This review is an integral part of the special issue for the 60 years of the journal Clinical Chemistry and Laboratory Medicine (CCLM). The aim of the review is to highlight the role of the clinical laboratory since the emergence of the “severe acute respiratory syndrome coronavirus 2” (SARS-CoV-2), which causes Coronavirus disease 2019 (COVID-19), with special focus on the contribution of the journal in generating knowledge in SARS-CoV-2 diagnosis. As of October 30, 2022, a total of 186 CCLM publications were dedicated to COVID-19. Of importance, major International Federation of Clinical Chemistry (IFCC) guidelines related to the diagnosis of COVID-19 were published in CCLM. Between early-2020 and late October 2022, COVID-19 publications represented around 27% of all articles in CCLM, highlighting the willingness of the editorial board to help the field in order to better describe and diagnose this new emerging disease. First launched in 1963 under the name “Zeitschrift für Klinische Chemie”, the Journal was entirely devoted to clinical chemistry in the strict sense. The various topics published in relation to COVID-19 including its diagnosis, its impact on biochemical or hematological measures, as well as biosafety measures, is the perfect example that shows that the journal has greatly diversified over time.
Coronavirus disease 2019 (COVID-19) is an infectious disease caused by a coronavirus, identified as “severe acute respiratory syndrome coronavirus 2” (SARS-CoV-2) . The disease emerged in Wuhan, China, in December 2019 and rapidly became a pandemic. As of November 14, 2022, there have been more than 631 million confirmed cases of COVID-19 and nearly 6.6 million deaths around the globe . In order to control the pandemic, major efforts have been made to produce and clinically validate new COVID-19 vaccines at an unprecedented speed. As of November 14, 2022, nearly 13 billon COVID-19 vaccine doses have been administered worldwide , 49 vaccines are approved and 239 are currently in clinical development or awaiting clinical validation .
This article is an integral part of the special issue for the 60 years of the journal Clinical Chemistry and Laboratory Medicine (CCLM). The journal was launched in 1963 under the name “Zeitschrift für Klinische Chemie” and was entirely devoted to clinical chemistry in the strict sense. COVID-19 permits to appreciate the large diversity of fields that are now addressed in CCLM, demonstrating how the journal has considerably diversified its interests over time. This article is also an opportunity to look back at the achievements of CCLM since the emergence of SARS-CoV-2 in 2020.
The aim of the review is to highlight the role of the clinical laboratory since the emergence of SARS-CoV-2, with a special interest regarding the contribution of CCLM in generating knowledge in COVID-19 diagnosis and clinical monitoring. The first CCLM article about COVID-19 was published online on the March 3rd, 2020 by Lippi and Plebani . Soon after, a special issue was devoted to COVID-19 (volume 58, issue 7). Major guidelines from the International Federal of Clinical Chemistry (IFCC) were also published in CCLM , , , , , , , , , . Details about the types of submission is given in Figure 1. A total of 186 publications on COVID-19 have been published so far (last update October 30, 2022). This represented 27.4% of all the publications of CCLM during the same period. The main topics discussed were the measurement of antibodies against SARS-CoV-2 (n=59), the impact of COVID-19 on biochemistry (n=56) or hematological (n=45) measures, followed by SARS-CoV-2 diagnosis by means of molecular assays (n=29) (Figure 2).
The diagnosis of a SARS-CoV-2 infection can be performed by different approaches. The present review will mainly focus on the diagnosis through direct methods including nucleic acid amplification tests (NAATs), antigen and antibody testing, that represent a total of 97 publications in CCLM. Indirect methods that rely on biochemical and/or hematological measurands or their variable combination for predicting SARS-CoV-2 infection (i.e., Corona-Score) are characterized by much lower accuracy for use in a time when testing availability was limited and will not be discussed in detail [7, 15], , .
