CDC 2019-Novel Coronavirus (2019-nCoV)
Real-Time RT-PCR Diagnostic PanelFor Emergency Use Only

Instructions for Use

Catalog # 2019-nCoVEUA-01
1000 reactions
For In-vitro Diagnostic (IVD) Use
Rx Only

Centers for Disease Control and Prevention
Division of Viral Diseases
1600 Clifton Rd NE
Atlanta GA 30329

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Table of Contents
Intended Use ............................................................................................................................. 3
Summary and Explanation ....................................................................................................... 4
Principles of the Procedure ..................................................................................................... 4
Materials Required (Provided) ................................................................................................. 6
Materials Required (But Not Provided) ................................................................................... 7
Warnings and Precautions .................................................................................................... 10
Reagent Storage, Handling, and Stability ............................................................................. 12
Specimen Collection, Handling, and Storage....................................................................... 12
Specimen Referral to CDC ..................................................................................................... 13
Reagent and Controls Preparation........................................................................................ 13
General Preparation ............................................................................................................... 14
Nucleic Acid Extraction.......................................................................................................... 14
Assay Set Up ........................................................................................................................... 16
Create a Run Template on the Applied Biosystems 7500 Fast Dx Real-time PCR
Instrument (Required if no template exists) ......................................................................... 20
Defining the Instrument Settings .......................................................................................... 26
Running a Test ........................................................................................................................ 28
Interpretation of Results and Reporting ............................................................................... 34
2019-nCoV rRT-PCR Diagnostic Panel Results Interpretation Guide ................................ 36
Quality Control ........................................................................................................................ 37
Limitations .............................................................................................................................. 37
Conditions of Authorization for the Laboratory ................................................................... 38
Performance Characteristics ................................................................................................. 40
Disposal................................................................................................................................... 50
References .............................................................................................................................. 50
Revision History ..................................................................................................................... 51
Contact Information, Ordering, and Product Support ......................................................... 52
Appendix A: Heat Treatment Alternative to Extraction ....................................................... 53
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Appendix B: Pooled Specimen Preparation and Processing ............................................. 57
Appendix C: Implementation and Monitoring of Pooled Specimen Testing...................... 66

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Intended Use
The CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel is a real-time RT-PCR
test intended for the qualitative detection of nucleic acid from SARS-CoV-2 in upper and lower
respiratory specimens (such as nasopharyngeal or oropharyngeal swabs, sputum, lower respiratory tract
aspirates, bronchoalveolar lavage, and nasopharyngeal wash/aspirate or nasal aspirate) collected fromindividuals suspected of COVID-19 by their healthcare provider 1.
This test is also for use with anterior nasal swab specimens that are self-collected at home 2 or in ahealthcare setting by individuals using the authorized HealthPulse@home or HealthPulse@home
Blueprint kit manufactured for Audere (healthpulsenow.org) when determined to be appropriate by ahealthcare provider.

This test is also for the qualitative detection of nucleic acid from the SARS-CoV-2 in pooled samples
containing up to four of the individual upper respiratory swab specimens (nasopharyngeal (NP),oropharyngeal (OP), NP/OP combined, or nasal swabs) that were collected using individual vials
containing transport media from individuals suspected of COVID-19 by their healthcare provider.
Negative results from pooled testing should not be treated as definitive. If a patient’s clinical signs andsymptoms are inconsistent with a negative result or results are necessary for patient management, thenthe patient should be considered for individual testing. Specimens included in pools with a positive,
inconclusive, or invalid result must be tested individually prior to reporting a result. Specimens with low
viral loads may not be detected in sample pools due to the decreased sensitivity of pooled testing.
Testing is limited to laboratories certified under the Clinical Laboratory Improvement Amendments of1988 (CLIA), 42 U.S.C. § 263a, that meet the requirements to perform high complexity tests.
Results are for the identification of SARS-CoV-2 RNA. SARS-CoV-2 RNA is generally detectable in upperand lower respiratory specimens during infection. Positive results are indicative of active infection with
SARS-CoV-2 but do not rule out bacterial infection or co-infection with other viruses. The agent detected
may not be the definite cause of disease. Laboratories within the United States and its territories are
required to report all results to the appropriate public health authorities.
Negative results do not preclude SARS-CoV-2 infection and should not be used as the sole basis fortreatment or other patient management decisions. Negative results must be combined with clinicalobservations, patient history, and epidemiological information.
Testing with the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel is intended for use by trainedlaboratory personnel who are proficient in performing real-time RT-PCR assays. The CDC 2019-Novel
Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel is only for use under a Food and Drug
Administration’s Emergency Use Authorization.

For this EUA, a healthcare provider includes, but is not limited to, physicians, nurses, pharmacists, technologists, laboratory
directors, epidemiologists, or any other practitioners or allied health professionals.
2
For this EUA, home is defined as any non-healthcare setting.
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Summary and Explanation
An outbreak of pneumonia of unknown etiology in Wuhan City, Hubei Province, China was initiallyreported to WHO on December 31, 2019. Chinese authorities identified a novel coronavirus (2019-nCoV,
also referred to as SARS-CoV-2), which has resulted in millions of confirmed human infections globally.
Cases of asymptomatic infection, mild illness, severe illness, and deaths have been reported.
The CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel is a molecular in vitro diagnostic test that aids inthe detection and diagnosis of SARS-CoV-2 infection and is based on widely used nucleic acidamplification technology. The product contains oligonucleotide primers and dual-labeled hydrolysis
probes (TaqMan®) and control material used in real-time reverse transcription polymerase chain
reaction (rRT-PCR) for the in vitro qualitative detection of 2019-nCoV RNA in respiratory specimens.
The term “qualified laboratories” refers to laboratories in which all users, analysts, and any person
reporting results from use of this device should be trained to perform and interpret the results from thisprocedure by a competent instructor prior to use.

Principles of the Procedure
The oligonucleotide primers and probes for detection of 2019-nCoV were selected from regions of thevirus nucleocapsid (N) gene. The panel is designed for specific detection of SARS-CoV-2 (twoprimer/probe sets). An additional primer/probe set to detect the human RNase P gene (RP) in control
samples and clinical specimens is also included in the panel.
RNA isolated and purified from upper and lower respiratory specimens is reverse transcribed to cDNAand subsequently amplified in the Applied Biosystems 7500 Fast Dx Real-Time PCR Instrument with SDS
version 1.4 software. In the process, the probe anneals to a specific target sequence located between
the forward and reverse primers. During the extension phase of the PCR cycle, the 5’ nuclease activity of
Taq polymerase degrades the probe, causing the reporter dye to separate from the quencher dye,generating a fluorescent signal. With each cycle, additional reporter dye molecules are cleaved fromtheir respective probes, increasing the fluorescence intensity. Fluorescence intensity is monitored ateach PCR cycle by Applied Biosystems 7500 Fast Dx Real-Time PCR System with SDS version 1.4 software.
Detection of viral RNA not only aids in the diagnosis of illness but also provides epidemiological andsurveillance information.

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Summary of Preparation and Testing Process

Upon receipt ofrRT-PCR Panelreagents

Resuspend
primer/probe mix,
aliquot and store at≤ -20°C

Upon obtaining
sampleResuspend andaliquot nCoVPC,
store at -70°C

Extract sample RNA
and HSC RNA

Prepare master mix(15 µL)

Prepare rRT-PCR
plate (5 µL RNA)Run assay on
ABI 7500Fast Dx

Analyze data

Report results

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Materials Required (Provided)
Note: CDC will maintain on its website a list of commercially available lots of primer and probe sets
and/or positive control materials that are acceptable alternatives to the CDC primer and probe set
and/or positive control included in the Diagnostic Panel. Only material distributed through the CDC
International Reagent Resource and specific lots of material posted to the CDC website are acceptablefor use with this assay under CDC’s Emergency Use Authorization.
This list of acceptable alternative lots of primer and probe materials and/or positive control materials
will be available at:https://www.cdc.gov/coronavirus/2019-nCoV/lab/virus-requests.html
Primers and Probes:
Catalog #2019-nCoVEUA-01 Diagnostic Panel Box #1:
Reagent Label
2019-nCoV_N1
2019-nCoV_N2
RPPart #
RV202001
RV202015
RV202002
RV202016
RV202004
RV202018

Quantity /TubeReactions /Tube2019-nCoV_N1 Combined Primer/Probe Mix

22.5 nmol

1000

2019-nCoV_N2 Combined Primer/Probe Mix

22.5 nmol1000

Human RNase P Combined Primer/Probe Mix

22.5 nmol

1000

Description

Positive Control (either of the following products is acceptable):
Catalog #2019-nCoVEUA-01 Diagnostic Panel Box #2:
Reagent
LabelPart #

Description

Quantity

Notes

nCoVPC

RV202005

2019-nCoV Positive Control (nCoVPC)For use as a positive control with the CDC 2019nCoV Real-Time RT-PCR Diagnostic Panelprocedure. The nCoVPC contains noninfectious
positive control material supplied in a dried stateand must be resuspended before use. nCoVPC
consists of in vitro transcribed RNA. nCoVPC will
yield a positive result with each assay in the
2019-nCoV Real-Time RT-PCR Diagnostic Panelincluding RP.

4 tubes

Provides(800) 5 µL
test reactions

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Catalog #VTC-04 CDC 2019-nCoV Positive Control (nCoVPC)
Reagent
LabelPart #

Description

Quantity

Notes

nCoVPC

RV202005

2019-nCoV Positive Control (nCoVPC)For use as a positive control with the CDC 2019nCoV Real-Time RT-PCR Diagnostic Panelprocedure. The nCoVPC contains noninfectious
positive control material supplied in a dried stateand must be resuspended before use. nCoVPC
consists of in vitro transcribed RNA. nCoVPC will
yield a positive result with each assay in the
2019-nCoV Real-Time RT-PCR Diagnostic Panelincluding RP.

4 tubes

Provides(800) 5 µL
test reactions

Materials Required (But Not Provided)
Human Specimen Control (HSC)

Description
Manufactured by CDC. For use as a nucleic acid extraction procedural
control to demonstrate successful recovery of nucleic acid as well as
extraction reagent integrity. The HSC consists of noninfectious (betaPropiolactone treated) cultured human cell material supplied as a liquidsuspended in 0.01 M PBS at pH 7.2-7.4.

Quantity

CDC Catalog No.

10 vials x 500 µL

KT0189Acceptable alternatives to HSC:
• Negative human specimen material: Laboratories may prepare a volume of human specimenmaterial (e.g., human sera or pooled leftover negative respiratory specimens) to extract andrun alongside clinical samples as an extraction control. This material should be prepared in
sufficient volume to be used across multiple runs. Material should be tested prior to use as
the extraction control to ensure it generates the expected results for the HSC listed in these
instructions for use.
• Contrived human specimen material: Laboratories may prepare contrived human specimenmaterials by suspending any human cell line (e.g., A549, Hela, or 293) in PBS. This material
should be prepared in sufficient volume to be used across multiple runs. Material should betested prior to use as the extraction control to ensure it generates the expected results for
the HSC listed in these instructions for use.
CDC will maintain on its website a list of commercially alternative extraction controls, if applicable,that are acceptable for use with this assay under CDC’s Emergency Use Authorization, at:https://www.cdc.gov/coronavirus/2019-nCoV/lab/virus-requests.html

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rRT-PCR Enzyme Mastermix Options
Reagent

Quantity

Catalog No.

100 x 20 μL rxns
(1 x 1 mL)
2000 x 20 μL rxns
(1 x 20 mL)
500 x 20 μL rxns
(5 x 1 mL)
100 x 20 µL rxns
(500 µL)
500 x 20 μL rxns
(5 x 500 µL)
1000 x 20 μL rxns
(1 x 5 mL)
200 x 20 μL rxns
(2 mL)
1250 x 20 μL rxns
(12.5 mL)

Quantabio qScript XLT One-Step RT-qPCR ToughMix

Quantabio UltraPlex 1-Step ToughMix (4X)

Promega GoTaq® Probe 1- Step RT-qPCR System

Thermofisher TaqPath™ 1-Step RT-qPCR Master Mix, CG

95132-100
95132-02K
95132-500
95166-100
95166-500
95166-01K
A6120
A6121

1000 reactions

A152992000 reactions

A15300RNA Extraction Options
For each of the kits listed below, CDC has confirmed that the external lysis buffer is effective forinactivation of SARS-CoV-2.
Instrument/Manufacturer
QIAGEN

Extraction Kit

Catalog No.

2

QIAmp DSP Viral RNA Mini Kit

50 extractions (61904)

QIAamp Viral RNA Mini Kit

50 extractions (52904)
250 extractions (52906)

2

48 extractions (62724)
Buffer AVL (19073 or 19089)

EZ1 DSP Virus Kit

2

EZ1 Advanced XL DSP Virus Card (9018703)

QIAGEN EZ1 Advanced XL

48 extractions (955134)
Buffer AVL (19073 or 19089)

EZ1 Virus Mini Kit v2.0

2

EZ1 Advanced XL Virus Card v2.0 (9018708)

Roche MagNA Pure 24

Roche MagNA Pure 96
Roche MagNA Pure LC

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96 extractions (07 658 036 001)

MagNA Pure 24 Total NA
Isolation Kit

2External Lysis Buffer (06 374 913 001, 12 239 469
103, 03 246 779 001 or 03 246 752 001)

DNA and Viral NA Small Volume
Kit

2

576 extractions (06 543 588 001)
External Lysis Buffer (06 374 913 001, 12 239 469
103, 03 246 779 001 or 03 246 752 001)
192 extractions (03 038 505 001)

Total Nucleic Acid Kit

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Instrument/Manufacturer

Extraction Kit

Catalog No.

Nucleic Acid Isolation Kit I

32 extractions (03 730 964 001)

Maxwell® RSC Viral Total
Nucleic Acid Purification Kit

48 extractions (AS1330)

QIAmp DSP Viral RNA Mini Kit

50 extractions (61904)

1

Roche MagNA Pure Compact

2

Promega Maxwell® RSC 48
and Maxwell® CSC 48

3

2

QIAGEN QIAcube

1

QIAamp Viral RNA Mini Kit

2

144 extractions (ASB1330)

50 extractions (52904)
250 extractions (52906)
EasyMAG® Magnetic Silica (280133)

bioMérieux NucliSENS®
easyMAG®
and1, 3
bioMérieux EMAG®
(Automated magnetic
extraction reagents soldseparately. Both instruments
use the same reagents anddisposables, with the
exception of tips.)
1, 3

EasyMAG® Lysis Buffer (280134)
EasyMAG® Lysis Buffer, 2 mL (200292)
EasyMAG® Wash Buffers 1,2, and 3(280130, 280131, 280132)
EasyMAG® Disposables (280135)
Biohit Pipette Tips (easyMAG® only) (280146)
EMAG®1000μL Tips (418922)

Equivalence and performance of these extraction platforms for extraction of viral RNA were demonstrated with the CDC
Human Influenza Virus Real-Time RT-PCR Diagnostic Panel (K190302). Performance characteristics of these extraction
platforms with 2019-nCoV (SARS CoV-2) have not been demonstrated.
2
CDC has confirmed that the external lysis buffer used with this extraction method is effective for inactivation of SARS-CoV2.
3
CDC has compared the concentration of inactivating agent in the lysis buffer used with this extraction method and has
determined the concentration to be within the range of concentrations found effective in inactivation of SARS-CoV-2.
1

Alternative to Extraction:
If a laboratory cannot access adequate extraction reagents to support testing demand due to theglobal shortage of reagents, CDC has evaluated a heat treatment procedure for upper respiratory
specimens using the Quantabio UltraPlex 1-Step ToughMix (4X), CG. Though performance wascomparable, this method has been evaluated with a limited number of clinical specimens and apotential reduction in sensitivity due to carryover of inhibitory substances or RNA degradation cannot
be ruled out. It should only be used when a jurisdiction determines that the testing need is great
enough to justify the risk of a potential loss of sensitivity. Heat-treated specimens generating
inconclusive or invalid results should be extracted with an authorized extraction method prior toretesting. Details and procedure for the heat treatment alternative to extraction may be found in
Appendix A.

Equipment and Consumables Required (But Not Provided)

Vortex mixer
Microcentrifuge
Micropipettes (2 or 10 μL, 200 μL and 1000 μL)
Multichannel micropipettes (5-50 μL)

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














Racks for 1.5 mL microcentrifuge tubes
2 x 96-well -20°C cold blocks
7500 Fast Dx Real-Time PCR Systems with SDS 1.4 software (Applied Biosystems; catalog#4406985 or #4406984)
Extraction systems (instruments): QIAGEN EZ1 Advanced XL, QIAGEN QIAcube, Roche MagNA
Pure 24, Roche MagNA Pure 96, Promega Maxwell® RSC 48, Roche MagNA Pure LC, Roche
MagNA Pure Compact, bioMérieux easyMAG, and bioMérieux EMAG
Molecular grade water, nuclease-free
10% bleach (1:10 dilution of commercial 5.25-6.0% hypochlorite bleach)
DNAZapTM (Ambion, cat. #AM9890) or equivalent
RNase AWAY™ (Fisher Scientific; cat. #21-236-21) or equivalent
Disposable powder-free gloves and surgical gowns
Aerosol barrier pipette tips
1.5 mL microcentrifuge tubes (DNase/RNase free)
0.2 mL PCR reaction plates (Applied Biosystems; catalog #4346906 or #4366932)
MicroAmp Optical 8-cap Strips (Applied Biosystems; catalog #4323032)

Qualifying Alternative Components:
If a laboratory modifies this test by using unauthorized, alternative components (e.g., extraction
methods or PCR instruments), the modified test is not authorized under this EUA. FDA’s Policy for
Diagnostic Tests for Coronavirus Disease-2019 during the Public Health Emergency, updated May 11,2020, does not change this. As part of this policy, FDA does not intend to object when a laboratory
modifies an EUA-authorized test, which could include using unauthorized components, withoutobtaining an EUA or EUA amendment, where the modified test is validated using a bridging study to the
EUA-authorized test.

