key: cord-0792242-ahu7lda5
authors: Case, James Brett; Bailey, Adam L.; Kim, Arthur S.; Chen, Rita E.; Diamond, Michael S.
title: Growth, detection, quantification, and inactivation of SARS-CoV-2
date: 2020-06-13
journal: Virology
DOI: 10.1016/j.virol.2020.05.015
sha: cb27f97386cf60885e4cfe6b2264f902b64f1d5f
doc_id: 792242
cord_uid: ahu7lda5

Severe acute respiratory syndrome coronavirus (SARS-CoV)-2 is the agent responsible for the coronavirus disease 2019 (COVID-19) global pandemic. SARS-CoV-2 is closely related to SARS-CoV, which caused the 2003 SARS outbreak but disappeared rapidly. Although numerous reagents were developed to study SARS-CoV infections, few have been applicable to evaluating SARS-CoV-2 infection and immunity. Current limitations in studying SARS-CoV-2 include few validated assays with fully replication-competent wild-type virus. We have developed protocols to propagate, quantify, and work with infectious SARS-CoV-2. Here, we describe: (1) virus stock generation, (2) RT-qPCR quantification of SARS-CoV-2 RNA; (3) detection of SARS-CoV-2 antigen by flow cytometry, (4) quantification of infectious SARS-CoV-2 by focus-forming and plaque assays; and 5) validated protocols for virus inactivation. Collectively, these methods can be adapted to a variety of experimental designs, which should accelerate our understanding of SARS-CoV-2 biology and the development of effective countermeasures against COVID-19.

Severe acute respiratory syndrome coronavirus (SARS-CoV)-2 is an enveloped virus with a single-stranded positive-sense RNA genome. Zoonotic transmission of SARS-CoV-2 from an as yet unidentified animal reservoir occurred in late 2019, resulting in human-to-human transmission by respiratory droplets that has grown into the ongoing Coronavirus disease 2019 pandemic (Wu et al., 2020b; Zhou et al., 2020; Zhu et al., 2020) . The rapid spread and relatively high case fatality rate of COVID-19 has led to an urgent need to develop diagnostics, therapeutics, and vaccines.

The SARS-CoV-2 genome is comprised of approximately 30,000 nucleotides. The first two-thirds of the genome encodes for nonstructural proteins in open reading frames 1a and 1ab that principally facilitate genome replication and viral RNA synthesis. The remaining one-third is comprised of genes encoding structural proteins such as spike (S), envelope (E), membrane (M), and nucleocapsid (N), which form the virion, and accessory proteins that regulate host cellular responses. Whole-genome phylogenetic analysis identified the SARS-like bat CoV (GenBank MG772933) as the closest known relative of SARS-CoV-2. Bats also are the reservoir host for SARS-CoV (Wu et al., 2020a) . Alignment of SARS-CoV-2 to the consensus sequence of SARS-like CoV revealed 380 amino acid differences including 27 amino acid differences in the S protein and six substitutions in the receptor binding domain (RBD) (Wu et al., 2020a) .

SARS-CoV entry is mediated by initial engagement of the RBD of the S protein with the human ACE2 receptor Li et al., 2003) , and recent studies have established that SARS-CoV-2 utilizes the same receptor for entry (Letko et al., 2020) . The S protein also is a key target for neutralizing antibodies and vaccine strategies (Rockx et al., 2008; Sui et al., 2005; Zhu et al., 2007) . Although the S protein of SARS-CoV and SARS-CoV-2 are structurally similar Walls et al., 2020; Wrapp et al., 2020) , genetically similar (Walls et al., 2020) , and use the same receptor (Lei et al., 2020; Li et al., 2003) , neutralizing anti-SARS-CoV RBD antibodies (Abs) generally lack cross-reactivity to SARS-CoV-2 (Wrapp et al., 2020) . However, polyclonal sera from mice immunized with recombinant SARS-CoV RBD protein inhibits SARS-CoV-2 infection (Walls et al., 2020) . Recent studies have identified cross-reactive, nonneutralizing monoclonal Abs (mAbs) against SARS-CoV and SARS-CoV-2, which were isolated previously using phage display or hybridoma fusion screens (Joyce et al., 2020; ter Meulen et al., 2006; Tian et al., 2020; Tripp et al., 2005; Yuan et al., 2020) . Competition binding studies show that two of these mAbs, CR3022 and 240CD, both recognize the SARS-CoV-2 RBD. A co-crystal structure revealed that CR3022 binds an epitope on the RBD distal to the binding site of ACE2 and SARS-CoV neutralizing antibodies (Yuan et al., 2020) .

