Large-Scale SARS-CoV-2 Testing Utilizing Saliva and Transposition Sample Pooling

Zusammenfassung

The protocol is intended to serve as a blueprint for universities and other organizations considering large-scale testing for SARS-CoV-2 or developing preparedness plans for future viral outbreaks.

Zusammenfassung

Identification and isolation of contagious individuals along with quarantine of close contacts, is critical for slowing the spread of COVID-19. Large-scale testing in a surveillance or screening capacity for asymptomatic carriers of COVID-19 provides both data on viral spread and the follow-up ability to rapidly test individuals during suspected outbreaks. The COVID-19 early detection program at Michigan State University has been utilizing large-scale testing in a surveillance or screening capacity since fall of 2020. The methods adapted here take advantage of the reliability, large sample volume, and self-collection benefits of saliva, paired with a cost-effective, reagent conserving two-dimensional pooling scheme. The process was designed to be adaptable to supply shortages, with many components of the kits and the assay easily substituted. The processes outlined for collecting and processing SARS-CoV-2 samples can be adapted to test for future viral pathogens reliably expressed in saliva. By providing this blueprint for universities or other organizations, preparedness plans for future viral outbreaks can be developed.

Einleitung

The COVID-19 pandemic, caused by the SARS-CoV-2 virus, has caused the deaths of over 6.2 million people to date, with numbers rising every day¹. The gold standard of testing for SARS-CoV-2 is quantitative real-time (RT-q) PCR, with primers designed to target the viral genome, such as nucleocapsid, envelope, spike, and RNA-dependent RNA polymerase genes². At the beginning of the pandemic, sufficient capacity for SARS-CoV-2 testing was severely lacking. It arose from a lack of validated assays, testing components, clinical personnel, and an infrastructure unprepared to rapidly expand to accommodate pandemic-level, mass testing. Due to shortages, testing centers often required a physician's referral to be test eligible. These shortages resulted in delays for testing approval, long lines for uncomfortable nasopharyngeal sample collection, and lengthy wait times for results. Additionally, because of these constraints, testing efforts could not accommodate pre-symptomatic, mild, or asymptomatic carriers unknowingly spreading the SARS-CoV-2. The lack of easily accessible, widespread testing likely contributed to the uncontrolled spread of COVID-19.

Large-scale interval testing can be performed either as surveillance or screening. Both can be used to monitor local positivity rates in high-density or high-risk transmission areas and can be utilized to make public health decisions. Surveillance testing is intended to monitor the incidence and prevalence of disease in a population and is not used for individual diagnostics³. Surveillance results typically are de-identified and not returned to participants; laboratories conducting surveillance testing need not be clinically certified, nor are they required to use an FDA-authorized assay. Screening allows for results to be returned to individual participants, but screening laboratories in the United States must have a Clinical Laboratory Improvement Amendments (CLIA) certificate and meet all applicable CLIA requirements.

The Michigan State University (MSU) early detection program began in September 2020 and has processed over 350,000 samples. The program arose out of a research group's efforts to design a highly sensitive SARS-CoV-2 assay that did not require high demand testing supplies^4,5,6,7,8. The goals were to aid clinical labs to increase capacity and develop flexible processes to accommodate supply shortages while also developing a screening strategy to establish a return-to-work plan for the MSU College of Human Medicine. The initial efforts focused on alternative collection, extraction, and quantitation methods for SARS-CoV-2. High demand and subsequent shortages of nasopharyngeal swabs led to the evaluation of anterior nares samples collected with buccal swabs, and reagent shortages resulted in development of a sample extraction method adapted from early reports out of Wuhan, China⁹. To increase sensitivity for detecting SARS-CoV-2 in anterior nares samples, droplet digital PCR was substituted for RT-qPCR^6,7. Though droplet digital PCR is highly sensitive and can provide absolute values with an endpoint readout, it was determined that its use was not feasible for large-scale testing due to the lack of reliable high-throughput instrumentation for the technology. Additionally, self-collection of anterior nares samples based on levels of human RNase P was extremely variable, suggesting that it was not sufficiently reliable for mass testing.

An alternative to nasopharyngeal and anterior nares swabs is the collection of saliva. Respiratory viruses such as SARS-CoV, H1N1, and MERS were all historically detected in saliva^10,11,12,13. This was subsequently proven true for SARS-CoV-2^14,15,16,17. Direct comparison between saliva and nasopharyngeal samples showed saliva yields higher viral titers than nasopharyngeal swabs in matched samples, and that saliva is less variable with repeated sample collection¹⁴. Saliva has also been reported to be more sensitive in certain variants, such as Omicron, compared to Delta¹⁶. Added benefits to saliva collection are the relative ease of off-site self-collection without high-demand supplies, the ability to repeatedly retest the sample if needed, the elimination of on-site staffing requirements for sample collection, and the avoidance of participant queues which could increase the potential for viral transmission. The lab-assembled saliva kit was developed as a collaboration among lab assay developers, experts in the school of packaging, university branding experts, safety officers, and external manufacturing partners that produced the box and labeling system.

