Resilient SARS-CoV-2 diagnostics workflows including viral heat inactivation.

There is a worldwide need for reagents to perform SARS-CoV-2 detection. Some laboratories have implemented kit-free protocols, but many others do not have the capacity to develop these and/or perform manual processing. We provide multiple workflows for SARS-CoV-2 nucleic acid detection in clinical samples by comparing several commercially available RNA extraction methods: QIAamp Viral RNA Mini Kit (QIAgen), RNAdvance Blood/Viral (Beckman) and Mag-Bind Viral DNA/RNA 96 Kit (Omega Bio-tek). We also compared One-step RT-qPCR reagents: TaqMan Fast Virus 1-Step Master Mix (FastVirus, ThermoFisher Scientific), qPCRBIO Probe 1-Step Go Lo-ROX (PCR Biosystems) and Luna® Universal Probe One-Step RT-qPCR Kit (Luna, NEB). We used primer-probes that detect viral N (EUA CDC) and RdRP. RNA extraction methods provided similar results, with Beckman performing better with our primer-probe combinations. Luna proved most sensitive although overall the three reagents did not show significant differences. N detection was more reliable than that of RdRP, particularly in samples with low viral titres. Importantly, we demonstrated that heat treatment of nasopharyngeal swabs at 70°C for 10 or 30 min, or 90°C for 10 or 30 min (both original variant and B 1.1.7) inactivated SARS-CoV-2 employing plaque assays, and had minimal impact on the sensitivity of the qPCR in clinical samples. These findings make SARS-CoV-2 testing portable in settings that do not have CL-3 facilities. In summary, we provide several testing pipelines that can be easily implemented in other laboratories and have made all our protocols and SOPs freely available at https://osf.io/uebvj/.

Introduction

“Test, test, test”–this was the message from the World Health Organization’s (WHO) Head Tedros Adhanom Ghebreyesus on the 16th of March 2020. This message is still current, more than a year after the pandemic was declared. To fight the exponential spread of SARS-CoV-2, measures of social distancing have been imposed in many countries worldwide, while others are now in a phase of de-escalation. Social distancing and lockdown measures have resulted in stagnant or dropping numbers of new infections. However, appearance of outbreaks has proven inevitable in places where measures have been relaxed. COVID-19 immunisation has decreased transmission in certain countries; however, these are few nations and the spread of new variants makes testing as important as before. Test, Trace and Isolate have been essential to halt SARS-CoV-2 infection. Non-PCR tests such as lateral flow tests have proven useful particularly in the case of symptomatic testing [1]; however, large scale PCR-based testing is essential to contain and prevent outbreaks due to its high sensitivity. This is particularly relevant in asymptomatic individuals and should also be central in implementing an ‘exit strategy’ plan.

In order to increase testing capacity, many countries rely on centralised efforts to build large diagnostic centres. However, the involvement of smaller academic or commercial laboratories has proven helpful and necessary too [2-5]. These decentralised laboratories can repurpose existing equipment and technical expertise for SARS-CoV-2 testing, for example by comparing methods of extraction vs extraction-free methods or samples treated with heat [6-11], combining heat with proteinase K treatments to improve sensitivity [12] or establishing sensitivity of primer-probe pairs [13,14]. The UK government document “Guidance for organisations to seek supporting the COVID-19 testing programme” published on the 9th of April 2020, by the Department of Health and Social Care, clearly welcomed academic institutions to increase testing capacities within the UK, referred to here as NHS-helper labs. However, due to global high demand of the kits and reagents used in the WHO, CDC (Centres for Disease Control, US), ECDC (European Centre for Disease Prevention and Control) and PHE (Public Health England) ratified testing strategies, the NHS-helper labs were encouraged to use alternative strategies that would not interfere with the reagent demand of larger testing facilities. Moreover, helper laboratories could provide their research expertise and experimental validation of other kits enabling clinical labs to benefit from their results. We set out to perform this task.

Here we describe different strategies for SARS-CoV-2 PCR-based detection by employing reagents that are not currently used in the NHS setting. We also performed heat inactivation employing dry bead baths of SARS-CoV-2, for both the original variant and the more recent alpha (B 1.1.7) variant and present data supporting good limit of detection (LoD) for both variants after heat treatment. Within the UK, the NHS agrees on the use of alternative RNA isolation and qPCR protocols, providing these have been internally validated and discussed with the local NHS partner. To increase visibility of these alternative strategies, we have created a webpage under the Open Science Framework platform (https://osf.io/uebvj/) that we hope will stimulate exchange between smaller laboratory facilities, increase confidence in tested alternative routes of RNA isolation and viral RNA detection and thereby expedite the establishment of smaller academic testing centres.

Materials and methods

All materials with their catalogue numbers are available at https://osf.io/uebvj/.

Heat inactivation

Swab tubes containing Viral Transport Medium (VTM) were checked for cracks to ensure no viral material had leaked. Swab tubes were then transferred to a water bath (Grant) containing dry metallic beads (Starlab) preheated to 70°C or 90°C, ensuring the entire swab tube (including lid) was covered by the beads. Samples were incubated in the following conditions: 70°C for 10 mins, 70°C for 30 mins, 90°C for 10 mins, or 90°C for 30 mins, then transferred back to Class I MSC and allowed to cool to room temperature prior to RNA extraction.

RNA extraction

Serial dilutions were done in Hank balanced salt solution (HBBS) and 1% Bovine Serum Albumin (BSA) to closely mimic viral transport media.

Qiagen QIAamp Viral RNA Mini Kit

From swab tube, 140 μl sample was transferred to 1.5 mL screw-cap microcentrifuge tube and treated with 560 μl AVL, containing carrier RNA, followed by 560 μl molecular-grade 100% ethanol (Fisher Scientific). Samples were then taken out of the Class I MSC and CL-3 lab as AVL is known to inactivate SARS-CoV-2, transferred into QIAamp mini spin columns (Qiagen) and centrifuged according to manufacturer’s instructions. Two wash steps were performed, with 714 μl buffer AW1 and 714 μl buffer AW2 (both Qiagen). RNA was then eluted from the columns with 40 μl RNase-free water (Ambion), followed by a second 40 μl elution to maximise RNA yield and giving a final RNA sample volume of 80 μl.

Beckman Coulter Agencourt RNAdvance Blood Total RNA Kit

Reagents were prepared prior to RNA extraction according to manufacturer’s instructions. The protocol was conducted in a Class I MSC in a CL-3 lab. From a swab tube, 140 μl were transferred to a Zymo-Spin I-96 Plate (Zymo Research). 7 μl of Proteinase K/PK buffer and 105 μl of Lysis buffer was added to each sample, mixed and incubated at room temperature for 15 minutes. Following incubation, 143 μl of Bind1/Isopropanol was added to each sample, mixed, and the samples were left to incubate at room temperature for 5 min. The Zymo-Spin I-96 Plate was placed on ZR-96 MagStand (Zymo Research), and the magnetic beads left to form a pellet. The supernatant was removed, and the magnetic beads washed three times, first, with 280 μl of Wash buffer (Beckman Coulter), followed by two washes with 70% ethanol. Following the wash steps, RNA was eluted from the columns with 80 μl RNase-free water (Ambion).