SARS-CoV-2 research must be performed in a biosafety level 3 laboratory by personnel equipped with a powered air-purifying respirator (PAPR). This limitation has compelled the development of many in vitro assays that utilize heterologous pseudotyped viruses expressing the SARS-CoV-2 S protein (Lei et al., 2020; Letko et al., 2020) . However, this approach only can be used to study cellular and antibody interactions involving the S protein that principally affect attachment and entry. Here, we developed or adapted multiple methodologies to quantify SARS-CoV-2 infection in vitro using a patient isolate of SARS-CoV-2: 1) RT-qPCR quantification of viral RNA; 2) detection of viral antigen by flow cytometry; 3) focus-forming assay through immunostaining of the S protein and 4) plaque assay. We also have identified and validated chemical and heat treatment methods to inactivate replication-competent virions, which are compatible with downstream quantification assays. Together, the methodologies can be used to examine SARS-CoV-2 pathogenesis and antibody responses, and to screen for potential inhibitors of infection.

Isolates of SARS-CoV-2 from patients or animals often need to be propagated to generate high-titer virus stocks. We have tested several cell types and found African Green Monkey cell lines and derivatives thereof to be most permissive to SARS-CoV-2 infection. These include Vero-CCL81 (ATCC-CCL81), Vero-furin (Mukherjee et al., 2016) , Vero E6 (ATCC-CRL1586), Vero-TMPRSS2 (Matsuyama et al., 2020) , and MA104

(ATCC-CRL-2378.1) cells. Each cell type is sufficient to propagate SARS-CoV-2 using the protocol detailed below. All procedures should be completed only after appropriate safety training is obtained and using aseptic technique within a certified biosafety cabinet under BSL-3 containment.

Chosen cell type (Vero-CCL81, Vero-furin, Vero E6, Vero-TMPRSS2, and MA104 cells)

Standard media for chosen cell type (see Recipes)

Infection media (see Recipes) SARS-CoV-2 seed stock 150 cm 2 (T150) tissue culture flasks 15mL disposable polystyrene conical tubes with screw caps (e.g., Falcon) 50mL disposable polystyrene conical tubes with screw caps (e.g., Falcon) 1.5mL or 0.5-mL O-ring tubes 1.) In a standard BSL2 laboratory, plate cells for infection one day prior into two T150 flasks in standard media for the chosen cell type. One flask serves as a mock-infected control and the other for infection. Plate cells so they will be ~80-90% confluent the following day. *For instance, plate 1 x 10 7 Vero CCL81 cells per T150 flask. Place flasks in a humidified 37°C incubator with 5% CO 2 overnight.

2.) Transfer flasks into BSL3 facility the following day. Rapidly thaw a SARS-CoV-2 stock at 37 o C. Calculate the volume of virus needed to infect at the desired multiplicity of infection (MOI) using the following formula: Real-time PCR assay for SARS-CoV-2 detection. Detection of viral RNA by reversetranscription quantitative polymerase chain reaction (RT-qPCR) using a TaqMan probe is a highly-sensitive and specific method for measuring viral burden in a variety of specimens.

Because CoVs generate subgenomic RNAs as a template for translation, the abundance of viral RNA varies for each gene and depends upon the gene position within the genome. Genes located closer to the 3' end of the (+) sense genome will have a greater abundance of transcripts than those located at the 5' end of the (+) sense genome. This should be considered when designing primer/probe combinations, as "N gene" transcripts will be more abundant than genomic RNA copies, which can be quantified by targeting sequences within the ORF1a gene.

Many primer/probe combinations have been designed and validated, several of which are used in clinical diagnosis ((CDC), 2020; Corman et al., 2020) . In the clinical setting, precise copynumber quantitation of viral RNA is not necessary and instead sensitivity is paramount.

However, quantitative assays are desirable for research applications, and may have utility in longitudinal studies of infected human subjects. RT-qPCR cycle threshold (Ct) values can be converted to transcript or genome copy number equivalents by generating an RNA standard curve, the design and production of which is described below.

The CoV replication strategy should be considered when designing a RT-qPCR assay.

