# Assessment of inactivation procedures for SARS-CoV-2

Heidi Auerswald, Sokhoun Yann, Sokha Dul, Saraden In, Philippe Dussart, Nicholas J. Martin, Erik A. Karlsson, Jose A. Garcia-Rivera

► To cite this version:

Heidi Auerswald, Sokhoun Yann, Sokha Dul, Saraden In, Philippe Dussart, et al. Assessment of inactivation procedures for SARS-CoV-2. Journal of General Virology, 2021, 102 (3), pp.001539. 10.1099/jgv.0.001539. pasteur-03221742

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# Assessment of inactivation procedures for SARS-CoV-2

Heidi Auerswald¹, Sokhoun Yann¹, Sokha Dul², Saraden In¹, Philippe Dussart¹, Nicholas J. Martin³, Erik A. Karlsson¹,* and
Jose A. Garcia-Rivera²,*

## Abstract

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the causative agent of coronavirus disease 2019 (COVID-19), presents a challenge to laboratorians and healthcare workers around the world. Handling of biological samples from individuals infected with the SARS-CoV-2 virus requires strict biosafety measures. Within the laboratory, non-propagative work with samples containing the virus requires, at minimum, Biosafety Level-2 (BSL-2) techniques and facilities. Therefore, handling of SARS-CoV-2 samples remains a major concern in areas and conditions where biosafety for specimen handling is difficult to maintain, such as in rural laboratories or austere field testing sites. Inactivation through physical or chemical means can reduce the risk of handling live virus and increase testing ability especially in low-resource settings due to easier and faster sample processing. Herein we assess several chemical and physical inactivation techniques employed against SARS-CoV-2 isolates from Cambodia. This data demonstrates that all chemical (AVL, inactivating sample buffer and formaldehyde) and heat-treatment (56 and 98 °C) methods tested completely inactivated viral loads of up to 5 log₁₀.

## INTRODUCTION

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the causative agent of coronavirus disease 2019 (COVID-19) has rapidly spread across the world. On 30 January 2020, the World Health Organization (WHO) declared the outbreak a Public Health Emergency of International Concern and upgraded it to a pandemic on 11 March 2020 [1]. As of 20 August 2020, there have been over 22.2 million laboratory-confirmed cases and greater than 780000 deaths reported globally [2]. Extensive testing is necessary to ensure accurate diagnosis, contact tracing and mitigate SARS-CoV-2 spread through isolation and quarantine procedures. Additionally, extensive testing facilitates the global public health response against COVID-19, providing critical information regarding the effectiveness of mitigation efforts.

Given the limited specific treatment options and the lack of an approved vaccine, SARS-CoV-2 isolates need to be handled according to strict biosafety measures [3, 4]. Extreme care in handling live samples prevents occupational exposure and requires extensive technical training and appropriate

containment devices wearing recommended personal protective equipment (PPE). Therefore, the need for extensive testing in areas and conditions where biosafety for specimen handling is difficult to maintain remains a major concern. Indeed, scenarios still exist globally where there is need to sample outside of designated testing centres, conduct field investigations in difficult locales, transport non-secure samples, and, especially, testing being conducted in resource strained, underequipped and/or under-maintained laboratories. Inactivation through physical or chemical means reduces the risk from handling live samples and increases testing ability especially in low-resource settings due to easier and faster sample processing.

Cambodia is a tropical, resource poor, least developed country (LDC) in Southeast Asia with a large socio-economic dependence on tourism [5]. Cambodia is also a major hotspot of endemic and emerging infectious disease [6]. One particular, but not unique, issue faced in LDCs is the expansion of testing capacity due to a scarcity of testing laboratories, especially in remote provincial health centres. Therefore, SARS-CoV-2

Received 19 August 2020; Accepted 25 November 2020; Published 08 January 2021

**Author affiliations:** ¹Virology Unit, Institut Pasteur du Cambodge, Institut Pasteur International Network, Phnom Penh, Cambodia; ²Naval Medical Research Unit TWO, Phnom Penh, Cambodia; ³Naval Medical Research Unit TWO, Singapore.