While saliva samples offer ample genetic starting material and RT-qPCR provides sensitive, reliable outcomes, the cost of reagents (primer/probes and master mix) made large-scale testing of individual samples a costly endeavor on an individual, per sample basis. Since the primer/probes and master mix are the most expensive components of the process, the goal was to seek solutions that would stretch their use and therefore decrease the per sample cost. Systemically optimizing sample pool size based upon incidence in the community and assay sensitivity has been proposed for SARS-CoV-2 testing¹⁸. However, when pools of any size indicate presence of SARS-CoV-2, all participants in the pool must be retested, resulting in lost time and increased opportunities for spread. To address these limitations, a two-dimensional pooling method was employed, like the process proposed by Zilinskas and others¹⁹ to conduct a first pass under the strictures of surveillance testing. In this process, 96 individual samples are placed in a 96-well plate consisting of 12 columns and eight rows. Each sample is included in a pool of eight and a pool of 12 on two different reaction plates. This results in every sample being uniquely represented with the two pools. Deconvolution of the pools based on the coordinates identifies potentially positive samples. Samples in pools where SARS-CoV-2 was not detected, do not move from surveillance testing to screening. Meanwhile, samples from individuals testing positive in the surveillance process are re-extracted through a CLIA-approved screening process. If confirmed positive, individuals are given their result, referred to the university physician's office, contact tracing is initiated, and the health department is notified. In total, an individual's sample is tested in three separate reactions before being declared positive, twice in surveillance pools and once as a single confirmatory screen, reducing the chances of a false positive. Sample pooling uses ~80% less reagents than running samples individually, resulting in a cost of ~$12 per sample.

Beyond the saliva kit, pooling strategy, and assay development process, the team also developed a logistics plan for distribution of kits, collection of samples, and reporting of results. Participants in the program pick up their kit, register the unique alphanumeric code on their tube, produce their sample and deposit it in one of the many drop-off bins where they are picked up daily and transported to the lab. The lab processes the samples, the technical supervisor reviews and uploads results, and participants are notified to check the results portal. This process has a turnaround time of 24-48 h from the time a sample is deposited. The collaboration from all parts of the institution were key for a successful large-scale implementation of this hybrid surveillance and screening process. The following procedures and descriptions of the testing program and infrastructure needed are intended as blueprints on how to scale-up testing for future surveillance and/or screening purposes.

Protokoll

Studies performed to optimize the methods for the Early Detection Program were approved by the Michigan State University Institutional Review Board. All figures were reproduced with contrived samples and are representative of the observed human results. No data, information, or results shown in the manuscript are from any participant in the Michigan State University Early Detection Program.

1. Kit production

NOTE: During kit assembly, wear masks, gloves, eye protection, and lab coats at all times to prevent kit component contamination.

1. Preparation of kit components
   1. Using a permanent marker, draw a line denoting 1 mL on the 25 mL conical tube. Using a permanent marker, draw a line denoting 1 mL on the transfer pipet.
   2. Add four ceramic beads to the 5 mL sample tube. Pipet 1 mL of RNA stabilization solution into the sample tube and close the tube. Attach a barbell sticker consisting of a unique identifying code to sample tube with the data matrix barcode on the top and the identifier repeated in alphanumeric code on the side.
      NOTE: The ceramic beads are RNase/DNase-free and do not need to be further sterilized.
2. Place a 25 mL conical tube, 5 mL sample tube, transfer pipet, and small biohazard bag into the kit box (Supplementary Figure 1).
   ​NOTE: Conical tubes do not need to be DNase/RNase-free. Integrity of the sample is preserved by the RNA stabilization solution.

2. Kit uses for self-sample collection

1. As per the kit instructions, ask the participants not to eat or drink 30 min prior to providing a sample. Ask the participants to spit into the 25 mL conical tube until 1 mL of saliva is collected (marked line).
2. Using the pipet, transfer 1 mL of saliva (up to the marked line) into the 5 mL sample tube containing RNA stabilization solution and ceramic beads. Close and shake the sample tube for 15 s.
3. Ask the participants to register the alphanumeric code assigned to sample on the designated website. Once done, ask them to place the sample tube in small biohazard bag and deposit into a collection bin.