Nucleic acid amplification tests (NAATs)
On January 10, 2020, the first SARS-CoV-2 genetic sequence was uploaded to the Global Initiative on Sharing Avian Influenza Data (GISAID) platform . Rapidly, diagnostic companies and manufacturers developed NAATs to detect SARS-CoV-2 ribonucleic acid (RNA) in various clinical specimens . NAATs are still considered the gold standard for identification of SARS-CoV-2 in clinical specimens . The most common type of NAAT is real-time reverse transcription polymerase chain reaction (rRT-PCR), which is used by both the Centers for Disease Control and Prevention (CDC)-developed assay and the World Health Organization (WHO)-endorsed assays . Upper (URT) and lower respiratory tract (LRT) specimens collected during the acute phase of infection are booth deemed suitable for detecting SARS-CoV-2 with NAATs . Saliva samples could be considered as an alternative when other samples cannot be collected from symptomatic patients and only with validated NAATs or laboratory-based antigen immunoassays [5, 19]. As for other measurands , , , important pre-analytical issues have been related to inadequate procedures for collection, handling, transport and storage of specimens (especially using URT specimens) [23, 24]. It is estimated that 20–30% of false negative results may occur with URT specimens, which is potentially due to sample collection issues . The utilization of LRT specimens could hence be more desirable for molecular testing, but this may not be clinically realistic due to the invasive nature of this type of sampling . Therefore, the NP swab is still considered the gold standard sample matrix for detecting SARS-CoV-2 with molecular tests . Interfering substances present in the specimens, sample contamination, and pipetting errors are additional important preanalytical issues [11, 24].
The main gene targets employed by currently available NAATs include the nucleocapsid (N), envelope (E), spike (S), RNA-dependent RNA polymerase (RdRP) and open reading frame 1 ab (ORF1ab) genes . In order to minimize the risk of false negative test results, it has been recommended to use at least two SARS-CoV-2 gene targets. The IFCC guidelines also proposed a methodology for verification or validation of the analytical and clinical performance of SARS-CoV-2 assays . It is also better to report the name and target genes used for the NAATs. Importantly, test positivity must be defined based on manufacturer’s recommended cut-off, whilst a thoughtful clinical validation must be conducted when a different cutoff will be used.
Determination of infectiousness status represents an important need to identify patients requiring isolation. The proposed reference standard for establishing infectiousness is viral culture (i.e., absence of viral culture generally implies absence of contagiousness). However, this technique is challenging to perform and requires high biosafety measures that preclude its use in clinical routine . Values of cycle threshold (Ct) of rRT-PCR assays have been proposed as a potential surrogate of viral culture results in clinical samples and for predicting illness severity [5, 26]. Compared to viral culture, Ct values are easier to estimate from amplification curves. However, the suitability of the Ct value to estimate contagiousness remained limited, since Ct results varies between analyzers and laboratories, and direct comparison of values obtained with different techniques must be avoided. The standardization of this measure remains therefore an unresolved issue to date [5, 27, 28]. An illustration of this lack of standardization is the range of Ct cut-offs values reported for contagiousness, which may vary from 24 to over 35 . Furthermore, it also depends on the sample matrix, specimen collection and processing. Finally, a single test does not allow to identify whether patients with low viral load are at the early onset of symptoms or are instead recovering from their infection, thus being in the declining phase of viral load .
The clinical context of the patient is paramount for interpretation of SARS-CoV-2 NAAT results. In symptomatic subjects, the viral RNA can be detected at symptom onset and peaks within the first week. A NAAT can still be interpreted as positive several weeks after the onset of symptoms to subsequently becoming undetectable in most patients . A negative NAAT therefore does not rule out SARS-CoV-2 infection when the test is performed too early or too late during the acute infection . As well as for pre-analytical issues, analytical and post-analytical errors may also be associated with false negative test results.
In patients with signs and symptoms suggestive of SARS-CoV-2 infection (i.e., high-risk patients), the diagnosis carried out with laboratory-based molecular assays should not be confirmed, due to the high positive and negative predictive values of these tests. If a negative result is obtained in a high-risk patient, a repeated test within 24–48 h is recommended, using a different laboratory-based molecular assay if possible. In a patient with a low risk of infection (i.e., hospital admission, contact tracing), the utilization of point-of-care (POC) molecular assays [8, 30] has been reported as a valuable alternative to routine laboratory-based assays, since the diagnostic sensitivity and specificity of several of these tests is only marginally lower (i.e., as high as 95 and 99%, respectively). These methods have low throughput but can provide timely results on a very short timeframe, allowing efficient patient triage compared to laboratory-based molecular assays with turnaround time (TAT) generally ranging between 4 and 12 h .
