key: cord-0870867-4zhhijvw
authors: Mestdagh, Pieter; Gillard, Michel; Dhillon, Sharonjit K.; Pirnay, Jean-Paul; Poels, Jeroen; Hellemans, Jan; Hutse, Veronik; Vermeiren, Celine; Boutier, Maxime; De Wever, Veerle; Soentjens, Patrick; Djebara, Sarah; Malonne, Hugues; André, Emmanuel; Arbyn, Marc; Smeraglia, John; Vandesompele, Jo
title: Evaluating diagnostic accuracy of saliva sampling methods for SARS-CoV-2 reveals differential sensitivity and association with viral load
date: 2021-08-03
journal: J Mol Diagn
DOI: 10.1016/j.jmoldx.2021.07.017
sha: 9bd7bda3e3f5954993c88bf25a7bed8d8e0b2b91
doc_id: 870867
cord_uid: 4zhhijvw

Nasopharyngeal swabs are considered the preferential collection method for SARS-CoV-2 diagnostics. Alternative sampling procedures that are less invasive and do not require a healthcare professional, such as saliva collection, would be more preferable. We compared saliva specimens and nasopharyngeal (NP) swabs with respect to sensitivity in detecting SARS-CoV-2. We obtained a nasopharyngeal and two saliva specimens (collected by spitting or oral swabbing) from more than 2500 individuals. All samples were tested by RT-qPCR detecting RNA of SARS-CoV-2. We compared the test sensitivity on the two saliva collections with the nasopharyngeal specimen for all subjects and stratified by symptom status and viral load. Of the 2850 patients for which all three samples were available, 105 were positive on NP, whereas 32 and 23 were also positive on saliva spitting and saliva swabbing samples, respectively. The sensitivity of the RT-qPCR to detect SARS-CoV-2 among NP-positive patients was 30.5% (95% CI=1.9%-40.2%) for saliva spitting, and 21.9% (95% CI=14.4%-31.0%) for saliva swabbing. However, when focusing on subjects with medium to high viral load, sensitivity on saliva increased substantially: 93.9% (95% CI=79.8%-99.3%) and 76.9% (95% CI=56.4-91.0) for spitting and swabbing, respectively, regardless of symptomatic status. Our results suggest that saliva cannot readily replace nasopharyngeal sampling for SARS-CoV-2 diagnostics but may enable identification of the most contagious cases with medium to high viral loads.

Massive RT-qPCR based testing for the presence of SARS-CoV-2 RNA is a key element in the strategy to control the current COVID-19 pandemic. At present, collecting samples from the upper respiratory tract is recommended for diagnostic testing by the World Health Organization and (American and European) Centers for Disease Control and Prevention, with nasopharyngeal (NP) swabs being considered the standard collection procedure 1,2 (https://www.cdc.gov/coronavirus/2019-ncov/lab/guidelines-clinical-specimens.html). While extremely sensitive, this sampling procedure is relatively invasive, causing discomfort and anxiety in individuals undergoing the procedure, and relies on trained healthcare workers wearing full personal protective equipment (PPE) to obtain samples. Before the outbreak of SARS-CoV-2, several studies have reported on the utility of saliva as a diagnostic specimen for testing respiratory viruses [3] [4] [5] [6] . Additionally, studies related to SARS-CoV-2 have shown that the virus binds to angiotensin-converting enzyme (ACE2) receptors that are present in epithelial cells of the oral mucosa suggesting the use of saliva as a potential sample for SARS-CoV-2 detection. The non-invasive nature of saliva collection in a simple container makes this specimen a valuable biomaterial. Besides, saliva sampling could be a solution in resource-limiting settings with respect to healthcare personnel, and reduce the amount of contact required between a healthcare provider and the patient, lowering the risk of transmission and PPE use. As saliva sampling is patient-friendly, it could also be of value when testing in children during a SARS-CoV-2 outbreak in schools. While several recent studies have documented the potential utility of saliva for diagnostic testing of SARS-CoV-2 [7] [8] [9] [10] [11] [12] [13] , these studies suffer from one or more limitations, i.e. non-paired study design, small cohorts and testing in biased populations such as previously confirmed positive cases and/or hospitalized patients. Here, we set out to prospectively evaluate the potential use of saliva samples for diagnostic testing of SARS-CoV-2 using a large population of more than 2500 individuals in triage centers in Belgium. Individuals were sampled using two saliva collection devices and a matching nasopharyngeal swab, and samples were analyzed by two test laboratories to independently verify conclusions ( Figure 1 ).

