

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Fig. 2. a–d: Harmonisation of BK virus assay estimates through the use of the WHO International standard. Plots depict direct estimates of each sample E (2a), C (2b), F (2c) and G (2d) in log10 copies/mL and then relative potency calculations in log10 IU/mL shown alongside using the WHO IS (Sample D). Relative potency calculations using a nominal log10 IU/mL value are also depicted using the candidate standard (B) and the synthetic reference preparation (A). The individual laboratory estimates of log10 potency estimates, calculated relative to each calibrator are provided in supplementary Table 3 for quantitative assays performed.

manage disease. However, the absence of a primary calibrant has contributed to the wide variability in copy number estimates. Since the establishment of the 1st WHO IS for CMV and EBV in 2009 for use as primary biological calibrants, numerous reports describing their utility by clinical laboratories have been invaluable in uncovering the complexities of standardising NAT assays for these clinical infections [18–21]. Our current study sought to evaluate the utility of candidate materials for the calibration of BKV NAT assays and secondly to establish a potency value for unitage assignment. Our multicentre international collaborative study included 33 laboratories which provided a wide representation of diagnostic end-users, and a breadth of assay methodologies for the evaluation of the candidate materials. This study exemplified the current need for standardisation because whilst the intra-laboratory variability of results demonstrated excellent assay reproducibility within each laboratory, high inter-laboratory variation highlighted the need for a common calibrator shared between laboratories. These observations concur with those of the College of American Pathologists proficiency study [9] and the BKV French study [8] that report variation of more than 3.5 log10 copies/mL in estimates of a common clinical sample. Similar variation in submitted estimates were observed for samples included in this study. In some respects, the variability between estimates of Sample A, the plasmid sample, is intriguing given that fewer steps were required to evaluate this purified DNA sample. However given the construct encodes a genotype 1a sequence which, is most prevalent in Africa, the possibility of some sequence mismatch between some assay primers and probes may have contributed to the variability [22,23]. Sanger

Table 3 Study sample overall mean log10 potencies relative to the BKV WHO IS in IU/ mL (or nominal IU/mL-relative to B or A) for quantitative assays. Sample

Relative to

n

Mean

SD

GCV

Min

Max

C C C

D B A

34 34 33

6.73 6.69 6.76

0.54 0.39 0.67

248% 146% 364%

5.13 5.08 5.00

7.73 7.33 8.62

E E E

D B A

35 35 34

7.24 7.19 7.24

0.48 0.48 0.72

199% 205% 428%

5.35 5.00 4.93

7.87 8.32 8.81

F F F

D B A

28 28 28

2.74 2.64 2.69

0.63 0.63 0.73

327% 323% 432%

1.12 1.46 1.41

4.28 3.88 4.28

G G G

D B A

28 28 27

2.96 2.95 3.14

0.46 0.39 0.62

187% 147% 313%

1.80 2.08 2.23

3.93 3.72 5.08

Data excluding laboratory 13a.

4. Discussion Opportunistic reactivation of latent viruses following alterations in the immune status of immunocompromised hosts, present clinicians with challenges in the management of transplant recipients. Detecting the reactivation of BK virus in patients relies on NAT quantification, using a universal viral load cut-off which has been recommended to 80

