The current pandemic caused by novel coronavirus SARS-CoV-2, first detected in late 2019 in the province of Wuhan, China, has rapidly spread worldwide, infecting more than 10 million individuals as of 29 June 2020 [1] [2] [3] . Infection with SARS-CoV-2 can lead to development of COVID-19, a disease associated with severe acute respiratory syndrome, responsible for hundreds of thousands of deaths globally. Controlling the spread of SARS-CoV-2 relies on the ability of healthcare systems to quickly identify infected individuals, which has mainly relied on RT-qPCR for viral RNA detection 4 . International competition for commercial kits and reagents has negatively impacted the ability of many countries to scale up testing capacity to deal with the increased demand caused by rampant infection. Implementing RT-qPCR testing programs requires specialised laboratory equipment and reagents, presenting additional challenges.
In an effort to increase the diagnostic capacity for SARS-CoV-2 infection in the UK, the Francis Crick Institute, a biomedical research institute based in London, rapidly repurposed its staff and facilities in late March 2020 to serve as a clinical diagnostic testing facility through a partnership between a major local healthcare provider (University College London Hospitals National Health Services Trust) and an accredited clinical diagnostic laboratory (Health Services Laboratories, HSL), termed the CRICK COVID-19 Consortium (CCC) 5 . The pipeline utilises a series of in-house buffers to first inactivate patient samples received from care homes and hospitals, and to then extract RNA before using a CE marked commercial kit to detect SARS-CoV-2 by RT-qPCR. Patient results are reported through a custom online web portal that interfaces with the reference laboratory 5 . In order to avoid dependence on any singular testing methodology, to continue increasing testing capacity, and to provide a potential means to deliver diagnostics at the point-of-care, the CCC was also tasked with developing and validating alternative SARS-CoV-2 testing strategies.
Herein, we describe the use of loop mediated isothermal amplification PCR coupled with reverse transcription (RT-LAMP) as a robust method for SARS-CoV-2 detection in clinical specimens 6 . The entire strand displacement and amplification procedure is carried out at a single temperature in less than 30 minutes, alleviating the need for a thermocycler qPCR system. Diagnostic tests utilising this technique have been developed for RNA viruses and other pathogens [7] [8] [9] , and there are now several preprints focusing on the use of RT-LAMP to detect SARS-CoV-2 10-13 . We set up a SARS-CoV-2 RT-LAMP assay using the WarmStart Colorimetric LAMP 2X Mastermix commercialised by New England Biolabs (NEB), which allows for visual assessment of DNA amplification. Alternatively, DNA amplification can be quantified on a real-time PCR machine by complexing the reaction with the DNA dye SYTO 9 10 . We made use of primers developed by Zhang et al. targeting the nucleocapsid All rights reserved. No reuse allowed without permission.
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The copyright holder for this preprint this version posted July 1, 2020. . https://doi.org/10.1101/2020.06.29.20142430 doi: medRxiv preprint phosphoprotein (N gene) of SARS-CoV-2 10 and, in a parallel reaction, primers that detect human 18S rRNA to control for specimen quality 9 .
Our results demonstrate that within the CCC pipeline, RT-LAMP can readily replace RT-qPCR as a means for detecting SARS-CoV-2 transcripts within RNA extracted from nosethroat swabs and endotracheal secretions/bronchoalveolar lavage fluid. RT-LAMP for the N gene shows absence of non-specific amplification and cross-reactivity with other human coronaviruses or respiratory viruses, and displays a sensitivity threshold almost equivalent to the gold standard RT-qPCR. Switching to RT-LAMP translates into a ten-fold decrease in total reagent cost and a potential four-fold increase in pipeline output. Additionally, we provide preliminary data suggesting that RT-LAMP can be performed without prior RNA extraction, allowing rapid and cost-effective testing that could potentially be extended to point-of-care.
The entire workflow was validated under extended governance by public health authorities during the pandemic and inspected by a qualified UKAS assessor against GenQA guidelines to verify compliance to ISO 15189:2012 equivalent standard (US equiv. CAP/CLIA). As such, the standard operating procedure (SOP) developed here is ready to be deployed for diagnostic testing of SARS-CoV-2.