*Correspondence: Erik A. Karlsson, ekarlsson@pasteur-kh.org; Jose A. Garcia-Rivera, jose.a.garciarivera.mil@mail.mil

**Keywords:** biosafety; COVID-19; chemical inactivation; heat inactivation; SARS-CoV-2.

**Abbreviations:** AVL, viral lysis buffer; BSL-2, biosafety Level 2; COVID-19, coronavirus disease 2019; CPE, cytopathic effect; Ct, cycle threshold; DMEM, Dulbecco's modified Eagle medium; FBS, fetal bovine serum; GITC, Guadinium thiocyanate; LDC, least developed country; MERS, Middle East Respiratory Syndrome coronavirus; PEG, Polyethylene glycol; p.i., post-inoculation; PPE, personal protective equipment; RT-PCR, reverse transcription polymerase chain reaction; SARS, Severe Acute Respiratory Syndrome coronavirus; SARS-CoV-2, Severe acute respiratory syndrome coronavirus 2; TCID50, tissue culture infectious dose 50%; USCDC, United States Centers for Disease Control; WHO, World Health Organization. 001539

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samples from these rural health centres requires safe but rapid transportation to designated testing sites. Aside from active training and great care when handling live specimens from suspected cases, transportation of potentially infectious material requires increased protective equipment and packaging, often in reduced supply or of poor quality, even in the best of scenarios. Therefore, simple and effective inactivation of suspected samples that can be conducted onsite can greatly decrease risk of exposure during transportation, handling and testing, as well as reduce demand for protective equipment and supplies at a current global scarcity.

Herein, we evaluated the efficiency of various thermal and chemical inactivation methods on SARS-CoV-2 utilizing three separate SARS-CoV-2 isolates collected in Cambodia as part of the national surveillance and response to determine their effect on viral infectivity and RNA integrity tested via real-time reverse transcription polymerase chain reaction (RT-PCR).

# METHODS

## Ethics statement

All isolates were obtained as part of the first-line testing, analysis and preparedness of suspected cases through the national outbreak response. No primary clinical specimens or identifying information or any other individual-specific information was utilized in these studies or manuscript.

## Cell lines

African green monkey kidney cells (Vero; ATCC CCL-81) were used for the isolation and culture of SARS-CoV-2 isolates. Vero E6 cells were used for the titration of infectious virus via TCID₅₀. Both cell lines were cultivated in Dulbecco's modified Eagle medium (DMEM; Sigma-Aldrich, Steinheim, Germany) supplemented with 10% foetal bovine serum (FBS; Gibco, Gaithersburg, MD, USA) and 100 U ml⁻¹ penicillin-streptomycin (Gibco) at 37 °C and 5% CO₂ atmosphere.

## Virus culture and titration

Three SARS-CoV-2 isolates (designated hCoV-19/ Cambodia/1775/2020, 1775; hCoV-19/Cambodia/2018/2020, 2018; and, hCoV-19/Cambodia/2310/2020, 2310) were obtained and cultured in Vero cells. Upon infection with SARS-CoV-2 the culture medium was replaced by infection medium containing DMEM, 5% FBS, antibiotics, 2.5 µg ml⁻¹ Amphotericin B (Gibco) and 16 µg ml⁻¹ TPCK-trypsin (Gibco). Virus-containing supernatants, as determined by the presence of cytopathic effect (CPE), were harvested 6 days after infection by centrifugation at 1500 r.p.m. for 10 min. The concentration of viable virus was measured by TCID₅₀ assay on Vero E6 cells in 96-well plates (TPP, Trasadingen, Switzerland) [7]. Briefly, serial dilutions of viral culture supernatant were inoculated onto cells using infection medium. After 4 days of incubation, plates were inactivated with 4% formaldehyde for 20 min then stained with 1% crystal violet solution in PBS for 20 min. Titre of viable virus was calculated applying the Spearman-Karber formula [8].