Since the identification of the initial strain in Wuhan, the wild-type variant, several mutations occurred in the SARS-CoV-2 genome, leading to appearance of different viral lineages . This is the typical consequence of a natural pressure to which all viruses are subjected. In this context, gene sequencing of SARS-CoV-2 is notably essential for the monitoring of emerging lineages that may impact human health . Five lineages have been designated as a variant of concern (VOC) by the WHO so far, namely the Alpha, Beta, Gamma, Delta and Omicron variants. First identified in November 2021, the Omicron lineage is to date the leading variant all over the world. This variant is characterized by a huge number of dominant mutations in the S protein, nearly half of which are located within the sequence of the receptor-binding domain (RBD), thus conferring increased transmissibility and considerable immune escape from acquired protection through COVID-19 vaccination or previous infection with other (different) SARS-CoV-2 variants. Currently, Omicron is largely dominant and several subvariants have emerged including BA.2, BA.2.12.1, BA.2.75.2, BA.4 and BA.5, BQ.1 and XBB.1. All these sublineages also demonstrated considerable escape to acquired immunity [32, 33]. Recently, the Omicron subvariant BA.4.6 (a sublineage of BA.4) has also been identified and presents an increase immune escape compared to BA.4-BA.5 .
The choice of gene targets and primers used by manufacturers should be reviewed to ensure they considered robustness to at least the most common mutant strains and are targeted to highly conserved regions [8, 27, 35]. Each assay must hence be validated against newly emerged SARS-CoV-2 variants, to prevent the risk of generating false negative test results [5, 11]. The Omicron lineages BA.1, BA.1.1 and BA.3 are paradigmatic examples of variants that may generate specific test failures. The emergence of a 69–70del mutation in these variants causes a 6-nucleotide deletion (21,765–21,770) in the S gene of SARS-CoV-2, which results in deletion of two amino acids located between positions 69 (histidine) and 70 (valine) of the S protein, thus impairing the probe annealing in certain assays, ultimately leading to S-gene target failure (SGTF) . More recently, an in silico evaluation found that the sublineages BA.4 and BA.5 may potentially result in false negative test results using four distinct laboratory-based molecular assays .
Compared to NAATs that detect SARS-CoV-2 RNA, antigen assays are aimed at identifying (and possibly measuring) the presence of viral antigens to indicate current viral infection . Various collected specimens (i.e., nasal, nasopharyngeal (NP), saliva, blood, urine) can be used but NP specimens have been most widely validated and used [9, 37], , . Available assay methodology mainly includes rapid detection tests (RDTs) (i.e., manual chromatographic immunoassays, also known as lateral flow immunoassays) used at the POC, as well as laboratory-based immunoassays (i.e., automated immunoassays) [9, 25, 29, 40, 41].
The viral N is the main target of the majority of antigen based RDTs . Compared to other SARS-CoV-2 antigens, the N antigen is the good choice for two main reasons [5, 9]. First, the protein is produced at higher levels compared to other viral SARS-CoV-2 proteins (i.e., S protein), leading to higher assay sensitivity . Second, the selective pressure placed by the increasing number of seropositive people worldwide (either post-infection or post-vaccination) is responsible for boosting higher viral mutations in the S gene, encoding the mature S protein, so that the use of other viral antigens is at least theoretically more advisable . The N antigen seems, however, less vulnerable to selective pressure to develop mutations and is not subjected to the risk of false positive test results in patients undergoing COVID-19 vaccination with vaccines encoding the S protein . As with NAAT, it remains crucial to validate each immunoassay against newly emerged SARS-CoV-2 variants, to prevent false negative test results [5, 42].
Proposed advantages for RDT in NP specimens include its widespread availability as decentralized testing, rapid TAT, patient stratification, overall lower cost, no use of specific equipment or highly trained staff, and preventative case identification. The time of positivization of self-performed RDTs was also reported to reflect the viral load in clinical samples . Nevertheless, major concerns regarding analytical performance persist . Based on available evidence, the sensitivity of RDTs is significantly lower compared to NAATs. Several factors should be considered when assessing RDTs performance: patient characteristics (clinical severity, type and time of onset of symptoms), viral load, and assay method . A Cochrane systematic review has recently evaluated 49 different commercial immunoassays in 155 cohorts totaling 100,462 unique samples . Compared to NAAT, average sensitivity was higher in symptomatic (73.0%, 95% CI 69.3–76.4%) compared to asymptomatic participants (54.7%, 95% CI 47.7–61.6%). Average sensitivity was higher in the first week after symptom onset (80.9%, 95% CI 76.9–84.4%) than in the second (53.8%, 95% CI 48.0–59.6%). Average specificity was similarly high for symptomatic (99.1%) or asymptomatic (99.7%) participants. The sensitivity varied widely according to the different devices (from 34.3 to 91.3% in symptomatic and from 28.6 to 77.8% in asymptomatic participants) . The selection of the assay is hence of upmost importance. In this context, the WHO has set a minimum performance requirement compared to a reference NAAT of ≥80% diagnostic sensitivity and ≥97% diagnostic specificity, respectively .
