Laboratory Studies in Coronavirus Disease 2019 (COVID-19) 

Updated: Nov 12, 2020
  • Author: James J Dunn, PhD, D(ABMM), MT(ASCP); more...
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Laboratory Studies

Because the signs and symptoms of coronavirus disease 2019 (COVID-19) may overlap with those of other respiratory pathogens, it is important to perform laboratory testing to specifically identify symptomatic individuals infected with severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). Moreover, it is estimated that up to 40% of people with SARS-CoV-2 infection may be asymptomatic (subclinical infection) or presymptomatic, and still potentially capable of transmitting the virus to others. [1, 2]  Therefore, in certain cases, individuals without obvious signs or symptoms of SARS-CoV-2 infection also require testing.

Currently, there are three basic types of tests to determine if an individual has been infected with SARS-CoV-2: viral nucleic acid (RNA) detection, viral antigen detection, and detection of antibodies to the virus. Viral tests (nucleic acid or antigen detection tests) are used to assess acute infection, whereas antibody tests provide evidence of prior infection with SARS-CoV-2. (The US Food and Drug Administration [FDA] has not authorized the use of antibody tests for the diagnosis of acute infection.)

Cell culture isolation of SARS-CoV-2 is possible, but the Centers for Disease Control and Prevention (CDC) recommends that clinical laboratories not attempt this unless it is performed in a biosafety level 3 (BSL-3)–certified laboratory.

With any type of laboratory test, the clinical accuracy or reliability depends on performance characteristics such as sensitivity and specificity, as well as the pretest probability that a person has SARS-CoV-2 infection and the prevalence of COVID-19 in the local community. Taken together, these parameters determine whether a positive or negative result should be interpreted as correct.

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Specimen Selection, Collection, and Transport

Collecting the appropriate specimen at the right time and transporting it to the laboratory under proper conditions are critical pre-analytic components of the testing process. The type of specimen collected will depend to some extent on the authorized SARS-CoV-2 viral test used; the instructions for use (IFU) provided by the manufacturer for FDA Emergency Use Authorization (EUA) will detail the approved sample types. Initial direct viral detection is typically performed using an upper respiratory tract (URT) specimen. Such specimens may include healthcare personnel (HCP)–collected nasopharyngeal (NP), oropharyngeal (OP), mid-turbinate (MT), or anterior nares swabs, as well as NP or nasal wash/aspirate specimens. [3] Swabs should be composed of synthetic fibers and have plastic or wire shafts. Wooden shafts or calcium alginate swabs may contain substances that inhibit some viruses and nucleic acid detection reactions. If both NP and OP swabs are collected, they can be combined in a single tube to maximize test sensitivity and conserve transport devices.

In certain situations, it may be acceptable for patients to collect their own nasal or mid-turbinate swab or saliva (1-5 mL), as long as they are given a clear, step-by-step protocol of the process. [4] Swabs are typically placed in 1.5-3 mL of viral transport media (VTM) prior to transport to the laboratory. In some cases, for point-of-care tests, the manufacturer's IFU may call for direct testing of the swab without dilution in VTM. Collection of lower respiratory tract (LRT) specimens such as sputum, bronchoalveolar lavage (BAL), or tracheal aspirate (in mechanically ventilated patients) may be warranted in certain cases. However, in hospitalized patients with LRT disease, the Infectious Diseases Society of America (IDSA) recommends first obtaining a URT specimen for testing; if it is negative for SARS-CoV-2, an LRT specimen can be collected and tested. [4] HCP collecting specimens and in close patient contact should use recommended personal protective equipment (PPE) and maintain good infection control practices. Other HCP handling specimens after collection should follow standard precautions.

Viral tests (nucleic acid or antigen) are recommended to diagnose acute infection of symptomatic and asymptomatic individuals. [4, 5]  The results can then be used to guide contact tracing, treatment options, and isolation requirements. Asymptomatic individuals who have had direct contact with a known or suspected case of COVID-19 while not wearing appropriate PPE can be tested, but testing is not recommended in asymptomatic, otherwise healthy, non-immunocompromised individuals seeking healthcare services in communities where the prevalence of COVID-19 is less than 2% (if they are not undergoing emergent surgery or aerosol-generating procedures). [4] In contrast, testing is recommended for these same patients if they are hospitalized in communities with a high prevalence (ie, ≥10%) of COVID-19.