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collaborative study (see Fig. 2a–d). These findings concur with the observation from the collaborative study to establish the 1st WHO IS for CMV DNA [12]. Even though the plasmid construct included in that study was genetically identical with the cell culture derived candidate materials, the plasmid failed to act as an effective reference standard. It is proposed that the limited value of plasmid nucleic acid to act as a reference standard for harmonising data from clinical materials arises from its inability to control for the extraction processes as well as the amplification step. The inclusion of the two clinical samples in the evaluation panel provided an early indication of the reference standards’ ability to perform in a manner that would enable it to effectively harmonise data generated from clinical samples. Whilst these data indicate that the candidate International Standard is suitable for this purpose, a more extensive assessment is required to determine whether these materials are fully commutable. These types of studies are not straightforward as they require large volumes of clinical materials which is not always possible to acquire. Furthermore, commutability can only be demonstrated for the assays in which both the clinical samples and the reference have been tested and is subject to considerable bias depending on multiple variables including the statistical methodology employed [24]. For many analytes this is very challenging and alternative approaches may be required. For example, through the inclusion of the WHO International standard in external quality assurance panels in a suitable format and dilution. The design of the BK candidate materials in an “universal buffer” is common to that of other recent International Standards which are designed to harmonise diagnostic assays in which a test sample may be supplied in several different clinical matrices. At the same time, it provides an opportunity for laboratories to investigate the impact of extraction from different clinical matrices on the efficiency of detection and quantification of different assays. In this collaborative study a significantly lower mean estimate was observed for the WHO BKV IS across laboratories when urine was used as the matrix compared with plasma. However, further commutability studies are required to investigate this. Ninety years of biological standardisation has led the WHO ECBS to develop robust guidelines for materials which are to be considered for establishment as International Standards. Since these standards are to be designed to last at least 10 years, then sufficient starting material must be sourced at an appropriate titre to allow a calibration curve to be prepared with a 3–4 log10 point dilution range. As a result, it was decided to use cell culture propagated BK virus as the starting material. It has been reported that genomic sequence variability across the BKV subtypes can impact viral quantitation particularly with BK genotypes III and IV, leading to under quantification in patients [22]. Both IS candidates described are genotype 1 sub-type b, selected since genotype I is predominant in most geographical regions with a prevalence of 46–82% throughout the world. Furthermore, since only one candidate can be established as a WHO IS, further evaluation of alternate genotypes was not considered. Therefore, primers targeting the variable VP1 region should be avoided to prevent under quantification of relatively uncommon genotypes. Based on the collaborative study data, both candidates demonstrated suitability as effective biological standards, performing equally in harmonising agreement of sample E and F. Whilst candidate B marginally out-performed candidate D in the harmonisation of sample C and G, based on the subtype determination, Sample D (subtype 1b-2) was thought to provide a more representative genotype for the majority of diagnostic assays and was therefore subsequently recommended as the optimal candidate material. The WHO Expert Committee for Biological Standardisation reviewed these data and established sample D as the 1st International Standards for BK virus DNA at a unitage of 7.2 log10 IU/mL. This unit was calculated by rounding up the mean of the quantitative datasets only, omitting the minority representation of qualitative estimates, because of the primary clinical requirement to

Fig. 3. The impact of assay methodology on potency estimates for WHO IS. Overall quantitative mean log10 copies/mL estimates are presented with 95% CI for the mean, comparing (a) automated (Mean 7.14, SD 0.57, Min-Max 5.69–8.33, n = 27) vs manual (Mean 7.26, SD 0.72, Min-Max 5.84–8.19, n = 9) extraction methods and (b) commercial amplification assays (Mean 7.19, SD 0.43, Min-Max 6.63–8.33, n = 24) vs laboratory developed tests (LDT) (Mean 7.13, SD 0.87, Min-Max 5.69–8.19, n = 12).

sequence analyses of the candidate IS materials samples B and D have identified them to be genotypes BKV 1b-1 and 1b-2 respectively. Nevertheless, this is unlikely to account for the broad inter-laboratory variability in initial estimates. Intriguingly dissection of the collaborative study data identified the key factors associated with high variability were the application of qualitative assays relying on an end-point dilution analyses and in particular quantitative assays targeting the VP1 region. Despite clear initial differences in the performance and or calibration of different assays, it was possible to use the candidate materials as a reference to calculate the relative potency of liquid preparations of viruses supplied to participants of the collaborative study. This process harmonised the data from laboratories using a broad range of assays. This was observed even when the candidate standards were used to assess the relative potencies of clinical materials. Greater concordance of data was obtained for the plasma sample compared with the urine sample. This difference was not a statistical artefact as the same number of quantitative estimates was obtained for both samples and the viral loads in both samples were comparable. Further work is required to determine whether either the sequence of the clinical sample or the matrix itself may account for the difference in performance. It is widely believed that a plasmid provides a useful reference sample for assay calibration. As a result, one of the samples included in this study was a plasmid derived from a BKV1a genotype isolate. Using this sample as a reference standard resulted in only minimal improvement in concordance of the mean estimates of samples in the 81