Warnings and Precautions•••

For in vitro diagnostic use (IVD).
• This test has not been FDA cleared or approved; this test has been authorized by FDA underan EUA for use by authorized laboratories certified under CLIA, 42 U.S.C. § 263a, that meetrequirements to perform high complexity tests.
• This test has been authorized only for the detection of nucleic acid from SARS CoV-2, notfor any other viruses or pathogens.
• This test is only authorized for the duration of the declaration that circumstances exist
justifying the authorization of emergency use of in vitro diagnostic tests for detection
and/or diagnosis of COVID-19 under Section 564(b)(1) of the Federal Food, Drug andCosmetic Act, 21 U.S.C. § 360bbb-3(b)(1), unless the authorization is terminated or revokedsooner.
Follow standard precautions. All patient specimens and positive controls should be consideredpotentially infectious and handled accordingly.
Do not eat, drink, smoke, apply cosmetics or handle contact lenses in areas where reagents andhuman specimens are handled.

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•
•
•
•
•
•
•

•
•

Handle all specimens as if infectious using safe laboratory procedures. Refer to Interim
Laboratory Biosafety Guidelines for Handling and Processing Specimens Associated with 2019nCoV https://www.cdc.gov/coronavirus/2019-nCoV/lab-biosafety-guidelines.html.
Specimen processing should be performed in accordance with national biological safety
regulations.
If infection with 2019-nCoV is suspected based on current clinical and epidemiological screening
criteria recommended by public health authorities, specimens should be collected withappropriate infection control precautions.
Performance characteristics have been determined with human upper respiratory specimens andlower respiratory tract specimens from human patients with signs and symptoms of respiratory
infection.
Perform all manipulations of live virus samples within a Class II (or higher) biological safetycabinet (BSC).
Use personal protective equipment such as (but not limited to) gloves, eye protection, and labcoats when handling kit reagents while performing this assay and handling materials including
samples, reagents, pipettes, and other equipment and reagents.
Amplification technologies such as PCR are sensitive to accidental introduction of PCR productfrom previous amplifications reactions. Incorrect results could occur if either the clinicalspecimen or the real-time reagents used in the amplification step become contaminated by
accidental introduction of amplification product (amplicon). Workflow in the laboratory should
proceed in a unidirectional manner.
 Maintain separate areas for assay setup and handling of nucleic acids.
 Always check the expiration date prior to use. Do not use expired reagents. Do notsubstitute or mix reagents from different kit lots or from other manufacturers.
 Change aerosol barrier pipette tips between all manual liquid transfers.
 During preparation of samples, compliance with good laboratory techniques is essential
to minimize the risk of cross-contamination between samples and the inadvertent
introduction of nucleases into samples during and after the extraction procedure. Proper
aseptic technique should always be used when working with nucleic acids.
 Maintain separate, dedicated equipment (e.g., pipettes, microcentrifuges) and supplies(e.g., microcentrifuge tubes, pipette tips) for assay setup and handling of extracted
nucleic acids.
 Wear a clean lab coat and powder-free disposable gloves (not previously worn) whensetting up assays.
 Change gloves between samples and whenever contamination is suspected.
 Keep reagent and reaction tubes capped or covered as much as possible.
 Primers, probes (including aliquots), and enzyme master mix must be thawed andmaintained on a cold block at all times during preparation and use.
 Work surfaces, pipettes, and centrifuges should be cleaned and decontaminated withcleaning products such as 10% bleach, DNAZap™, or RNase AWAY™ to minimize risk of
nucleic acid contamination. Residual bleach should be removed using 70% ethanol.
RNA should be maintained on a cold block or on ice during preparation and use to ensurestability.
Dispose of unused kit reagents and human specimens according to local, state, and federal
regulations.
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Reagent Storage, Handling, and Stability
•

•
•
•
•
•

Store all dried primers and probes and the positive control, nCoVPC, at 2-8°C until re-hydrated foruse. Store liquid HSC control materials at ≤ -20°C.
Note: Storage information is for CDC primer and probe materials obtained through theInternational Reagent Resource. If using commercial primers and probes, please refer to the
manufacturer’s instructions for storage and handling.
Always check the expiration date prior to use. Do not use expired reagents.
Protect fluorogenic probes from light.
Primers, probes (including aliquots), and enzyme master mix must be thawed and kept on a cold
block at all times during preparation and use.
Do not refreeze probes.
Controls and aliquots of controls must be thawed and kept on ice at all times during preparation
and use.

Specimen Collection, Handling, and Storage
Inadequate or inappropriate specimen collection, storage, and transport are likely to yield false test
results. Training in specimen collection is highly recommended due to the importance of specimen
quality. CLSI MM13-A may be referenced as an appropriate resource.
The following recommendations for specimen collection, handling, and storage are not for use with
self-collected swabs. Refer to the HealthPulse@home or HealthPulse@home Blueprint instructions
for self- collection, transport, storage, receipt, and accessioning procedures.
 Collecting the Specimen• Refer to Interim Guidelines for Collecting, Handling, and Testing Clinical Specimens for COVID19 https://www.cdc.gov/coronavirus/2019-nCoV/guidelines-clinical-specimens.html• Follow specimen collection device manufacturer instructions for proper collection methods.
• Swab specimens should be collected using only swabs with a synthetic tip, such as nylon or
Dacron®, and an aluminum or plastic shaft. Calcium alginate swabs are unacceptable andcotton swabs with wooden shafts are not recommended. Place swabs immediately into
sterile tubes containing 1-3 mL of appropriate transport media, such as viral transport media(VTM).
 Transporting Specimens• Specimens must be packaged, shipped, and transported according to the current edition of
the International Air Transport Association (IATA) Dangerous Goods Regulation. Follow
shipping regulations for UN 3373 Biological Substance, Category B when sending potential
2019-nCoV specimens. Store specimens at 2-8°C and ship overnight to CDC on ice pack. If a
specimen is frozen at -70°C or lower, ship overnight to CDC on dry ice.
 Storing Specimens• Specimens can be stored at 2-8°C for up to 72 hours after collection.
• If a delay in extraction is expected, store specimens at -70°C or lower.
• Extracted nucleic acid should be stored at -70°C or lower.
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Specimen Referral to CDC
For state and local public health laboratories:• Ship all specimens overnight to CDC.
• Ship frozen specimens on dry ice and non-frozen specimens on cold packs.
• Refer to the International Air Transport Association (IATA - www.iata.org) for requirements
for shipment of human or potentially infectious biological specimens. Follow shipping
regulations for UN 3373 Biological Substance, Category B when sending potential 2019-nCoV
specimens.
• Prior to shipping, notify CDC Division of Viral Diseases (see contact information below) thatyou are sending specimens.
• Send all samples to the following recipient:Centers for Disease Control and Prevention
c/o STATT
Attention: Unit 66
1600 Clifton Rd., Atlanta, GA 30329-4027
Phone: (404) 639-3931
The emergency contact number for CDC Emergency Operations Center (EOC) is
770-488-7100.

All other laboratories that are CLIA certified and meet requirements to perform high complexity
testing:• Please notify your state and/or local public health laboratory for specimen referral andconfirmatory testing guidance.

Reagent and Controls Preparation
NOTE: Storage information is for materials obtained through the CDC International Reagent Resource.
If using commercial products for testing, please refer to the manufacturer’s instructions for storage,
handling, and preparation instructions.
Primer and Probe Preparation:1) Upon receipt, store dried primers and probes at 2-8°C.
2) Precautions: These reagents should only be handled in a clean area and stored at appropriate
temperatures (see below) in the dark. Freeze-thaw cycles should be avoided. Maintain cold
when thawed.
3) Using aseptic technique, suspend dried reagents in 1.5 mL of nuclease-free water and allow
to rehydrate for 15 min at room temperature in the dark.
4) Mix gently and aliquot primers/probe in 300 μL volumes into 5 pre-labeled tubes. Store asingle, working aliquot of primers/probes at 2-8°C in the dark. Store remaining aliquots at
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≤ -20°C in a non-frost-free freezer. Do not refreeze thawed aliquots (stable for up to 4 months
at 2-8°C).
2019-nCoV Positive Control (nCoVPC) Preparation:
1) Precautions: This reagent should be handled with caution in a dedicated nucleic acid handlingarea to prevent possible contamination. Freeze-thaw cycles should be avoided. Maintain on
ice when thawed.
2) Resuspend dried reagent in each tube in 1 mL of nuclease-free water to achieve the proper
concentration. Make single use aliquots (approximately 30 μL) and store at ≤ -70°C.
3) Thaw a single aliquot of diluted positive control for each experiment and hold on ice until
adding to plate. Discard any unused portion of the aliquot.
Human Specimen Control (HSC) (not provided):1) Human Specimen Control (HSC) or one of the listed acceptable alternative extraction controls
must be extracted and processed with each specimen extraction run.
2) Refer to the Human Specimen Control (HSC) package insert for instructions for use.
No Template Control (NTC) (not provided):
1) Sterile, nuclease-free water2) Aliquot in small volumes3) Used to check for contamination during specimen extraction and/or plate set-up

General Preparation
Equipment Preparation
Clean and decontaminate all work surfaces, pipettes, centrifuges, and other equipment prior to use.
Decontamination agents should be used including 10% bleach, 70% ethanol, and DNAZap™, or RNase
AWAY™ to minimize the risk of nucleic acid contamination.
Self-Collected Specimens
If using specimens self-collected with HealthPulse@home or HealthPulse@home Blueprint kits, referto the instructions for use for procedures on rehydration before nucleic acid extraction.

Nucleic Acid Extraction
NOTE: The extraction instructions below are for use when testing individual specimens ONLY. Whenpooling specimens, refer to Appendix B for modified extraction instructions. Procedures for poolingspecimens cannot be performed with self-collected specimens.
Performance of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel is dependent upon the amountand quality of template RNA purified from human specimens. The following commercially available

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RNA extraction kits and procedures have been qualified and validated for recovery and purity of RNA
for use with the panel:
Qiagen QIAamp® DSP Viral RNA Mini Kit or QIAamp® Viral RNA Mini Kit
Recommendation(s): Utilize 100 μL of sample and elute with 100 μL of buffer or utilize 140 μL of
sample and elute with 140 μL of buffer.
Qiagen EZ1 Advanced XL
Kit: Qiagen EZ1 DSP Virus Kit and Buffer AVL (supplied separately) for offboard lysis
Card: EZ1 Advanced XL DSP Virus Card
Recommendation(s): Add 120 μL of sample to 280 μL of pre-aliquoted Buffer AVL (total input sample
volume is 400 μL). Proceed with the extraction on the EZ1 Advanced XL. Elution volume is 120 μL.
Kit: Qiagen EZ1 Virus Mini Kit v2.0 and Buffer AVL (supplied separately) for offboard lysis
Card: EZ1 Advanced XL Virus Card v2.0
Recommendation(s): Add 120 μL of sample to 280 μL of pre-aliquoted Buffer AVL (total input sample
volume is 400 μL). Proceed with the extraction on the EZ1 Advanced XL. Elution volume is 120 μL.
Roche MagNA Pure 96
Kit: Roche MagNA Pure 96 DNA and Viral NA Small Volume Kit
Protocol: Viral NA Plasma Ext LysExt Lys SV 4.0 Protocol or Viral NA Plasma Ext Lys SV Protocol
Recommendation(s): Add 100 μL of sample to 350 μL of pre-aliquoted External Lysis Buffer (supplied
separately) (total input sample volume is 450 μL). Proceed with the extraction on the MagNA Pure 96.
(Internal Control = None). Elution volume is 100 μL.
Roche MagNA Pure 24
Kit: Roche MagNA Pure 24 Total NA Isolation Kit
Protocol: Pathogen 1000 2.0 Protocol
Recommendation(s): Add 100 µL of sample to 400 µL of pre-aliquoted External Lysis Buffer (supplied
separately) (total input sample volume is 500 µL). Proceed with the extraction on the MagNA Pure 24.
(Internal Control = None). Elution volume is 100 µL.
Promega Maxwell® RSC 48 and Maxwell® CSC 48
Kit: Promega Maxwell® Viral Total Nucleic Acid Purification Kit
Protocol: Viral Total Nucleic Acid
Recommendation(s): Add 120 µL of sample to 330 µL of pre-aliquoted External Lysis Buffer (300 µL
Lysis Buffer plus 30 µL Proteinase K; supplied within the kit) (total input volume is 450 µL). Proceedwith the extraction on the Maxwell® RSC 48. Elution volume is 75 µL.
Equivalence and performance of the following extraction platforms were demonstrated with the CDC
Human Influenza Virus Real-Time RT-PCR Diagnostic Panel (K190302) and based on those data are
acceptable for use with the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel.

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QIAGEN QIAcube
Kit: QIAGEN QIAamp® DSP Viral RNA Mini Kit or QIAamp® Viral RNA Mini Kit
Recommendations: Utilize 140 μL of sample and elute with 100 μL of buffer.
Roche MagNA Pure LC
Kit: Roche MagNA Pure Total Nucleic Acid Kit
Protocol: Total NA External_lysis
Recommendation(s): Add 100 μL of sample to 300 μL of pre-aliquoted TNA isolation kit lysis buffer(total input sample volume is 400 μL). Elution volume is 100 μL.
Roche MagNA Pure Compact
Kit: Roche MagNA Pure Nucleic Acid Isolation Kit I
Protocol: Total_NA_Plasma100_400
Recommendation(s): Add 100 μL of sample to 300 μL of pre-aliquoted TNA isolation kit lysis buffer(total input sample volume is 400 μL). Elution volume is 100 μL.
bioMérieux NucliSENS® easyMAG® Instrument
Protocol: General protocol (not for blood) using “Off-board Lysis” reagent settings.
Recommendation(s): Add 100 μL of sample to 1000 μL of pre-aliquoted easyMAG lysis buffer (total
input sample volume is 1100 μL). Incubate for 10 minutes at room temperature.
Elution volume is 100 μL.
bioMérieux EMAG® Instrument
Protocol: Custom protocol: CDC Flu V1 using “Off-board Lysis” reagent settings.
Recommendation(s): Add 100 μL of samples to 2000 μL of pre-aliquoted easyMAG lysis buffer (total
input sample volume is 2100 μL). Incubate for 10 minutes at room temperature. Elution volume is 100
μL. The custom protocol, CDC Flu V1, is programmed on the bioMérieux EMAG® instrument with theassistance of a bioMérieux service representative. Installation verification is documented at the time of
installation. Laboratories are recommended to retain a record of the step-by-step verification of the
bioMérieux custom protocol installation procedure.
Manufacturer’s recommended procedures (except as noted in recommendations above) are to befollowed for sample extraction. HSC must be included in each extraction batch.
Disclaimer: Names of vendors or manufacturers are provided as examples of suitable product sources.
Inclusion does not imply endorsement by the Centers for Disease Control and Prevention.

Assay Set Up
Reaction Master Mix and Plate Set Up
Note: Plate set-up configuration can vary with the number of specimens and workday
organization. NTCs and nCoVPCs must be included in each run.
1) In the reagent set-up room clean hood, place rRT-PCR buffer, enzyme, and primer/probes on
ice or cold-block. Keep cold during preparation and use.
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2) Mix buffer, enzyme, and primer/probes by inversion 5 times.
3) Centrifuge reagents and primers/probes for 5 seconds to collect contents at the bottom of
the tube, and then place the tube in a cold rack.
4) Label one 1.5 mL microcentrifuge tube for each primer/probe set.
5) Determine the number of reactions (N) to set up per assay. It is necessary to make excess
reaction mix for the NTC, nCoVPC, HSC (if included in the rRT-PCR run), and RP reactions andfor pipetting error. Use the following guide to determine N:• If number of samples (n) including controls equals 1 through 14, then N = n + 1• If number of samples (n) including controls is 15 or greater, then N = n + 2
6) For each primer/probe set, calculate the amount of each reagent to be added for eachreaction mixture (N = # of reactions).
Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix
Step #

Vol. of Reagent Added
per Reaction

Reagent

1

Nuclease-free Water

N x 8.5 µL

2Combined Primer/Probe Mix

N x 1.5 µL

3TaqPathTM 1-Step RT-qPCR Master Mix (4x)N x 5.0 µL

Total Volume

N x 15.0 µL

Promega GoTaq® Probe 1- Step RT-qPCR System
Step #

Vol. of Reagent Added
per Reaction

Reagent

1

Nuclease-free Water

N x 3.1 µL

2Combined Primer/Probe Mix

N x 1.5 µL

3GoTaq Probe qPCR Master Mix with dUTP

N x 10.0 µL

4Go Script RT Mix for 1-Step RT-qPCR

N x 0.4 µL

Total Volume

N x 15.0 µL

Quantabio qScript XLT One-Step RT-qPCR ToughMix
Step #

Vol. of Reagent Added
per Reaction

Reagent

1

Nuclease-free Water

N x 3.5 µL

2Combined Primer/Probe Mix

N x 1.5 µL

3qScript XLT One-Step RT-qPCR ToughMix(2X)

N x 10.0 µL

Total Volume

N x 15.0 µL

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Quantabio UltraPlex 1-Step ToughMix (4X)
Step #

Vol. of Reagent Added
per Reaction

Reagent

1

Nuclease-free Water

N x 8.5 µL

2Combined Primer/Probe Mix

N x 1.5 µL

3UltraPlex 1-Step ToughMix (4X)

N x 5.0 µL

Total Volume

N x 15.0 µL

7) Dispense reagents into each respective labeled 1.5 mL microcentrifuge tube. After addition of
the reagents, mix reaction mixtures by pipetting up and down. Do not vortex.
8) Centrifuge for 5 seconds to collect contents at the bottom of the tube, and then place thetube in a cold rack.
9) Set up reaction strip tubes or plates in a 96-well cooler rack.
10) Dispense 15 µL of each master mix into the appropriate wells going across the row as shown
below (Figure 1):
Figure 1: Example of Reaction Master Mix Plate Set-Up
1

2

3

4

5

6

7

8

9

10

11

12

A

N1

N1

N1

N1

N1

N1

N1

N1

N1

N1

N1

N1

B

N2

N2

N2

N2

N2

N2

N2

N2

N2

N2

N2

N2

C

RP

RP

RP

RP

RP

RP

RP

RP

RP

RP

RP

RP

D
E
F
G
H

11) Prior to moving to the nucleic acid handling area, prepare the No Template Control (NTC)
reactions for column #1 in the assay preparation area.
12) Pipette 5 µL of nuclease-free water into the NTC sample wells (Figure 2, column 1). Securely
cap NTC wells before proceeding.
13) Cover the entire reaction plate and move the reaction plate to the specimen nucleic acidhandling area.
Nucleic Acid Template Addition
1) Gently vortex nucleic acid sample tubes for approximately 5 seconds.
2) Centrifuge for 5 seconds to collect contents at the bottom of the tube.
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3) After centrifugation, place extracted nucleic acid sample tubes in the cold rack.
4) Samples should be added to columns 2-11 (column 1 and 12 are for controls) to the specific

5)
6)
7)
8)

9)

assay that is being tested as illustrated in Figure 2. Carefully pipette 5.0 µL of the first sample
into all the wells labeled for that sample (i.e. Sample “S1” down column #2). Keep othersample wells covered during addition. Change tips after each addition.
Securely cap the column to which the sample has been added to prevent cross contaminationand to ensure sample tracking.
Change gloves often and when necessary to avoid contamination.
Repeat steps #4 and #5 for the remaining samples.
If necessary, add 5 µL of Human Specimen Control (HSC) extracted sample to the HSC wells(Figure 2, column 11). Securely cap wells after addition. NOTE: Per CLIA regulations, HSC
must be tested at least once per day.
Cover the entire reaction plate and move the reaction plate to the positive template control
handling area.