## Inactivation of SARS-CoV-2 isolates

Inactivation was performed in triplicate using 140 µl aliquots of SARS-CoV-2 isolates (1775, 2018, and 2310; passage 3 from Vero cells). Chemical inactivation was performed following manufacturers' recommendation and included: (i) adding 560 µl viral lysis buffer (AVL buffer including carrier RNA; AVL buffer) from the QIAamp Viral RNA Mini Kit (Qiagen, Hilden, Germany) for 10 min at room temperature according to the manufacturer's recommendations; (ii) 200 µl inactivating sample buffer (GeneReach, Taichung City, Taiwan) containing 50% guanidinium thiocyanate (GITC) and 6% t-Octylphenoxypolyethoxyethanol (Triton X-100) for 15 min at room temperature; or (iii) 140 µl 4% Formaldehyde in PBS (General Drugs House, Bangkok, Thailand; end concentration 2%) for 15 min at room temperature. Thermal inactivation similarly performed on 140 µl aliquots of fresh virus culture: (iv) 56 °C for 30 min; (v) 56 °C for 60 min; and (vi) 98 °C for 2 min in a thermo-block (Eppendorf, Hamburg, Germany). Sterile DMEM treated in the similar methods served as negative controls, and untreated SARS-CoV-2 isolates as positive controls. All samples and controls were handled in 1.5 ml PP reaction tubs (Greiner bio-one, Kremsmünster, Austria) during the treatment.

## Analysis for viable virus post-inactivation

To determine if any viable virus remained post-inactivation, 50% Polyethylene glycol 8000 (Sigma-Aldrich, St. Louis, USA) in PBS was added (1/5 of total sample volume) to an aliquot from each sample condition and incubated overnight at 4 °C. Following incubation, the virus was recovered by centrifugation at 1500 r.p.m. for 1 h. Precipitates were washed twice with sterile PBS, re-constituted with infection medium, and used for infecting the TCID₅₀ on Vero E6 cells and recovery cultures on Vero cells. TCID₅₀ and virus cultures plates were analysed for CPE 7 days post-inoculation (p.i.). Negative controls were treated the same way to examine cytotoxicity of possible remaining traces of inactivation solutions.

## SARS-CoV-2 real-time RT-PCR

Following inactivation, RNA from one aliquot per condition per virus isolate and negative control was immediately extracted with the QIAamp Viral RNA Mini Kit (Qiagen) and stored at -80 °C until further processing. Real-time RT-PCR assays for SARS-CoV-2 RNA detection were performed in duplicate using the Charité Virologie algorithm (Berlin, Germany) to detect both E and RdRp genes [9]. In brief, real-time RT-PCR was performed using the SuperScript III One-Step RT-PCR System with Platinum Taq DNA Polymerase (Invitrogen) on the CFX96 Touch Real-Time PCR Detection System (BioRad, Hercules, CA, USA) ABI. Each 25 µl reaction mixture contained 5 µl of RNA, 3.1 µl of RNase-free water, 12.5 µl of 2X PCR buffer, 1 µl of SuperScriptTM III RT/ Platinum Taq Mix, 0.5 µl of each 10 µM forward and reverse primer, and 0.25 µl of probe (E_Sarbeco_P1 or RdRP_SARSrP2) using the following thermal cycling conditions; 10 min at 55 °C for reverse transcription, 3 min at 94 °C for PCR initial activation, and 45 cycles of 15 s at 94 °C and 30 s at 58 °C.
<table><caption>**Table 1.** SARS-CoV-2 isolates used for inactivation</caption><thead><tr><th>SARS-CoV-2 isolate</th><th>Before treatment TCID<sub>50</sub> ml<sup>-1</sup></th><th>After PEG precipitation (positive control) TCID<sub>50</sub> ml<sup>-1</sup></th><th>Post inactivation (all methods) TCID<sub>50</sub> ml<sup>-1</sup></th></tr></thead><tbody><tr><td>1775</td><td>2.11E+05</td><td>4.10E+04</td><td>Not detected</td></tr><tr><td>2018</td><td>1.19E+04</td><td>6.09E+03</td><td>Not detected</td></tr><tr><td>2310</td><td>6.67E+05</td><td>1.22E+05</td><td>Not detected</td></tr></tbody></table>

## Statistical analysis

Statistics were performed using GraphPad Prism for Windows, version 7.02 (GraphPad Software, La Jolla, CA, USA). Analysis of variance was performed comparing mean Ct values for each inactivation method. Difference between standard (AVL) and each specific inactivation method was determined using Dunnett's test for many-to-one comparison. A P-value of less than 0.05 was considered to indicate statistical significance. Agreement, including bias and 95% confidence interval, between Ct values following inactivation by AVL and other methods was assessed using a method described by Bland and Altman [10]. The mean Ct value of AVL and the other inactivation method assessed was plotted on the X-axis. The difference between the two values was plotted on the Y-axis. Cut-off values of 2 and -2 are plotted.