Primer/probe combinations targeting the N gene are most sensitive; those targeting the spike gene can also be used to titer spike-containing pseudoviruses; those targeting the ORF1a gene provide genome equivalents; and those targeting the leader sequence can give an estimation of the total number of viral transcripts ( Table 1) . For a given viral gene target, a template (~500-1000 bp) for in vitro transcription can be generated by RT-PCR using primers that flank the intended target, with the forward (F) primer also including a 5' T7 promoter sequence (Vogels et al., 2020) . If multiple targets are desired, a single dsDNA fragment can be synthesized to include concatenated gene fragments, each of which spans the entirety of the target amplicons.

This strategy also can be used to quantify host genes of interest (e.g., ACE2).

1. (Day 1) The DNA fragment/amplicon containing the primer/probe targets to be used in the RT-qPCR assay should be introduced into a vector containing a T7 (or other DNAdependent RNA-polymerase) promoter sequence using Gibson Assembly, restriction enzyme cloning, blunt-end ligation, or gene synthesis. These vectors should be transformed into competent E. coli (e.g., DH5α) for antibiotic selection.

2. (Day 2) Pick clones and amplify to miniprep scale. We nornally pick 6 to 12 clones to ensure proper cloning.

3. (Day 3) Purify plasmid from clones, and identify a clone with the proper insert using restriction enzyme digestion and/or Sanger sequencing.

4. (Day 4) Linearize ~2-4 μg of the DNA in preparation for in vitro transcription by performing an overnight restriction digest using a high-fidelity restriction enzyme that cuts each plasmid only once in a position 3′ to the insert. The distance between the T7 transcriptional start-site and the 3′ end restriction site should be ~500-1500 nucleotides.

be apparent relative to the non-linearized plasmid. Extract and cleanup the linearized product with a commercially-available gel-extraction (e.g., Qiagen) kit.

6. Perform in vitro transcription using a commercially available kit (e.g., MEGAscript T7).

Note: to prevent contamination of PCR workstations with transcribed RNA, all steps hereafter should be performed in a contained hood/workspace that is separate from the area where PCR reaction setup is performed.

7. Digest DNA, then perform RNA cleanup using a commercially available kit (e.g., MEGApure).

8. Quantify the RNA using a spectrophotometer (e.g., Nanodrop or Qubit) by diluting the RNA with RNase-free water until the concentration is within the analytical measurement range of the spectrophotometer. 10. Dilute the transcript with RNase-free water containing 1% of added ribonuclease inhibitor (e.g., RNaseOUT) to obtain 1 to 2 mL of standard at a 1×10 10 copies/μL. Mix by pipette.

11. Aliquot the diluted RNA transcript into PCR strip tubes (with individual caps) in aliquots of 6 to 12 μL/aliquot.

12. Freeze at -80 o C. The remaining concentrated RNA can be frozen and re-quantified later as needed.

The RNA standard is concentrated and poses a risk for contamination of reagents and specimens. Follow best-practices for PCR preparation (Standards Unit, 2010) and only handle RNA standards after all reagents and specimens have been stored. Appropriate no-template controls must be used to eliminate and track possible contamination. Wipe down work areas and pipettes with 10% bleach followed by 70% ethanol. Bleach pipette tips.

1. Create a 20x stock of primer/probe mix by diluting primers to a concentration of 10 μM and probe to a concentration of 2 μM.

2. For "n" number of reactions, create a master-mix for n+1 by combining one-step RT-qPCR reaction buffer, primer/probe mix, and reverse-transcriptase enzyme at the appropriate concentration/volumes. Aliquot master-mix into wells of a RT-qPCRcompatible plate.

3. Separate a single tube containing the RNA standard from the stock. Work quickly to avoid thawing other aliquots in the adjacent strip tubes.

4. Thaw the aliquot and briefly centrifuge to collect contents at the bottom of the tube. 5. Dilute the standard into a volume of RNase-free water to obtain 1.0 x 10 9 RNA copies per reaction. Mix gently but thoroughly by pipette. Change gloves.

6. Make 10-fold serial dilutions in a PCR strip-tube by transferring 10 μL into 90 μL of RNase-free water. Mix each dilution thoroughly with a p100 pipette set to 70 μL. Discard tips between each dilution.

Note: When testing a new RNA standard, perform serial dilutions several-fold below 1 copy per reaction. Reactions containing less that 1-10 copies/well should fail to amplify.

7. Transfer the appropriate volume of RNA standard from each dilution into the reaction plate using a multichannel pipette.