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Fig. 4. The impact of target sequence on potency estimates (a) BKV IS Sample D (b) candidate IS Sample B. Overall quantitative mean estimates presented with 95% CI for the mean separated according to laboratory target designation.

quantify BK viral load. As a primary standard, the assigned unitage does not carry an uncertainty associated with its calibration. The uncertainty may therefore be considered, to be the variance of the vial content. Whilst the assigned potency is taken from the reported copy number

estimates, any conversion to copies/mL must be empirically determined by each individual laboratory method following the recommendations provided in the WHO guidance document on the calibration of secondary references materials [25]. 82

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Fig. 5. The impact of clinical matrix on the detection and quantification of BKV. Overall quantitative mean log10 copies/mL estimates presented with 95% CI for the mean for (a) Sample D BKV IS and (b) Sample B candidate IS diluted in either plasma (n = 21) or urine (n = 13) across all laboratories.

Fig. 6. Intra-laboratory assessment of impact of clinical matrix on BKV quantification. Overall quantitative mean estimates presented with 95% CI for the mean for (a) Sample D BKV IS and (b) Sample B candidate IS diluted in either whole blood (n = 4), plasma (n = 4) or urine (n = 4). Each data point per matrix was obtained using the same extraction platform and amplification method.

External quality assurance reports as well as this study have demonstrated that quantitative diagnostic NAT assays vary between laboratories. A recent appraisal by Hayden et al. reports on the merits of WHO international quantitative standards and in particular the 1st IS for EBV DNA for improving concordance of reported data between laboratories, through the adoption of relative potencies measured in international units compared with copy number estimates [26]. Laboratories detecting, and quantifying BK viral load may wish to consider the implementation of the same principles and measure BK viral DNA in international units in order to provide a more reliable approach for establishing and comparing treatment thresholds and strategies for clinical management. Since its establishment in 2015, next generation sequence analysis of the 1st IS for BK virus DNA has revealed that the cell culture derived material is heterogeneous, containing a sub-population of virus with deletions particularly across the Large T antigen [27]. We have complementary sequence data that concurs with this report (Manuscript in preparation). Prior to the establishment of this material we saw little evidence of this discrepancy, possibly obscured due to the relatively small dataset available for detailed comparisons of viral load estimates across the various target regions, as well as the inherent variability when comparing data from non-standardised multi-centre assays. We have since conducted further analyses in collaboration with the National Institute of Standards and Technology, to assess the impact on viral load detection comparing both digital droplet PCR and qPCR. Using a single laboratory approach a significant difference in quantification was observed when targeting the LTAg region compared with the VP2 region (Manuscript in preparation). Consequently, the calibration of secondary reference materials to the WHO IS would be prudently performed outside of the LTAg region. Nevertheless, the data presented here demonstrate that even an imperfect whole virus-based reference,

harmonised the participant data far better than a sequence perfect plasmid standard. Therefore, caution is needed in the face of recommendations that synthetic nucleic acid materials provide superior standards [28]. To conclude we have developed a stable reference material that has been evaluated in a multi-centre collaborative study that demonstrated the utility of the 1st WHO BKV IS as a common calibrant for diagnostic BKV qPCR assays, exemplified by improved agreement across laboratories. The use and evaluation of this material by the wider clinical community offers the potential for improving inter-assay comparability [29], which in turn should lead to improvements in the consistency of patient management. Acknowledgements We gratefully acknowledge the important contributions of the collaborative study participants: Dr Seweryn Bialassiewicz, Sir Albert Sakzewski Virus Research Centre, Australia; Dr Marijke Reynders, AZ Sint-Jan, Belgium; Jaclyn Ugulini, Norgen Biotek Corp, Canada; Martina Salakova, Institute of Haematology and Blood Transfusion, Czech Republic; Dr Jana Zdychova, PLM-OKI/IKEM, Czech Republic; Dr Claus Bohn Christiansen M.D, Rigshospitalet, Denmark; Dr David Boutolleau, University Hospital Pitie-Salpetriere, France; Dr Céline Bressollette and Dr Marina Illiaquer, University Hospital Nantes, France; Dr Samira Fafi-Kremer, University Hospital Strasbourg , France; Dr Catherine Mengelle and Dr Jean-Michel Mansuy, Federative Institute of Biology, France; Dr Matthieu Vignoles BioMerieux, France; Dr Karin Rottengatter and Dr Waldemar Fischer, Altona Diagnostics GmbH, Germany; Dr Steffi Silling, National Reference Center for Papilloma83