Recently, Kessler et al. found that a RDT correctly identified all rRT-PCR positive samples with Ct < 25 and that inoculation of cell cultures of samples that were RDT-/rRT-PCR+ did not generate cytopathic effects, presuming the absence of contagiousness when RDT result is negative . The population size was, however, low and some reports showed that RDT could miss specimens with Ct<25 . Along with the limitations of using the Ct derived from rRT-PCR described above, using RDT for identification of contagious patients should be interpreted with caution, especially in the light of new and highly mutated SARS-CoV-2 variants. There is also a higher risk of pre-analytical errors that need to be taken into account, as well as the risk of misuse [9, 29, 45].
Compared to RDTs, laboratory-based assays represented a valuable alternative, with overall higher sensitivities (82.0–88.5%) and high specificities (93.0–99.5%), depending on studies, but still present lower performance compared to NAATs [5, 29, 46], , , . The sensitivity increased to 92.5–100% if considering only samples with higher viral loads (i.e., Ct<25–30) [40, 46, 47, 49].
Given the risk of missing true positive patients compared to NAATs, the place of antigen-based assays (especially RDT) should be considered carefully. In the light of their lower sensitivity, current guidelines recommend performing a laboratory-based molecular assay in patients with high pre-test probability of SARS-CoV-2 infection . When antigen-based tests are used, it is recommended to confirm negative test results by NAAT-based testing in high pre-test probability settings, since negative test result does not definitively exclude the presence of active infection . In low or moderate pre-test probability settings, positive results obtained by RDT should be confirmed by NAAT-based testing . These confirmatory processes can present some organizational challenges, but deserve to be considered for an appropriate patient management.
As earlier discussed, the optimal sample for diagnosing SARS-CoV-2 is an URT specimen, though the use of saliva has also be considered as a reliable option for some antigen immunoassays . Ren et al.  found 92% sensitivity and 100% specificity using an ECLIA (electrochemiluminescence immunoassay) targeting the N protein in saliva samples. Aita et al. also reported 90% sensitivity but lower specificity (92%) using another automated platform. The sensitivity rises to 100% when considering samples with Ct<30 . Although this matrix is interesting, especially in the ambulatory setting, it is important to consider the lower performance of RAD test using a saliva specimen.
The possibility of measuring SARS-CoV-2 antigens in blood has also been explored [37, 51], , . Compared to NAAT, the clinical sensitivity ranged from 85.2 to 93.0% considering studies that included patients who developed symptoms up to a maximum of 2 weeks. The sensitivity increased to 94.2–100% with samples collected within the first days since symptom onset. However, clinical sensitivity significantly decreased after 2 weeks (43.2–74.5%) and after 4 weeks since symptom onset (from 0 to 34.2%). Given that the peak of the N antigen is typically reached after 7 days, as for the viral load in NP samples, and that a continuous decline is observed afterwards, the timing since symptoms is of paramount information for the evaluation of antigenemia. The mean clinical specificity is 90.9% . Of note, the performance of the S antigen assay (sensitivities of 64–85.3%) in blood was lower compared that of the N antigen assay [37, 54]. Higher concentrations of N and S antigens were observed in more severe patients [37, 53], as well as positive correlations with inflammatory biomarker levels (i.e., CRP or IL-6) . Moreover, compared to rRT-PCR , it may provide useful information about illness severity [37, 52, 53]. Finally, such measurements may facilitate patient triage to optimize better intensive care utilization in patients presenting early after symptom onset . The measurement of SARS-CoV-2 antigens in blood is, however, not widely used, though it displays better performance for predicting disease severity compared to NAATs.