Assay Control Addition
1) Pipette 5 µL of nCoVPC RNA to the sample wells of column 12 (Figure 2). Securely cap wells
after addition of the control RNA.
NOTE: If using 8-tube strips, label the TAB of each strip to indicate sample position. DO NOTLABEL THE TOPS OF THE REACTION TUBES!
2) Briefly centrifuge reaction tube strips for 10-15 seconds. After centrifugation return to coldrack.
NOTE: If using 96-well plates, centrifuge plates for 30 seconds at 500 x g, 4°C.
Figure 2. 2019-nCoV rRT-PCR Diagnostic Panel: Example of Sample and Control Set-up
1

2

3

4

5

6

7

8

9

10

11a

12

A

NTC

S1

S2

S3

S4

S5

S6

S7

S8

S9

S10

nCoV PC

B

NTC

S1

S2

S3

S4

S5

S6

S7

S8

S9

S10

nCoV PC

C

NTC

S1

S2

S3

S4

S5

S6

S7

S8

S9

S10

nCoV PC

D
E
F
G
H
a

Replace the sample in this column with extracted HSC if necessary

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Create a Run Template on the Applied Biosystems 7500 Fast Dx Real-time PCR
Instrument (Required if no template exists)
If the template already exists on your instrument, please proceed to the RUNNING A TEST section.
1) Launch the Applied Biosystems 7500 Fast Dx Real-time PCR Instrument by double clicking on the
Applied Biosystems 7500 Fast Dx System icon on the desktop.
2) A new window should appear, select Create New Document from the menu.
Figure 3. New Document Wizard Window

Make sure to change
Run Mode to
STANDARD 7500

3) The New Document Wizard screen in Figure 3 will appear. Select:
a. Assay: Standard Curve (Absolute Quantitation)b. Container: 96-Well Clear
c. Template: Blank Documentd. Run Mode: Standard 7500
e. Operator: Your Name
f. Comments: SDS v1.4
g. Plate Name: Your Choice4) After making selections click Next at the bottom of the window.

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Figure 4. Creating New Detectors

NOTE: ROX is the default passive reference. This will be changed to “none” in step 12.

5) After selecting next, the Select Detectors screen (Figure 4) will appear.
6) Click the New Detector button (see Figure 4).
7) The New Detector window will appear (Figure 5). A new detector will need to be defined foreach primer and probe set. Creating these detectors will enable you to analyze each primer andprobe set individually at the end of the reaction.

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Figure 5. New Detector Window

8) Start by creating the N1 Detector. Include the following:a. Name: N1
b. Description: leave blankc. Reporter Dye: FAM
d. Quencher Dye: (none)e. Color: to change the color of the detector indicator do the following:⇒ Click on the color square to reveal the color chart⇒ Select a color by clicking on one of the squares⇒ After selecting a color click OK to return to the New Detector screen
f. Click the OK button of the New Detector screen to return to the screen shown in Figure 4.
9) Repeat step 6-8 for each target in the panel.

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Reporter Dye

Quencher Dye

N1

FAM

(none)

N2

FAM(none)

RP

FAM(none)

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10) After each Detector is added, the Detector Name, Description, Reporter and Quencher fields
will become populated in the Select Detectors screen (Figure 6).
11) Before proceeding, the newly created detectors must be added to the document. To add the
new detectors to the document, click ADD (see Figure 6). Detector names will appear on theright-hand side of the Select Detectors window (Figure 6).
Figure 6. Adding New Detectors to Document

12) Once all detectors have been added, select (none) for Passive Reference at the top right-handdrop-down menu (Figure 7).
Figure 7. Select Passive Reference

Passive reference should be set to “(none)” as described above.

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13) Click Next at the bottom of the Select Detectors window to proceed to the Set Up Sample Plate
window (Figure 8).
14) In the Set Up Sample Plate window (Figure 8), use your mouse to select row A from the lower
portion of the window, in the spreadsheet (see Figure 8).
15) In the top portion of the window, select detector N1. A check will appear next to the detector
you have selected (Figure 8). You will also notice the row in the spreadsheet will be populatedwith a colored “U” icon to indicate which detector you’ve selected.
16) Repeat step 14-15 for each detector that will be used in the assay.
Figure 8. Sample Plate Set-up17) Select Finish after detectors have been assigned to their respective rows. (Figure 9).
Figure 9. Finished Plate Set-up

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18) After clicking “Finish”, there will be a brief pause allowing the Applied Biosystems 7500 Fast Dx
to initialize. This initialization is followed by a clicking noise. Note: The machine must be turned
on for initialization.
19) After initialization, the Plate tab of the Setup (Figure 10) will appear.
20) Each well of the plate should contain colored U icons that correspond with the detector labelsthat were previously chosen. To confirm detector assignments, select Tools from the file menu,
then select Detector Manager.
Figure 10. Plate Set-up Window

21) The Detector Manager window will appear (Figure 11).
Figure 11. Detector Manager Window

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22) Confirm all detectors are included and that each target has a Reporter set to FAM and theQuencher is set to (none).
23) If all detectors are present, select Done. The detector information has been created andassigned to wells on the plate.

Defining the Instrument Settings
1) After detectors have been created and assigned, proceed to instrument set up.
2) Select the Instrument tab to define thermal cycling conditions.
3) Modify the thermal cycling conditions as follows (Figure 12):
Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix, CG
a.
b.
c.
d.
e.
f.
g.
h.
i.

In Stage 1, Set to 2 min at 25°C; 1 Rep.
In Stage 2, Set to 15 min at 50°C; 1 Rep.
In Stage 3, Set to 2 min at 95°C, 1 Rep.
In Stage 4, Step 1 set to 3 sec at 95°C.
In Stage 4, Step 2 set to 30 sec at 55.0°C.
In Stage 4, Reps should be set to 45.
Under Settings (Figure 12), bottom left-hand box, change volume to 20 µL.
Under Settings, Run Mode selection should be Standard 7500.
Step 2 of Stage 4 should be highlighted in yellow to indicate data collection (see Figure
12).

OR
Quantabio qScriptTM XLT One-Step RT-qPCR ToughMix or UltraPlex 1-Step ToughMix (4X)
a.
b.
c.
d.
e.
f.
g.
h.

In Stage 1, Set to 10 min at 50°C; 1 Rep.
In Stage 2, Set to 3 min at 95°C, 1 Rep.
In Stage 3, Step 1 set to 3 sec at 95°C.
In Stage 3, Step 2 set to 30 sec at 55.0°C.
In Stage 3, Reps should be set to 45.
Under Settings (Figure 12), bottom left-hand box, change volume to 20 µL.
Under Settings, Run Mode selection should be Standard 7500.
Step 2 of Stage 3 should be highlighted in yellow to indicate data collection (see Figure
12).

OR
Promega GoTaq® Probe 1-Step RT-qPCR Systema. In Stage 1, Set to 15 min at 45°C; 1 Rep.
b. In Stage 2, Set to 2 min at 95°C, 1 Rep.
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c.
d.
e.
f.
g.
h.

In Stage 3, Step 1 set to 3 sec at 95°C.
In Stage 3, Step 2 set to 30 sec at 55.0°C.
In Stage 3, Reps should be set to 45.
Under Settings (Figure 12), bottom left-hand box, change volume to 20 µL.
Under Settings, Run Mode selection should be Standard 7500.
Step 2 of Stage 3 should be highlighted in yellow to indicate data collection (see Figure
12).

Figure 12. Instrument Window

4) After making changes to the Instrument tab, the template file is ready to be saved. To save thetemplate, select File from the top menu, then select Save As. Since the enzyme options havedifferent instrument settings, it is recommended that the template be saved with a nameindicating the enzyme option.
5) Save the template as 2019-nCoV Dx Panel TaqPath or 2019-nCoV Dx Panel Quanta or 2019nCoV Dx Panel Promega as appropriate in the desktop folder labeled “ABI Run Templates” (youmust create this folder). Save as type should be SDS Templates (*.sdt) (Figure 13).

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Figure 13. Saving Template

Running a Test
1) Turn on the ABI 7500 Fast Dx Real-Time PCR Instrument.
2) Launch the Applied Biosystems 7500 Fast Dx Real-time PCR System by double clicking on the7500 Fast Dx System icon on the desktop.
3) A new window should appear, select Open Existing Document from the menu.
4) Navigate to select your ABI Run Template folder from the desktop.
5) Double click on the appropriate template file (2019-nCoV Dx Panel TaqPath or 2019-nCoV DxPanel Quanta or 2019-nCoV Dx Panel Promega)6) There will be a brief pause allowing the Applied Biosystems 7500 Fast Dx Real-Time PCR
Instrument to initialize. This initialization is followed by a clicking noise. Note: The machine mustbe turned on for initialization.

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Figure 14. Plate Set-up Window

7) After the instrument initializes, a plate map will appear (Figure 14). The detectors and controls
should already be labeled as they were assigned in the original template.
8) Click the Well Inspector iconfrom the top menu.
9) Highlight specimen wells of interest on the plate map.
10) Type sample identifiers to Sample Name box in the Well Inspector window (Figure 15).

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Figure 15. Labeling Wells

11) Repeat steps 9-10 until all sample identifiers are added to the plate setup.
12) Once all specimen and control identifiers are added click the Close button on the Well Inspector
window to return to the Plate set up tab.
13) Click the Instrument tab at the upper left corner.
14) The reaction conditions, volumes, and type of 7500 reaction should already be loaded (Figure16).

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Figure 16. Instrument Settings

15) Ensure settings are correct (refer to the Defining Instrument Settings).
16) Before proceeding, the run file must be saved; from the main menu, select File, then Save As.
Save in appropriate run folder designation.
17) Load the plate into the plate holder in the instrument. Ensure that the plate is properly alignedin the holder.
18) Once the run file is saved, click the Start button. Note: The run should take approximately 1 hourand 20 minutes to complete.

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Data Analysis
1) After the run has completed, select the Results tab at the upper left corner of the software.
2) Select the Amplification Plot tab to view the raw data (Figure 17).
Figure 17. Amplification Plot Window

b
c
d
e

a

c

c

3) Start by highlighting all the samples from the run; to do this, click on the upper left-hand box (a)of the sample wells (Figure 17). All the growth curves should appear on the graph.
4) On the right-hand side of the window (b), the Data drop down selection should be set to Delta
Rn vs. Cycle.
5) Select N1 from (c), the Detector drop down menu, using the downward arrow.
a. Please note that each detector is analyzed individually to reflect different
performance profiles of each primer and probe set.
6) In the Line Color drop down (d), Detector Color should be selected.
7) Under Analysis Settings select Manual Ct (e).
b. Do not change the Manual Baseline default numbers.
8) Using the mouse, click and drag the red threshold line until it lies within the exponential phaseof the fluorescence curves and above any background signal (Figure 18).

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Figure 18. Amplification Plot

Exponential
PCR PhaseThreshold adjusted
to fall within the
PCR exponential
phase.

Background noise9) Click the Analyze button in the lower right corner of the window. The red threshold line will turnto green, indicating the data has been analyzed.
10) Repeat steps 5-9 to analyze results generated for each set of markers (N1, N2, RP).
11) Save analysis file by selecting File then Save As from the main menu.
12) After completing analysis for each of the markers, select the Report tab above the graph todisplay the Ct values (Figure 19). To filter report by sample name in ascending or descendingorder, simply click on Sample Name in the table.
Figure 19. Report

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Interpretation of Results and Reporting
Extraction and Positive Control Results and Interpretation
No Template Control (NTC)
The NTC consists of using nuclease-free water in the rRT-PCR reactions instead of RNA. The NTC
reactions for all primer and probe sets should not exhibit fluorescence growth curves that cross thethreshold line. If any of the NTC reactions exhibit a growth curve that crosses the cycle threshold, sample
contamination may have occurred. Invalidate the run and repeat the assay with strict adherence to theguidelines.
2019-nCoV Positive Control (nCoVPC)The nCoVPC consists of in vitro transcribed RNA. The nCoVPC will yield a positive result with thefollowing primer and probe sets: N1, N2, and RP.
Human Specimen Control (HSC) (Extraction Control)
When HSC is run with the CDC 2019-nCoV rRT-PCR Diagnostic Panel (see previous section on Assay Set
Up), the HSC is used as a nucleic acid extraction procedural control to demonstrate successful recoveryof nucleic acid as well as extraction reagent integrity. The HSC control consists of noninfectious cultured
human cell (A549) material. Purified nucleic acid from the HSC should yield a positive result with the RP
primer and probe set and negative results with all 2019-nCoV markers.
Expected Performance of Controls Included in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Control
Type

External
Control
Name

Used toMonitor2019
nCoV_N1

2019
nCoV_N2

RP

Expected Ct
Values+++< 40.00 Ct

Positive

nCoVPC

Substantial
reagent failure
includingprimer andprobe integrity

Negative

NTC

Reagent and/or
environmental
contamination

-

-

-

None
detected

HSC

Failure in lysis
and extraction
procedure,
potential
contamination
duringextraction

-

-

+

< 40.00 Ct

Extraction

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If any of the above controls do not exhibit the expected performance as described, the assay may havebeen set up and/or executed improperly, or reagent or equipment malfunction could have occurred.
Invalidate the run and re-test.
RNase P (Extraction Control) All clinical samples should exhibit fluorescence growth curves in the RNase P reaction that crossthe threshold line within 40.00 cycles (< 40.00 Ct), thus indicating the presence of the human
RNase P gene. Failure to detect RNase P in any clinical specimens may indicate:− Improper extraction of nucleic acid from clinical materials resulting in loss of RNA and/or
RNA degradation.
− Absence of sufficient human cellular material due to poor collection or loss of specimen
integrity.
− Improper assay set up and execution.
− Reagent or equipment malfunction.
 If the RP assay does not produce a positive result for human clinical specimens, interpret as
follows:− If the 2019-nCoV N1 and N2 are positive even in the absence of a positive RP, the result
should be considered valid. It is possible, that some samples may fail to exhibit RNase P
growth curves due to low cell numbers in the original clinical sample. A negative RP signal
does not preclude the presence of 2019-nCoV virus RNA in a clinical specimen.
− If all 2019-nCoV markers AND RNase P are negative for the specimen, the result should beconsidered invalid for the specimen. If residual specimen is available, repeat the extraction
procedure and repeat the test. If all markers remain negative after re-test, report theresults as invalid and a new specimen should be collected if possible.
2019-nCoV Markers (N1 and N2)• When all controls exhibit the expected performance, a specimen is considered negative if all
2019-nCoV marker (N1, N2) cycle threshold growth curves DO NOT cross the threshold line
within 40.00 cycles (< 40.00 Ct) AND the RNase P growth curve DOES cross the threshold line
within 40.00 cycles (< 40.00 Ct).
• When all controls exhibit the expected performance, a specimen is considered positive for 2019nCoV if all 2019-nCoV marker (N1, N2) cycle threshold growth curves cross the threshold line
within 40.00 cycles (< 40.00 Ct). The RNase P may or may not be positive as described above, but
the 2019-nCoV result is still valid.
• When all controls exhibit the expected performance and the growth curves for the 2019-nCoV
markers (N1, N2) AND the RNase P marker DO NOT cross the cycle threshold growth curve within
40.00 cycles (< 40.00 Ct), the result is invalid. The extracted RNA from the specimen should be retested. If residual RNA is not available, re-extract RNA from residual specimen and re-test. If there-tested sample is negative for all markers and RNase P, the result is invalid and collection of a
new specimen from the patient should be considered.
• When all controls exhibit the expected performance and the cycle threshold growth curve for any
one marker (N1 or N2, but not both markers) crosses the threshold line within 40.00 cycles (<40.00 Ct) the result is inconclusive. The extracted RNA should be retested. If residual RNA is notavailable, re-extract RNA from residual specimen and re-test. If the same result is obtained,
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•report the inconclusive result. Consult with your state public health laboratory or CDC, asappropriate, to request guidance and/or to coordinate transfer of the specimen for additional
analysis.
If HSC is positive for N1 or N2, then contamination may have occurred during extraction or
sample processing. Invalidate all results for specimens extracted alongside the HSC. Re-extract
specimens and HSC and re-test.

2019-nCoV rRT-PCR Diagnostic Panel Results Interpretation Guide
The table below lists the expected results for the 2019-nCoV rRT-PCR Diagnostic Panel. If a laboratory
obtains unexpected results for assay controls or if inconclusive or invalid results are obtained and cannot
be resolved through the recommended re-testing, please contact CDC for consultation and possible
specimen referral. See pages 13 and 53 for referral and contact information.
2019
nCoV_N1

2019
nCoV_N2

RP

ResultInterpretationa

ReportActions++±

2019-nCoV
detected

Positive 2019-nCoV

Report results to CDC andsender.