Omega Bio-tek Mag-Bind® Viral DNA/RNA kit

Reagents were prepared prior to RNA extraction according to manufacturer’s instructions. The protocol was conducted in a Class I MSC in a CL-3 lab. From a swab tube, 140 μl sample was transferred to a Zymo-Spin I-96 Plate (Zymo Research). 369.5 μl of Lysis mastermix was added to each sample, mixed, and incubated at room temperature for 10 minutes. Following incubation, 7 μl of Mag-Bind® Particles CNR and 7 μl of Proteinase K solution was added to each sample, mixed and incubated at room temperature for 5 minutes. The Zymo-Spin I-96 Plate was placed on ZR-96 MagStand (Zymo Research), and the magnetic beads left to form a pellet. The supernatant was removed, and the magnetic beads washed three times, first, with 280 μl of VHB buffer (Omega Bio-tek), followed by two washes with 350 μl SPR Wash Buffer (Omega Bio-tek). Following the wash steps, RNA was eluted from the columns with 80 μl RNase-free water (Ambion).

One-step RT-qPCR

qPCRBIO Probe 1-Step Go Lo-ROX (PCR Biosystems)

Reactions were done with 5 μL RNA, 5 μL 2x qPCRBIO Probe 1-Step Go mix, 1.2 μL forward primer RdRP_SARSr-F2 (10 μM), 1.6 μL reverse primer RdRP_SARSr-R1 (10 μM), and 0.2 μL probe RdRP_SARSr-P2 (10 μM), 2 μL of 20x RTase Go, and completed with RNase-free water to 20 μL. The samples were incubated in a QuantStudio 5 (Applied Biosystems/ThermoFisher Scientific). Reverse transcription was performed for 10 minutes at 45°C. The DNA polymerase was activated for 2 minutes at 95°C and the samples underwent 50 cycles of denaturation (5 seconds at 95°C) and annealing/extension (30 seconds at 60°C). A plate read was included at the end of each extension step. Each sample was run in duplicate.

TaqMan Fast Virus 1-Step Master Mix (Applied Biosystems)

Reactions were performed with 5 μL RNA, 5 μL TaqMan Fast Virus 1-Step master mix, with probes and water making the 20 μL reaction. For Charité/WHO/PHE primers, 1.2 μL forward primer RdRP_SARSr-F2 (10 μM), 1.6 μL reverse primer RdRP_SARSr-R1 (10 μM), and 0.2 μL probe RdRP_SARSr-P2 (10 μM), and 7 μL RNase-free water were used. For CDC primers (EUA kit IDT), 1.5 μL of each primer-probe premixture (N1, N2 or RNAseP) and 8.5 μL water were used. The samples were run in a QuantStudio 5 (Applied Biosystems/ThermoFisher Scientific) using the “Fast” cycling mode. Reverse transcription was performed for 5 minutes at 50°C. The reverse-transcriptase was then inactivated for 20 seconds at 95°C and the samples underwent 50 cycles of denaturation (3 seconds at 95°C) and annealing/extension (30 seconds at 60°C). A plate read was included at the end of each extension step. Each sample was run in duplicate except for Fig 5C and 5D where singlets (mimicking testing) were employed.

Fig 5

Primer comparisons and heat inactivation of nasopharyngeal swab samples.

(A) A different set of six positive samples were used to compare directly non treated with heat treated at 70°C for 30 min. RNA extraction was done with QIAamp and RT-qPCR with N1 primers and NEB Luna mix. (B) Three additional positive samples were subjected to different temperatures and incubation times as indicated, with RNA extracted by QIAamp. All three primer-probe sets from panel A were used, together with Taqman FastVirus master mix. (C) 30 additional samples, 14 from positive donors (1–14) and 16 from negative donors (15–30) were used to compare directly non treated with heat treated at 70°C for 30 min. RNA extraction was done with Beckman and RT-qPCR with N1 primer-probes and Luna Master Mix. Paired t-test was employed to compare the effect of heat. (D) 93 additional samples were non treated or heat treated at 90°C for 10 min. RNA extraction was done with Beckman and RT-qPCR with N1, N2 and RNAseP primer-probes and FastVirus Master Mix. Samples were run in singlets.

Luna Universal Probe One-Step RT-qPCR (NEB)

Reactions were performed with 5 μL RNA, 10 μL 2x Luna Universal Probe One-Step reaction mix, 1 μL Luna WarmStart RT enzyme mix, 1.5 μL of each CDC primer-probe premixture (N1, N2 or RNAseP), and 2.5 μL RNase-free water. The samples were incubated in a QuantStudio 5 (Applied Biosystems/ThermoFisher Scientific) using the “Fast” cycling mode. Reverse transcription was performed for 10 minutes at 55°C. The samples were denatured for 1 minute at 95°C and then underwent 50 cycles of denaturation (10 seconds at 95°C) and annealing/extension (30 seconds at 60°C). A plate read was included at the end of each extension step. Each sample was run in duplicate except for Fig 5C where singlets (mimicking testing) were employed.

Primer and probe sequences are supplied in Supporting Information. S1 and S2 Tables in S1 File show the volume reaction and cycling conditions.

Virus work

Original SARS-CoV-2 Strain England 2 (England 02/2020/407073) was obtained from Public Health England and 2 lineage B 1.1.7 (VOC 2 2020212/01) was kindly provided by W. Barclay (Imperial College London). The infectious virus titre was determined by plaque assay in Vero-E6 cells. Limit of detection was 40 plaque forming units (pfu)/mL. Experiments were performed n = 3.

Vero-E6 cells were kindly provided by W. Barclay (Imperial College London). Cells were maintained in complete DMEM GlutaMAX (Gibco) supplemented with 10% foetal bovine serum (FBS; Gibco), 100 U/mL penicillin and 100μg/mL streptomycin and incubated at 37°C with 5% CO2.

Study approval

This study was approved by Guy’s and St Thomas’ NHS Trust, REC Ref 18/NW/0584; and Service Delivery for King’s College Hospital.

Statistical analysis

Normality was firstly assessed prior to performing either paired t-tests (parametric) or Wilcoxon matched-pairs signed rank test in Figs comparing two variables. Serial dilutions in Figs 2A, 3B and 4B were analysed employing a semi-log regression to calculate the coefficient of determination (R2). Data in Fig 2C was analysed using a Shapiro-Wilk test for normality assessment prior to analysis employing ANOVA (parametric data) or Friedman test (non-parametric data).