3. Sample intake and preparation for RNA isolation

NOTE: All steps take place in a biosafety cabinet or centrifuge bucket with a biocontainment lid.

1. Disinfect and visually inspect samples for quality control while still in the biohazard bag.
2. Reject the sample if (Figure 1): sample is of non-natural color (i.e., green, blue) or contains food particles or blood; sample is leaking or missing beads; sample has incorrect expected volume (<1.5 mL or >3 mL); sample does not have a barcode or alphanumeric code attached.
   1. If the sample is rejected, scan and record the sample into a rejection file with a note as to why the sample was rejected. If sample is rejected because it has no identifiers, log the sample as No Barcode in the rejection file.
3. If sample is not rejected, cut open the biohazard bag with scissors and remove the sample tube. Vortex the tube for 15 s then place the sample tube into a tube adaptor for a swing-bucket centrifuge.
4. Once tube adapters are full, secure the biocontainment lid over the centrifuge bucket and transfer to the centrifuge. Centrifuge samples at 4,100 x g for 2 min and return samples to the biosafety cabinet.
   NOTE: Centrifugation is used to pellet debris and force more viscous material in saliva to the bottom of the tube. This eliminates the need for proteinase K in the process. A full run consists of 768 samples (eight 96-well plates full of sample after RNA isolation).
5. Without disturbing the pelleted material, transfer sample tubes to a 96-slot tube rack pre-labeled with the Run ID which includes the rack number (1-8), date, and the corresponding group the samples will be pooled/assessed with (an identifier to denote the group, i.e., group A is the first run of the day).
6. When 96-slot tube racks are full or no samples remain, using a handheld barcode scanner, scan tubes into a plate map file (Supplementary File 1 for full pools; Supplementary File 2 for half size pools). In the event a barcode does not work, use the alphanumeric code on the tube to log the sample.
   NOTE: A mask is required to prevent the scanning of adjacent barcodes. A mask can be made by cutting a hole in opaque paper, then laminating it. Barcodes scanned into the plate map file will appear in the same orientation as in the 96-slot rack.
7. Spray the sample tubes in the rack with 70% ethanol and dab dry with a paper towel.
   NOTE: Wiping the tubes instead of dabbing can damage the barcodes.
8. Label a 96-deep-well plate of 2 mL well capacity with the Run ID. Transfer 200 µL of saliva sample from each sample tube to the corresponding well of the 96-deep-well plate. If samples are too viscous to transfer, vortex and centrifuge again. Reject sample if problem remains.
   NOTE: Saliva samples can be stringy and leave a snail-trail across the plate during transfer if the technician is not careful. This can lead to initial false positives and more samples that will eventually need to be re-run in the confirmation step (see section 6).
9. When all samples are transferred, cover the 96-deep-well plate with a silicone mat and store at room temperature. Save and store the 96-slot racks with remaining samples for confirmation of potentially positive samples.
   ​NOTE: The protocol can be paused here.

4. RNA isolation

1. Label a 96-well RNA isolation cartridge with the Run ID and place it on top of an empty collection plate.
   NOTE: Plates containing samples can be removed from the biosafety cabinet at this point.
2. Remove the silicone mat from the 96-deep-well plate. Transfer 400 µL of viral loading buffer into each well of the 96-deep-well plate.
3. Mix each sample by gently pipetting up and down two to four times and transfer the entire sample to the corresponding column in the 96-well RNA isolation cartridge.
   NOTE: Mixing large volumes will create bubbles that may lead to contamination of neighboring wells if performed haphazardly. Mixing in a deliberate fashion can lower this risk and protect the integrity of downstream processing.
4. Centrifuge the RNA isolation cartridge on top of the collection plate at 4,100 x g for 10 min to load samples onto the cartridge. Transfer the RNA isolation cartridge to a clean collection plate.
5. Add 500 µL of wash buffer to every well of the RNA isolation cartridge and centrifuge at 4,100 x g for 5 min. Transfer RNA isolation cartridge to a clean collection plate.
6. Add 500 µL of wash buffer to every well of the RNA isolation cartridge and centrifuge at 4,100 x g for 5 min. Transfer RNA isolation cartridge to a clean collection plate.
7. Add 500 µL of 100% ethanol to every well of the RNA isolation cartridge and centrifuge at 4,100 x g for 5 min. Transfer RNA isolation cartridge to a clean collection plate.
   NOTE: If the sample has not completely gone through the column at this point, the sample is too viscous or small pieces of debris in the sample are clogging the column. The sample needs to be removed completely from the cartridge as to not contaminate the downstream pooling process, and the sample number added to the rejection file.
8. Centrifuge the RNA isolation cartridge on top of the collection plate at 4,100 x g for 5 min to dry the columns. Transfer the RNA isolation cartridge onto a clean elution plate labeled with the Run ID and the word row, then place both on top of a clean collection plate.
9. Add 25 µL of DNase/RNase-free molecular grade water to each well of the RNA isolation cartridge. Centrifuge at 4,100 x g for 10 min to elute the RNA. If sample does not completely elute, re-centrifuge the sample for 5 min.
10. Transfer the elution plates onto pre-chilled cold plates and cover. Immediately proceed onto sample pooling and RT-qPCR.