If only one of the twotargets is positive±

Inconclusive
ResultInconclusive

-

-

+

2019-nCoV notdetected

Not Detected

-

-

-

Invalid Result

Invalid

Repeat testing of nucleic acidand/or re-extract and repeatrRT-PCR. If the repeated result
remains inconclusive, contact
your State Public Health
Laboratory or CDC for
instructions for transfer of thespecimen or further guidance.
Report results to sender.
Consider testing for otherrespiratory viruses.b
Repeat extraction and rRT-PCR.
If the repeated result remains
invalid, consider collecting a
new specimen from the patient.

aLaboratories should report their diagnostic result as appropriate and in compliance with their specific reporting

system.

bOptimum specimen types and timing for peak viral levels during infections caused by 2019-nCoV have not been

determined. Collection of multiple specimens from the same patient may be necessary to detect the virus. The
possibility of a false negative result should especially be considered if the patient’s recent exposures or clinicalpresentation suggest that 2019-nCoV infection is possible, and diagnostic tests for other causes of illness (e.g.,other respiratory illness) are negative. If 2019-nCoV infection is still suspected, re-testing should be considered
in consultation with public health authorities.

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Quality Control
•

•
•
•
•
•

Quality control requirements must be performed in conformance with local, state, and federal
regulations or accreditation requirements and the user’s laboratory’s standard quality control
procedures. For further guidance on appropriate quality control practices, refer to 42 CFR
493.1256.
Quality control procedures are intended to monitor reagent and assay performance.
Test all positive controls prior to running diagnostic samples with each new kit lot to ensure all
reagents and kit components are working properly.
Good laboratory practice (cGLP) recommends including a positive extraction control in eachnucleic acid isolation batch.
Although HSC is not included with the 2019-nCov rRT-PCR Diagnostic Panel, the HSC extraction
control must proceed through nucleic acid isolation per batch of specimens to be tested.
Always include a negative template control (NTC) and the appropriate positive control (nCoVPC)
in each amplification and detection run. All clinical samples should be tested for human RNase P
gene to control for specimen quality and extraction.

Limitations
•
•

•

•
•
•
•

All users, analysts, and any person reporting diagnostic results should be trained to perform thisprocedure by a competent instructor. They should demonstrate their ability to perform the test
and interpret the results prior to performing the assay independently.
Performance of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel has only been establishedin upper and lower respiratory specimens (such as nasopharyngeal or oropharyngeal swabs,
sputum, lower respiratory tract aspirates, bronchoalveolar lavage, and nasopharyngeal
wash/aspirate or nasal aspirate).
Negative results do not preclude 2019-nCoV infection and should not be used as the sole basis
for treatment or other patient management decisions. Optimum specimen types and timing forpeak viral levels during infections caused by 2019-nCoV have not been determined. Collection ofmultiple specimens (types and time points) from the same patient may be necessary to detect
the virus.
A false-negative result may occur if a specimen is improperly collected, transported or handled.
False-negative results may also occur if amplification inhibitors are present in the specimen or if
inadequate numbers of organisms are present in the specimen.
Positive and negative predictive values are highly dependent on prevalence. False-negative test
results are more likely when prevalence of disease is high. False-positive test results are more
likely when prevalence is moderate to low.
Do not use any reagent past the expiration date.
The Performance of this test was established based on the evaluation of a limited number ofclinical specimens. The clinical performance has not been established in all circulating variants
but is anticipated to be reflective of the prevalent variants in circulation at the time and locationof the clinical evaluation. Performance at the time of testing may vary depending on the variants
circulating, including newly emerging strains of SARSCoV-2 and their prevalence, which changeover time.

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• If the virus mutates in the rRT-PCR target region, 2019-nCoV may not be detected or may be
detected less predictably. Inhibitors or other types of interference may produce a false-negative
result. An interference study evaluating the effect of common cold medications was not
performed.
• Test performance can be affected because the epidemiology and clinical spectrum of infection
caused by 2019-nCoV is not fully known. For example, clinicians and laboratories may not know
the optimum types of specimens to collect, and, during the course of infection, when these
specimens are most likely to contain levels of viral RNA that can be readily detected.
• Detection of viral RNA may not indicate the presence of infectious virus or that 2019-nCoV is thecausative agent for clinical symptoms.
• The performance of this test has not been established for monitoring treatment of 2019-nCoV
infection.
• The performance of this test has not been established for screening of blood or blood products
for the presence of 2019-nCoV.
• This test cannot rule out diseases caused by other bacterial or viral pathogens.

Conditions of Authorization for the Laboratory
The CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel Letter of Authorization, along with theauthorized Fact Sheet for Healthcare Providers, the authorized Fact Sheet for Patients, andauthorized labeling are available on the FDA website: https://www.fda.gov/medicaldevices/coronavirus-disease-2019-covid-19-emergency-use-authorizations-medical-devices/vitrodiagnostics-euas.
However, to assist clinical laboratories using the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel(“your product” in the conditions below), the relevant Conditions of Authorization are listed below:•

Authorized laboratories using your product will include with test result reports, all authorized
Fact Sheets available on the CDC website. Under exigent circumstances, other appropriate
methods for disseminating these Fact Sheets may be used, which may include mass media.

•

Authorized laboratories using your product will use your product as outlined in the authorizedlabeling available on the CDC website. Deviations from the authorized procedures, including theauthorized RT-PCR instruments, authorized extraction methods, authorized clinical specimen
types, authorized control materials, authorized other ancillary reagents and authorized materials
required to use your product are not permitted under this authorization.

•

Authorized laboratories that receive the commercially manufactured and distributed primer andprobe sets identified as acceptable on the CDC website for use with your product, and are notable to obtain the authorized Human Specimen Control and authorized Positive Control for 2019nCoV (nCoVPC) materials described in your product’s authorized labeling, may use appropriatematerials identified as acceptable materials on the CDC website for use with your product.

•

Authorized laboratories when testing authorized specimens self-collected using home-collection
kits authorized for use with your product must have in place a suitable specimen receipt and

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accessioning protocol SOP and/or follow any specimen accessioning protocol provided with theself-collection kit when accepting specimens for testing.
•Authorized laboratories that receive your product will notify the relevant public healthauthorities of their intent to run your product prior to initiating testing.

•Authorized laboratories using your product will have a process in place for reporting test results
to healthcare providers and relevant public health authorities, as appropriate.

•

Authorized laboratories will collect information on the performance of your product and report
to DMD/OHT7-OIR/OPEQ/CDRH (via email: CDRH-EUA-Reporting@fda.hhs.gov) and CDC(respvirus@cdc.gov) any suspected occurrence of false positive or false negative results andsignificant deviations from the established performance characteristics of the test of which they
become aware.

•

Authorized laboratories using specimen pooling strategies when testing patient specimens withyour product will include with negative test result reports for specific patients whose specimen(s)were the subject of pooling, a notice that pooling was used during testing and that “Patient
specimens with low viral loads may not be detected in sample pools due to the decreased
sensitivity of pooled testing.”•

Authorized laboratories implementing pooling strategies for testing patient specimens must use
the “Implementation and Monitoring of Pooled Specimen Testing” available in the authorizedlabeling to evaluate the appropriateness of continuing to use such strategies based on therecommendations in the protocol.

•

Authorized laboratories will keep records of specimen pooling strategies implemented including
type of strategy, date implemented, and quantities tested, and test result data generated as partof the Protocol for Monitoring of Specimen Pooling Testing Strategies. For the first 12 months
from the date of their creation, such records will be made available to FDA within 48 businesshours (2 business days) for inspection upon request, and will be made available within areasonable time after 12 months from the date of their creation.

•

Authorized laboratories will report adverse events, including problems with your products
performance or results, to MedWatch by submitting the online FDA Form 3500
(https://www.accessdata.fda.gov/scripts/medwatch/index.cfm?action=reporting.home) or bycalling 1-800-FDA-1088.

•

All laboratory personnel using the test must be appropriately trained in RT-PCR techniques anduse appropriate laboratory and personal protective equipment when handling this kit and use
the test in accordance with the authorized labeling.

•

CDC, IRR, manufacturers and distributors of commercial materials identified as acceptable on the
CDC website, and authorized laboratories using your product will ensure that any records
associated with this EUA are maintained until otherwise notified by FDA. Such records will bemade available to FDA for inspection upon request.

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Performance Characteristics
Analytical Performance:
Limit of Detection (LoD):
LoD studies determine the lowest detectable concentration of 2019-nCoV at which approximately 95%
of all (true positive) replicates test positive. The LoD was determined by limiting dilution studies using
characterized samples.
The analytical sensitivity of the rRT-PCR assays contained in the CDC 2019 Novel Coronavirus (2019nCoV) Real-Time RT-PCR Diagnostic Panel were determined in Limit of Detection studies. Since noquantified virus isolates of the 2019-nCoV were available for CDC use at the time the test was developed
and this study conducted, assays designed for detection of the 2019-nCoV RNA were tested withcharacterized stocks of in vitro transcribed full length RNA (N gene; GenBank accession: MN908947.2) of
known titer (RNA copies/µL) spiked into a diluent consisting of a suspension of human A549 cells andviral transport medium (VTM) to mimic clinical specimen. Samples were extracted using the QIAGEN EZ1
Advanced XL instrument and EZ1 DSP Virus Kit (Cat# 62724) and manually with the QIAGEN DSP Viral
RNA Mini Kit (Cat# 61904). Real-Time RT-PCR assays were performed using the Thermo Fisher Scientific
TaqPath™ 1-Step RT-qPCR Master Mix, CG (Cat# A15299) on the Applied Biosystems™ 7500 Fast Dx RealTime PCR Instrument according to the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel instructions
for use.
A preliminary LoD for each assay was determined testing triplicate samples of RNA purified using eachextraction method. The approximate LoD was identified by extracting and testing 10-fold serial dilutionsof characterized stocks of in vitro transcribed full-length RNA. A confirmation of the LoD was determinedusing 3-fold serial dilution RNA samples with 20 extracted replicates. The LoD was determined as thelowest concentration where ≥ 95% (19/20) of the replicates were positive.
Table 4. Limit of Detection Confirmation of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panelwith QIAGEN EZ1 DSP
Targets
2019-nCoV_N1
2019-nCoV_N2
1
0.5
0.0
-0.5
0.5
RNA Concentration
10
10
10
10
10 0.0 10 -0.5

1

Positives/Total
Mean Ct2
Standard Deviation
(Ct)

20/20
32.5

19/20
35.4

13/20
NA

20/20
35.8

17/20
NA

9/20
NA

0.5

0.8

NA

1.3

NA

NA

Concentration is presented in RNA copies/µL

2 Mean Ct reported for dilutions that are ≥ 95% positive. Calculations only include positive results.

NA not applicable

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Table 5. Limit of Detection Confirmation CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel withQIAGEN QIAmp DSP Viral RNA Mini Kit
2019-nCoV_N1
2019-nCoV_N2
Targets
RNA Concentration1 10 0.5 10 0.0 10 -0.5 10 0.5 10 0.0 10 -0.5 10 -1.0
20/20 20/20 6/20 20/20 20/20 20/20 8/20
Positives/Total
32.0
32.8
NA
33.0
35.4
36.2
NA
Mean Ct2
Standard Deviation
0.7
0.8
NA
1.4
0.9
1.9
NA
(Ct)
1

Concentration is presented in RNA copies/µL

2 Mean Ct reported for dilutions that are ≥ 95% positive. Calculations only include positive results.

NA not applicable

Table 6. Limit of Detection of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Limit of Detection (RNA copies/µL)
Virus
Material
QIAGEN EZ1
QIAGEN DSP Viral
Advanced XL
RNA Mini Kit
2019 Novel
N Gene RNA
100.5
100
Coronavirus
Transcript
FDA Sensitivity Evaluation: The analytical sensitivity of the test will be further assessed by evaluating an
FDA-recommended reference material using an FDA developed protocol if applicable and/or whenavailable.
FDA SARS-CoV-2 Reference Panel Testing
The evaluation of sensitivity and MERS-CoV cross-reactivity was performed using reference material (T1),blinded samples and a standard protocol provided by the FDA. The study included a range finding study
and a confirmatory study for LoD. Blinded sample testing was used to establish specificity and to confirm
the LoD. Samples were extracted using the QIAGEN EZ1 Advanced XL with the QIAGEN EZ1 DSP Virus Kit.
Extracted samples were then tested using the 2019-nCoV Real-Time RT-PCR Diagnostic Panel on the
Applied BioSystems 7500 Fast Dx Real-Time PCR Instrument using the ThermoFisher TaqPath™ 1-Step
RT-qPCR Master Mix. The results are summarized in Table 7.
Table 7: Summary of LoD Confirmation Result using the FDA SARS-CoV-2 Reference Panel
Reference Materials Provided by FDA
SARS-CoV-2
MERS-CoVNDU/mL = RNA NAAT detectable units/mL
N/A: Not applicable
ND: Not detected

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Specimen Type

Product LoD

CrossReactivity

NP swab1.8x104 NDU/mL
N/A

N/A
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In Silico Analysis of Primer and Probe Sequences:
The oligonucleotide primer and probe sequences of the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel were evaluated against 615,472 high-quality genome sequences available in Global
Initiative on Sharing All Influenza Data (GISAID, https://www.gisaid.org) as of March 18, 2021 todemonstrate the predicted inclusivity of the 2019-nCoV real-time RT-PCR Diagnostic panel.
Nucleotide mismatches in the primer/probe regions with frequency of >0.1% are shown in the tablebelow. Mismatch frequency of the N1 reverse primer was less than 0.1% and is therefore not listedon the table. Frequency of mismatches for all the individual primers and probes was <1% andvariable in location indicating that the mismatches were sporadic and not indicative of a consistent
mutation. One hundred fifty-eight sequences out of 615,472 had two nucleotide mismatches within
the same primer/probe sequence.
In summary, the assessment of homology between available sequences of SARS CoV-2 as of March
18, 2021 and the CDC panel assay primers and probes shows that the risk of significant loss inreactivity and false negative results is very low due to the absence of significant numbers of
mismatches. The design of the primers and probes, with melting temperatures of >60°C and an
annealing temperature of 55°C, can tolerate up to two mismatches depending on location with nosignificant loss in assay sensitivity.
2019-nCoV primer and probe mismatches with frequency >0.1% among 615,472 high-quality
genome sequences available from GISAID as of March 18, 2021.

Table 8. In Silico Inclusivity Analysis of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel Among
615,472 Genome Sequences Available from GISAID as of March 18, 2021
Primer/probe
N1 forward
N1 probe
N2 forward
N2 probe
N2 reverse
5
6
14
2
3
5
13
4
8
16
7
10
13
4
13
Location (5'>3')
Mismatch Nucleotide C>T C>A G>T C>A/C>T C>T C>T G>A/G>T C>T C>T G>T T>C C>T C>T C>A G>A
994 1036 2554 1394/996 3694 690 860/1026 1452 924 5277 1966 3292 1444 4499 733
Mismatch No.
0.16
0.17 0.41 0.23/0.16 0.60 0.11 0.14/0.17 0.24 0.15 0.86 0.32 0.53 0.23 0.73 0.12
Mismatch Frequency(%)
Specificity/Exclusivity Testing: In Silico Analysis
BLASTn analysis queries of the 2019-nCoV rRT-PCR assays primers and probes were performed against
public domain nucleotide sequences. The database search parameters were as follows: 1) The nucleotide
collection consists of GenBank+EMBL+DDBJ+PDB+RefSeq sequences, but excludes EST, STS, GSS, WGS,
TSA, patent sequences as well as phase 0, 1, and 2 HTGS sequences and sequences longer than 100Mb;
2) The database is non-redundant. Identical sequences have been merged into one entry, while
preserving the accession, GI, title and taxonomy information for each entry; 3) Database was updated on
10/03/2019; 4) The search parameters automatically adjust for short input sequences and the expect
threshold is 1000; 5) The match and mismatch scores are 1 and -3, respectively; 6) The penalty to create
and extend a gap in an alignment is 5 and 2 respectively.

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2019-nCoV_N1 Assay:
Probe sequence of 2019-nCoV rRT-PCR assay N1 showed high sequence homology with SARS
coronavirus and Bat SARS-like coronavirus genome. However, forward and reverse primers showed nosequence homology with SARS coronavirus and Bat SARS-like coronavirus genome. Combining primers
and probe, there is no significant homologies with human genome, other coronaviruses or humanmicroflora that would predict potential false positive rRT-PCR results.
2019-nCoV_N2 Assay:
The forward primer sequence of 2019-nCoV rRT-PCR assay N2 showed high sequence homology to Bat
SARS-like coronaviruses. The reverse primer and probe sequences showed no significant homology withhuman genome, other coronaviruses or human microflora. Combining primers and probe, there is no
prediction of potential false positive rRT-PCR results.
In summary, the 2019-nCoV rRT-PCR assay N1 and N2, designed for the specific detection of 2019-nCoV,
showed no significant combined homologies with human genome, other coronaviruses, or humanmicroflora that would predict potential false positive rRT-PCR results.
In addition to the in silico analysis, several organisms were extracted and tested with the CDC 2019-nCoV
Real-Time RT-PCR Diagnostic Panel to demonstrate analytical specificity and exclusivity. Studies were
performed with nucleic acids extracted using the QIAGEN EZ1 Advanced XL instrument and EZ1 DSP
Virus Kit. Nucleic acids were extracted from high titer preparations (typically ≥ 105 PFU/mL or ≥ 106
CFU/mL). Testing was performed using the Thermo Fisher Scientific TaqPath™ 1-Step RT-qPCR Master
Mix, CG on the Applied Biosystems™ 7500 Fast Dx Real-Time PCR instrument. The data demonstrate thatthe expected results are obtained for each organism when tested with the CDC 2019-nCoV Real-Time RTPCR Diagnostic Panel.
Table 9. Specificity/Exclusivity of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Virus
Human coronavirus
Human coronavirus
Human coronavirus
Human coronavirus
MERS-coronavirus
SARS-coronavirus
bocavirus
Mycoplasma pneumoniae
Streptococcus
Influenza A(H1N1)
Influenza A(H3N2)
Influenza B
Human adenovirus, type 1
Human metapneumovirus
respiratory syncytial virus
rhinovirus
parainfluenza 1
parainfluenza 2
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StrainSource

229E
OC43
NL63
HKU1

IsolateIsolateclinical specimen
clinical specimen
IsolateIsolateclinical specimen
IsolateIsolateIsolateIsolateIsolateIsolateIsolateIsolateIsolateIsolateIsolate-

Ad71
Long A
C35
Greer

43

2019nCoV_
N1
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3

2019nCoV_
N2
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3
0/3

CDC/DDID/NCIRD/ Division of Viral Diseases

Final
Result
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.
Neg.