In response to SARS-CoV-2 infection, the humoral immunity will lead to generation of several types of immunoglobulins (Ig) against the pathogen (especially IgA, IgG and IgM), which have a primary function to protect the body mucosae from virus penetration (i.e., secretory IgG and dimeric IgA), as well as to neutralize the virus once it has colonized the body and/or entered the circulation (especially IgG) . The measurement of antibodies is classically performed in serum or plasma obtained from a venipuncture, but the use of whole blood  or dried blood spots represent a valuable alterative [56, 57]. The use of saliva has also been explored, but requires further investigation .
Manufacturers have mostly focused on developing immunoassays against IgG or total antibodies (sum of all isotypes) rather than IgA and IgM . The detection of IgG or total antibodies (i.e., seroconversion) occurs approximately within 7–14 days after symptom onset [11, 59], , . The IgA response develops early, coinciding with that of IgM, peaks after 18–21 days, and appears to be even stronger and more persistent than the IgM . Performance of IgM assays was found to be lower compared to IgG and total antibodies for detecting previously infected individuals , , , . In convalescent patients, antibodies can still be measured in most individuals after a long period of time (>8–10 months post-infection), with total antibodies being even more persistent compared to IgG , , , , , . However, a slow decay is observed over time.
Available assays could target either the S protein, the RBD of the Subunit one of the S protein, or the N protein of SARS-CoV-2 . Antibodies against the N protein will only be generated in infected patients or in those receiving attenuated vaccines, while anti-S/RBD antibodies are generated in both infected and vaccinees. Some multiplex methods are able to measure several different targets within a run [73, 74]. Immunoassays can use different technologies (LFA, ELISA, CLIA), and can be quantitative, semi-quantitative or qualitative , , , . Given the different types of assays, their fit for purpose properties need to be validated, and the IFCC provides an ad hoc evaluation protocol . These evaluations are important since the performance of antibody assays are not equivalent, especially considering the use of rapid tests [77, 78]. The use of an orthogonal testing strategy has been for example proposed to avoid false positive results, especially in low prevalence settings [10, 73, 79].
As compared to binding antibodies, neutralizing antibodies (NAbs) represent the first line of response of adaptive immunity against SARS-CoV-2. They are of particular importance because they inhibit the binding of the RBD at the surface of the S protein to the human angiotensin-converting enzyme 2 (ACE2) receptor, thus hampering host cell penetration [32, 33, 80]. NAbs represents the best correlate of immunity against symptomatic infections [32, 80, 81]. It is important to highlight that current serology immunoassays available in the diagnostic market (i.e., binding antibody assays) do not provide definitive information regarding patient immunity. For this analysis, a neutralization assay obtained in a cell culture system is needed to determine the presence of active antibodies and relative protection against future infection [11, 32, 33, 82]. NAbs are only modestly correlated against commercial assay targeting S or RBD directed antibodies [32, 65, 69, 83], , , , , . Binding antibodies are therefore not suited to reliably reflect the presence of NAbs. These latter can be measured using live virus neutralization test, pseudovirus neutralization test or by a surrogate assay [65, 69, 86]. Considering the cumulative evidence from the scientific literature showing the modest correlation between binding antibodies and neutralization tests, manufacturers of binding assays need to rethink their current commercial proposal (e.g., by a modification of antigen and epitopes, or by modifying the testing method) to design assays capable of predicting neutralizing activity against emerging and highly mutated SARS-CoV-2 variants such as Omicron. In such case, in fact, antibodies titers do not efficiently reflect the neutralizing capacity of serum despite the availability of Omicron-based standards for calibrating the assay (i.e., the real problem here is, again, the monoclonal antibodies used for biding a different antigen, not the calibrator). This would also avoid misinterpretation (i.e., high protection in case of high-binding antibody titers). Current methods used to measure NAbs present a low throughput, are time-consuming, need skillful operators, and require high levels of biosafety (especially for live virus neutralization assay). It would therefore be easier to use commercial assays that can be a surrogate of these reference methods [32, 33]. As for NAATs and antigen testing, there is a need for harmonization in serological assays, including specific concerns on quality controls .