Fig 2

Comparison between different RNA extraction methods.

(A) Dilution curve of the positive control provided by IDT (plasmid containing SARS-CoV-2 N gene) using N1 or N2 primer-probe sets with the Taqman FastVirus mix. A semi-log regression was used to calculate the coefficient of determination (R2). (B, C) A set of four swab samples were used for RNA extraction with the indicated kit. RT-qPCR was run with N1 and N2 primer-probe sets employing the FastVirus (B) or the Luna Master mixes (C). Shapiro-Wilk test was used for normality assessment prior to analysis employing ANOVA (parametric data) or Friedman test (non-parametric data). These samples were previously classified as positive (CPS) by the diagnostics lab, the number indicates different donors. Dots represent each individual RT-qPCR technical duplicate, line connects average of replicates and statistics were performed in average duplicates.

Fig 3

Sensitivity of qPCR detection by serial dilutions of extracted RNA or swab samples.

(A) RNA from sample CPS83 was serially diluted and extracted with three different methods. RT-qPCR was run with N1 and N2 primer-probe sets with Luna Master Mix. (B) Three distinct positive swab samples (CPS) were serially diluted followed by RNA extraction by the indicated method. RT-qPCR was run with N1 and N2 primer-probe sets with Luna Master Mix. Dots represent each individual technical duplicate. A semi-log regression was used to calculate the coefficient of determination (R2).

Fig 4

Sensitivity of qPCR detection by serial dilutions of viral stocks of the new B.1.1.7 variant.

B.1.1.7 SARS-CoV-2 viral stocks were serially diluted in viral transport medium, extracted employing Beckman and assessed employing N1 and N2 primer-probe sets using the FastVirus Master Mix. Samples were heat treated for 30 minutes with either 70°C (A) or 90°C (B). Dots represent the mean of the qPCR technical duplicates. A semi-log regression was used to calculate the coefficient of determination (R2). (C). Results of plaque assays for heat treatment of cultured SARS-CoV-2 B.1.1.7 variant (n = 3).

Results and discussion

Our pipelines are adaptable for both manual and automatic handling; we also employed heat inactivation of virus within the swabs for easier processing. We compared three RNA extraction methods, one column-based and two magnetic beads-based that can be automatized. As a benchmark, we used the QIAamp Viral RNA Mini Kit (QIAGEN) as their proprietary buffer AVL inactivates SARS-CoV-2 according to CDC guidelines. We also validated three different one-step RT-qPCR kits. We used the CDC recommended N1 and N2 primer-probe sets [15] and compared these against RdRP_SARSr primers [16]. We did not test efficiency of the reverse transcription (RT) step, as we had no access to in vitro transcribed RNA. For these validations we received clinical swab material from St Thomas’ Hospital and King’s College Hospital (London, UK) and compared our results with their diagnostics clinical pipelines. Negative swabs were diagnosed as such in the clinical laboratories (not pre-pandemic). Detailed step-to-step standard operating procedures (SOPs) can be found at https://osf.io/uebvj/.

Comparison of RNA extraction kits

We have created a flowchart of the different processing steps and combinations in our pipeline (Fig 1), which we subsequently explain in more detail.

Fig 1

Representation of our workflow.

We employed heat inactivation vs non heat inactivation [17]; compared three different RNA extraction kits (blue) followed by three RT-qPCR mixes and three sets of primers (green).

To test the efficiency and detection range of the CDC-recommended N1 and N2 primer-probes, we amplified serial dilutions of plasmids encoding the N SARS-CoV-2 gene (positive controls provided by Integrated DNA Technologies, IDT) using the TaqMan™ Fast Virus 1-Step Master Mix (Fig 2A), FastVirus hereafter. Good linearity could be achieved up to 10 copies of DNA molecules. Using the N1 and N2 primer-probes, we compared the efficiency of RNA recovery between the column-based QIAamp Viral RNA Mini Kit (QIAGEN, QIAmp herein) endorsed by the CDC, and two magnetic bead extraction kits: the RNAdvance Blood (now RNAdvance Viral) (Beckman hereinafter) and Mag-Bind Viral DNA/RNA 96 Kit (Omega Bio-tek, Omega herein), starting from the same material (140 μL). RNA isolation from four different coronavirus positive samples (CPS) with all three kits rendered comparable cycle thresholds (Cts) when amplified with the primer-probes N1 and N2. This was the case for two different RT-qPCR Master Mixes, FastVirus (Fig 2B) or Luna® Universal Probe One-Step RT-qPCR Kit (NEB, Luna hereafter) (Fig 2C).

Comparison of one-step RT-qPCR kits

We also compared different one-step RT-qPCR kits to amplify swab material purified using the QIAamp viral RNA mini kit which we considered our ‘benchmark’ given CDC guidelines on buffer AVL inactivating SARS-CoV-2. S1 Fig in S1 File shows RNA from ten different positive donors amplified with Luna, FastVirus and qPCRBIO Probe 1-Step Go Lo-ROX (PCR Biosystems, qPCRBio). All Master Mixes detected comparable amounts of RNA using primer-probes against N1 primer-probe, with the exception of donor CPS_101 which had borderline Ct values of 38 in both FastVirus and Luna and was undetectable using qPCRBio Master Mix (ANOVA p = 0.1278). Tukey’s multiple comparisons showed Luna performing better than FastVirus (p adjusted = 0.007) with no samples missed by FastVirus. Thus, three different RNA extraction kits, and three different one-step RT-qPCR kits achieve almost comparable detection of viral RNA within swab material.

As a diagnostic assay, it is paramount to be able to detect very low viral loads in swab samples. To determine the efficiency of the RT-qPCR we serially diluted the RNA from a confirmed positive swab (CPS83) isolated with each one of the three different kits used in this study. All dilutions were assessed with the N1 and N2 primer-probe amplification employing the Luna Master Mix. Fig 3A shows that the RT-qPCR reaction remained linear over a 105-fold RNA dilution range. To ensure that the RNA from samples with low viral loads could be reliably extracted with each one of these extraction methods, we prepared serial dilutions of swab material from three different CPS donors in Hank’s Balanced Salt Solution (HBSS) + 1% bovine serum albumin (BSA) since viral transport medium contains these only with the addition of amphotericin and gentamicin. Viral RNA was isolated from these diluted swabs with the three RNA isolation kits from Fig 2. Fig 3B shows that all three kits recovered viral RNA over a wide range of concentrations, with the N gene being reliably amplified with the N1 and N2 primer-probe sets and the Luna Master Mix. CPS21 10−1 dilution was excluded from the r calculations. The N2 primer-probe set in the donor CPS79 extracted with Omega showed poor linearity, possibly related to initial variation in the non-diluted sample.