This study has been reported using the Standards for Reporting of Diagnostic Accuracy Studies (STARD) guidelines 2015 14 . As part of the Belgian national testing platform, we prospectively enrolled asymptomatic and symptomatic individuals suggestive of coronavirus disease 2019 (COVID-19) at centralized triage centers in Belgium. More than 2500 individuals were tested in these triage centers from June 2020 to July 2020, at the end of the first infection wave. All individuals aged 18 years or older that presented at triage centers were considered eligible.

Study samples were collected by trained mobile teams. Individuals were sampled using 3 different procedures: (1) a nasopharyngeal swab sample representing the standard comparator for SARS-CoV-2 diagnostics, (2) a saliva sample collected through self-sampling with a commercial saliva spitting device (Saliva RNA Collection and Preservation Device, Norgen Biotek, Canada) and (3) a saliva sample collected through self-sampling with a commercial oral swabbing device (Oracollect RNA, DNA Genotek, Canada). For the saliva spitting device (Norgen Biotek), the collected volume of saliva was 2 ml. For the saliva swabbing device (DNA Genotek), the collected volume of saliva was ~300 µl. NP sample collection was performed using iClean NP swabs (Rhino Diagnostics). All samples were collected in a transport buffer that inactivates the virus and stabilizes the RNA (Norgen Biotek or DNA Genotek buffer for the respective saliva samples and 2 ml of DNA/RNA shield buffer (Zymo Research), custom prefilled in Vacuette tubes (Greinder Bio-One) for the nasopharyngeal samples). Participants in the study were asked not to eat, drink, smoke or use chewing gum 30 minutes preceding saliva sampling. Saliva sample were collected according to the manufacturer's instructions. These instructions were available as an instruction sheet with each saliva collection device. Instructions were communicated to each participant by a healthcare professional prior to sampling. Participants were not instructed to produce deep throat saliva or gargle prior to saliva collection. For the swabbing device, participants were instructed to place the swab between the right cheek and gum, swab 10 times, and repeat for the left cheek. After sample collection, a short survey was completed and data was collected on age group, ease of use of the saliva devices, comfort of saliva sampling versus nasopharyngeal sampling and symptomatic status. To enquire about symptomatic status, the case definition of Sciensano, the Belgian Institute for Public Health was used. The case definition stated that a possible case of COVID-19 had at least one of the following main symptoms that occurred acutely without other plausible cause: cough, dyspnea, thoracic pain, anosmia or dysgeusia; or at least two of the following symptoms that occurred without other plausible cause: fever, muscle strain, fatigue, rhinitis, sore throat, headache, anorexia, watery diarrhea, acute confusion, sudden fall; or worsening of chronic respiratory symptoms (COPD, asthma, chronic cough) without other plausible cause. This study was approved by the ethical review committee of the University Hospital of Leuven on May 29, 2020 as S64125.

Test methods SARS-CoV-2 testing was performed by two independent test laboratories, applying different RNA extraction and RT-qPCR workflows (see below). Note that, because of logistics reasons, not all samples were analyzed by both laboratories. After sample collection, samples were shipped to one of the laboratories where the required volume of sample for RNA extraction was removed from the sample collection tube. Sample collection tubes were subsequently shipped to the other test laboratory for analysis. Prior to sample transfer, samples were vortexed and centrifuged. For highly viscous samples, aspiration was performed at low speed.