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All the samples processing through the CCC pipeline was done in accordance with SOPs described recently 5 . A redefined reaction, running, and reporting SOP can be found in the Supplemental Methods.
RT-LAMP reaction was performed in a total volume of 15 µL, mixing 7.5 µL WarmStart Colorimetric LAMP 2X Master Mix (New England Biolabs, M1800), 1.5 µL of 10X primer mix, 1.5 µL SYTO 9 Green Fluorescent Nucleic Acid Stain (ThermoFisher Scientific, S34854), 1.5 µL of Nuclease-free water (Ambion, AM9932) and 3 µL of sample unless stated otherwise.
10X SYTO 9 solution at 5 µM in nuclease free water was prepared for a final concentration of 0.5 µM in the final RT-LAMP reaction. 10X primer mix was prepared from 100 µM desalted DNA primers obtained from Sigma Custom Oligos Service. 10X primer mixes for N gene and 18S contained FIP and BIP primers at 16 µM, F3 and B3 at 2 µM, LF and LB at 4 µM. RT-LAMP was ran following a Standard Operating Procedure (cf. Appendix) on a 7500, 7500 Fast, QuantStudio 3, 5, or 7 Real-Time PCR System (Applied Biosystems). A negative no template control (NTC) and a positive control (supplied by HSL, SARS-CoV-2 clinical sample) were included on every run. Experiments utilising laboratory grown SARS-CoV-2 were performed in containment level 3 at the Francis Crick Institute (FCI) by trained personnel according to health and safety guidelines.
We found that the assembled reaction mix was unstable when kept at 4 o C for more than a few hours, and sensitive to freeze-thaw when kept at -20 o C for more than a day. The RT-LAMP should be performed using freshly prepared reaction mix. The individual components should be stored at -20 o C until use and avoid repeated freeze/thaw cycles.
As outlined recently 5 , the CCC was formed in partnership with HSL, a UCLH UKAS accredited lab, who already had a COVID-19 RT-qPCR test in scope. All samples were received and communicated by HSL under their accreditation and the CCC RT-LAMP assay was validated against their existing RT-qPCR test and the CCC's validated RT-qPCR test, which uses a CE marked commercial kit (BGI). Given the urgent timeframe required to implement testing, it was All rights reserved. No reuse allowed without permission.
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The copyright holder for this preprint this version posted July 1, 2020. . https://doi.org/10.1101/2020.06.29.20142430 doi: medRxiv preprint not possible to secure official clinical laboratory accreditation for the Francis Crick Institute.
However, full measures were taken to ensure that the CCC RT-LAMP test was evaluated, verified and performed for diagnostic use in an environment that adhered to equivalent international standards (ISO 15189:2012, US equiv. CAP/CLIA), overseen and audited by HSL. These measures were implemented under the advice and oversight of registered professionals from existing nearby ISO accredited medical laboratories, and included writing and following clinical diagnostic SOPs for every stage of the pipeline from sample reception, processing to result reporting by qualified clinical scientists prior to results being communicated to patients by HSL. Additional SOPs were followed for sample storage, disposal of materials, batch certification of reagents and incident reporting. Appropriate risk assessments, training and competency assessment procedures were established and documented. Record sheets were created to document the receipt, batch acceptance testing, and start/end of use dates for key reagents and consumables. An inventory of all key equipment was compiled which, where appropriate, included details of service and calibration records. Systems were also established for the control of all key documents (version implementation, distribution and acknowledgement), audit trailing (what samples were tested when, by whom, with what equipment and using which consumable/reagent batches), and the recording of all untoward incidents/issues (thus facilitating appropriate investigation, rectification and recurrence prevention). Samples were barcoded and tracked using the Crick Clarity library information management system (LIMS). All key documents are available at https://www.crick.ac.uk/research/covid-19/covid19-consortium. NHS governance was extended by a specific memorandum of understanding for diagnostic testing between UCLH and the Francis Crick Institute enabled by NHS England. Assurance of the pipeline was performed in collaboration with GenQA, following their checklist for non-accredited laboratories and the lab and CCC workflow were inspected by a qualified UKAS assessor against the GenQA guidelines to verify compliance to IS015189:2012 equivalent standard.