Heat inactivation comparison

One major limitation for many academic and commercial laboratory settings is the lack of available CL-3 laboratory space and/or Class I MSCs required to handle/open the potentially infectious swabs. Moreover, samples with high viral load pose a risk of infection for the handler. Heat treatment of viral particles has been shown effective in inactivating SARS-CoV-2 with 70°C 5min treatment rendering viral infectivity undetectable employing Vero E6 cells (limit of detection of TCID50 assay was 100 TCID50/mL) [18-20]. Other heat treatment protocols have also been demonstrated, with variable effects on PCR sensitivity [21]. We set out to establish the effect of heat inactivation on the sensitivity of SARS-CoV-2 detection.

We firstly assessed different temperature and time conditions for heat treatment of both SARS-CoV-2 original and B 1.1.7 variants. The novel variant of SARS-CoV-2 B.1.1.7 was originally described in December 2020 in the UK, firstly detected in samples as early as 20th September 2020 [22]. Since then, it has spread to many other countries where it is the predominant variant, together with the recent delta. We thus evaluated the effect of heat on detecting of B.1.1.7. We performed serial dilutions of cultured virus in viral transport medium, extracted RNA employing Beckman and assessed the presence of N using the N1 and N2 primer-probe combinations. Fig 4 shows that heat at either 70°C (Fig 4A) or 90°C (Fig 4B) for 30 minutes did not alter the detection of viral RNA (red vs blue). Fig 4C and S2A Fig in S1 File show that treatment of B.1.1.7 with 70°C or 90°C for 10 and 30 mins inactivates this variant of SARS-CoV-2 virus. S2B Fig in S1 File shows inactivation of the original variant at all temperatures and times (limit of detection 40 pfu/mL). Importantly, we observed a reduction in infectivity, and not inactivation (according to our 40 pfu/mL limit), when temperatures between 60 and 70 degrees were applied for 30 min (S3 Fig in S1 File), making the use of well-calibrated thermometers or use of higher temperatures/longer time periods critical.

We then assessed if heat treating nasopharyngeal swab material could be a method of treating samples within their original unopened collection tubes without compromising RT-qPCR results. Data shown were obtained employing QIAamp RNA extraction. We first assessed heating sample aliquots at 70°C for 30min with different samples and performed RT-qPCR with N1 primer-probes and Luna Master Mix. As Fig 5A shows, we did not observe any change in Ct values upon heat treatment of the sample (t-test p = 0.1946).

We then set out to test a wider range of heat-inactivation conditions on six confirmed positive samples using two extraction methods. We treated aliquots of the same sample with no heat, 70°C for 10min, 70°C for 30min, 90°C for 10min or 90°C for 30min and extracted RNA employing the QIAamp kit. We employed a dry metallic bead bath to heat the sample tubes, since water baths are not allowed in Cat-3 laboratories due to the risk of spillage. Our results showed that none of the heat conditions altered the Ct values (ANOVA for N1 p = 0.3656 and Friedman test for N2 p = 0.3469, Fig 5B). Both the N1 and N2 primer-probe sets gave reliable and near-identical amplification of viral RNA; however, we noticed that the RdRP primer-probe set failed to amplify viral samples with high Ct values. These results of high Ct values for RdRP were also observed when we used the MagMax kit (ThermoFisher Scientific) as an extra RNA extraction method (S4 Fig in S1 File). Comparison of the different primer-probe combination for RdRP rendered similar results (S5 Fig in S1 File) and as previously shown [13].

To confirm the reproducibility of our results, we employed another distinct set of samples, assessing both positive and negative samples. We aliquoted swab material, warmed it at 70°C for 30min, extracted their RNA using QIAmp and performed RT-qPCR using Luna Master Mix using primer-probe N1. Ct values did not change upon heat inactivation as observed previously (t-test p = 0.5578). S6 Fig in S1 File shows the data for N2 and RNAseP. We detected one previously diagnosed sample as negative (sample 27) in which we could amplify N1 at a Ct of 37.489, which is at the limit of detection. All our water controls (no template and water template) yielded no amplification. RNAse P controls are in S6 Fig in S1 File.

To further test a higher temperature we employed 90°C for 10min in 93 samples, extracted their RNA using Beckman and performed RT-qPCR using FastVirus Master Mix using primer-probes for N1, N2 and RNAseP. Fig 5D shows the results for those samples where we detected positive amplification in either of the treatments (88 for N1 and 90 for N2). More samples were lost upon heating, although statistical analysis of Ct ranges remained similar between heat vs non-heat (S3 Table in S1 File). N2 primer-probe appeared more heat-resistant, as 89 samples were detected vs 88 with N1 primer-probes. Interestingly, we observed no clear trend for samples with high Ct values lost upon heat treatment (S7 Fig in S1 File).

Our work shows different workflows for nasopharyngeal swab processing for SARS-CoV-2 detection employing different combinations of inactivation, extraction and amplification. We present data for the validation of two viral RNA purification kits (Beckman and Omega) as alternatives to the QIAamp viral RNA mini kit. We have also tested three alternative, commercially available one step RT-qPCR kits (FastVirus, Luna and PCRBio) and assessed different recommended primer-probe sets (N1, N2, RdRP) which can currently detect all circulating SARS-CoV-2 variants to date.

With regards to extraction methods, our data in Fig 2 suggest that Beckman performed better as seen for sample CPS_84. This is also supported by data in Fig 3 where CPS_21 and CPS_79 samples did not show good linearity when diluted and extracted with Omega. When analysed together, we did not observed any statistically significant difference between Luna, FastVirus and PCRBio, although one sample was not detected employing PCRBio and Luna appeared to perform better than FastVirus (S1 Fig in S1 File). The difference in Ct values between Luna and FastVirus was very small and did not alter overall sensitivity (mean Ct difference 0.8335).