Effective: 03/07/2023

parainfluenza 3parainfluenza 4

C-43
M-25

IsolateIsolate

0/3
0/3

0/3
0/3

Neg.
Neg.

Endogenous Interference Substances Studies:
The CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel uses conventional well-established nucleic acidextraction methods and based on our experience with CDC’s other EUA assays, including the CDC Novel
Coronavirus 2012 Real-time RT-PCR Assay for the presumptive detection of Middle East Respiratory
Syndrome Coronavirus (MERS-CoV) and the CDC Human Influenza Virus Real-Time RT-PCR Diagnostic
Panel-Influenza A/H7 (Eurasian Lineage) Assay for the presumptive detection of novel influenza A (H7N9)
virus that are both intended for use with a number of respiratory specimens, we do not anticipate
interference from common endogenous substances.
Specimen Stability and Fresh-frozen Testing:To increase the likelihood of detecting infection, CDC recommends collection of lower respiratory andupper respiratory specimens for testing. If possible, additional specimen types (e.g., stool, urine) should
be collected and should be stored initially until decision is made by CDC whether additional specimen
sources should be tested. Specimens should be collected as soon as possible once a PUI is identified
regardless of symptom onset. Maintain proper infection control when collecting specimens. Store
specimens at 2-8°C and ship overnight to CDC on ice pack. Label each specimen container with thepatient’s ID number (e.g., medical record number), unique specimen ID (e.g., laboratory requisition
number), specimen type (e.g., nasal swabs) and the date the sample was collected. Complete a CDC
Form 50.34 for each specimen submitted.
Clinical Performance:
As of February 22, 2020, CDC tested 2071 respiratory specimens from persons under investigation (PUI)
in the U.S. using the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel. Specimen types include
bronchial fluid/wash, buccal swab, nasal wash/aspirate, nasopharyngeal swab, nasopharyngeal/throatswab, oral swab, sputum, oropharyngeal (throat) swab, swab (unspecified), and throat swab.

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Table 10: Summary of CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel
Data Generated by Testing Human Respiratory Specimens Collected from PUI Subjects in the U.S.
Specimen TypeBronchial
fluid/wash
Buccal swab
Nasal
wash/aspirate
Nasopharyngeal
swab
Nasopharyngeal
swab/throatswab
Oral swab
Pharyngeal(throat) swab
SputumSwab(unspecified)1
Tissue (lung)Total

2019 nCoV
Negative2019 nCoV
Positive

Inconclusive

Invalid

Total

2

0

0

0

2

5

1

0

0

6

6

0

0

0

6

927

23

0

0

950

4

0

0

0

4

476

9

0

0

485

363

10

0

1

374

165

5

0

0

170

71

1

0

0

72

2
2021

0
49

0
0

0
1

2
2071

1Actual swab type information was missing from these upper respiratory tract specimens.

Two thousand twenty-one (2021) respiratory specimens of the 2071 respiratory specimens tested
negative by the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel. Forty-nine (49) of the 2071
respiratory specimens tested positive by the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel. Only
one specimen (oropharyngeal (throat) swab) was invalid. Of the 49 respiratory specimens that tested
positive by the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel, seventeen (17) were confirmed bygenetic sequencing and/or virus culture (positive percent agreement = 17/17, 95% CI: 81.6%-100%)
During the early phase of the testing, a total of 117 respiratory specimens collected from 46 PUI subjects
were also tested with two analytically validated real-time RT-PCR assays that target separate andindependent regions of the nucleocapsid protein gene of the 2019-nCoV, N4 and N5 assays. The
nucleocapsid protein gene targets for the N4 and N5 assays are different and independent from thenucleocapsid protein gene targets for the two RT-PCR assays included in the CDC 2019-nCoV Real-Time
RT-PCR Diagnostic Panel, N1 and N2. Any positive result from the N4 and/or the N5 assay was further
investigated by genetic sequencing.
Performance of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel testing these 117 respiratory
specimens was estimated against a composite comparator. A specimen was considered comparator
negative if both the N4 and the N5 assays were negative. A specimen was considered comparator
positive when the N4 and/or the N5 assay generated a positive result, and the comparator positive
result(s) were further investigated and confirmed to be 2019-nCoV RNA positive by genetic sequencing.

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Table 11: Percent Agreement of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel with theComposite Comparator
CDC 2019-nCoV
Composite Comparator Result
Panel Result
Positive
Negative
1
Positive
13
0Inconclusive00
Negative
0104
1Composite comparator results were available for 13 of 49 CDC 2019-nCoV Panel positive specimens
only.
Positive percent agreement = 13/13 = 100% (95% CI: 77.2% - 100%)
Negative percent agreement = 104/104 = 100% (95% CI: 96.4% - 100%)
Enzyme Master Mix Evaluation:
The limit of detection equivalence between the Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix
and the following enzyme master mixes was evaluated: Quantabio qScript XLT One-Step RT-qPCR
ToughMix, Quantabio UltraPlex 1-Step ToughMix (4X), and Promega GoTaq® Probe 1- Step RT-qPCR
System. Serial dilutions of 2019 novel coronavirus (SARS CoV-2) transcript were tested in triplicate with
the CDC 2019-nCoV Real-time RT-PCR Diagnostic Panel using all four enzyme master mixes. Bothmanufactured versions of oligonucleotide probe, BHQ and ZEN, were used in the comparison. The lowest
detectable concentration of transcript at which all replicates tested positive using the Quantabio qScriptXLT One-Step RT-qPCR ToughMix and Quantabio UltraPlex 1-Step ToughMix (4X) was similar to thatobserved for the Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix. The lowest detectableconcentration of transcript when using the Promega GoTaq® Probe 1- Step RT-qPCR System was one
dilution above that observed for the other candidates when evaluated with the BHQ version of the CDC
assays. The candidate master mixes all performed equivalently or at one dilution below the Thermo
Fisher TaqPath™ 1-Step RT-qPCR Master Mix when evaluated with the ZEN version of the CDC assays.
Table 12: Limit of Detection Comparison for Enzyme Master Mixes – BHQ Probe Summary Results
Thermo Fisher
TaqPath™ 1-Step RTqPCR Master Mix

Quantabio qScript XLT
One-Step RT-qPCR
ToughMix

Quantabio UltraPlex 1Step ToughMix (4X)

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

102 copies/µL

3/3

3/3

3/3

3/3

3/3

3/3

3/3

3/3

101 copies/µL

3/3

3/3

3/3

3/3

3/3

3/3

3/3

3/3

100 copies/µL

3/3

3/3

3/3

3/3

3/3

3/3

3/3

2/3

10-1 copies µL

2/3

0/3

1/3

1/3

1/3

1/3

0/3

0/3

Copy Number

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Promega GoTaq® Probe
1- Step RT-qPCR System

Effective: 03/07/2023

Table 13: Limit of Detection Comparison for Enzyme Master Mixes – ZEN Probe Summary Results
Thermo Fisher
TaqPath™ 1-Step RTqPCR Master Mix

Quantabio qScript XLT
One-Step RT-qPCR
ToughMix

Quantabio UltraPlex 1Step ToughMix (4X)

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

2019nCoV_N1

2019nCoV_N2

102 copies/µL

3/3

3/3

3/3

3/3

3/3

3/3

3/3

3/3

101 copies/µL

3/3

3/3

3/3

3/3

3/3

3/3

3/3

3/3

100 copies/µL

3/3

2/3

3/3

3/3

3/3

2/3

3/3

3/3

10-1 copies µL

1/3

1/3

0/3

0/3

0/3

1/3

1/3

1/3

Copy Number

Promega GoTaq® Probe
1- Step RT-qPCR System

Retrospective positive (18) and negative (17) clinical respiratory specimens were extracted using the
QIAGEN EZ1 Advanced XL instrument and EZ1 DSP Virus Kit and were tested with the CDC 2019-nCoV
Real-time RT-PCR Diagnostic Panel using the Quantabio qScript XLT One-Step RT-qPCR ToughMix,
Quantabio UltraPlex 1-Step ToughMix (4X), and Promega GoTaq® Probe 1- Step RT-qPCR System mastermixes. All three enzyme master mixes performed equivalently, demonstrating 100% positive and 100%
negative agreement with expected results and a 95% confidence interval of 82.4%-100% and 81.6%100%, respectively.
Table 14: Clinical Comparison – Retrospective Study Summary Results
CDC 2019-nCoV
Real-time RTPCR Diagnostic
Panel Result
Positive
Negative

Quantabio qScript XLT
One-Step RT-qPCR
ToughMix
Positive
Negative180017Quantabio UltraPlex 1-Step
ToughMix (4X)

Promega GoTaq® Probe 1Step RT-qPCR System

Positive
18
0

Positive
18
0

Negative
0
17

Negative
017

Roche MagNA Pure 24 and MagNA Pure 96 Extraction Platform Evaluation:
Performance of the 2019-CoV Real-time RT-PCR Diagnostic Panel using the Roche MagNA Pure 24 and
MagNA Pure 96 extraction platforms was compared to performance with an authorized extraction
method. Serial dilutions of quantified inactivated SARS-CoV-2 virus (USA-WA1/2020; 100 RNA copies/µL)
in lysis buffer were added to pooled negative upper respiratory tract specimen matrix. Five samples ofeach dilution were extracted in parallel with the QIAGEN EZ1 Advanced XL (EZ1 DSP Virus Kit Cat# 62724)
and the Roche MagNA Pure 24 (MagNA Pure 24 Total NA Isolation Kit Cat# 07658036001) and Roche
MagNA Pure 96 (MagNA Pure 96 DNA and Viral Nucleic Acid Small Volume Kit Cat# 06543588001)
extraction platforms and evaluated using the 2019-nCoV Real-Time RT-PCR Diagnostic Panel and
ThermoFisher TaqPath™ 1-Step RT-qPCR Master Mix. The observed LoD was defined as the lowest
concentration at which 100% (5 out of 5 total) of all replicates tested positive for both primer/probe sets
(N1 and N2) in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel. The acceptance criteria for
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equivalence were defined as demonstrating an observed LoD either at the same endpoint or within a 3fold dilution. The results showed that both the MagNA Pure 24 and MagNA Pure 96 extraction platforms
performed equivalently or within one 3-fold dilution of the LoD observed when using the QIAGEN EZ1
Advanced XL extraction platform.
Table 15. Limit of Detection Comparison between the QIAGEN EZ1 Advanced XL, Roche MagNA Pure96, and Roche MagNA Pure 24 Extraction Platforms using the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel
Platform

Parameter

2019-nCoV_N1 Assay

2019-nCoV_N2 Assay

Observed
LoD1

RNA copies/µL
101.0
100.5
100.0
101.0
100.5
100.0
# pos./total
5/5
5/5
5/5
5/5
5/5
3/5
QIAGEN EZ1
100.5
Advanced XL
Mean Ct2
34.0
35.0
36.3
33.9
36.6
NA
Std. Deviation
0.2
0.8
0.2
0.4
0.9
NA
RNA copies/µL
101.0
100.5
100.0
101.0
100.5
100.0
# pos./total
5/5
5/5
5/5
5/5
5/5
2/5
Roche MagNA Pure
100.5
96
Mean Ct2
33.3
34.6
36.1
33.2
35.7
NA
Std. Deviation
0.5
0.5
0.3
0.3
0.4
NA
1.0
0.5
0.0
1.0
0.5
RNA copies/µL
10
10
10
10
10
100.0
# pos./total
5/5
3/5
3/5
5/5
5/5
5/5
Roche MagNA Pure
101.0
2
24
Mean Ct
34.4
NA
NA
35.2
36.9
36.2
Std. Deviation
0.6
NA
NA
0.5
1.0
0.8
1
Concentration is presented in RNA copies/µL. The observed LoD is the lowest concentration where both assays showed 100%
positive detection.
2
Mean Ct reported for dilutions that show 100% positivity. Calculations only include positive results.
NA = not applicable

Previously characterized clinical remainder specimens (14 positive and 15 negative) were extracted using
both the Roche MagNA Pure 96 and MagNA Pure 24 extraction platforms and evaluated using the 2019nCoV Real-Time RT-PCR Diagnostic Panel and Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix.
Acceptance criteria for clinical equivalence was defined as demonstrating 100% concurrence withqualitative results shown with the authorized comparator method (QIAGEN EZ1 Advanced XL). Resultsfrom this study showed 100% concurrence with the comparator method for both the Roche MagNA Pure
96 and Roche MagNA Pure 24 extraction platforms when used with the CDC 2019-nCoV Real-Time RTPCR Diagnostic panel.
Table 16. Clinical Comparison Results – Retrospective Study Results
Test
Platform
ResultPositive
Roche MagNA
Negative
Pure 96
Positive
Roche MagNA
Pure 24
Negative
1
CI = 95% confidence interval
Test Platform

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QIAGEN EZ1
Advanced XL Result
Positive Negative
14
0
0
15
14
0
0
15

Positive % Agreement (CI)1

Negative % Agreement (CI)1

100.0 (78.5 – 100.0)

100.0 (79.6 – 100.0)

100.0 (78.5 – 100.0)

100.0 (79.6 – 100.0)

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Promega Maxwell® RSC 48 Extraction Platform Evaluation:
Performance of the 2019-CoV Real-time RT-PCR Diagnostic Panel using the Promega Maxwell® RSC 48
extraction platform was compared to performance with an authorized extraction method. Serial
dilutions of quantified inactivated SARS-CoV-2 virus (USA-WA1/2020; 100 RNA copies/µL) in VTM wereadded to pooled negative upper respiratory tract specimen matrix. Five samples of each dilution wereextracted in parallel with the QIAGEN EZ1® Advanced XL (EZ1 DSP Virus Kit Cat# 62724) and the Promega
Maxwell® RSC 48 (Promega Maxwell® Viral Total Nucleic Acid Purification Kit Cat# AS1330) extraction
platforms and evaluated using the 2019-nCoV Real-Time RT-PCR Diagnostic Panel and ThermoFisher
TaqPath™ 1-Step RT-qPCR Master Mix. The observed LoD was defined as the lowest concentration atwhich 100% (5 out of 5 total) of all replicates tested positive for both primer/probe sets (N1 and N2) in
the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel. The acceptance criteria for equivalence were
defined as demonstrating an observed LoD either at the same endpoint or within a 3-fold dilution. The
results showed that the performance of the Maxwell® RSC 48 extraction platform performed
equivalently or within one 3-fold dilution of the LoD observed when using the QIAGEN EZ1® Advanced XL
extraction platform.
Table 17. Limit of Detection Comparison Between the QIAGEN EZ1® Advanced XL and Promega
Maxwell® RSC 48 Extraction Platforms Using the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Platform

Parameter

2019-nCoV_N1 Assay

2019-nCoV_N2 Assay

Observed
LoD1

RNA copies/µL
100.5
100.0
10-0.5100.5
100.0
10-0.5QIAGEN EZ1®
# pos./total
5/5
5/5
0/5
5/5
5/5
3/5
100.0
Advanced XL
Mean Ct2
32.27
33.80
NA
35.13
36.41
NA
Std. Deviation
0.81
0.40
NA
0.81
0.40
NA
RNA copies/µL
100.5
100.0
10-0.5100.5
100.0
10-0.5
Promega Maxwell®
# pos./total
5/5
5/5
3/5
5/5
5/5
5/5
100.0
2
RSC 48Mean Ct
31.11
32.97
NA
31.89
33.95
35.17
Std. Deviation
0.24
0.34
NA
0.24
0.35
0.65
1
Concentration is presented in RNA copies/µL. The observed LoD is the lowest concentration where both assays showed 100%
positive detection.
2
Mean cycle threshold (Ct) reported for dilutions that show 100% positivity. Calculations only include positive results.
NA = not applicable

Previously characterized clinical remainder specimens (15 positive and 15 negative) were extracted using
the Promega Maxwell® RSC 48 extraction platform alongside the currently authorized QIAGEN EZ1®
Advanced XL extraction platform and evaluated using the 2019-nCoV Real-Time RT-PCR Diagnostic Paneland Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix. Results from the Maxwell® RSC 48 werecompared with the QIAGEN EZ1® Advanced XL extraction performed in parallel showing 100% (15/15)
qualitative concurrence on positive samples and 93.3% (14/15) qualitative concurrence on negative
samples. This evaluation showed that two originally negative (QIAGEN QIAamp® DSP Viral RNA Mini Kit)
specimens (Specimens 16 and 24) yielded an inconclusive result after extraction using the QIAGEN EZ1®
Advanced XL. Repeat of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel resolved one of the two
specimens (Specimen 24, negative result). The second specimen (Specimen 16) remained inconclusive.
Both these specimens yielded a negative result on the Maxwell® RSC 48.

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Table 18. Clinical Comparison Results – Retrospective Study Results
Promega Maxwell® RSC 48

Test Platform

QIAGEN EZ1®
Advanced XL
1ResultPositive

Negative

Inconclusive

Positive

15

0

0

Negative0140Inconclusive

0

1

0

Positive %Agreement (CI)1
100.0(79.6-100.0)

Negative %Agreement (CI)1
93.3(70.2-98.9)

CI = 95% confidence intervalHome Collection Kits
The CDC 2019-Novel Coronavirus (2019-nCov) Real-Time RT-PCR Diagnostic Panel can be used with the
HealthPulse@home and HealthPulse@home Blueprint kits. Audere has granted CDC a right of reference
to the performance data that supports the use of these authorized home collection kits. Refer to the
HealthPulse@home Emergency Use Authorization Summary for specific data [EUA link].

Disposal
Dispose of hazardous or biologically contaminated materials according to the practices of yourinstitution.

References
1. Ballew, H. C., et al. “Basic Laboratory Methods in Virology,” DHHS, Public Health Service 1975
(Revised 1981), Centers for Disease Control and Prevention, Atlanta, Georgia 30333.
2. Clinical Laboratory Standards Institute (CLSI), “Collection, Transport, Preparation and Storage ofSpecimens for Molecular Methods: Proposed Guideline,” MM13-A
3. Lieber, M., et al. "A Continuous Tumor Cell Line from a Human Lung Carcinoma with Properties ofType II Alveolar Epithelial Cells." International Journal of Cancer 1976, 17(1), 62-70.