8. Perform real-time PCR using the following thermocycling parameters:

1. 48 º C for 15 minutes 2. 95 º C for 10 minutes 3. 95 º C for 15 seconds 4. 60 º C for 1 minute -Acquire Signal 5. Go to "step 3" 49x (i.e., 50 cycles) Note: these parameters may vary depending on the specific RT-qPCR kit used; our parameters have been tested using the TaqMan RNA-to-CT 1-step kit (Applied Biosystems) on the QuantStudio 6 flex Real-time PCR system (Applied Biosystems).

9. Upon completion of the run, examine your standard curve. Approximately 3.3 Ct should separate each dilution, which corresponds to a change of one log 10 copies for a reaction that is >90% efficient.

Quantification of SARS-CoV-2 by plaque assay. The plaque assay is the gold standard test for quantifying infectious virus in a sample. The plaque assay measures "plaques," which describe the zone of cellular death that occurs after one infectious unit has entered a cell and spread to adjacent cells over the time period of incubation (Fig 2) . The assay does not rely on the use of any virus-specific reagents, which is beneficial when reagents are unavailable. As this cell-based assay typically is performed in 6-well plates, it is relatively low-throughput, labor-intensive, and may not be reliable when the samples themselves are cytotoxic (e.g., homogenate from certain tissues) or when the virus is poorly cytopathic in a given cell type.

Thus, it is important to choose a highly permissive cell type (e.g., Vero E6 cells) for which SARS-CoV-2 causes substantive cell death. 1. Plate approximately 7.5 x 10 5 Vero E6 or Vero-furin cells/well into 6-well plates. Plate enough wells to test each dilution in duplicate (starting from 10 -1 to 10 -6 ; 10-fold dilutions). Incubate cells overnight (12-18 h) at 37°C.

*12-well tissue culture plates also will work. Plate approximately 2.5 x 10 5 cells/well.

2. Dilute samples to be titered in infection media in 96-well U-bottom plates. Make a 10fold dilution series, providing enough volume to add 200 µL per 6-well plate.

3. Remove existing cell culture media from 6-well plates. Add 200 uL of each dilution to one well of a 6-well plate (200uL to 12-well plate) starting with most diluted so the same pipette tip can be used up the dilution series.

4. Incubate 6-well plates at 37°C in 5% CO 2 for 1 h, rocking plates every 15 min to prevent cells from drying out. 5. Meanwhile, mix 2X MEM + 4% FBS with 2% methylcellulose in a 1:1 ratio. Place in 37°C incubator while plates are incubating to decrease viscosity of the solution.

6. After 1 h incubation, add 2 mL of MEM:methylcellulose mixture to each well of the 6-well plates (1 mL to 12-well plate). 7. Incubate plates at 37°C in 5% CO 2 for 3 days. 

A focus-forming assay is similar to a plaque assay in that it detects infectious virus in a sample. A "foci" describes the zone of cells that have become infected from a single infectious unit. These foci of cells express high amounts of viral antigen, which can be detected using a virus-specific antibody that is directly conjugated to a colorimetric readout (e.g. peroxidase) or through use of secondary antibodies (Fig 3) . This approach adds specificity to the assay, but also increases the number of processing steps post-infection. However, because the focus-forming assay captures infected foci before the cells die and develop into plaques, this assay typically requires shorter incubation times than the plaque assay. It also can be performed in 96-well plate format, which can increase throughput. 14. Add 50 µL primary antibody/well in Perm wash. Incubate at 4°C overnight with no rocking or 2 h at room temperature with rocking.

*These conditions are optimized for using CR3022 (Yuan et al., 2020) 22. Tap plate dry on a paper towel and image with CTL Immunospot plate reader. 2. Add cellular growth media and centrifuge in a swinging bucket rotor for 5 min at 500

x g.

Note: this must be performed in an aerosol-tight bucket with gasketed lid. 10. Analyze cells on a flow cytometer (Fig 4) .

To evaluate many aspects of COVID-19 biology, methods for inactivating SARS-CoV-2 infectivity are needed so that samples can be worked with safely outside of the BSL-3. To test whether a specific method or treatment completely inactivates SARS-CoV-2, a virus outgrowth assay should be used. This type of assay is highly sensitive in that it allows for the outgrowth of as little as a single infectious unit. However, it is not quantitative and must be adapted to the application in question. Alternate agents and methods are benchmarked against "gold-standard" methods of inactivation of SARS-CoV-2 ( Table 2) . We describe methods that we have tested and validated, although before use individual Institutional Biosafety Committees likely will need to review data before providing clearance.