We found that N1 and N2 primer-probe are more sensitive than RdRP (Fig 5B), as reported by others [13]. This may likely be due to higher amount of sub-genomic RNAs encoding for N [23]. With regards to inactivation, most recommended viral inactivation protocols use a combination of guanidinium thiocyanate (GTC) and Triton X-100. GTC became scarce due to its wide use, it is quite toxic and not compatible with some kits, and the use of chemical inactivation protocols after sample collection inherently requires opening of the swab sample, with the consequent risk of exposure for lab staff. Our data show that heat, which is an economic and fast way of inactivating the virus, inactivates the original and B 1.1.7 variants at 70°C or 90°C for 10 and 30 mins (Fig 4 and S2 Fig in S1 File). Importantly, we observed that high viral concentrations using inadequate temperatures between 60 and 70°C were not fully inactivated (S3 Fig in S1 File). We observed this in early experiments where were relied on a heat block thermometer, which was proven inaccurate and set at a temperature of ~62–63°C instead. This highlights the need to accurately measure temperatures when performing inactivation of clinical samples–or the use of temperatures above 70°C. Importantly, temperatures of the swabs must be considered when performing heat inactivation, since swabs kept in the fridge will take longer to reach a certain temperature vs swabs kept at room temperature. Our data also highlight that adequate titrations employing high viral loads of SARS-CoV-2 are required to establish if full inactivation has been achieved. 70°C during 30 min appeared to have no effect on sensitivity (Fig 5C) while 90°C during 10 min appeared to decrease sensitivity (Fig 5D). Our data supports heating at temperatures below 90°C, method that may be used to reduce the need for CL-3 laboratory and to speed up sample processing given the chemical inactivation methods are labour-intensive increasing the risk of exposure to the lab staff. Some laboratories have implemented dry heat in ovens [4] to inactivate samples; we propose the use of dry heat with beads as it allows for both high and low throughputs and is safe against possible sample leaks (beads can be disinfected at the moment of leakage). Our pipeline can therefore be implemented in places that only have CL-2 facilities to detect SARS-CoV-2. Our results differ from those observed by others with regards to samples with low viral loads being lost upon heating [21] since samples with low viral loads were still detected and some samples were only detected upon heat treatment (S7 Fig in S1 File). Our higher temperatures employed may possibly denature RNAses and/or facilitate viral RNA denaturation while preserving enough integrity for detection with the N1 and N2 primer-probe sets. Regardless, we advise the use of 70°C over 90°C when possible.

Together these data highlight the need for performing cross-validations of RNA extraction kits and primer-probe pairs prior to implementing in diagnostics, with an emphasis on the need of using clinical samples (rather than diluted RNA or plasmid DNA) to establish ‘real-world’ data that better relate to clinical samples. Swab material and inherent inhibitors will perform variably with different workflows and we thus highlight the need to assess their performance–a task in which diagnostics laboratories can collaborate with academic institutions to speed up the establishment of new protocols. Moreover, establishing limits of detection at each laboratory purchasing international standards such as those provided by the WHO, or viral cultures as directed and established by their Local Health Authorities, is essential to ensure good practice and implementation of diagnostics.

Conclusions

Based on the above, and understanding that including RT-qPCR duplicates may decrease the number of samples a diagnostic laboratory can process (particularly if employing 96 well plates), we suggest to:

employ heat (70°C preferably to 90°C) for 10-30min. Ensure temperature is at least 70°C;

preferably employ N1 and N2 primer-probes vs RdRP;

test samples without RT-qPCR technical replicates to increase the testing throughput;

run duplicates in case of borderline ≥36Ct [24] and always check amplification curves of samples. If 1) amplification is shown reproducibly consider it a positive sample with low viral load 2) amplification unclear (one replicate positive, one negative) for these donors to be re-tested as soon as possible to confirm positive or negative detection of SARS-CoV-2 regardless of symptoms. Although we acknowledge the limitations, if possible, re-swabbing individuals with unclear or discrepant results is highly recommended as a first option.

Supporting information contains Supporting information for materials and methods, S1-S7 Figs and S1-S3 Tables.

(PDF)

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21 Apr 2021

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10 Jun 2021

PONE-D-21-12761

Resilient SARS-CoV-2 diagnostics workflows including viral heat inactivation

PLOS ONE

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1. Is the manuscript technically sound, and do the data support the conclusions?

The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented.

Reviewer #1: No

Reviewer #2: Yes

Reviewer #3: Yes

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2. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: N/A

Reviewer #2: Yes

Reviewer #3: Yes

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3. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Yes

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4. Is the manuscript presented in an intelligible fashion and written in standard English?

PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.

Reviewer #1: Yes

Reviewer #2: Yes

Reviewer #3: Yes

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5. Review Comments to the Author

Reviewer #1:

Lista et al. pursue two main goals with this study. On the one hand they evaluate and compare different extraction methods and PCR chemistry and primersets, on the other they try to validate heat-inactivation protocols for SARS-CoV-2 in clinical samples. The title is not well chosen and does not represent what is actually done in this work. This manuscript has been reviewed before by other reviewers and revised by the authors.

I will address a few general points about this study:

1. Claims about “complete inactivation” of SARS-CoV-2 by heat inactivation in this study are not supported by the data. I am missing a lot of details in the methods about the exact methodology used for cell culture experiments, as well as the number of repeats carried out for the experiments, etc.. In general, claims of “complete inactivation” of infectivity are held to very high standards and there are good studies in the literature about other viruses that show the amount of scrutiny necessary to guarantee that a certain procedure will with certainty abolish all infectivity in samples (e.g. Smither et al. “Buffer AVL Alone Does Not Inactivate Ebola Virus in a Representative Clinical Sample Type.” JCM). There have been good studies about this for SARS-CoV-2 in the past, see e.g. Pastorino et al. “Evaluation of heating and chemical protocols for inactivating SARS-CoV-2”. Instead of declaring complete inactivation, authors should state a reduction of infectivity, in relation to their limit of detection in plaque forming units.

2. The authors assess sensitivity of different extraction workflows by serial dilution of clinical samples, with duplicate measurements, essentially performing hit-rate analysis without a quantitative reference and without giving detection limits (figure3). Additionally they perform serial dilution with plasmids as quantitative reference (figure2). All experiments should be described more in detail in the methods (incl. repeats performed).

In general, the methodology is inadequate to evaluate analytical performance of molecular methods. I would expect use of a quantitative standard (i.e. Qnostics, Acculex, etc, or WHO-Standard, which is now available for SARS-CoV-2) for serial dilutions near LoD with 8-21 repeats for each step and determination of LoD by probit analysis (log regression, 95% probability of detection)

The use of DNA-plasmids to simulate positive material of an RNA virus is several decades outdated and absolutely unacceptable. The authors state that they bought the plasmids from IDT; they could just as well have bought synthetic RNA, or done IVT on their DNA target, or bought a commercial reference standard, as stated above.

3. Why did authors choose to compare N1/N2 and Corman’s RdRp assay? It had been known very early that the Charite RdRp is a poor assay (i.e. Vogels et al.’s preprint on comparing LDTs in April 2020, later published in nature microbiology), making the comparison a foregone conclusion. Corman’s E-Sarbeco is substantially better and more commonly used for SCoV-2 detection, making it a fairer competitor, however, these kinds of experiments have already been done and published a year ago.