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Revision History
Revision #Effective Date
1
February 4, 2020
2
March 15, 2020

3

March 30, 2020

4

June 12, 2020

Summary of Revisions
Original Instructions for Use• Intended use update• Removal of N3 primer and probe set from Diagnostic Panel• Performance data update• Addition of alternative nucleic acid extraction platforms• Addition of acceptable alternatives to HSC and addition of
QIAGEN RUO extraction reagents• Positive results no longer presumptive. No confirmation of
positive results required• Addition of alternative enzyme master mix options
•
•
•
•
•

5

July 13, 2020

6

December 1, 2020

7

July 21, 2021

8

March, 7, 2023

CDC-006-00019, Revision: 08

•
•
•
•
•
•
•
•
•

Addition of MagNA Pure 24 extraction method
Addition of performance data for the MagNA Pure 96
extraction method with SARS-CoV-2
Addition of heat treatment alternative to specimen
extraction
Addition of Roche and QIAGEN external lysis buffer
alternatives
Acknowledgment of FDA policy permitting end users toqualify alternative components without seeking an EUA or
EUA amendment
Addition of Promega Maxwell® RSC 48 extraction method
Update to in silico inclusivity analyses
Addition of specimen pooling instructions and monitoringprocedure (Appendices B and C)
Addition of Promega Maxwell CSC 48 extraction instrument
Addition of data for CDC testing of FDA reference panel
Addition of Audere home collection kit
Update of in silico analysis of primer/probe sequences
Update to Warnings and Precautions language
Update to add additional limitation

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Contact Information, Ordering, and Product Support
For technical and product support, contact the CDC Division of Viral Diseases directly.
Send email to: respvirus@cdc.gov
Note: If your laboratory is using reagents sourced from someone other than the CDC International
Reagent Resource, please refer to the manufacturer’s instructions provided with the commercial
materials. Technical and product support related to the HealthPulse@home and HealthPulse@home
Blueprint kits should be directed to the kit manufacturer, Audere at healthpulsenow.org.

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Appendix A: Heat Treatment Alternative to Extraction
UltraPlex 1-Step ToughMix (4X)This procedure is only for use by public health laboratories.
Purpose:In response to a global shortage of nucleic acid extraction reagents causing significant delays in testing,
the CDC has investigated the use of a heat treatment method requiring minimal reagents as a specimen
processing alternative to nucleic acid extraction for use with the 2019-nCoV Real-Time RT-PCR Diagnostic
Panel.
Where possible, laboratories should use qualified RNA or total nucleic acid extraction methods forprocessing of specimens for subsequent testing by the CDC 2019-nCoV Real-Time RT-PCR Diagnostic
Panel. Extraction removes inhibitory substances from specimens that could negatively impact PCR
performance.
This procedure for use of heat treatment for specimen processing is only recommended when ashortage of qualified extraction reagents is a limiting factor in a laboratory’s ability to meet urgent SARSCoV-2 testing demand.
Precautions/Warnings/Limitations:••••

CDC has evaluated this heat treatment process and has determined that this process is effective
for inactivation of SARS-CoV-2 in patient specimens.
Performance was evaluated with only upper respiratory specimens. Heat treatment of lower
respiratory specimens for subsequent testing by the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel has not been evaluated.
This procedure for heat treatment of specimens is only for use with the Quantabio UltraPlex 1Step ToughMix (4X).
Heat treatment should only be conducted when a lab is ready to test the specimens by PCR.
Testing of heat-treated specimens must be conducted the same day.

Acceptable Specimens:•

Upper respiratory specimens
Note: Do not use heat treatment to process specimens that appear bloody or that containparticulate matter. Such specimens should be extracted using a qualified RNA or TNA extraction
method prior to testing.

Materials Required (not provided):••••

70% ethanol
10% bleach, freshly prepared
96-well PCR reaction plates (Applied Biosystems catalog # 4346906, 4366932, 4346907, orequivalent)
Optical strip caps (Applied Biosystems 4323032, or equivalent)

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•
•
•
•
•
•
•
•
•

1.5 mL Sarstedt tubes or equivalent
Aerosol resistant micropipette tips
Micropipettes
96-well cold block
Cold blocks for 1.5 mL - 2.0 mL tubes
Vortex mixer
96-well plate centrifuge or equivalent
Thermal cycler or equivalentClass II Biological Safety Cabinet (BSC)

Procedure:
Sample Preparation
1) Decontaminate BSC with 10% bleach followed by 70% ethanol.
2) If samples are frozen, thaw on ice or at 4°C. Wipe the outside of the sample tube with 70%
ethanol. Place thawed sample on cold rack or ice in BSC.
3) Pulse vortex each sample and briefly spin down in a centrifuge to collect the liquid at thebottom of the tube.
Heat Treatment
1) Place a thermal cycler in the BSC, turn on, and program for 95°C for 1 min followed by 4°C
hold.
2) Place a 96-well PCR plate onto a cold rack or ice in the BSC.
3) Transfer 100 µL of each sample to the 96-well PCR plate and securely cap each well using
optical strip caps.
NOTE: Ensure that an HSC extraction control is included in each batch run as required under
CLIA.
4) Place this 96-well PCR plate on the pre-heated thermal cycler and start run. Leave plate on
thermal cycler at 4°C, or place on ice or a cold block.
5) Remove plate and centrifuge for 1 minute at 500 x g to pellet cellular debris.
6) Place plate on a cold rack or ice and proceed to testing the supernatant by rRT-PCR.
7) Testing of heat-treated specimens must be conducted the same day heat treatment isperformed. For long term storage, keep the original specimen at ≤-70°C.
Special Testing Considerations for Heat Treated Specimens:
Enzyme Master Mix• Testing of specimens that have been processed with heat treatment should be conductedwith the Quantabio UltraPlex 1-Step ToughMix (4X), which demonstrated the best
performance with heat treated specimens. PCR testing of heat-treated specimens should
follow the instructions in the main body of this Instructions for Use document.

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Resolution of Inconclusive and Invalid Results• Retesting of heat-treated specimens that generated an inconclusive or invalid result must
include extraction of the original specimen with a qualified RNA or total nucleic acid (TNA)extraction method, if available. Do not re-test the heat-treated specimen material to resolveinconclusive or invalid test results.
Verification:
CDC recommends performance of verification studies for the heat treatment method prior to diagnostic
use that includes side-by-side preparation of a panel of positive and negative clinical specimens using aqualified extraction method and this heat treatment method with subsequent testing by the CDC 2019nCoV Real-Time RT-PCR Diagnostic Panel.
Performance Characteristics:
Quantabio UltraPlex 1-Step ToughMix (4X)
Limit of Detection Comparison
Serial dilutions of inactivated SARS-CoV-2 [SARS-CoV-2 USA-WA1/2020] were prepared in simulated
specimen material (human A549 cells suspended in viral transport medium). Each concentration was
prepared side-by-side five times by both EZ1 extraction and by heat treatment. Each extracted or heattreated sample was subsequently tested by the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel using
the Quantabio UltraPlex 1-Step ToughMix (4X) on the Applied Biosystems 7500 Fast Dx instrument.
Observed detection was similar between the two specimen preparation methods.
Table A1: UltraPlex Limit of Detection Comparison between QIAGEN EZ1 Advanced XL extraction and heat
treatment (95°C for 1 min) method – Summary Results

QIAGEN
EZ1
Advanced
XL

Platform

Heat
Treatment
95°C
for 1 minQuantabio UltraPlex 1Step ToughMix (4X)
5 µL Template Addition

EnzymeParameter

2019-nCoV_N1 Assay

Observed
LoD1

2019-nCoV_N2 Assay

RNA copies/µL

101.0

100.5

100.0

10-0.5 10-1.0

101.0# pos./total
Mean Ct2

5/5
5/5
34.11 34.59

4/5
NA

4/5
NA

3/5
NA

5/5
5/5
5/5
32.97 33.76 34.70

2/5
NA

2/5
NA

Std. Deviation

0.75

0.99

NA

NA

NA

0.33

0.72

0.98

NA

NA

RNA copies/µL

101.0

100.5

100.0

10-0.510-1.0

101.0

100.5

100.0

10-0.510-1.0# pos./total

5/5

5/5

4/5

5/5

1/5

5/5

5/5

4/5

2/5

1/5

Mean Ct2
Std. Deviation

33.41 34.32
0.62 0.40

NA
NA

36.73
0.82

NA
NA

33.45 35.25
0.40 0.80

NA
NA

NA
NA

NA
NA

1Concentration is presented in RNA copies/µL.

positive detection.

100.5

100.0

10-0.5 10-1.0
100.5

100.5

The observed LoD is the lowest concentration where both assays showed 100%

2Mean Ct reported for dilutions that show 100% positivity. Calculations only include positive results.

NA = not applicable

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Clinical Comparison
A panel of 39 upper respiratory specimens were tested side-by-side using extraction with the Qiagen EZ1
extraction instrument and heat treatment. Extracted and heat-treated specimens were subsequently
tested with the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel using the Quantabio UltraPlex 1-Step
ToughMix (4X). Qualitative results were compared to demonstrate agreement.
Table A2: Clinical Comparison Results Summary – Heat Treatment versus QIAGEN EZ1 Advanced XL
Test Result
Positive
QIAGEN EZ1
Inconclusive
Advanced XL
Negative
Total
1 CI = 95% confidence interval

Heat Treatment
Positive
180018Inconclusive
10
0
1

Negative
0
0
20
20

Total
19
0
20
39

Positive %Agreement (CI)1

Negative %Agreement (CI)1

94.7 (75.4-99.1)

100 (83.9-100)

Questions and Comments:
If you have questions or comments about this procedure, please send by email to: respvirus@cdc.gov

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Appendix B: Pooled Specimen Preparation and Processing
Purpose:In response to strong demand for higher throughput testing approaches as well as a global shortage of
nucleic acid extraction reagents causing significant delays in testing, the CDC has evaluated specimen
pooling and determined that pooling of up to 4 specimens is suitable for use with the 2019-nCoV RealTime RT-PCR Diagnostic Panel.
This pooled specimen preparation and processing procedure cannot be used with specimens selfcollected at home or elsewhere using either the HealthPulse@home or the HealthPulse@home
Blueprint kit.
Specimen pooling may cause a slight reduction in test sensitivity and therefore may be most appropriate
for screening or diagnostic testing when laboratory staff, equipment or reagents are insufficient toaccommodate testing demand. Specimen pooling only presents a throughput advantage when the
disease prevalence is low. Therefore, laboratories should monitor specimen positivity rates over time todetermine if pooling of specimens continues to provide a test throughput advantage over individual
specimen testing. Please see Appendix C for more detail on implementation considerations.
While this procedure describes the process to prepare, process and test a pool size of up to 4 specimens,
specimen pool sizes from 2-4 are authorized for use with the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel. When using a pool size of less than 4 specimens, please use the following instructions
as a model. Pooled specimen input volume and the pooled specimen to lysis buffer volume ratios mustremain as prescribed below (not a lower proportion of lysis buffer) to ensure inactivation of SARS-CoV-2
in patient specimens. An N-pool specimen approach should include equal volumes of each of the N
specimens pooled together to create the total pooled specimen input volume required under the below
pooled specimen extraction instructions.
Precautions/Warnings/Limitations:
•
•
•

•

Pooling cannot be used for specimens self- collected using either the HealthPulse@home or the
HealthPulse@home Blueprint kit.
Pooling of specimens has the potential to decrease sensitivity. The specimens in a pooled
procedure are diluted, which could result in a low concentration of viral genetic material below
the limit of detection of a given test.
When pooling specimens, the laboratory cannot ensure the diagnostic integrity of an individual
specimen because it is combined with other specimens before testing. Specimen integrity can beaffected by the quality of swab specimen collection, which could result in some swabs having
limited amounts of viral genetic material for detection. Inadequate individual specimens,
including those with limited amounts of viral genetic material, might not be eliminated from thepooled specimen before testing and may be reported as negative through this process.
Performance of this specimen pooling process was evaluated with nasopharyngeal (NP) swabs.
Pooling of other types of specimens for subsequent testing with the CDC 2019-nCoV Real-Time
RT-PCR Diagnostic Panel has not been evaluated, however evidence suggests the Ct value

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•
•
•
•
•

distribution for other upper respiratory swab specimens is similar to that observed for NP swab
specimens.
Pooling should only be implemented in laboratories with a minimum of 10 days experience using
the CDC 2019-nCoV Real-time RT-PCR Diagnostic Panel for diagnostic testing in the population of
specimens being considered for pooling.
This procedure for pooling of specimens has only been evaluated for use with the QIAGEN EZ1
Advanced XL, Roche MagNA Pure 96, and the Promega Maxwell RSC 48 instruments. Poolingusing other extraction methods is not authorized under this EUA.
Sufficient volume of specimens must be available to allow subsequent extraction and testing ofindividual specimens should the specimen pool test positive, inconclusive or invalid.
Appropriate interpretation, reporting and next steps may be different for surveillance testing andare not limited by the diagnostic interpretation instructions below.
Laboratories are encouraged to revisit the question of whether pooling continues to make sensewhen testing demand no longer exceeds laboratory capacity and/or when testing reagents are nolonger in short supply.

Acceptable Specimen Types:•

Upper respiratory swabs (e.g., nasopharyngeal (NP) swabs, oropharyngeal (OP) swabs, NP/OP
swabs combined, nasal swabs) collected by or under the supervision of a healthcare provider
Note: Specimens must have adequate volume to support pooled specimen testing and any
subsequent individual re-testing.

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Overview of Pooled Specimen Testing:This flow chart provides an overview of the steps involved in pooled specimen testing and location of the
instructions for each step within the document. For those steps that are identical for both individual
specimen testing and for pooled specimen testing, instructions are found in the main body of the
instructions for use. For steps in which pooled specimen testing differs from individual specimen testing,
instructions specific to pooled specimen testing are presented in this Appendix.

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Procedure:Specimen Pool Preparation
1) Decontaminate BSC with 10% bleach followed by 70% ethanol.
2) If specimens are frozen, thaw on ice or at 4°C. Wipe the outside of the specimen tube with 70%
ethanol. Place thawed specimen on cold rack or ice in BSC.
3) Pulse vortex each specimen and briefly spin down in a centrifuge to collect the liquid at thebottom of the tube.
4) For each specimen pool, add an equal volume of each specimen (depending on total volume
needed for extraction platform) to be included into the pool, into a sterile, nuclease-free tube.
5) Pulse vortex each specimen pool and briefly centrifuge to collect the liquid at the bottom of the
tube.
6) Proceed to extraction using the modified extraction parameters below.
Extraction Instructions
NOTE: These pooled specimen extraction instructions have been modified to optimize
recovery of nucleic acid when processing pooled specimens. When testing individual
specimens, please follow the individual specimen extraction instructions (p. 14-16).
QIAGEN EZ1 Advanced XL• Kit: Qiagen EZ1 DSP Virus Kit and Buffer AVL (supplied separately) for offboard lysis• Card: EZ1 Advanced XL DSP Virus Card
• Instructions: Add 200 μL of pooled specimen to 200 μL of pre-aliquoted Buffer AVL (total
input sample volume is 400 μL). Proceed with the extraction on the EZ1 Advanced XL. Elution
volume is 60 μL.
Roche MagNA Pure 96• Kit: Roche MagNA Pure 96 DNA and Viral NA Small Volume Kit• Protocol: Viral NA Plasma Ext LysExt Lys SV 4.0 Protocol or Viral NA Plasma Ext Lys SV Protocol• Instructions: Add 225 μL of pooled specimen to 225 μL of pre-aliquoted MagNA Pure 96
External Lysis Buffer (supplied separately) for total input sample volume of 450 μL. Proceedwith the extraction on the MagNA Pure 96. (Internal Control = none).
• Elution volume is 50 μL.
Promega Maxwell RSC 48 or Maxwell CSC 48• Kit: Promega Maxwell® Viral Total Nucleic Acid Purification Kit• Protocol: Viral Total Nucleic Acid
• Instructions: Add 240 μL of pooled specimen to 660 μL of pre-aliquoted External Lysis Buffer(600 μL Lysis Buffer plus 60 μL Proteinase K; supplied within the kit) (total input volume is 900μL). Proceed with the extraction on the Maxwell® RSC 48 or Maxwell CSC 48.
• Elution volume is 50 μL.

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Testing, Interpretation, and Reporting Instructions
1) After extraction, continue PCR testing using the steps outlined in the following sections of theInstructions for Use: Assay Set Up (p. 16-20), Running a Test (p. 29-32) and Data Analysis(p. 33-34).
2) Interpret controls and PCR test results as outlined in the following sections of the Instructions
for Use: Interpretation of Results and Reporting (p. 35-37), 2019-nCoV rRT-PCR Diagnostic
Panel Results Interpretation Guide (p. 38). All extraction and PCR run controls must perform
as expected.
A. If a specimen pool generates negative results, report “Not Detected” for each specimen
included in the pool. The test result report should include a statement that the specimen
was tested in a pooled format. No further testing is required.
B. If a specimen pool generates a positive result, do not report the result. Test eachspecimen included in the pool individually, according to the Instructions for Use. Report
the individual result for each specimen.
C. If a specimen pool generates an inconclusive result, do not report the result. Test eachspecimen included in the pool individually, according to the Instructions for Use. Report
the individual result for each specimen.
D. If a specimen pool generates an invalid result, do not report the result. Test eachspecimen within the pool individually, according to the Instructions for Use. Report theindividual result for each specimen.
Verification:
Verification should be conducted by all laboratories prior to implementing pooling for diagnostic testing.
In addition to method verification studies, verification must include a validation of the accuracy andfunction of the laboratory’s process of specimen and pool labeling, tracking and reporting through thelaboratory information management system(s).
Performance Characteristics:
Pooling Validation
A panel of 20 positive, previously characterized nasopharyngeal (NP) specimens, with 25% (5)representing the weak positive range (Ct 36.00-39.99), were used for evaluation of specimen poolingwith the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel; 80 negative NP specimens were also
included. These specimens were used to create twenty positive 4-specimen pools, by combining one
positive specimen with three negative specimens. Twenty negative pools were also created by
combining equal volumes of four negative specimens. These pools were tested and qualitative resultsfrom pooled-testing were compared with the results from individual-testing to determine the positive
and negative percent agreement of the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel for testing
using 4-specimen pools. The performance of 4-specimen pooling was evaluated using the QIAGEN EZ1®
Advanced XL, the Promega Maxwell RSC 48, or the Roche MagNA Pure 96 platforms. Modifications weremade to the input and output volume for each extraction platform, for pooling patient specimen, tocompensate for the dilution effect due to pooling.