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RT-LAMP was performed with 3 µL of RNA in a total reaction volume of 15 µL unless stated otherwise. Each sample was tested in two separate reactions, one with primers targeting the N gene of SARS-CoV-2, and the second targeting human 18S rRNA to control for specimen integrity and quality ( Figure 1A ). The RT-LAMP mastermix contains a colorimetric pH indicator that turns from pink to yellow upon DNA amplification ( Figure 1B ). In addition, we benchmarked the RT-LAMP method by measuring DNA amplification using a SYBR based dye in a real-time PCR machine. When measuring fluorescence every minute ('cycle'), double stranded DNA accumulation follows a characteristic exponential amplification phase that eventually plateaus ( Figure 1C and 1D ).
An initial characterization of the technique was performed using 24 RNA samples purified from patient nasopharyngeal swabs, 12 of which were positive for SARS-CoV-2 as assessed by the CCC pipeline via RT-qPCR 5 . RT-LAMP could detect SARS-CoV-2 in the 12 positive samples with no amplification detected in all 12 negative samples, displaying 100% concordance to our current clinical diagnostic platform ( Figures 1C and 1E ). Internal control signal was detected in all samples ( Figure 1D ).
The background signal of the SARS-CoV-2 RT-LAMP assay was assessed by running RNA elution buffer (7 samples) or nuclease free water (7 samples) no template controls (NTCs) (which was not certified by peer review) is the author/funder, who has granted medRxiv a license to display the preprint in perpetuity.
The copyright holder for this preprint this version posted July 1, 2020. Figure 3B ). With the N gene RT-LAMP, no signal was observed with any of the 14 distinct virus containing specimens tested, confirming that the assay is highly specific for SARS-CoV-2 ( Figure 3B ).
In order to ascertain the sensitivity of the N gene RT-LAMP assay, RNA from a SARS-CoV-2 quantified gene copy number standard obtained from the UK National Institute for Biological Standards and Control (NIBSC) was extracted and assessed by limiting dilution.
The results indicate that the limit of reliable detection of the N gene RT-LAMP assay is between 500-1000 copies of N gene ( Figure 4A -B). This was confirmed using 10-fold serial dilutions of RNA extracted from laboratory-grown SARS-CoV-2 quantified using the NIBSC standard ( Figure 4C ). Notably, a highly linear response was observed, even in the presence of 1% Triton-X 100, which has been proposed as a means of inactivating SARS-CoV-2 14 .
The RT-LAMP assay reproducibility and precision were determined by extracting RNA 5 times from a confirmed COVID-19 positive patient sample through the CCC pipeline and assessing by N gene and 18S RT-LAMP in 5 independent experiments, performed by two different operators ( Figure 4D ). The coefficient of variation was 0.03151 for the N gene RT-LAMP analysis and 0.03595 for the 18S rRNA internal control. We also verified that RT-LAMP can be performed in a 384-well plate format using a QuantStudio 5 real-time PCR machine with equivalent results ( Figure S3 ). All rights reserved. No reuse allowed without permission.
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The RT-LAMP assay was benchmarked against the RT-qPCR methods used by the reference clinical diagnostic laboratory, HSL, and that used by the CCC validated clinical diagnostic platform by assessing 37 clinical samples processed in parallel by both laboratories with duplicate RT-qPCR analyses. RNA extracted by the CCC pipeline was then tested by RT-LAMP in duplicate runs. RT-LAMP detected 16 positives in both experiments, which was 100% concordant with results obtained by the CCC's RT-qPCR assay using a CE marked kit from BGI and HSL's N gene RT-qPCR assay. However, positives with Ct > 35 near the limit of detection (Ct = 37) of the BGI RT-qPCR assay, which were termed 'borderline positives', 
Lastly, we asked if RT-LAMP could be performed on dry swabs without the need for RNA extraction. Recent work shows that 0.5% Triton X-100 inactivates SARS-CoV-2 14 , and we observed that performing RT-LAMP with samples diluted in 1% Triton X-100 does not All rights reserved. No reuse allowed without permission.