Nucleic acid extraction and RT-qPCR in laboratory 1 RNA extraction was performed using the Total RNA Purification Kit (Norgen Biotek #24300) according to the manufacturer's instructions using 200 µl viral transport medium (for the nasopharyngeal swab, i.e. DNA/RNA Shield (#R1100-250, Zymo Research) or 200 µl saliva collected with the spitting or swabbing device (i.e. saliva mixed with transport buffer) as input in the 96-well filter plate. Samples were supplemented with 200 µl lysis buffer, 200 µl ethanol, 4 µl of a proprietary 700 nucleotides spike-in control RNA (5000 copies, produced through in vitro transcription) and carrier RNA (200 ng of yeast tRNA (Roche #10109517001). Filter plates were further processed with a centrifuge (5810R with rotor A-4-81, both from Eppendorf). RNA was eluted from the filter plates using 50 µl elution buffer (nuclease-free water), resulting in approximately 45 µl eluate. RNA extractions were simultaneously performed for 94 patient samples and 2 negative controls (nuclease-free water). To the eluate of one of the negative control wells, 7500 (digital PCR value assigned) RNA copies of positive control RNA (Synthetic SARS-CoV-2 RNA Control 2, Twist Biosciences #102024) were added to serve as positive PCR control.

Six µl of RNA eluate was used as input for a 20 µl duplex RT-qPCR reaction in a CFX384 qPCR instrument using 10 µl iTaq one-step RT-qPCR mastermix (Bio-Rad #1725141) according to the manufacturer's instructions. Reactions were set up using 400 nM final concentration of primers and 250 nM of a hydrolysis probe. Primers and probes were synthesized by Integrated DNA Technologies using clean-room GMP production. For detection of the SARS-CoV-2 virus, the Charité E gene assay was used (FAM) 15 ; for the internal control, a proprietary hydrolysis probe assay (HEX) was used. Cq values were generated using the FastFinder software v3.300.5 (UgenTec). The FastFinder software was also used to call a sample positive or negative for SARS-CoV-2. Only batches with a clean negative control and a positive control in the expected range were approved. Proper RNA extraction and RT-qPCR was confirmed by observing spikein RNA signal in each sample well in the expected range.

Nucleic acid extraction and RT-qPCR in laboratory 2 RNA extraction was performed using a magnetic bead-based RNA extraction method developed by University of Liège (CoRNA kit) and according to the recommended protocol. For nasopharyngeal swab samples, 200 µl of sample was transferred to 11 µl of a proteinase K solution (20 ml/ml). For both saliva devices, 100 µl of saliva was transferred to 175 µl of a lysis buffer mix (11:164 (vol/vol) of proteinase K solution (20 mg/ml) + lysis buffer respectively). All samples were spiked with the MS2 phage as internal control (10µl, conc proprietary info from supplier Thermofisher Scientific kit A47814), and in presence of carrier RNA (10µl, 20ng/µl, Merck/Roche #10109517001) to increase RNA extraction efficiency. The multiplex RT-qPCR was performed on 5 µl of RNA eluate using TaqPath COVID-19 Combo Kit (comprising ORF1ab, N gene, S gene, and MS2 as internal control, #A47814, Thermo Fisher), TaqPath positive control kit (containing a stock of 10 4 copies SARS-CoV-2 /µl, #A47816, Thermo Fisher) and TaqPath 1-Step Multiplex Master Mix (no ROX) (#A28523, Thermo Fisher), following the manufacturer's instructions. Positive control was diluted to 25 copies/µl in control dilution buffer and 2 µl (50 copies) was further diluted in 3 µl nuclease free water which was added to the well of the RT-PCR plate. Cq values were generated using the FastFinder software v3.300.5 (UgenTec). The FastFinder software was also used to call a sample positive or negative for SARS-CoV-2. Results were approved when a clean negative control and a positive control in the expected range were obtained. Correct RNA extraction and RT-qPCR setup was also confirmed by controlling MS2 amplification in each sample well (applying an MS2 Cq-cutoff of 33).