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QIAGEN EZ1® Advanced XL
Individual positive panel members and positive and negative pools were extracted using the QIAGEN
EZ1® Advanced XL platform for subsequent testing on the Applied Biosystems 7500 Fast
Dx with the Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix, CG. Results obtained for twenty, 4specimen pools were compared with the results from individual testing. Pools containing a positive
specimen were considered to be in agreement with results from individual testing if the pool
generated positive or inconclusive results (i.e., at least one of the SARS-CoV-2-specific primer and probe
sets generated a Ct value less than 40). Negative specimen pools were considered to be in agreementwith expected results if the pooled specimen result was negative (i.e., neither SARS-CoV-2-specificprimer and probe set generated positive results).
A comparison of qualitative test results from individual testing of panel members with pooled specimen
testing is summarized below in Table B1, along with testing agreement with results from individual
testing for positive and negative specimen pools. Of the five positive specimen panel members in the
weak positive category (determined by original diagnostic results): one returned an inconclusive result
when tested individually, but returned a positive result when tested in a 4-specimen pool; one returneda negative result when tested individually, but returned a positive result when tested in a 4-specimen
pool; and three were in agreement when tested individually and in 4-specimen pools.
Table B1 -Summary of Pooled vs. Individual-testing - QIAGEN EZ1® Advanced XL
Individual testing result
Positive Pools
Test Platform
Percent
Agreement (CI)1
ResultPositive Inconclusive† Negative
Positive
4-specimen
Inconclusive‡
Pooling Result
Negative

17

1

1

1

0

0

0

0

20

100%
(82.4-100)

Negative Pools
Percent Agreement (CI)1
95.2%(77.3-99.2)

CI = 95% confidence interval†Inconclusive individually-tested results are not included in the final performance calculations.
‡Inconclusive pooled results are considered in agreement with the positive individually-tested results for the final performance calculations.
1

Promega Maxwell RSC 48
Individual positive panel members and positive and negative pools were extracted using the Promega
Maxwell RSC 48 platform for subsequent testing on the Applied Biosystems 7500 Fast
Dx with the Thermo Fisher TaqPath™ 1-Step RT-qPCR Master Mix, CG. Results obtained for twenty, 4specimen pools were compared with the results from individual testing. Pools containing a positive
specimen were considered to be in agreement with the results from individual testing if the pooled
specimen generated positive or inconclusive results (i.e., at least one of the SARS-CoV-2-specific primer
and probe sets generated a Ct value less than 40). Twenty negative specimen pools (made from 4
known negative specimens) were considered to be in agreement with expected results if the pooled
specimen result was negative (i.e., neither SARS-CoV-2-specific primer and probe set generated positive
results).
A comparison of qualitative test results from individual testing of panel members with pooled specimen
testing is summarized below in Table B2, along with testing agreement with results from individual62
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testing for positive and negative specimen pools. Of the five positive specimen panel members in the
weak positive category (determined by original diagnostic results): one returned a positive result whentested individually, but returned a negative result when tested in a 4-specimen pool; and four were inagreement when tested individually and in 4-specimen pools. Additionally, one moderate to highpositive panel member returned an inconclusive result when tested individually but returned a positive
result when tested in a 4-specimen pool.
Table B2 -Summary of Pooled vs. Individual-testing Results – Promega Maxwell RSC 48

Test Platform

Individual-testing result
ResultPositive Inconclusive† Negative

Positive
4-specimen Inconclusive‡
Pooling Result Negative

16

1

0

2

0

0

1

0

20

Positive Pools
Percent Agreement (CI)1

Negative Pools
Percent Agreement (CI)1

94.7%(75.4-99.1)

100%(83.9-100)

CI = 95% confidence interval

1†Inconclusive individually-tested results are not included in the final performance calculations.

‡Inconclusive pooled results are considered in agreement with the positive individually-tested results for the final performance

calculations.

MagNA Pure 96
Individual positive panel members and positive and negative pools were extracted on the MagNA Pure
96 platform for subsequent testing on the Applied Biosystems 7500 Fast Dx using the Thermo
Fisher TaqPath™ 1-Step RT-qPCR Master Mix, CG. Results obtained for twenty, 4-specimen pools werecompared with the results from individual testing. Pools containing a positive specimen wereconsidered to be in agreement with the results from individual testing if the pooled specimen generated
positive or inconclusive results (i.e., at least one of the SARS-CoV-2-specific primer and probe sets
generated a Ct value less than 40). Twenty negative specimen pools (made from 4 known negative
specimens) were considered to be in agreement with expected results if the pooled specimen result was
negative (i.e., neither SARS-CoV-2-specific primer and probe set generated positive results).
A comparison of qualitative test results from individual testing of panel members with pooled specimen
testing is summarized below in Table B3, along with testing agreement with results from individual
testing for positive and negative specimen pools. Of the five positive specimen panel members in the
weak positive category (determined by original diagnostic results): one returned an inconclusive result
when tested individually, but returned a positive result when tested in a 4-specimen pool; and four werein agreement when tested individually and in 4-specimen pools.

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Table B3 -Summary of Pooled vs. Individual-testing Results – Roche MagNA Pure 96
Individual-testing result
Positive Pools
Test Platform
Percent
Agreement (CI)1†
ResultPositive Inconclusive Negative
Positive
4-specimen
Inconclusive‡
Pooling Result
Negative

18

1

0

1

0

0

0

0

20

Negative Pools
Percent Agreement (CI)1

100%(83.2-100)

100%(83.9-100)

CI = 95% confidence interval

1†Inconclusive individually-tested results are not included in the final performance calculations.

‡Inconclusive pooled results are considered in agreement with the positive individually-tested results for the final performance

calculations.

In Silico Sensitivity
An in silico analysis was conducted to evaluate the effect of 4-sample pooling on the clinical sensitivity of
the CDC 2019-Novel Coronavirus Real-Time RT-PCR Diagnostic Panel using the QIAGEN EZ1® Advanced
XL extraction platform. This analysis was conducted by performing a Passing-Bablok regression using the“Pooling Validation” data to calculate the Ct shifts resulting from the dilution effect of 4-specimen pools(1 positive specimen combined with 3 negative specimens) for each target. In the regression analysis,
the X-axis displayed individual Ct values for positive specimens and the Y-axis displayed Ct values for thecorresponding pools with one positive specimen and 3 negative specimens. The regression analysis wasused to calculate an interval of Ct values [X*, 40] where individual specimens with Ct values within thisinterval would have negative results in 4-specimen pools (1 positive and 3 negative).
The results from individually-tested NP swab specimens processed at the CDC using the QIAGEN EZ1®
Advanced XL extraction platform (n= 381), were analyzed to determine an in silico PPA for 4-specimen
pooling. The number of individual specimens with Ct values ranging from [X*, 40] was determined. The
X* value for the N1 target was 37.2 and the X* value for the N2 target was 37.5. The results, summarized
Table B4, show that 97.1% (370/381 95% CI 94.9-98.3%) of the specimens would not have negative
results when combined into 4-specimen pools.
Table B4. in silico Sensitivity for Pooled Specimens Extracted using the QIAGEN EZ1® Advanced XLn

N1 Interval[X*, 40]

Number ofsamples with
N1 Ct values
in the interval

381[37.2, 40]

16

N2 Interval[X*, 40]

Number ofsamples with
N2 Ct values in
the interval

Number of
Samples withboth target Ct
values inintervals

Neg

Inc

Pos

% Positive
Percent
Agreement*[37.5, 40]

63

11

11

57

313

97.1

*

Since any pool that is not negative is re-tested as individual samples, the Positive Percent Agreement includes all pools that were notnegative.

The specimens submitted to CDC for testing included many that had generated inconclusive results in
the hands of public health laboratories and were forwarded to CDC for further testing. Thus, specimens
received by CDC for testing overrepresent the weak positive category. The CDC also obtained the resultsof individually-tested NP swab specimens from three additional geographic locations: state lab 1 (n=2217), state lab 2 (n= 6559), and state lab 3 (n= 2315). These data sets were analyzed with regard to the
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percent of weak positives (Ct 36.00-39.99) collected at each site. The results, summarized in Table B5,
suggest that the percent of weak positive NP specimens received at State Health Department
laboratories is smaller than that obtained at CDC. Therefore, the in silico analysis, summarized in Table
B4, presents a PPA in a scenario where 4-specimen pooling is applied to testing populations with agreater than average number of weak positive specimens.
Table B5. Percent of Weak Positive Specimens at CDC Compared with Three Geographic Locations
Ct < 36.0

Ct 36.0 - 39.99

n

N1

N2

N1

N2

CDC

381

314 (82.4 %)

254 (66.7 %)

67 (17.6 %)

127 (33.3 %)State 1

2217

2121 (95.7 %)2021 (91.2 %)

150 (6.8 %)

250 (11.3%)

State 26559

6343 (96.7 %)5883 (89.7 %)

216 (3.3%)

676 (10.3%)

State 32315

2073 (89.5 %)1884 (81.4 %)

242 (10.5 %)431 (18.6 %)References:
1. Abdalhamid, B., et al. “Assessment of Specimen Pooling to Conserve SARS CoV-2 Testing
Resources.” Am J Clin Path 2020, 153, 715-718.
Questions and Comments:
If you have questions or comments about this procedure, please send by email to: respvirus@cdc.gov

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Appendix C: Implementation and Monitoring of Pooled Specimen Testing
Purpose:
These instructions are intended to assist laboratories and jurisdictions to implement and monitor theefficiency and effectiveness of pooled specimen testing over time. A laboratory should have a minimumof 10 days experience performing the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel for diagnostic
testing within the intended population prior to implementation of a pooling strategy.
Laboratories should weigh the potential risks associated with pooling as described in Appendix B against
the benefits of pooling for their laboratory and patient population. When pooling has been implemented
to address overwhelming testing demands and/or scarcity of testing materials, laboratories should
consider a return to individual specimen testing when laboratory capacity and resources permit.
Pooling of specimens cannot be performed with specimens self-collected using either the
HealthPulse@home or the HealthPulse@home Blueprint kit.
Prior to Implementation of a Specimen Pooling Strategy:•

Determine the individual specimen positivity rate (PIndividual) for your testing population.
Before implementation of specimen pooling, evaluate test data from the testing population forthe previous 7-10 days to estimate the positivity rate (PIndividual) which is the number of positive
results divided by the total number of specimens tested during these 7-10 days. PIndividual will be
used to determine if pooling should be considered and will be used to monitor the performanceof your pooling strategy.
Note: To calculate the efficiency of 4-sample pooling, using PIndividual, apply the formula F=1/(1+1/n-(1- PIndividual)n), where F is the efficiency and n is the pool size. For example, when PIndividual is1%, the efficiency, F, is 3.46 for n = 4. This means that 1,000 tests can cover testing of 3,460
patients on average, which translates to a reduction in test volume of 71%. Using this formula, apool size of n = 3 produces a greater efficiency than n = 4 at PIndividual ≥ 13%.

•

Identify your positivity thresholds.
Table C1: Association Between Individual Specimen Positivity Rates and Reduction in Test Volume with
PoolingSpecimen
Positivity
Rate (P)No. tests with
4 specimen
pooling for
300 patients

Reduction in
testing
volume with
4 specimen
pooling

No. tests with
3 specimen
pooling for
300 patients

Reduction in
testing
volume with
3 specimen
pooling

No. tests with
2 specimen
pooling for
300 patients

Reduction in
testing
volume with
2 specimen
pooling

1%
2%
3%
4%
5%
6%
8%
10%

87
98
109
120
131
141
160
178

71%
67%
64%
60%
56%
53%
47%
41%

109
118
126
135
143
151
166
181

64%
61%
58%
55%
52%
50%
45%
40%

156
162
168
174
179
185
191
207

48%
46%
44%
42%
40%
38%
36%
31%

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Specimen
Positivity
Rate (P)No. tests with
4 specimen
pooling for
300 patients

Reduction in
testing
volume with
4 specimen
pooling

No. tests with
3 specimen
pooling for
300 patients

Reduction in
testing
volume with
3 specimen
pooling

No. tests with
2 specimen
pooling for
300 patients

Reduction in
testing
volume with
2 specimen
pooling

11%
12%
13%
14%
15%
16%
17%
18%
19%
20%

187
195
203
211
218
226
233
239
246
252

38%
35%
32%
30%
27%
25%
22%
20%
18%
16%

189
196
202
209
216
222
228
235
241
246

37%
35%
33%
30%
28%
26%
24%
22%
20%
18%

212
218
223
228
233
238
243
248
253
258

29%
27%
26%
24%
22%
21%
19%
17%
16%
14%

Note: Table C1 does not account for running the positive, negative, and extraction controls with each plate.
Note: Unshaded cells indicate the optimal reduction in testing volume compared to other the pool sizes.

Table C1 presents the estimated increase in efficiency achieved through pooling at different ratesof specimen positivity. Laboratories are encouraged to use this table and factor in additional
testing necessary to monitor the effectiveness of pooling to determine the specimen positivity
thresholds (PThreshold) at which (1) four-specimen pooling is appropriate, (2) smaller specimen
pools are appropriate, and (3) the threshold at which specimen pooling offers no testing
capacity/reagent consumption advantages. These thresholds should be documented for yourlaboratory’s pooling monitoring process.
Example: a laboratory might consider a 30% reduction in test volume as the minimum required
reduction in test volume to be considered beneficial (this is subjective). Thus, the laboratory
would set a positivity threshold (PThreshold) at 12% 3 for 4 specimen pooling, 14% for 3 specimen
pooling, and 10% or less for 2 specimen pooling.
NOTE: It is recommended that PThreshold should not be greater than 20% for any pool size.
Moreover, laboratories may choose a threshold positivity rate lower than 20% for when to
consider a pooling strategy.
•

Define and document a specimen pooling strategy/process for your laboratory.

•

Verify laboratory performance of pooled specimen testing (CLIA requirement).

•

Establish and implement a pooling monitoring process appropriate to assure the quality of results
generated through the laboratory’s pooling strategy. The pooling monitoring process should
document the laboratory’s PThreshold for each pool size, PIndividual and the laboratory’s process toaddress the monitoring requirements below.

3

At PIndividual ≥ 13%, n=3 offers a greater efficiency than n=4, therefore it is recommended that PThreshold should not be greaterthan 12% for n=4.

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•Please note that information from the monitoring process described in this Appendix may not be theonly information useful to monitor for pooling performance issues or to aid in decision-making forpooling strategy. Thus, laboratories are encouraged to monitor other sources of COVID-19
prevalence or test positivity rate information for their target population, comparing those trends totheir pooling testing trends, as part of the laboratory’s pooling monitoring process in addition to theapproach below.

Monitoring of the Pooling Strategy
At regular intervals (weekly at a minimum, continual analysis with a moving 7-10 day average
preferred), laboratories must monitor the positivity rate of specimen pools as described below.

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Figure C1: Overview of Pooling Monitoring Process to be Conducted at Defined Intervals
Assess current testing demand and reagent availability.
Will laboratory continue pooled specimen testing?

No

Yes
Calculate PPool for the interval.

Is PPool lower than PThreshold for the numberof specimens being pooled?

Stop pooling.

No

Reduce pool sizeor stop pooling.

No

Stop pooling.

Yes
No

Is PPool less than 0.85 x PIndividual?
Yes
Conduct re-assessment of pooling.
Did re-assessment indicate pooling is still
appropriate?
Yes
Set new PIndividual based on re-assessment
data. Document for future monitoring
activities.

Continue pooled
specimen testing.

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Data Monitoring Steps to Be Conducted at Each Interval
1. Evaluate need for pooled specimen testing going forward.
Evaluate test demand and testing resource availability. When testing capacity and resource
availability are sufficient to meet testing demand without the use of specimen pooling, consider
whether the risks of reduced test sensitivity with pooling and extra effort of pooling outweigh thebenefits of pooled specimen testing.
2. Calculate Pooled Percent Positive (PPool) for the interval.
PPool =

Specimens tested under pooling strategy generating positive results
Total specimens tested under pooling strategy

3. Check Interval PPool Against Positivity Threshold (PThreshold) for the pool size in use.
Compare the PPool for the current interval to the specimen positivity threshold established duringpooling implementation.
•

If PPool is greater than PThreshold for the pool size in use, consider a reduction in pool size or
discontinuation of specimen pooling until specimen positivity rates decrease in yourtesting population.

•

If PPool is less than PThreshold, proceed to step 4.

4. Check Interval PPool Against Re-assessment Criteria
Compare PPool to PIndividual.
• If PPool is equal to or greater than 85% of PIndividual. (PPools ≥ 0.85 × PIndividual), thenmonitoring checks are complete for this interval. Proceed with the current poolingstrategy.
• If PPool is less than 85% of PIndividual. (PPools < 0.85 × PIndividual), re-assessment is necessary to
determine if pooled-specimen testing process is still acceptable. Please proceed to theRe-assessment of Pooling section below.
5. Update PIndividual if Indicated by Re-assessment.
If re-assessment is conducted and indicates n-specimen pooling is still acceptable, re-establish
PIndividual in your laboratory using the data generated during re-assessment.
If 10 positive specimens were tested during re-assessment:
PIndividual =10 positive specimens used for re-assessment
The total number of specimens tested to reach 10 positive specimens

X

10
11

X

20
21

If 20 positive specimens were tested during re-assessment:
20 positive specimens used for re-assessment

PIndividual = The total number of specimens tested to reach 20 positive specimens
This updated new positivity rate should be used as PIndividual in the future monitoring.