## RESULTS

### Inactivation efficiency

All chemical and thermal inactivation methods resulted in the reduction of viable SARS-CoV-2 to undetectable levels. Untreated virus isolates had a concentration of viable virus up to $6.67 \times 10^5$ (isolate 2310) before treatment (Table 1). Therefore, the reduction of viable virus across inactivation levels was at least $5 \log_{10}$. Precipitation of virus and complete removal of inactivation solution before infecting Vero E6 cells

for TCID$_{50}$ titration ensured that cell death was not induced by chemical products used in the inactivation procedure. Successful recovery of virus post-PEG precipitation was determined by RT-PCR on the same samples used for TCID$_{50}$. All attempts to recover viable virus post-inactivation on Vero cells were unsuccessful up to day 6 p.i.

### Effect of inactivation procedure on RT-PCR

As all samples were incubated with AVL buffer, this condition serves as the standard for the comparison of the diverse inactivation methods. There were significant differences between the Ct values for the RdRp (ANOVA; *P*<0.0001) and E (ANOVA; *P*<0.0001) genes. Following many-to-one comparison of AVL to all other forms of inactivation used in this study (Fig. 1), only formaldehyde inactivation was significantly different for the RdRp (Dunnet's test; *P*=0.0016) and E (Dunnet's test; *P*=0.0007) genes. In order to demonstrate the agreement in Ct values for the inactivation methods compared to the standard AVL, Bland-Altman plots were compared. Samples inactivated by formaldehyde were the only ones where the absolute bias Ct value for all samples was greater than two compared to AVL for RdRp (–20.32±1.75) and E (–19.80±1.17) genes. All other inactivation methods resulted in absolute bias Ct values of less than one except for the RdRp gene following inactivation at 56°C for 30 min (–1.15±1.08), though this was still within the limits of agreement.

## DISCUSSION

Following the rapid global spread of SARS-CoV-2 and the need for universal testing, more and more individuals are exposed to live virus samples, thereby increasing the chances of occupational infection. The WHO and United States Centers for Disease Control (US CDC) have released laboratory guidelines to mitigate risk of exposure during diagnostic and research procedures [3, 4]. Despite recommendations for handling within contained biosafety cabinets,

<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Comparison of Ct values of SARS-CoV-2 (a) E gene and (b) RdRp gene for three isolates (1775; circles, 2018; squares, 2310; triangles) inactivated by different methods. Inactivation with 2% formaldehyde for 15 min at room temperature results in significantly elevated Ct values for both genes (***P=0.0001, one-way ANOVA comparison to AVL inactivation).</figcaption></figure>
individuals working with these samples are still required to
handle potentially live virus at the initial steps of acquiring
and preparing the suspected samples prior testing, thereby
increasing the risk of exposure. Potential exposure greatly
increases in situations requiring large numbers of samples
to be processed under harsh conditions, in underequipped
or poorly maintained laboratories, and even within the
sample transportation system, such as found in developing
or rural areas of the world. Therefore, the continued need for
COVID-19 testing worldwide requires utilization of simple
and effective inactivation techniques.

Previous studies have been conducted on the effectiveness of
chemical inactivation techniques on SARS-CoV-2 [11–16],
the majority of these based on the experiences with infectious
agents of concern such as Ebola [17] and SARS and MERS
coronaviruses [18]. As with other viruses, the primary step in
the molecular detection of SARS-CoV-2 is viral lysis to begin
the extraction of nucleic acids. The buffers used in this lysis
step yield varying results [11, 13, 16, 17, 19, 20]; however, the
study by Pastorino et al. [11] found that AVL buffer alone
was successfully able to fully inactivate up to 4–5 log₁₀ of
virus from three different primary isolates of SARS-CoV-2.
Other studies using higher vial litres in their inactivation
experiments [11, 13, 16] observed only incomplete inactiva-
tion with AVL buffer alone nevertheless confirming the 5–6
log₁₀ decrease of infectivity. Apart from differences in isolates
utilized and a slight reduction in titre, it is unclear as to the
reasons why AVL buffer fully inactivated in this study versus
others, but further work is warranted to determine the exact
effectiveness of this step alone. Moreover, the impact of the
inactivation method on various clinical specimen should be
investigated as we utilized virus stock for this study, and the
complex composition of clinical samples might influence the
efficacy of inactivation as well as virus detection.