Digital PCR was done on a QX200 instrument (Bio-Rad) using the One-Step RT-ddPCR Advanced Kit for Probes (Bio-Rad #1864022) according to the manufacturer's instructions. Briefly, 22 µl pre-reactions were prepared consisting of 5 µl 4x supermix, 2 µl reverse transcriptase, 6 µl positive control RNA (see further), 15 mM dithiothreitol, 900 nM of each forward and reverse primer and 250 nM E-gene hydrolysis probe (FAM) 15 . 20 µl of the pre-reaction was used for droplet generation using the QX200 Droplet Generator, followed by careful transfer to a 96-well PCR plate for thermocycling: 60 min 46 °C reverse transcription, 10 min 95 °C enzyme activation, 40 cycles of 30 sec denaturation at 95 °C and 1 min annealing/extension at 59 °C, and finally 10 min 98 °C enzyme deactivation. Droplets were analyzed by the QX200 Droplet Reader and QuantaSoft software. With an input of 4000 RNA copies per reaction (Armored RNA Quant SARS-CoV-2 Panel, Asuragen #52036), the digital PCR result was 875 cDNA copies (or 21.88% of the expected number). Of note, no RNA extraction was done on the Armored RNA material; instead, a short heat release of RNA was done per the manufacturer's instructions. With an RNA input of 750 copies per reaction (Synthetic SARS-CoV-2 RNA Control 2, Twist Biosciences #102024), the digital PCR result was 150 cDNA copies (or 20% of the expected number.

The Cq value cut-off for viral load classification was determined based on the Cq correlation between NP and saliva samples. The cut-off represents the NP viral load above which saliva samples show highest sensitivity for SARS-CoV-2 detection in NP positive samples, and below which sensitivity for SARS-CoV-2 detection in saliva drops to almost 0%. This analysis resulted in an E-gene Cq-cutoff of 24.5 and N-gene Cq-cutoff of 25.5 for laboratory 1 and 2, respectively. To convert the laboratory 1 Cq value cut-off to SARS-CoV-2 RNA copies/ml of viral transport medium, we created a 6-point 10-fold serial dilution of Armored RNA in triplicate (from 2.19E8 to 2.19E3 digital PCR value assigned copies/ml), followed by RNA extraction and RT-qPCR (using the laboratory 1 method). Based on the slope of -3.381, the y-intercept of 45.387 and the r 2 value of 0.995, the laboratory 1 Cq value Cq cut-off corresponds to 1.51E6 copies/ml viral transport medium. This viral copy number corresponds to a viral load that is typically associated with infectious individuals in literature. For instance, van Kampen et al. demonstrated that the probability of isolating infectious SARS-CoV-2 was less than 5% when the viral load was below 6.63 log10 RNA copies/mL 16 . Therefore, we decided to refer to this as 'high viral load'.

The sample size was computed for assessment of a hypothesis on non-inferior SARS-CoV-2 positivity on saliva compared to on nasopharyngeal specimen in paired testing as proposed by Tang 17 , using target values from a systematic review 18 . We accepted a confidence of 95%, a power of 80%, a sensitivity of the test in NP samples of 95%, a proportion of saliva-/NP+ samples of 5% and 0.90 as benchmark for the relative positivity rate (saliva/NP), which yielded 84 SARS-CoV-2+ subjects needed. These could be found in a study population of 841 to 8410 subjects assuming a prevalence of 1% to 10%. Given the substantially larger contrast in test positivity between saliva and NP specimens, study enrolment was stopped after reaching 2850 inclusions.

All results presented in the manuscript are based on data generated by laboratory 1, unless stated otherwise. Only patients with available results for the three specimens were included. Patient paired data was used to construct 2x2 contingency tables. The sensitivity of SARS-CoV-2 testing was defined by the proportion of saliva positive patients (index+) among those that were positive on NP (reference+). We also computed the test positivity ratio as the proportion J o u r n a l P r e -p r o o f with a positive index test over the proportion with a positive reference or comparator test. Ninety five percent confidence intervals for binomial data were computed as well as for ratios of paired proportions. Three separate analyses were performed: one comparing spit samples to NP samples, a second comparing swab samples to NP samples, and a third comparing spit and swab samples. NP samples were considered the standard comparator or reference. Additionally, the estimations were stratified by viral load (categorized as high and low) and symptoms (categorized as symptomatic and asymptomatic). All statistical analyses were performed using Stata statistical software version 14.2 (Stata, College Station, TX, USA). Statistical significance was defined at p < 0.05.