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Explanation: This calculation attempts to estimate the individual testing positivity rate in the
population from which the 10 (or 20) individual positive samples were collected for reassessment. Since the total number of samples (N*) that needed to be tested to obtain 10 (or20) consecutive positive samples for re-assessment is stopped at the 10th (or 20th) positive
sample, then the positivity rate of 10/N* (or 20/N*) may be overestimated. PIndividual is correctedin the above calculations by including an appropriate multiplier: estimated positivity rate for 10
positive sample re-assessment data is (10/N*)×(10/11) and for 20 samples is (20/N*)×(20/21).
Re-assessment of Pooling:Re-assessment of pooling should be conducted as indicated by monitoring activities. Two possible
approaches are presented below.
Option 1
Incoming patient specimens should be tested individually until the collection contains 10 positive
specimens.
•

Using these specimens, 10 n-specimen pools should be created and tested, each with 1 positiveand (n-1) negative specimens.

•

Calculate the PPA between individual testing results and pooled testing results.
Note: For the calculation of the PPA, all pools that generate positive or inconclusive result areconsidered in agreement with the individually-tested positive result.

Option 2
Incoming patient specimens can continue to be tested in pools.
•

Re-assessment study should start from time T0 and should consist of individual sample testing inparallel with the pooled testing. However, since all non-negative sample pools require individual
testing of all individual samples included in the pool as a part of the n-sample pooling anddeconvoluting workflow, the re-assessment study essentially consists of testing individual
samples from the negative n-sample pools.

•

Re-assessment study may pause at time T1 when a minimum of 10 consecutive positive
individual results are obtained, including both positive individual results generated fromindividual testing of samples from the non-negative sample pools following the n-sample poolingand deconvoluting workflow, and positive individual results obtained from individual testing ofsamples from the negative sample pools for the time period from T0 to T1 [T0, T1].

•

Considering that number of positive individual sample results among negative pools is K, PPA
between testing n-sample pools and assaying single specimens using the candidate test should be
calculated as PPA (EUA Test pool vs. EUA Test individual) = 100% x (10-K)/10. It is critical that all
consecutive positive samples from time period [T0, T1] are included in the PPA calculations. With
regard to calculating the PPA, all non-negative results testing pooled samples should be counted
as in agreement with positive individually tested results.

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Re-assessment Acceptance Criteria for Option 1 and Option 2•

If the PPA between pooled-testing results and individual-testing results is ≥ 90% (9 out of 10 or 10
out of 10), then continuation of testing using n-specimen pooling is acceptable.

•

If the PPA between pooled-testing results and individual-testing results is less than 85%:
o If PPA ≤ 70% (7 out of 10), reduce the pool size (consider n = 3) and repeat the reassessment testing with the new pool size until PPA of pooled compared to individual
testing is ≥ 90% OR consider cessation of pooling patient specimens.
o If PPA is 80% (8 out of 10), collect an additional 10 consecutive individually positive
samples. Then, calculate the PPA from the combined data of 20 samples, between pooled
testing results and individual testing results. If the PPA is ≥ 85%, then implementation of
testing using n-sample pooling is acceptable. Or, to compensate for lost sensitivity, reduce
the pool size (consider n = 3) and repeat the re-assessment testing with the new pool size
until PPA of pooled compared to individual testing is ≥ 85%.

•

If PPA of at least 85% cannot be reached for any pool size, cease pooling patient specimens.

•

If n-sample pooling is acceptable, return to step 5 in “Data Monitoring” to re-establish PIndividual.

Additional Resources:
1. Abdalhamid, et al., Assessment of Specimen Pooling to Conserve SARS CoV-2 Testing resources,
Amer J Clin Pathol., Vol 153, June 2020, Pages 715–718. https://doi.org/10.1093/ajcp/aqaa064.
2. Advisory on feasibility of using pooled samples for molecular testing of COVID-19.
https://www.icmr.gov.in/pdf/covid/strategy/Advisory_on_feasibility_of_sample_pooling.pdf
3. CDC, Interim Guidance for Use of Pooling Procedures in SARS-CoV-2 Diagnostic, Screening and
Surveillance Testing. https://www.cdc.gov/coronavirus/2019-ncov/lab/pooling-procedures.html
Questions and Comments:
If you have questions or comments about this procedure, please send by email to: respvirus@cdc.gov.

CDC-006-00019, Revision: 08

72

CDC/DDID/NCIRD/ Division of Viral Diseases

Effective: 03/07/2023

Division of Viral Diseases / Respiratory Viruses Branch

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel – Verification
Requirements*** DO NOT DISCARD: Important product-specific information ***CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel –Verification Requirements
Please consult the following guidance from the Centers for Medicare & Medicaid Services(CMS) regarding diagnostic tests under Emergency Use Authorization (EUA):
https://www.cms.gov/Medicare/Provider-Enrollment-andCertification/SurveyCertificationGenInfo/Policy-and-Memos-to-States-and-RegionsItems/QSO18-19-CLIA
INTENDED USE
The CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel is a realtime RT-PCR test intended for the qualitative detection of nucleic acid from SARS-CoV-2 inupper and lower respiratory specimens (such as nasopharyngeal or oropharyngeal swabs,
sputum, lower respiratory tract aspirates, bronchoalveolar lavage, and nasopharyngeal
wash/aspirate or nasal aspirate) collected from individuals suspected of COVID-19 by theirhealthcare provider 1.
This test is also for the qualitative detection of nucleic acid from the SARS-CoV-2 in pooled
samples containing up to four of the individual upper respiratory swab specimens(nasopharyngeal (NP), oropharyngeal (OP), NP/OP combined, or nasal swabs) that werecollected using individual vials containing transport media from individuals suspected of
COVID-19 by their healthcare provider. Negative results from pooled testing should not betreated as definitive. If a patient’s clinical signs and symptoms are inconsistent with a negative
result or results are necessary for patient management, then the patient should be considered
for individual testing. Specimens included in pools with a positive, inconclusive, or invalid
result must be tested individually prior to reporting a result. Specimens with low viral loads
may not be detected in sample pools due to the decreased sensitivity of pooled testing.
Testing is limited to laboratories certified under the Clinical Laboratory Improvement
Amendments of 1988 (CLIA), 42 U.S.C. § 263a, that meet the requirements to perform highcomplexity tests.
Results are for the identification of SARS-CoV-2 RNA. SARS-CoV-2 RNA is generally detectablein upper and lower respiratory specimens during infection. Positive results are indicative ofactive infection with SARS-CoV-2 but do not rule out bacterial infection or co-infection withother viruses. The agent detected may not be the definite cause of disease. Laboratories
within the United States and its territories are required to report all results to the appropriate
public health authorities.
Negative results do not preclude SARS-CoV-2 infection and should not be used as the solebasis for treatment or other patient management decisions. Negative results must be
combined with clinical observations, patient history, and epidemiological information.
Testing with the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel is intended for use bytrained laboratory personnel who are proficient in performing real-time RT-PCR assays. The
CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel is only for use
under a Food and Drug Administration’s Emergency Use Authorization.
REQUIRED MATERIALSThe 2019 novel coronavirus positive control (nCoVPC) is provided with the CDC 2019-nCoV
Real-Time RT-PCR Diagnostic Panel and should be prepared according to the Instructions for

For this EUA, a healthcare provider includes, but is not limited to, physicians, nurses, pharmacists,
technologists, laboratory directors, epidemiologists, or any other practitioners or allied healthprofessionals.

1Document #: CDC-006-00005

Revision #: 06

Effective Date: 12/01/2020

Page 1 of 5

Division of Viral Diseases / Respiratory Viruses Branch

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel – Verification
Requirements*** DO NOT DISCARD: Important product-specific information ***Use. The nCoVPC consists of an RNA transcript of the 2019-nCoV N gene as well as human
RNase P gene segment. nCoVPC will yield a positive result with the following primer and probe
sets: 2019-nCoV_N1, 2019-nCoV_N2, and RP.
Approximately 2 mL of an upper respiratory specimen (e.g. nasopharyngeal swab (NPS) in
transport media) are needed for testing. Specimens may be pooled if less than 2 mL of one
specimen is available.
Refer to CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel package insert (manufacturer
instructions) for additional reagents, materials, and instructions.
PRECAUTIONSThis reagent should be handled in an approved biosafety level 2 (BSL-2) handling area to
avoid contamination of laboratory equipment and reagents that could cause false positive
results. This product is an RNA transcript and is non-infectious. However, the nCoVPC should
be handled in accordance with Good Laboratory Practices.
Store reagent at appropriate temperatures (see Instructions for Use) and hold on ice whenthawed.
Please use standard precautions when handling respiratory specimens.
INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE
QIAamp® DSP VIRAL RNA MINI KIT OR THE QIAamp® VIRAL RNA MINI KIT
•
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 560 µL of lysis buffer into each of nine tubes labeled 1-9.
Add 140 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 14 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 14 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 14 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE QIAGEN
EZ1® ADVANCED XL
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 280 µL of lysis buffer into each of nine tubes labeled 1-9.
Add 120 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.

Document #: CDC-006-00005

Revision #: 06

Effective Date: 12/01/2020

Page 2 of 5

Division of Viral Diseases / Respiratory Viruses Branch

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel – Verification
Requirements*** DO NOT DISCARD: Important product-specific information ***•Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE ROCHE
MagNA PURE TOTAL NUCLEIC ACID KIT OR THE ROCHE MagNA PURE NUCLEIC ACIDISOLATION KIT I
•
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 300 µL of lysis buffer into each of nine tubes labeled 1-9.
Add 100 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE ROCHE
MagNA PURE 24 AND TOTAL NUCLEIC ACID ISOLATION KIT
•
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 400 µL of lysis buffer into each of nine tubes labeled 1-9.
Add 100 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE ROCHE
MagNA PURE 96 DNA AND VIRAL NA SMALL VOLUME KIT
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 350 µL of lysis buffer into each of nine tubes labeled 1-9.
Add 100 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.

Document #: CDC-006-00005

Revision #: 06

Effective Date: 12/01/2020

Page 3 of 5

Division of Viral Diseases / Respiratory Viruses Branch

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel – Verification
Requirements*** DO NOT DISCARD: Important product-specific information ***••To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE
PROMEGA MAXWELL® RSC 48
•
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 330 µL of lysis buffer (300 µL of lysis buffer + 30 µL Proteinase K, included in thekit) into each of nine tubes labeled 1-9.
Add 120 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

INSTRUCTIONS FOR PREPARING SAMPLES BEFORE EXTRACTION WITH THE
BIOMÉRIEUX NucliSENS easyMAG OR THE BIOMÉRIEUX EMAG
•
•
•
•
•
•
•
•

Refer to the 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use forreconstitution of the materials for use. RNA should be kept cold during preparation and use.
Make a 1/10 dilution of nCoVPC by adding 5 µL of nCoVPC into 45 µL of nuclease-free wateror 10 mM Tris.
Aliquot 1000 μL or 2000 µL of pre-aliquoted easyMAG lysis buffer into each of nine tubes
labeled 1-9 for the easyMAG or eMAG, respectively.
Add 100 µL of upper respiratory specimen (e.g. NPS in viral transport media) into each of
the nine labeled tubes with lysis buffer.
To prepare samples at a moderate concentration, spike 12 µL of undiluted nCoVPC(rehydrated as described in the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Instructions for Use) into each tube labeled 1-3 containing lysis buffer and specimen.
To prepare samples at a low concentration, spike 12 µL of 1/10 dilution of nCoVPC into
each tube labeled 4-6 containing lysis buffer and specimen.
To prepare negative samples, spike 12 µL of nuclease-free water into each tube labeled 7-9
containing lysis buffer and specimen.
Perform extractions of all nine samples according to the CDC 2019-nCoV Real-Time RT-PCR
Diagnostic Panel Instructions for Use.

PROCEDURE
Follow the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel Instructions for Use for testing
the nine extracted samples at least once.
EXPECTED RESULTS
Moderate nCoVPC samples should be positive for 2019-nCoV.
Low nCoVPC samples should be positive for 2019-nCoV.
Negative upper respiratory samples should be negative for 2019-nCoV.
≥90% of test results should be in agreement with the expected results. If test results are<90% in agreement with expected results, contact CDC at respvirus@cdc.gov.

Document #: CDC-006-00005

Revision #: 06

Effective Date: 12/01/2020

Page 4 of 5

Division of Viral Diseases / Respiratory Viruses Branch

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel – Verification
Requirements*** DO NOT DISCARD: Important product-specific information ***LIMITATIONSThis test has not been FDA cleared or approved.
This test has been authorized by FDA under an EUA for use by authorized laboratories.
This test has been authorized only for the detection of nucleic acid from 2019-nCoV, not for
any other viruses or pathogens.
This test is only authorized for the duration of the declaration that circumstances exist
justifying the authorization of emergency use of in vitro diagnostics for detection and/or
diagnosis of SARS-CoV-2 under Section 564(b)(1) of the Federal Food, Drug, and Cosmetic
Act, 21 U.S.C. § 360bbb-3(b)(1), unless the authorization is terminated or revoked sooner.
QUESTIONS
Please send questions or comments by email to respvirus@cdc.gov.
DISTRIBUTION
Distributed to qualified laboratories by Centers for Disease Control and Prevention, 1600
Clifton Road, Atlanta, GA, 30329 USA

Document #: CDC-006-00005

Revision #: 06

Effective Date: 12/01/2020

Page 5 of 5

Division of Viral Diseases/Respiratory Viruses Branch

2019-nCoVEUA-01

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Product Information Sheet***DO NOT DISCARD: Important product-specific information***

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
For use under EMERGENCY USE AUTHORIZATION (EUA) only.
Rx only
CATALOG: 2019-nCoVEUA-01
KIT LOT:EXPIRATION DATE:INTENDED USE
The CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel is a real-time RT-PCR test intended
for the qualitative detection of nucleic acid from SARS-CoV-2 in upper and lower respiratory specimens (such as
nasopharyngeal or oropharyngeal swabs, sputum, lower respiratory tract aspirates, bronchoalveolar lavage, andnasopharyngeal wash/aspirate or nasal aspirate) collected from individuals suspected of COVID-19 by their healthcare
provider 1.
This test is also for the qualitative detection of nucleic acid from the SARS-CoV-2 in pooled samples containing up to
four of the individual upper respiratory swab specimens (nasopharyngeal (NP), oropharyngeal (OP), NP/OP combined,or nasal swabs) that were collected using individual vials containing transport media from individuals suspected of
COVID-19 by their healthcare provider. Negative results from pooled testing should not be treated as definitive. If apatient’s clinical signs and symptoms are inconsistent with a negative result or results are necessary for patientmanagement, then the patient should be considered for individual testing. Specimens included in pools with a positive,
inconclusive, or invalid result must be tested individually prior to reporting a result. Specimens with low viral loads may
not be detected in sample pools due to the decreased sensitivity of pooled testing.
Testing is limited to laboratories certified under the Clinical Laboratory Improvement Amendments of 1988 (CLIA), 42
U.S.C. § 263a, that meet the requirements to perform high complexity tests.
Results are for the identification of SARS-CoV-2 RNA. SARS-CoV-2 RNA is generally detectable in upper and lower
respiratory specimens during infection. Positive results are indicative of active infection with SARS-CoV-2 but do notrule out bacterial infection or co-infection with other viruses. The agent detected may not be the definite cause of
disease. Laboratories within the United States and its territories are required to report all results to the appropriate
public health authorities.
Negative results do not preclude SARS-CoV-2 infection and should not be used as the sole basis for treatment or other
patient management decisions. Negative results must be combined with clinical observations, patient history, andepidemiological information.
Testing with the CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel is intended for use by trained laboratory personnel
who are proficient in performing real-time RT-PCR assays. The CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time
RT-PCR Diagnostic Panel is only for use under a Food and Drug Administration’s Emergency Use Authorization.
PACKAGE CONTENTSPACKAGING

Oligonucleotide
Box

Control Box

COMPONENT
2019-nCoV_N1 Combined
Primer/Probe Mix
2019-nCoV_N2 Combined
Primer/Probe Mix
RP Combined Primer/Probe
Mix
nCoVPC 2019-nCoV Positive
Control (non-infectious)PART
NUMBERCOMPONENTLOT NUMBER

VIALSPER KITQUANTITY
/VIAL

STATE

1

22.5 nmol

Dried

1

22.5 nmol

Dried

1

22.5 nmol

Dried

4

1 x 104
copies/µL

Dried

IVD
For this EUA, a healthcare provider includes, but is not limited to, physicians, nurses, pharmacists, technologists,
laboratory directors, epidemiologists, or any other practitioners or allied health professionals.

1Document #: CDC-006-00006

Revision #: 03

Effective Date: 12/01/2020

Page 1 of 2

Division of Viral Diseases/Respiratory Viruses Branch

2019-nCoVEUA-01

CDC 2019-nCoV Real-Time RT-PCR Diagnostic Panel
Product Information Sheet

STORAGE INSTRUCTIONS
Upon receipt, store at 2-8°C. Refer to the CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic
Panel Instructions for Use before opening and preparing reagents for use.
PROCEDURE/INTERPRETATION/LIMITATIONS
Users should refer to the CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel
Instructions for Use posted on the FDA website for all IVD products used under Emergency Use Authorization,
http://www.fda.gov/MedicalDevices/Safety/EmergencySituations/ucm161496.htm.
This test has not been FDA cleared or approved.
This test has been authorized by FDA under an EUA for use by authorized laboratories.
This test has been authorized only for the detection of nucleic acid from 2019-nCoV, not for any other viruses orpathogens.
This test is only authorized for the duration of the declaration that circumstances exist justifying the authorization of
emergency use of in vitro diagnostics for detection and/or diagnosis of SARS-CoV-2 under Section 564(b)(1) of theFederal Food, Drug, and Cosmetic Act, 21 U.S.C. § 360bbb-3(b)(1), unless the authorization is terminated or revokedsooner.
PRECAUTIONSThis reagent should be handled in an approved BSL-2 handling area to avoid contamination of laboratory equipment
and reagents that could cause false positive results. This product is non-infectious. However, this product should be
handled in accordance with Good Laboratory Practices.
REAGENT COMPLAINTS/QUESTIONS
If you have a question/comment about this product, please contact the CDC Division of Viral Diseases/Respiratory
Viruses Branch by email at respvirus@cdc.gov.
DISTRIBUTED BY
Manufactured by the Centers for Disease Control and Prevention, 1600 Clifton Road, Atlanta, Georgia, 30329, USA

IVD

Document #: CDC-006-00006

Revision #: 03

Effective Date: 12/01/2020

Page 2 of 2