Inactivating sample transport media, either made in-house or
commercially available, also presents an attractive way to inac-
tivate samples at the point of sampling to ensure safe handling
along the transport chain and within the laboratory. These
inactivating transport media include the key components of
many viral lysis buffers including chaotropic agents (GITC),
detergents (Triton X-100) and buffering agents (EDTA, Tris-
HCL) to inactivate a preserve viral RNA. Previous studies
have shown that GITC-lysis buffers like AVL are able to
inactivate SARS-CoV-2 samples [11–13, 16]; however, the
addition of Triton-X may be necessary for complete inactiva-
tion [11, 16]. In line with these studies, commercial sample
transport media containing both GITC and GITC/Triton-X
buffers were successfully able to inactivate up to 5 log₁₀ of
virus with no loss of molecular diagnostic sensitivity.

Apart from sample media and buffers utilized for diagnostic
testing, various disinfectant and inactivating chemicals are avail-
able for sample treatment. Formaldehyde has a long history of
use for inactivation against a number of viruses and in a number
of fixation techniques, including vaccine preparations [21, 22].
Formaldehyde has been shown to successfully inactivate both
SARS and MERS [18, 23, 24] and has been suggested to be a

viable alternative for disinfection and inactivation of SARS-
CoV-2 [13–15, 23, 24]. Formaldehyde treatment (end concentra-
tion 2%, 15 min at room temperature) did successfully inactive
up to 5 log₁₀ of virus; however, this treatment severely impacted
viral detection in subsequent molecular testing. This decreased
detection is not unexpected as formaldehyde treatment results
in RNA degradation and modification [25]. Therefore, formal-
dehyde treatment does not appear to be a solution for increased
molecular SARS-CoV-2 testing; however, it does remain a viable
alternative for sample inactivation or disinfection.

Perhaps the most studied technique thus far regarding SARS-
CoV-2 has been thermal inactivation at various times and
temperatures [11, 14, 16, 26–29]. Several previous studies have
shown heat to be an effective inactivation technique against other
coronaviruses, including SARS, MERS and human seasonal
strains [18, 27, 30]. Similar to previous studies, 56 °C heat treat-
ment for 30 or 60 min was fully able to inactivate up to 5 log₁₀ of
SARS-CoV-2 from three different isolates [11, 15, 16, 26, 31, 32].
Interestingly, while other studies utilized 95 °C for 5 to 10 min
for inactivation, heat treatment at 98 °C for only 2 min was also
able to completely inactivate up to 5 log₁₀ of virus. These results
are very promising as high heat treatment is extremely rapid and
may be a vital addition to the testing arsenal, as RT-PCR can
possibly be performed directly from these samples without the
need for nucleic acid extraction [33, 34]. It has to be considered,
that we used small volumes (140 µl) of virus culture samples
and inactivation of larger volumes and clinical specimen might
be more difficult. It was already shown that heat-inactivation
decreases the sensitivity for N and ORF gene RT-PCRs in throat
swab samples with low viral load [35, 36] and nasopharyngeal
swabs [37]. Interestingly, the shortened time period of high heat
treatment may mitigate some of the reduction in detection seen
in previous studies and make this technique more employable
[11, 16]. This high heat treatment might be also favourable as
specimen with high vial load (6 log₁₀ TCID) remain infectious
even after treatment at 56 °C for 30 min and 60 °C for 60 min [16]

Overall, the agreement and retained sensitivity amongst RT-PCR
results, combined with the fact that all methods resulted in 100%
virus inactivation up to a viral load of 5 log₁₀, suggests that any
of the tested methods, except formaldehyde, are useful to inac-
tivate SARS-CoV-2 samples. Given the WHO recommendation
to ‘test, test, test’, these data can help to optimize sample inactiva-
tion for austere or remote areas. Indeed, it may be possible to
use basic tools such as a stopwatch and boiling water to achieve
100% virus inactivation without compromising sample integ-
rity, significantly decreasing possible exposure during sample
transportation and handling, allowing for dissemination of
testing to labs with decreased biosafety capacity, and possibly
reducing the global demand for a dwindling supply of PPE.