5. Sample pooling and RT-qPCR

NOTE: The process is set up in a two-plate system (as seen in Figure 2). The elution RNA plate number corresponds to the produced column plate number and the row plate letter (A = 1, B = 2, C = 3, etc.). For deconvolution purposes, the column plate row letter will correspond to the row letter from the RNA elution plate, and the row plate column number will correspond to the RNA elution plate column number. For example, a sample in RNA plate #6 in position C4 will be found in the row reaction plate position F4, and in the column reaction plate position C6.

1. Label a new elution plate with the Run ID and the word row, and place on a pre-chilled cold plate. Transfer 10 µL of RNA from each well on the row elution plate to the corresponding row on the column elution plate, thus making a copy of the original elution plate (Figure 2).
   NOTE: Elution plates marked column will be pooled in the column direction and elution plates marked row will be pooled in the row direction.
2. For the column pooled plate, transfer the entire volume from each well across the plate into the column number that corresponds to the Run ID plate number. For the row pooled plate, transfer the entire volume from each well up or down the plate into the row number that corresponds to the Run ID plate number (1 = A, 2 = B, etc.; Figure 2).
3. Transfer 13 µL of each pooled sample into the corresponding column or row of a 96-well reaction plate (Figure 2).
4. For this protocol, use three probes against the nucleocapsid (N), ORF1ab, and spike (S) genes to detect SARS-CoV-2, and a fourth primer probe targeting MS2 phage as an internal control. In the pooled samples, spike an MS2 phage positive control into the master mix. In samples run individually, spike MS2 phage into the sample on the RNA isolation cartridge and use as an extraction control.
   NOTE: Master mix, primer probes, fluorescent dyes, and quenchers will vary based on the assay. Appropriate ratios and fluorescent channels should be determined by each lab.
5. Prepare the positive control by adding 2 µL of positive control solution and 11 µL of DNase/RNase-free molecular grade water to the positive control well on the reaction plate (Figure 2).
6. Prepare the no template control by adding 13 µL of DNase/RNase-free molecular grade water to the negative control well on the reaction plate.
7. Make master mix according to manufacturer's instructions and spike in MS2 phage positive control. Dispense 7 µL of master mix into each well of the 96-well reaction plates that contain samples or controls. Limit exposure to fluorescent light.
8. Cover reaction plates with optical film and vortex for 2 min to mix samples well. Centrifuge reaction plates at 650 x g for 5 min.
9. Transfer plates into the RT-PCR system and run with cycling parameters appropriate for the master mix. For master mix used here, the following cycles were run: 25 °C for 2 min, 53 °C for 10 min, 95 °C for 2 min, 40 cycles of 95 °C for 3 s, and 60 °C for 30s.
10. Once the samples are run, perform a deconvolution of the pools to identify potential positive samples manually (Figure 3) and via a R Script Shiny App.
    1. Export results from column and row reaction plates as spreadsheet files. Open the pool deconvolution R script Shiny app (source code available at: https://github.com/kochman1/JoVE-MSU-COVID-EDP).
    2. Drag and drop the plate map, column, and row files into the app and run the app. Save the list produced from the app.
       ​NOTE: The list produced from the app contains all of the barcodes entered into the plate map and whether they are likely positive or negative based on the pool deconvolution script. These samples that are flagged as potential positives are reprocessed individually.