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The copyright holder for this preprint this version posted July 1, 2020. . https://doi.org/10.1101/2020.06.29.20142430 doi: medRxiv preprint affect the sensitivity of the assay ( Figure 4C ). We reverse-engineered swab specimens by coating hospital-grade swabs in serial dilutions with HEK293T cell supernatant of SARS-CoV-2 crude suspension generated in the laboratory. Swabs were left to dry overnight, and placed in 0.5% Triton X-100 for 15-30 minutes at room temperature before assessment. The RT-LAMP test used 4.5 µL of the solubilised material to maximise the sensitivity of the assay ( Figure 6 ). In parallel, samples were processed by the CCC pipeline and interrogated by RT-qPCR with a calculated equivalent amount of RNA. 'RT-LAMP Pre' (without RNA purification) results were 94% concordant to those generated by the CCC pipeline using RT-qPCR, suggesting that RT-LAMP may potentially be performed without traditional RNA extraction.
However, the use of RT-LAMP on material from swabs without RNA extraction requires clinical validation using bona fide nasopharyngeal specimens to ascertain reliability and compatibility with various transport media.
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Rapid and reliable detection of SARS-CoV-2 is required to efficiently diagnose infected individuals and to provide governments and health systems with guidance for treatment and quarantine strategies to reduce the risk of transmission. The CCC was formed to address a critical testing shortage in the London area, especially for frontline healthcare workers, with the additional goal of rapidly validating and disseminating SOPs for others to scale up their own diagnostic programs. The majority of testing is currently performed by RT-qPCR amplification of viral RNA obtained from nasopharyngeal samples. In the face of continued global demand and competition for reagents and resources and to minimise reliance on any singular testing strategy, we have outlined here procedures to utilise RT-LAMP as a costeffective and high throughput alternative to RT-qPCR for detecting SARS-CoV-2 in clinical specimens.
Implementing this testing method would reduce the CCC's operational costs ten-fold when accounting for all consumables (data not shown), provide the ability to scale up our output further ( Figure S3 ), and decrease reporting turnaround time in comparison to our current method. We confirmed that the assay is highly specific for SARS-CoV-2 and displays lack of cross-reactivity with other respiratory viruses, including seasonal coronaviruses. Assay accuracy and robustness is denoted by an absence of false positives and the further introduction of a melt curve stage allows increased confidence in the identification and reporting of genuine positives. When performed with extracted RNA, the N gene RT-LAMP assay displays a sensitivity comparable to clinically approved RT-qPCR methods. Additionally, preliminary experiments demonstrate that RT-LAMP can be performed directly without RNA extraction, by inactivating virus-containing dry swabs in a detergent-based solution. However, these experiments did not make use of true nasopharyngeal swabs that are likely to contain material that can cause RNA degradation or inhibit the assay (unpublished observations).
Our methodology makes use of a real-time qPCR machine that measures DNA amplification using a SYBR based dye, allowing for real-time detection and standardised reporting. We did not test extensively if RT-LAMP could be assessed by the colorimetric indicator alone, although some of our data suggests that the colour change readout is concordant with the SYTO 9 dye results (Figures 1, 3 and 4) . If RT-LAMP were coupled with a colorimetric read-out and the need for RNA extraction could be obviated, the result would be a testing modality that tremendously reduces the cost and time for SARS-CoV-2 diagnostics and allow its application at point-of-care and in remote areas where sophisticated testing infrastructures currently do not exist. All rights reserved. No reuse allowed without permission.
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The copyright holder for this preprint this version posted July 1, 2020. The clinical call from the reference laboratory is indicated above the graph. (which was not certified by peer review) is the author/funder, who has granted medRxiv a license to display the preprint in perpetuity.
The copyright holder for this preprint this version posted July 1, 2020. in-house generated samples tested by RT-LAMP assay directly from dry swabs without traditional RNA extraction procedures (red squares) compared to Ct values derived from the CCC RT-qPCR BGI assay (dark and light blue dots) and the RT-LAMP assay following RNA extraction (yellow squares). The clinical call from the reference laboratory is indicated above the boxes. Ct values of 'undetermined' are plotted as "Ct = 40/25" for illustrative purposes.
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Reference left y-axis Reference right y-axis 37 All rights reserved. No reuse allowed without permission.
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The copyright holder for this preprint this version posted July 1, 2020. [4] Summary Table   0  4  8  12  16  20  24  28  32  36  40  44  48  52  56  60  64  68 All rights reserved. No reuse allowed without permission.
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