RT-qPCR data from test laboratory 1 and test laboratory 2 as well as the survey data are provided as Supplemental Tables 1,2 and 3.

In total, 2954 individuals were recruited between June 2020 and July 2020. We excluded data from 104 (3.5%) participants owing to missing NP results. 2268 individuals were sampled with a nasopharyngeal swab and both saliva collection methods whereas 2469 and 2649 matched samples were available for spit and swab samples, respectively. The median age group of participants was between 31 and 40 years old and symptomatic status data was available for 2071 individuals.

While saliva sampling was generally perceived as more comfortable than nasopharyngeal sampling (Supplemental Figure 1A) , study participants scored the ease of use of the saliva swabbing device significantly higher than that of the saliva spitting device (p<0.0001, Mann-Whitney test) (Supplemental Figure 1B) .

Out of 2850 nasopharyngeal swab samples analyzed by laboratory 1, 115 (4.0%) were SARS-CoV-2 positive (Figure 2 ). In positive NP samples, 30.4% (35/115) showed high viral load. There were 105/115 nasopharyngeal positive samples for which a matching saliva spitting sample was available, and 105/115 nasopharyngeal positive samples for which a matching saliva swabbing sample was available (Figure 2 ). We observed 32/105 (sensitivity = 30.5%; 95%CI= 21.9%-40.2%) in the spitting sample and 23/105 (sensitivity = 21.9%; 95%CI=14.4%-31.0%) in the swabbing samples that were SARS-CoV-2 positive, indicating reduced overall sensitivity in saliva for SARS-CoV-2 detection (Figure 2, Figure 3 , Table 1 ). However, we observed a significantly higher nasopharyngeal viral load in patients with a truepositive saliva sample compared to patients with a false-negative saliva sample (spitting device: log2 fold change = 14.89, p=3.79 x 10 -15 ; swabbing device: log2 fold change = 14.7, p=3.67 x 10 -12 , Mann-Whitney test) (Supplemental Figure 2) . Individuals with an E-gene Cq > 24.5 in the nasopharyngeal sample (corresponding to 1.51E6 copies/ml viral transport medium as determined by digital PCR and further referred to as low viral load) almost always presented with a negative saliva sample (sensitivity = 1.4%; CI=0.07%-7.5% and sensitivity = 1.3%; CI=0.07%-7.2% in the saliva spitting and saliva swabbing sample, respectively) ( Figure 3 ). In contrast, for individuals with a high viral load (E-gene Cq < 24.5 in the nasopharyngeal sample), concordance between the nasopharyngeal and matching saliva sample improved dramatically, especially for the saliva spitting device, resulting in high sensitivity in this subgroup (sensitivity = 95.5%; 95%CI=77.2%-99.9% and sensitivity = 77.3%; 95%CI=54.6%-92.2% for the saliva obtained by spitting and swabbing, respectively) ( Figure 3 , Table 1 ). In addition, we observed a significant positive correlation between E-gene Cq-values in the nasopharyngeal and saliva samples for those individuals with high viral load (spitting device: r = 0.53, p = 0.002; swabbing device: r = 0.54, p = 0.006; Pearson's correlation). Notably, similar findings were obtained based on test results generated by laboratory 2 (Supplemental Figure 3 and Supplemental Table 4 ). In the medium to high viral load subgroup, saliva spitting resulted in a sensitivity = 96.9% (95%CI=83.8%-99.9%) while saliva swabbing resulted in a sensitivity = 60.7% (95%CI=40.6%-78.5%).