Inactivation agent/method of choice Vero E6 cells Vero cell culture medium

Selecting the appropriate inactivation agent/method requires a detailed understanding of the project in question: specifically, the properties of the measurand (e.g., DNA, RNA, protein, and cells); the effect of the agent/method on the integrity of the measurand (e.g., fragmentation of DNA by formaldehyde or lysis of RNA by boiling); and the types of specimens that will be treated for inactivation (e.g., whole blood, plasma, cell culture media, cells, and tissues). No one reagent or method works for all applications, and ideally, the activity of each inactivation agent should be tested against each specimen type that will be used in the project. Validation of reagents/methods with a diverse array of applications may result in fewer hours spent performing validation of inactivation reagents.

Many commonly used chemical inactivation agents (e.g., chaotropic salts, detergents, and formaldehyde-based solutions) are toxic to cells (Fig 5A) . Because viruses require infection of a cell to replicate, this toxic effect of the inactivation agent must be diluted sufficiently after the sample has been treated to enable virus outgrowth. Longer incubation times may be necessary. Mark wells in which toxicity is no longer obvious, and test the cells in these wells for viability and total cell number using trypan blue staining (or a variety of other live/dead counting methods). Use the media-only wells in column 12 for comparison. Wells that have the same viability and number of cells (±10%) should be used to calculate the required dilution factor.

To determine the ability of the agent/method to fully inactivate infectious virus, several high-titer specimens should be identified. If these are not available, then these specimens can be created by spiking specimens with SARS-CoV-2 virus stock (1:10) or infected cells. Ideally, specimens with the range of characteristics that will be encountered during the project (e.g., icteric, hemolyzed, and lipemic serum specimens) should be tested.

non-toxic level when >1 μL of treated sample is added. The number of cells also should be such that they will reach approximately 50% confluency upon adhering.

For example, for diluting an agent 1:1000, plate 3-4×10 4 Vero E6 cells in 4 mL and add 4 uL of the inactivated sample.

2. For the specimen(s) to be tested, split into two equal aliquots. Subject one to inactivation and the other to mock-inactivation (e.g., with addition of saline or medium instead of inactivation reagent). This should be performed at the temperature and for the duration of time that will be used for inactivation of experimental specimens.

3. Add the appropriate volume of inactivated (and mock-inactivated) sample to the Vero cells.

4. Incubate at 37 o C and observe daily (Fig 5B) . Once obvious signs of CPE are observed, examine the cells and/or supernatant for infection using the flow cytometry and/or focus forming assay described above (Fig 5C) to confirm viral infection.

The emergence of SARS-CoV-2 and the resulting COVID-19 pandemic has strained biomedical resources throughout the world. Necessarily, pressure has been placed on the scientific community to deliver countermeasures for this continually evolving threat. Although new technologies are being applied to address this problem, classic virological methods, such as those presented here, remain important. Within a remarkably short period of time, the scientific community has built an infrastructure for studying SARS-CoV-2, especially given the biosafety concerns surrounding SARS-CoV-2 research. However, given the complex nature of COVID-19 pathophysiology, a critical need remains for developing new modalities for studying and combating this novel disease. Further optimization of assays will be required, and these will include a need to amplify high-titer virus stocks from low-passage patient isolates and develop new culture modesl to evaluate infectivity, host responses, and outcomes. New methods for SARS-CoV-2 inactivation will be developed, and these will require rigorous validation before wide-scale implementation. Issues regarding SARS-CoV-2 biosafety and biocontainment will continue to evolve as the pandemic progresses, and methods for safely working with and titrating SARS-CoV-2 will require further evaluation. Filter sterilize and store at 4°C until ready for use.

This study was supported by NIH contracts and grants (75N93019C00062 and R01 following treatment with an inactivation agent or PBS (mock). Cells were dissociated to singlecell suspension once the mock-treated culture displayed CPE consistent with SARS-CoV-2 infection. Viability staining with Zombie violet was performed prior to fixation. Antibody staining was performed on 4% paraformaldehyde-fixed and permeabilized cells using the CR3022 anti-SARS-CoV-2 spike antibody followed by anti-human IgG-BV421 labelled secondary antibody. 

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