6. Validation of positive samples

1. Label two 1.5 mL microcentrifuge tubes and an RNA extraction column for each sample that needs to be tested. Use one microcentrifuge tube for mixing sample, MS2 phage, and viral loading buffer, and the second microcentrifuge tube for the RNA elution step.
2. Add 5 µL of MS2 Phage RNA to the 1.5 mL microcentrifuge tube. Transfer 200 µL of the potentially positive sample to the tube and add 400 µL of viral loading buffer to the tube, mix by pipetting.
3. Transfer the entire contents to the pre-labeled RNA extraction column in a clean collection tube. Centrifuge at 10,000 x g for 2 min. Aspirate flow through from the collection tube.
4. Add 500 µL of wash buffer to the RNA extraction column and centrifuge at 10,000 x g for 2 min. Aspirate flow through from the collection tube.
5. Add 500 µL of wash buffer to the RNA extraction column and centrifuge at 10,000 x g for 2 min. Aspirate flow through from the collection tube.
6. Add 500 µL of 100% ethanol to the RNA extraction column and centrifuge at 10,000 x g for 1 min. Transfer the RNA extraction column to a clean collection tube and centrifuge at 10,000 x g for 1 min to dry the column.
7. Transfer the RNA extraction column to a pre-labeled 1.5 mL microcentrifuge tube for elution. Add 25 µL of DNase/RNase-free molecular grade water to the RNA extraction column. Centrifuge at 10,000 x g for 2 min to collect the RNA and transfer tube to wet ice.
8. Make master mix according to manufacturer's instructions. Dispense 7 µL of master mix into each well of a 96-well reaction plate that will contain the sample. Transfer 13 µL of each isolated RNA into the reaction plate.
   NOTE: The master mix is the same one used in the pooled sample protocol but does not include the MS2 phage RNA spike.
9. Prepare the positive control by adding 2 µL of positive control solution and 11 µL of DNase/RNase-free molecular grade water to the positive control well on the reaction plate. Prepare the no template control by adding 13 µL of DNase/RNase-free molecular grade water to the negative control well on the reaction plate.
10. Cover the reaction plates with optical film and vortex for 2 min to mix samples well. Centrifuge reaction plates at 650 x g for 5 min. Transfer plates into the RT-PCR system and run with cycling parameters appropriate for the master mix.
    1. For master mix stated here, use 25 °C for 2 min, 53 °C for 10 min, 95 °C for 2 min, 40 cycles of 95 °C for 3 s, and 60 °C for 30 s.

Ergebnisse

The vast majority of samples received by the lab to date have been accepted and passed the initial visual quality control step. The need to reject a sample is limited to reasons that can negatively influence sample processing and/or the overall results for the sample. Specifically, incorrect volume in the tube, consistency, or color not natural to saliva, an absence of ceramic beads used to aid in sample homogenization, and missing barcodes are all reasons to reject a sample (Figure 1).

...

Diskussion

During sample processing, there are steps requiring careful attention. The initial quality control step which looks at the sample volume, consistency, color, and presence of added beads is critical to the overall success of the process. Tubes with samples that do not contain the correct amount of saliva could produce a false negative, as too little saliva would result in not enough genetic material; conversely, too much saliva would not be in the correct ratio with the RNA buffer and RNA degradation could occur. In rare ...

Materialien

Name	Company	Catalog Number	Comments
1 Step MM, no ROX	Thermo Fisher	A28523
1.2 mlDeep well Plates	Fisher	AB0564
100 mL reagent reservoirs	Corning	4872
2.8 mm Ceramic Beads	OMNI	19-646
25 ml conical w/screw cap	VWR	76338-496
50mL V bottom reservoirs	Costar	4870
5430-High-Speed Centrifuge	Eppendorf	22620601
5ml Eppendorf Tube	Fisher	14282300
8 strip tubes for QuantStudio	life technologies	4316567
Beta Mercaptoethanol	Fisher	AC125472500
Ethanol 200 Proof, Molecular Biology Grade	Fisher	BP28184
Microamp Endura Optical 96-well fast clear reaction plate with barcode	life technologies	4483485
Microamp Fast Optical 96 well plate	Fisher	4346906
Mini Microcentrifuge	Corning Medical	6770
optical caps for strip tubes	life technologies	AB-1820
Optical Film	Thermo Fisher	4311971
PCR plate sealing film Non-optical	Fisher	AB-0558
PCR Plate semi-skirted	Fisher	14230244
QuantStudio 3 Real-Time PCR System, 96-well, 0.1 mL	Thermo Fisher	A28136
Quick RNA Viral Kit confirmation	Zymo	R1035
Reagent Reservoir, 100ml	DOT	229298
RNA Shield	Zymo	R1200-1L
Small Biohazard Bags	Fisher	180000
Taqpath RTPCR COVID19 kit	Thermo Fisher	A47814
Thermo Scientific Sorvall ST4R Plus Centrifuge	Thermo Fisher	75009525
Transfer Pipet	Fisher	22170404
Viral 96 Kit	Zymo	R1041
Vortex Mixer	Fisher	2215414