To assess whether the poor sensitivity in saliva for SARS-CoV-2 detection could be due to sampling issues during saliva collection, we quantified the human gene RPS18 using RT-qPCR on a representative set of RNA samples used for SARS-CoV-2 detection. No differences in RPS18 levels were observed between saliva positive and negative samples for any of the saliva sampling devices, suggesting that sampling issues do not explain the false-negative saliva samples (Supplemental Figure 4 A-B) .

We were able to register presence or absence of symptoms of COVID-19 in 2123 study participants. From these, 1376 (64.8%) were symptomatic, 695 (32.7%) asymptomatic and 52 (2.5%) indicated they experienced symptoms 2 weeks prior the test. The latter group was excluded from further analyses due to the limited number of individuals. The proportions of individuals that were SARS-CoV-2 positive in the nasopharyngeal sample were similar in the symptomatic and asymptomatic group, 3.7% and 4.2% respectively, however the symptomatic group was enriched with high viral load samples (E-gene Cq <24.5, p = 0.042, Fisher's exact test). As a result, sensitivity in saliva for SARS-CoV-2 detection was higher among symptomatic cases (sensitivity = 33.3%; CI=20.8%-47.9% and sensitivity = 25.5%; CI=14.3%-39.6% for spitting and swabbing saliva device respectively) compared to asymptomatic cases (sensitivity = 13.8%; 95%CI=3.9%-31.7% for both the spitting and swabbing saliva device) (Figure 4 ). Among individuals with high viral load in NP samples, the sensitivity in the saliva samples was high, irrespective of symptomatic status. Sensitivity was 94.4% [95%CI=77.2%-99.9%] and 100.0% [95% CI=39.8%-100%] in symptomatic and asymptomatic individuals in the spitting sample whereas the sensitivity from the swabbing saliva sample was 72.2%; [95% CI=46.5%-90.3%] in symptomatic subjects and 100.0% [95% CI=39.8%-100%] in asymptomatic subjects. The sensitivity and test positive ratio are summarized in Table 1 (and Supplemental Table 4 for lab 2).

The literature on the use of saliva for SARS-CoV-2 detection is rapidly evolving and expanding. Saliva sampling for COVID-19 diagnostics has been put forward as an alternative for nasopharyngeal sampling in a number of independent studies. A rapid review of the literature estimated a pooled sensitivity of SARS-CoV-2 testing on saliva versus nasopharyngeal samples as high as 97% 18 . More recently, a meta-analysis of 16 unique studies representing 5922 patients reported a pooled sensitivity for SARS-CoV-2 detection in saliva of 83.2% 19 . Here, we compared the sensitivity of two different saliva collection devices to the nasopharyngeal swab in over 2500 individuals that were sampled at different triage centers in Belgium.