Funding information

This work, including that of Dr Auerswald, was supported, in part, by
internal funding through Institut Pasteur in Cambodia. Dr Karlsson
is supported, in part, by the U.S. Department of Health and Human
Services, Office of the Assistant Secretary for Preparedness and
Response (Grant No. 1 IDSEP190051-01-00) and through internal
funding at Institut Pasteur in Cambodia. The findings and conclusions
in this report are those of the authors and do not necessarily represent the official views of the US Department of Health and Human Services. This work was also partially funded and supported by the Armed Forces Health Surveillance, Branch Global Emerging Infections Surveillance Section.

## Acknowledgements

We would like to acknowledge the patients, Cambodian Ministry of Health and Cambodian Center for Disease Control, Rapid Response Team members, and the doctors, nurses, and staff of the reference hospitals in Cambodia for their response to the COVID-19 pandemic. We thank all of the technicians and staff at Institut Pasteur du Cambodge in the Virology Unit and at NAMRU-2 in the Molecular Unit involved in this work.

## Conflicts of interest

The authors declare that there are no conflicts of interest.

## References

1. World Health Organization. 2020. Coronavirus disease (COVID-19) outbreak. https://www.who.int/westernpacific/emergencies/covid-19 [accessed May 13th 2020].

2. Dong E, Du H, Gardner L. An interactive web-based dashboard to track COVID-19 in real time. *Lancet Infect Dis* 2020;20:533–534.

3. United States Centers for Disease Control and Prevention. Interim laboratory biosafety guidelines for handling and processing specimens associated with coronavirus disease 2019 (COVID-19) 2020.

4. World Health Organization. Laboratory testing for coronavirus disease 2019 (COVID-19) in suspected human cases: interim guidance. *2 March 2020;2020.*

5. United Nations. Un list of least developed countries 2019.

6. Burgos S, Ear S. Emerging infectious diseases and public health policy: insights from Cambodia, Hong Kong and Indonesia. *Trans-bound Emerg Dis* 2015;6:96–101.

7. Flint SJE W, Racaniello VR, Skalka AM. Virological Methods. Principles of Virology. ASM Press; 2009.

8. Ramakrishnan MA. Determination of 50% endpoint titer using a simple formula. *World J Virol* 2016;5:85–86.

9. Corman VM, Landt O, Kaiser M, Molenkamp R, Meijer A et al. Detection of 2019 novel coronavirus (2019-nCoV) by real-time RT-PCR. *Eurosurveillance* 2020;25:2000045.

10. Bland JM, Altman DG. Statistical methods for assessing agreement between two methods of clinical measurement. *Lancet* 1986;1:307–310.

11. Pastorino B, Touret F, Gilles M, Lamballerie de X, Charrel RN. Evaluation of heating and chemical protocols for inactivating SARS-CoV-2. *bioRxiv* 2020.

12. Scallan MF, Dempsey C, MacSharry J, O'Callaghan I, O'Connor PM. Validation of a lysis buffer containing 4 M guanidinium thiocyanate (GITC)/ Triton X-100 for extraction of SARS-CoV-2 RNA for COVID-19 testing: comparison of formulated lysis buffers containing 4 to 6 M GITC. *OxRxiv* 2020.

13. Welch SR, Davies KA, Buczkowski H, Hettiarachchi N, Green N et al. Analysis of inactivation of SARS-CoV-2 by specimen transport media, nucleic acid extraction reagents, detergents, and fixatives. *J Clin Microbiol* 2020;58:e01713–01720.

14. Patterson EI, Prince T, Anderson ER, Casas-Sanchez A, Smith SL et al. Methods of inactivation of SARS-CoV-2 for downstream biological assays. *J Infect Dis* 2020;222:1462–1467.

15. Jureka AS, Silvas JA, Basler CF. Propagation, inactivation, and safety testing of SARS-CoV-2. *Viruses* 2020;12:622.

16. Pastorino B, Touret F, Gilles M, de Lamballerie X, Charrel RN. Heat inactivation of different types of SARS-CoV-2 samples: what protocols for biosafety, molecular detection and serological diagnostics? *Viruses* 2020;12:735.

17. Smither SJ, Weller SA, Phelps A, Eastaugh L, Ngugi S et al. Buffer AVR alone does not inactivate Ebola virus in a representative clinical sample type. *J Clin Microbiol* 2015;53:3148–3154.