Authors state that the RdRp assay will not detect subgenomic RNA, however, the E-gene is also abscent in the vast majority of subgenomic RNA (Alexandersen et al. 2020, Nature communications). To avoid detecting subgenomic RNA altogether, there are other RdRp assays that perform very well like e.g. Chan et al.’s RdRp/Hel-assay (Chan et al. 2020 JCM).

4. The number of samples for workflow comparison (largely in fig5) is too small to draw valid conclusions (e.g. 10 positive and 10 negative for extraction workflow comparison). Clinical validation experiments such as this should aim at 100 samples minimum; value of the dataset increases with more samples.

5. Fig.5C raises some questions, I believe this is what Reviewer#1 was referring to in their final question. Right now, it looks as if negative samples were positive for CDC-N1 in roughly 50% of measurements. If this is the case, this experiment should be discarded and repeated after decontamination. CDC-N1 does not produce unspecific false-positives. (Note: Oligos from IDT were consistently contaminated with N1/2 and E-Sarbeco positive material from the start of the pandemic until early Fall 2020. Authors state that they buy oligos from IDT.)

6. As alluded to in previous points, the methods section should be expanded with details about analytical (fig2-4) and clinical (fig5) evaluation and cell culture experiments. Right now, it contains details about execution of individual extraction and PCR-protocols. These could be moved to supplement or replaced with “carried out according to manufacturer’s instructions” where applicable. Some of the details I am referring to are mentioned in the figure legend, but the figure legend is an addition and does not replace the methods section.

On a general note, this manuscript gives the impression of containing two largely separate projects, one of them being the inactivation aspect, and the other being analytical and clinical validation of different extraction methods. Both are executed haphazardly and strung together in a somewhat confusing fashion. When looking at the general topics, I see clear value in the comparison of different extraction workflows, as these studies are instrumental for diagnostic labs to make informed decisions about which products to employ for SARS-CoV-2 detection. The heat-inactivation topic has been published extensively in early/mid 2020 and this work does not provide anything valuable to the existing literature. My recommendation would be to restructure the manuscript and focus largely on the validation data, add more datapoints to the existing experiments and either remove the inactivation topic entirely or reducing it merely to impact on PCR performance without making any claims about efficacy of inactivation.

Reviewer #2:

In the present study Lista et al. performs a detailed comparison of 3 commercial RNA extraction kits, 3 different RT-aPCR mixes and 3 separate target genes, for the detection of SARS-CoV-2 virus in nasopharyngeal swaps collected during the current SARS-CoV-2 pandemic. Moreover, Lista et al. assess the impact of heat-inactivation of the virus (to reduce the safety requirements) on subsequent virus detection.

In general, the revised manuscript is well written, the results are clearly presented, and the methods described in sufficient details. From their study Lista et al. shows:

• The 3 tested commonly used commercial RNA extraction kits provided similar results.

• The 3 RT-qPCR mixes tested, showed no significant differences

• Detection of the N gene was more reliable than that of RdRP (as published elsewhere)

• Heat-inactivation of the virus sample specimens have limited impact on the sensitivity of the RT-qPCR for detection of SARS-CoV-2 in clinical specimens.

The manuscript in its revised form includes the appropriate statistical tests that together with the additional results included in the revision clearly support the authors conclusions.

While mass testing for SARS-CoV-2 using large-scale kit-free procedures has been implemented in most industrialized countries, the use of small-scale kit-based protocols/work flows still may be useful in settings where access to highly advanced and automated systems are limited. As such, I find the results presented by Lista et al. an important contribution to the field.

Reviewer #3:

The manuscript presents some open protocols for SARS-CoV-2 detection intended to help diagnostic in cases of lack of reagents or equipments, to non-experienced laboratories or to laboratories with no access to diagnostic kits. The goal is interesting and might be very useful to many laboratories around the world. Interestingly, the authors provide standard operating procedures to all their methods through a web repository. However, the manuscript might be improved at several points:

- The continuous narrative style of the Results section makes difficult to follow what is being tested in each section, adding subsection titles might be helpful.

- Authors should not use the term strain to refer to B.1.1.7 variant or to the so called original strain. Moreover, the term original strain is not very informative, was it an EU1 isolate? Was it an earlier isolate? In any case the authors are not using strains, they are using clinical isolates belonging to one or another variant.

- Why do the authors use RdRp probe and primers set from the Charité protocol and not the gene E or gene N probe and primer sets? RdRp is known to be the less reliable among the Charité set. Moreover RdRp is not used systematically, but only in the last heat-inactivation section, so probably it might be better to delete it from the work.

- The authors use RNAseP to control sample quality, but these controls are mentioned only at the end of the Results section. Authors should introduce them in the first paragraph of the Results, when they introduce the primers and probe sets.

- What is the difference between Fig. 3A and Fig. 3B? If there is no difference it might be better to include both in a single panel.

- The heat inactivation experiments are an important part of the manuscript, yet they are shown as supplementary material. The main text shows a table, Fig. 4C, summarizing the results of the experiments shown in Supp. Fig. 2A, but in Fig. 4C the reader cannot appreciate the difficulties in evaluating Supp. Fig. 2A. The difference in the native controls between the original variant and the B.1.1.7 variant is huge. The authors state in the figure legend that B.1.1.7 plaques are smaller, but the fact is that plaques are seen only in the -1 dilution. Given the critical value of these data to assess safety more convincing images should be provided.

- First paragraph in page 5 are not results. Better move it to discussion section.

- In Fig. 5 the authors state that extractions in these experiments were done by QIAamp, this means that all the samples had been inactivated with AVL buffer and this should be stated in a more explicit manner in the text and the figure legend, otherwise the labels "Not inactivated" and "Heat Inactivated" may be misleading because all the samples treated with AVL are inactivated.

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Reviewer #1: Yes: Dominik Nörz

Reviewer #2: No

Reviewer #3: No

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27 Jul 2021

Point by Point answers

Reviewer #1:

Lista et al. pursue two main goals with this study. On the one hand they evaluate and compare different extraction methods and PCR chemistry and primersets, on the other they try to validate heat-inactivation protocols for SARS-CoV-2 in clinical samples. The title is not well chosen and does not represent what is actually done in this work. This manuscript has been reviewed before by other reviewers and revised by the authors.

I will address a few general points about this study:

1. Claims about “complete inactivation” of SARS-CoV-2 by heat inactivation in this study are not supported by the data. I am missing a lot of details in the methods about the exact methodology used for cell culture experiments, as well as the number of repeats carried out for the experiments, etc.. In general, claims of “complete inactivation” of infectivity are held to very high standards and there are good studies in the literature about other viruses that show the amount of scrutiny necessary to guarantee that a certain procedure will with certainty abolish all infectivity in samples (e.g. Smither et al. “Buffer AVL Alone Does Not Inactivate Ebola Virus in a Representative Clinical Sample Type.” JCM). There have been good studies about this for SARS-CoV-2 in the past, see e.g. Pastorino et al. “Evaluation of heating and chemical protocols for inactivating SARS-CoV-2”. Instead of declaring complete inactivation, authors should state a reduction of infectivity, in relation to their limit of detection in plaque forming units.