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and Polyomaviruses, Germany; Dr Rajesh Kannangai and Asha Mary Abraham, Christian Medical College, India; Dr Mauro G. Tognon, University of Ferrara, Italy; Dr Christiana Olivo, ELITtech Group SpA, Italy; Dr L. Rurenga-Gard and Prof. H.G.M Niesters, University Medical Center Groningen, Netherlands; Dr Rob Schuurman, Univeristy Medical Center Utrecht, The Netherlands; Dr Juan E. Echevarría, National Center of Microbiology, Institute of Health Carlos III, Spain; Dr Mounir Trimeche, CHU Farhat Hached of Sousse, Tunisia; Prof. Dr. Dilek Colak, Akdeniz University Hospital Central Microbiology Laboratory, Turkey; Dr Elif Akyuz, Anatolia Geneworks, Turkey; Dr Elaine McCulloch, QCMD, United Kingdom; Dr Anna Blacha, QIAGEN Manchester Ltd, United Kingdom; Dr Kathleen Stellrecht and Mr Shafiq Butt, Albany Medical Center, USA; Dr Kristin S Lowery, AthoGen/Ibis Biosciences, USA; Dr Midori Mitui and Dr Damon Lacey, Children's Medical Centre, USA; Dr Mayur S Ramesh, Henry Ford Hospital, USA; Dr Jianli Dong, University of Texas Medical Branch, U.S.A; Dr Catherine Jakubowski, Luminex Corporation, USA; Dr Parmjeet Randhawa, UPMC-Montefiore Hospital, USA; Dr Angela Caliendo, The Miriam Hospital, USA. For the provision of materials used for the preparation of the candidate standard (B) we thank Dr JL Murk, UMC Utrecht, The Netherlands. We also thanks Dr P Vallone, National Institute of Standards and Technology, USA for BKV plasmid and Dr CB Christiansen, Rigshospitalet Department Clinical Microbiology, Denmark for the provision of clinical samples. The work of the staff in the Centre for Biological Reference Materials (CBRM) located at NIBSC is also gratefully acknowledged.

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Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.biologicals.2019.04.004. References [1] Egli A, et al. Prevalence of polyomavirus BK and JC infection and replication in 400 healthy blood donors. J Infect Dis 2009;199(6):837–46. [2] Knowles WA, et al. Population-based study of antibody to the human polyomaviruses BKV and JCV and the simian polyomavirus SV40. J Med Virol 2003;71(1):115–23. [3] Hirsch HH. BK virus: opportunity makes a pathogen. Clin Infect Dis 2005;41(3):354–60. [4] Babel N, Volk HD, Reinke P. BK polyomavirus infection and nephropathy: the virusimmune system interplay. Nat Rev Nephrol 2011;7(7):399–406. [5] Dropulic LK, Jones RJ. Polyomavirus BK infection in blood and marrow transplant recipients. Bone Marrow Transplant 2008;41(1):11–8. [6] Kidney Disease: Improving Global Outcomes Transplant Work G. KDIGO clinical practice guideline for the care of kidney transplant recipients. Am J Transplant 2009;9(Suppl 3):S1–155. [7] Randhawa P, et al. Correlates of quantitative measurement of BK polyomavirus (BKV) DNA with clinical course of BKV infection in renal transplant patients. J Clin Microbiol 2004;42(3):1176–80. [8] Solis M, et al. Sequence variation in amplification target genes and standards influences interlaboratory comparison of BK virus DNA load measurement. J Clin Microbiol 2015;53(12):3842–52.

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