18. Darnell MER, Subbarao K, Feinstone SM, Taylor DR. Inactivation of the coronavirus that induces severe acute respiratory syndrome, SARS-CoV. *J Virol Methods* 2004;121:85–91.

19. Blow JA, Dohm DJ, Negley DL, Mores CN. Virus inactivation by nucleic acid extraction reagents. *J Virol Methods* 2004;119:195–198.

20. Ngo KA, Jones SA, Church TM, Fuschino ME, George KS et al. Unreliable inactivation of viruses by commonly used lysis buffers. *Applied Biosafety* 2017;22:56–59.

21. Sabbaghi A, Miri SM, Keshavarz M, Zargar M, Ghaemi A. Inactivation methods for whole influenza vaccine production. *Rev Med Virol* 2019;29:e2074.

22. Salk JE, Krech U, Youngner JS, Bennett BL, Lewis LJ et al. Formaldehyde treatment and safety testing of experimental poliomyelitis vaccines. *Am J Public Health Nations Health* 1954;44:563–570.

23. Henwood AF. Coronavirus disinfection in histopathology. *J Histotechnol* 2020;43:102–104.

24. Kampf G, Todt D, Pfaender S, Steinmann E. Persistence of coronaviruses on inanimate surfaces and their inactivation with biocidal agents. *J Hosp Infect* 2020;104:246–251.

25. Evers DL, Fowler CB, Cunningham BR, Mason JT, O'Leary TJ. The effect of formaldehyde fixation on RNA: optimization of formaldehyde adduct removal. *J Mol Diagn* 2011;13:282–288.

26. Batéjat C, Grassin Q, Manuguerra J-C, Leclercq I. Heat inactivation of the severe acute respiratory syndrome coronavirus 2. *bioRxiv* 2020.

27. Kampf G, Voss A, Scheithauer S. Inactivation of coronaviruses by heat. *J Hosp Infect* 2020;105:348–349.

28. Lista M, Page R, Sertkaya H, Matos P, Ortiz-Zapater E. Resilient SARS-CoV-2 diagnostics workflows including viral heat inactivation 2020.

29. Péré H, Podglajen I, Baillard J-L, Mboumba Bouassa R-S, Veyer D et al. Thermal inactivation and nucleic acid amplification-based testing for SARS-CoV-2. *J Clin Virol* 2020;131:104588.

30. Leclercq I, Batéjat C, Burguière AM, Manuguerra J-C. Heat inactivation of the middle East respiratory syndrome coronavirus. *Influenza Other Respir Viruses* 2014;8:585–586.

31. Wu Z-G, Zheng H-Y, Gu J, Li F, Lv R-L, Z-g W, H-y Z, R-I L et al. Effects of different temperature and time durations of virus inactivation on results of real-time fluorescence PCR testing of COVID-19 viruses. *Curr Med Sci* 2020;40:614–617.

32. Wang Y, Song W, Zhao Z, Chen P, Liu J et al. The impacts of viral inactivating methods on quantitative RT-PCR for COVID-19. *Virus Res* 2020;285:197988.

33. Grant PR, Turner MA, Shin GY, Nastouli E, Levett LJ. Extraction-free COVID-19 (SARS-CoV-2) diagnosis by RT-PCR to increase capacity for national testing programmes during a pandemic. *bioRxiv* 2020.

34. Fomsgaard AS, Rosenstierne MW. An alternative workflow for molecular detection of SARS-CoV-2 - escape from the NA extraction kit-shortage, Copenhagen, Denmark, March 2020. *EuroSurveill* 2020;25.

35. Pan Y, Long L, Zhang D, Yuan T, Cui S et al. Potential false-negative nucleic acid testing results for severe acute respiratory syndrome coronavirus 2 from thermal inactivation of samples with low viral loads. *Clin Chem* 2020;66:794–801.

36. Zou J, Zhi S, Chen M, Su X, Kang L et al. Heat inactivation decreases the qualitative real-time RT-PCR detection rates of clinical samples with high cycle threshold values in COVID-19. *Diagn Microbiol Infect Dis* 2020;98:115109.

37. Chen H, Wu R, Xing Y, Du Q, Xue Z et al. Influence of different inactivation methods on severe acute respiratory syndrome coronavirus 2 RNA copy number. *J Clin Microbiol* 2020;58:e00958–00920.