Thank you for the helpful suggestion, we agree with the reviewer and have included the limit of detection of our plaque assays in the Figure. We performed our assays n=3 times and we have now included that information too.

2. The authors assess sensitivity of different extraction workflows by serial dilution of clinical samples, with duplicate measurements, essentially performing hit-rate analysis without a quantitative reference and without giving detection limits (figure3). Additionally they perform serial dilution with plasmids as quantitative reference (figure2). All experiments should be described more in detail in the methods (incl. repeats performed). In general, the methodology is inadequate to evaluate analytical performance of molecular methods. I would expect use of a quantitative standard (i.e. Qnostics, Acculex, etc, or WHO-Standard, which is now available for SARS-CoV-2) for serial dilutions near LoD with 8-21 repeats for each step and determination of LoD by probit analysis (log regression, 95% probability of detection). The use of DNA-plasmids to simulate positive material of an RNA virus is several decades outdated and absolutely unacceptable. The authors state that they bought the plasmids from IDT; they could just as well have bought synthetic RNA, or done IVT on their DNA target, or bought a commercial reference standard, as stated above.

We performed our quantitative serial dilutions employing cultured virus (Figure 4) to assess linearity, in addition to showing dilution of clinical swabs which we agree reflect much better the quality of the data vs plasmids. While we appreciate that commercial supplies can be purchased to this end, there are examples in the literature that show linearity of performance using cultured virus [1]. We have included a phrase to make the reader aware in our Discussion: ‘Moreover, establishing limits of detection at each laboratory purchasing international standards such as those provided by the WHO, or viral cultures as directed and established by their Local Health Authorities, is essential to ensure good practice and implementation of diagnostics’. Our manuscript aims to determine resilient protocols and thus to provide guidance and data performance on reagents/protocols that are available in molecular laboratories. Testing laboratories may decide in accordance with the Local Health Authority to establish their limit of detection employing either in house viral cultures or international standards, which are required regardless of what a manufacturer of IVD CE products may provide as evidence. Beyond these comments and suggestions, we believe performing these experiments is out of the scope of our work.

3. Why did authors choose to compare N1/N2 and Corman’s RdRp assay? It had been known very early that the Charite RdRp is a poor assay (i.e. Vogels et al.’s preprint on comparing LDTs in April 2020, later published in nature microbiology), making the comparison a foregone conclusion. Corman’s E-Sarbeco is substantially better and more commonly used for SCoV-2 detection, making it a fairer competitor, however, these kinds of experiments have already been done and published a year ago. Authors state that the RdRp assay will not detect subgenomic RNA, however, the E-gene is also abscent in the vast majority of subgenomic RNA (Alexandersen et al. 2020, Nature communications). To avoid detecting subgenomic RNA altogether, there are other RdRp assays that perform very well like e.g. Chan et al.’s RdRp/Hel-assay (Chan et al. 2020 JCM).

We chose RdRp as it is still employed in several commercially available kits such as Primer Design or Viasure. We believe that adding more data to current literature does not harm our manuscript; if anything it shows reproducibility between laboratories, an essential task in diagnostics. With regards to detection of genomic/subgenomic RNA the implications for clinical management are not yet established, to our knowledge, and thus we only added the phrase as a comment.

4. The number of samples for workflow comparison (largely in fig5) is too small to draw valid conclusions (e.g. 10 positive and 10 negative for extraction workflow comparison). Clinical validation experiments such as this should aim at 100 samples minimum; value of the dataset increases with more samples.

We agree that more samples increase the power of the study and this is why in Figure 5D we provide nearly 100 samples as comparison. We have also modified Figure 5C including 14 positives and 16 negatives. The extraction workflow comparison was done employing serial dilutions to demonstrate the linearity of the extraction process, which we believe is the right approach and already shows differences when comparing extraction methods in our dataset.

5. Fig.5C raises some questions, I believe this is what Reviewer#1 was referring to in their final question. Right now, it looks as if negative samples were positive for CDC-N1 in roughly 50% of measurements. If this is the case, this experiment should be discarded and repeated after decontamination. CDC-N1 does not produce unspecific false-positives. (Note: Oligos from IDT were consistently contaminated with N1/2 and E-Sarbeco positive material from the start of the pandemic until early Fall 2020. Authors state that they buy oligos from IDT.)

Thank you for this observation. We can demonstrate that we did not have contamination (water negative controls as explained in our previous Supplemental Figure 6). We agree with the Reviewer that this panel added confusion; our intention was to present all our data as we obtained it. The Ct threshold that laboratories employ depends on the manufacturer and internal validations. In our case, we presented detection and Ct value, which was in most cases very high and thus may be deemed as negative in some settings. Having said this, we have repeated the comparison in a new Figure 5C and Supplemental Figure 6 with new samples, including 14 positives and 16 negatives.

6. As alluded to in previous points, the methods section should be expanded with details about analytical (fig2-4) and clinical (fig5) evaluation and cell culture experiments. Right now, it contains details about execution of individual extraction and PCR-protocols. These could be moved to supplement or replaced with “carried out according to manufacturer’s instructions” where applicable. Some of the details I am referring to are mentioned in the figure legend, but the figure legend is an addition and does not replace the methods section.

Thank you for this observation, we have now expanded the methods section as helpfully pointed out by the Reviewer. We preferred to leave the methods section with the detailed information as we provide certain tips about each protocol, which we have also uploaded as Standard Operating Procedures in the Open Science Framework for everyone’s access.

On a general note, this manuscript gives the impression of containing two largely separate projects, one of them being the inactivation aspect, and the other being analytical and clinical validation of different extraction methods. Both are executed haphazardly and strung together in a somewhat confusing fashion. When looking at the general topics, I see clear value in the comparison of different extraction workflows, as these studies are instrumental for diagnostic labs to make informed decisions about which products to employ for SARS-CoV-2 detection. The heat-inactivation topic has been published extensively in early/mid 2020 and this work does not provide anything valuable to the existing literature. My recommendation would be to restructure the manuscript and focus largely on the validation data, add more datapoints to the existing experiments and either remove the inactivation topic entirely or reducing it merely to impact on PCR performance without making any claims about efficacy of inactivation.