In the IFCC interim guidelines about the role of serology in COVID-19, several indications were formulated . First, it is obvious that compare to NAATs and antigen testing, the measurement of antibodies is not used for diagnosis of acute infection. Its measurement could, however, be useful as adjunct to molecular testing in patients presenting with suggestive clinical features (>14 days post symptom onset), but molecular/antigen testing for SARS-CoV-2 is negative, undetermined, or unavailable . It is also useful to serve as adjunct when persistently positive molecular tests occur in the absence of evidence of acute infection, such as late after resolving infection or to assist in the diagnostic workup of multisystem inflammatory syndrome in children (MIS-C) . These guidelines were published before vaccination programs and were therefore not able to make a statement about the use of antibody testing in this context. The measurement of antibodies is also useful for seroprevalence study purposes and for developing risk prediction models . Indeed, more severe and non-immunocompromised patients tend to develop higher levels of anti-SARS-CoV-2 antibodies [6, 68, 69, 86]. Other indications that deserve further validation may include identification of patients at higher risk to develop a breakthrough infection. Indeed, lower levels of binding antibodies and NAbs in peri-infection (or during the pre-booster period) were observed in breakthrough patients compared to those who did not develop infection , , .
It has been anticipated that anti-SARS-CoV-2 antibodies measurement may have a role in the vaccination campaign, especially in scenarios of vaccine scarcity or contexts of organizational complexity. In vaccinees, the kinetics of anti-S/RBD antibodies was found to be similar to that in infected patients (i.e., from 7 to 14 days for seroconversion) and most vaccinees had quantifiable levels of antibodies after 14–21 days , , . In previously infected individuals, a boost in antibodies was observed sooner [94, 95]. The peak response was observed around 1 month after the vaccination and a rapid antibody decay was already observed 3 months after completing a primary vaccination [94, 97]. After 6 months, vaccine efficacy (VE) against symptomatic diseases was significantly reduced and was associated with disappearance of NAbs and binding antibodies [83, 98], , , , , . Notably, the level of binding antibodies still remained very high while NAbs were considered below positivity cut-off for approximately 50% of patients. The administration of a third dose was therefore expected to boost the level of NAbs. The levels of both NAbs and binding antibodies increased rapidly after the first booster [32, 100, 102]. The VE quickly increased in the same manner as NAbs, but a waning of antibodies and VE was also observed after the first booster [32, 104].
The emergence and surge of VOCs that present a considerable escape to acquired immunity is also responsible for lower VE. Mutations may jeopardize the reliability of the currently available commercial immunoassays for detecting anti-SARS-CoV-2 S and RBD total or IgG antibodies [33, 82, 105]. A crucial question is: are immunoassays in current clinical use for detecting anti-SARS-Cov-2 antibodies (either anti-S trimeric or anti-RBD) still valid for detecting (and monitoring) the neutralizing activity against highly mutated SARS-CoV-2 variants? It is virtually unquestionable that some of these variants have introduced such a huge number of mutations in the S protein (i.e. >30–35 for Omicron sublineages), such that the antigen (either S or RBD) and its relative epitopes coated in the immunoassays may no longer reflect the original sequence and structure of the prototype SARS-CoV-2 lineage first detected in Wuhan in 2019 and used as the target antigen in almost all commercial serological immunoassays .
This review was the occasion to look back at what CCLM has achieved during the past two years in regard to COVID-19 diagnostics. A high number of high-impact articles have been published, including IFCC guidelines that helped the clinical community to deal with possible causes of false negative and positive results. Several challenges related to COVID-19 diagnosis, however, still need to be addressed. Major efforts are needed for harmonization/standardization of diagnostic methods that could, for example, allow easier identification of contagious patients. Continuous evaluation of the performance of assays (NAAT, antigen and antibodies) is a leading task of laboratory professionals, especially with rapid emergence of new variants that can be associated with detection failures. The role of the clinical laboratory for monitoring vaccine response should also be explored more deeply, especially with respect to the development of surrogate markers for the presence of NAbs, that are highly correlated to VE. Finally, cellular immunity should be the next target of research in laboratory medicine, since T cell function is emerging as perhaps the best predictor of natural and vaccine-elicited immunity against the risk of developing severe COVID-19 illness , , .
The authors would like to thank Heike Jahnke for its precious help in gathering all the publishing data about CCLM in the COVID-19 era.
Research funding: None declared.
Author contributions: All authors have accepted responsibility for the entire content of this manuscript and approved its submission.
Competing interests: Authors state no conflict of interest.
Informed consent: Not applicable.
Ethical approval: Not applicable.
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