In contrast to the current literature, we observed a substantially lower SARS-CoV-2 test positivity rate in saliva than in nasopharyngeal samples. However, when focusing on individuals with a high viral load (more than 1.51E6 copies/ml viral transport medium), sensitivity improved dramatically, especially in saliva samples produced through spitting. There are several potential reasons for the discrepancy between results presented here and current literature reports. First, the study population may be very different. Most of the studies reported in literature predominantly include symptomatic patients (whether or not hospitalized) that are more likely to have a high viral load. Our study included individuals visiting triage centers to obtain diagnostic testing for SARS-CoV-2, either because they presented with COVID-19 symptoms or they had been in close contact with an infected individual. These individuals were not critically ill and often did not have symptoms. Secondly, while our study compared two different devices for saliva collection, we cannot exclude the possibility that these devices are less suitable for saliva collection compared to what has been used in other studies. Of note, we observed differential sensitivity between both devices and conclude that saliva sampling through spitting is more sensitive than swabbing. Whether the order of saliva collection (participants were asked to first swab, then spit) could impact these results remains to be investigated. A study conducted in British Columbia collected samples from outpatient testing centers and evaluated saline mouth rinse/gargle, also known as swish and gargle approach, to collect saliva samples compared to neat saliva collection and found that the swish/gargle method had a higher sensitivity than neat saliva, 97.5% [95 CI: 86.9%-99.9%] versus 78.8% [95% CI: 61%-91%]. It is unclear why the authors observed a significant difference in the sensitivity, but the saliva collection method could indeed influence sensitivity of the test result. Note that, in our study, saliva samples were collected without gargling or throat clearing, actions that may further improve the sensitivity of SARS-CoV-2 detection in saliva. Finally, other co-variates, including time of the day of sampling, stage of infection at sampling, and timing relative to an epidemiological wave (i.e. varying reproduction number) may also impact results. With respect to the latter, patients were sampled at the end of the 1 st (spring) wave in Belgium, a period of low prevalence and low individual viral load (compared to the beginning of the 2 nd (autumn) wave 20 ) . More studies are required in order to further investigate the impact of these factors. Notably, testing different but biologically related samples from each patient (saliva and oral swab) provides an internal validation of our study results, as both sample types lead to highly similar conclusions. Our study also had some limitations. Firstly, more detailed clinical and demographic data would have been helpful to evaluate if other factors could explain the difference in sensitivity between saliva and the nasopharyngeal swab. In addition, the inclusion of hospitalized patients may have allowed a more in-depth analysis on the relation between disease severity and detection sensitivity in saliva. Finally, longitudinal saliva collections in positive individuals could shed light on the dynamics of SAR-CoV-2 detection rates in function of disease progression.

In summary, our study suggests that saliva sampling cannot replace the standard nasopharyngeal swab for the diagnosis of SARS-CoV-2 in the population we studied. Nevertheless, because of its ease of use and compatibility with self-sampling, saliva sampling could play a role in systematic screening campaigns that aim to identify asymptomatic cases with medium to high viral loads. However, based on results presented here, such screening campaigns would fail to identify low positives. Whether, and to what extent, these low positives are capable of spreading the virus requires further investigation. Overview of study design. Study participants were sampled at triage centers using a nasopharyngeal swab and two different saliva collection devices. Samples were processed at 2 different test laboratories using independent sample processing workflows. 

European Centre for Disease Prevention and Control: Considerations for the use of saliva as sample material for COVID-19 testing

Laboratory testing for coronavirus disease (COVID-19) in suspected human cases: interim guidance

Comparison between saliva and nasopharyngeal swab specimens for detection of respiratory viruses by multiplex reverse transcription-PCR

Comparison among nasopharyngeal swab, nasal wash, and oropharyngeal swab for respiratory virus detection in adults with acute pharyngitis

Comparison of Nasopharyngeal and Oropharyngeal swabs for the diagnosis of eight respiratory viruses by real-time reverse transcription-PCR assays

Use of throat swab or saliva specimens for detection of respiratory viruses in children

Saliva is a reliable tool to detect SARS-CoV-2

Saliva sample as a non-invasive specimen for the diagnosis of coronavirus disease 2019: a cross-sectional study

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Saliva as a noninvasive specimen for detection of sars-cov-2

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Comparison of SARS-CoV-2 detection in nasopharyngeal swab and saliva

Saliva or Nasopharyngeal Swab Specimens for Detection of SARS-CoV-2

STARD 2015 guidelines for reporting diagnostic accuracy studies: explanation and elaboration

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Evaluation of efficiency and sensitivity of 1D and 2D sample pooling strategies for SARS-CoV-2 RT-qPCR screening purposes

The authors want to address special thanks to the doctors, the nurses and the paramedics of the Belgian Armed Forces and other teams involved in the massive sampling campaigns in the different triage centers. The authors want to thank Anke Maertens for logistics support, Pierre Wattiau (GSK Biologicals) and Olivier Vandeputte (GSK Biologicals) for constructive discussions on study design and results and Els Dequeker (KU Leuven) for supervising the ethical approval and processing of the survey forms. 

 Table 1 . Sensitivity and test positivity ratios of SARS-CoV-2 testing on NP specimens vs saliva (collected by swabs or spitting) and on swab saliva samples vs spitting saliva samples by viral load level and symptom status (Laboratory 1)