We thank the reviewer for the thoughtful comments and suggestions on the structure of our manuscript. We believe that our protocols add value to the current needs of increasing testing capacity in light of the rapid expansion of new variants such as the delta variant now sweeping the UK and other countries. We believe that our heat inactivation methods add value to existing literature as we provide details about how to perform them and specific temperatures and timings, with validation about specificity and sensitivity. We have provided new data on Figure 5C and modified the text to correctly point out inactivation according to limit of detection, as well as new sub-headings which we believe help reading and structuring our manuscript. Our aim is to provide a plethora of different protocols for the users to combine the steps that best suit their setting and needs.

Reviewer #2:

In the present study Lista et al. performs a detailed comparison of 3 commercial RNA extraction kits, 3 different RT-aPCR mixes and 3 separate target genes, for the detection of SARS-CoV-2 virus in nasopharyngeal swaps collected during the current SARS-CoV-2 pandemic. Moreover, Lista et al. assess the impact of heat-inactivation of the virus (to reduce the safety requirements) on subsequent virus detection.

In general, the revised manuscript is well written, the results are clearly presented, and the methods described in sufficient details. From their study Lista et al. shows:

• The 3 tested commonly used commercial RNA extraction kits provided similar results.

• The 3 RT-qPCR mixes tested, showed no significant differences

• Detection of the N gene was more reliable than that of RdRP (as published elsewhere)

• Heat-inactivation of the virus sample specimens have limited impact on the sensitivity of the RT-qPCR for detection of SARS-CoV-2 in clinical specimens.

The manuscript in its revised form includes the appropriate statistical tests that together with the additional results included in the revision clearly support the authors conclusions. While mass testing for SARS-CoV-2 using large-scale kit-free procedures has been implemented in most industrialized countries, the use of small-scale kit-based protocols/work flows still may be useful in settings where access to highly advanced and automated systems are limited. As such, I find the results presented by Lista et al. an important contribution to the field.

We are very grateful to the Reviewer for their comments on our manuscript and thrilled they deem it appropriate and of value to the field – this is certainly our intention when we performed all experiments.

Reviewer #3:

The manuscript presents some open protocols for SARS-CoV-2 detection intended to help diagnostic in cases of lack of reagents or equipments, to non-experienced laboratories or to laboratories with no access to diagnostic kits. The goal is interesting and might be very useful to many laboratories around the world. Interestingly, the authors provide standard operating procedures to all their methods through a web repository. However, the manuscript might be improved at several points:

- The continuous narrative style of the Results section makes difficult to follow what is being tested in each section, adding subsection titles might be helpful.

We thank the Reviewer and have now added subtitles as suggested which we think help follow the manuscript

- Authors should not use the term strain to refer to B.1.1.7 variant or to the so called original strain. Moreover, the term original strain is not very informative, was it an EU1 isolate? Was it an earlier isolate? In any case the authors are not using strains, they are using clinical isolates belonging to one or another variant.

We apologise for the mistake; indeed these are different variants and come from different clinical isolates. We have now corrected ‘strain’ with ‘variant’. Thank you for spotting this.

- Why do the authors use RdRp probe and primers set from the Charité protocol and not the gene E or gene N probe and primer sets? RdRp is known to be the less reliable among the Charité set. Moreover RdRp is not used systematically, but only in the last heat-inactivation section, so probably it might be better to delete it from the work.

We tested the RdRp probe and primers to establish their performance and compare them to the more commonly used CDC N probes and primers. There are some commercial kits that still employ RdRp and, in line with the work of others (Vogels et al., 2020), we observe a decrease in sensitivity which we think important to report.

- The authors use RNAseP to control sample quality, but these controls are mentioned only at the end of the Results section. Authors should introduce them in the first paragraph of the Results, when they introduce the primers and probe sets.

We thank the reviewer for the suggestion; as we only present the data in the last Figure we have decided to leave the explanation there.

- What is the difference between Fig. 3A and Fig. 3B? If there is no difference it might be better to include both in a single panel.

Thank you for asking this, as we realised we did not explain this well in the text, which we have now hopefully clarified. Figure 3A employed one donor’s RNA that was extracted with different kits and serially diluted. Figure 3B shows original material from 3 donors serially diluted in viral transport medium, extracted and then assessed by RT-qPCR. The latter aims to establish linearity about the extraction methods.

- The heat inactivation experiments are an important part of the manuscript, yet they are shown as supplementary material. The main text shows a table, Fig. 4C, summarizing the results of the experiments shown in Supp. Fig. 2A, but in Fig. 4C the reader cannot appreciate the difficulties in evaluating Supp. Fig. 2A. The difference in the native controls between the original variant and the B.1.1.7 variant is huge. The authors state in the figure legend that B.1.1.7 plaques are smaller, but the fact is that plaques are seen only in the -1 dilution. Given the critical value of these data to assess safety more convincing images should be provided.

We thank the Reviewer for considering this an important part of our manuscript. Our B.1.1.7 experiments have an inherent limitation since the viral stocks available had lower pfu/mL than the original UK02 variant. Moreover, B.1.1.7 has less fitness in cultured Vero E6 as compared with UK02 – this has previously been observed by Prof Wendy Barclay’s group in their pre-print work [2] and shows smaller plaque assays, which we demonstrate down to dilution -2.

- First paragraph in page 5 are not results. Better move it to discussion section.

We thank the Reviewer for the suggestion. We have now moved the central part of this paragraph to discussion and only left a small section to introduce our datasets, which we believe now reads better.

- In Fig. 5 the authors state that extractions in these experiments were done by QIAamp, this means that all the samples had been inactivated with AVL buffer and this should be stated in a more explicit manner in the text and the figure legend, otherwise the labels "Not inactivated" and "Heat Inactivated" may be misleading because all the samples treated with AVL are inactivated.

We thank the reviewer for this observation and agree- we have now modified the labels to ‘Not Treated’ and ‘Heat Treated’.

References

1. Nörz D, Frontzek A, Eigner U, Oestereich L, Wichmann D, Kluge S, et al. Pushing beyond specifications: Evaluation of linearity and clinical performance of the cobas 6800/8800 SARS-CoV-2 RT-PCR assay for reliable quantification in blood and other materials outside recommendations. Journal of Clinical Virology. 2020;132:104650. doi: https://doi.org/10.1016/j.jcv.2020.104650.

2. Brown JC, Goldhill DH, Zhou J, Peacock TP, Frise R, Goonawardane N, et al. Increased transmission of SARS-CoV-2 lineage B.1.1.7 (VOC 2020212/01) is not accounted for by a replicative advantage in primary airway cells or antibody escape. bioRxiv. 2021. doi: https://doi.org/10.1101/2021.02.24.432576

Submitted filename: Response to Reviewers.pdf

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17 Aug 2021

Resilient SARS-CoV-2 diagnostics workflows including viral heat inactivation

PONE-D-21-12761R1

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25 Aug 2021

PONE-D-21-12761R1

Resilient SARS-CoV-2 diagnostics workflows including viral heat inactivation

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