Asymptomatic, immunosuppressed patients being admitted to the hospital or persons undergoing immunosuppressive procedures should be tested within 48-72 hours of admission.

The decision to test asymptomatic patients will often depend on the availability of testing resources and the timeliness of results. However, asymptomatic individuals requiring urgent surgery can be tested as close to the planned procedure as possible (eg, within 48-72 hours). The need for testing asymptomatic persons prior to aerosol-generating procedures will to some extent depend on the availability of appropriate PPE.

Among HCP, the CDC recommends that testing be considered in four situations [6] :

  • COVID-19–consistent signs or symptoms in HCP
  • Asymptomatic HCP in whom there has been known or suspected exposure to SARS-CoV-2
  • Asymptomatic HCP in nursing homes, in response to an outbreak in the facility
  • HCP previously diagnosed with SARS-CoV-2 infection, to determine if they are no longer infectious

The fourth recommendation, which requires serial tests and improvement in symptoms, could be considered as a means of allowing HCP to return to work earlier than would a symptom-based assessment. However, the period of time during which RNA may be detectable in URT specimens may make the period prior to return to work longer than would be experienced with the symptom-based strategy.

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Nucleic Acid Detection

The currently recommended modality for the diagnosis of acute or current SARS-CoV-2 infection is a nucleic acid amplification test (NAAT) that detects one or more RNA gene targets specific to the virus. [5]  Symptomatic patients, individuals with known or suspected COVID-19 exposure, and persons from an area with high disease prevalence can be tested with this method. [4] IDSA guidelines also suggest that patients who initially test negative, but in whom there is a high clinical suspicion of COVID-19, should receive an additional NAAT, which is estimated to increase the sensitivity of the test by 17%. [4]

Presently, there are five SARS-CoV-2 assays that have EUA approval for the testing of asymptomatic patients or for use as part of community surveillance. [7] It is important, however, to test only asymptomatic patients with high pretest probability of having SARS-CoV-2, since this decreases the frequency of false negatives by increasing the negative predictive value (NPV). [8] .

Gene targets and multiplex NAATs

Some of the most frequently tested gene targets for the detection of SARS-CoV-2 include the E, S, and N genes and the open reading frame ORF1a/1b. [9] While some gene targets may also detect SARS-CoV-1, which is no longer known to be circulating in the human population, there is little cross-reactivity with the endemic coronaviruses that are associated with the common cold. [9, 10]  Detection of the E gene appears to have the highest analytic sensitivity, [11]  with detection capabilities for this gene being far below the estimated viral load for SARS-CoV-2–positive patients; [12] NAAT assays can detect as few as 10 copies per reaction. [13] However, different NAAT systems vary in terms of clinical sensitivity, independent of the gene detected. [9] In addition to detecting SARS-CoV-2, many NAATs have been approved as multiplex assays, which can simultaneously detect other respiratory viruses such as influenza A and B, as well as bacteria that cause atypical pneumonia. [7]

Viral load and sensitivity of NAATs

The timing of patient testing in the course of the disease can also impact the sensitivity of nucleic acid analysis. Viral RNA present in URT specimens from patients infected with SARS-CoV-2 is highest at the onset of symptoms. [14, 15, 16]

The viral loads for pre-symptomatic patients and those with subclinical infections are currently a topic of intense research. Viral loads in patients who are pre-symptomatic (ie, who are asymptomatic at the time of the test but who subsequently develop symptoms) do not appear to be significantly different than those in individuals with subclinical or symptomatic infections. [17, 18, 19] This would likely indicate that the sensitivity of nucleic acid testing would be the highest earlier in the course of the infection and should be equally effective prior to or at the onset of symptoms, though how soon after exposure to the virus that an NAAT can readily detect SARS-CoV-2 RNA is in question. [20]

After the first week of symptoms, which correlates with diminishing severity of symptoms in patients with a mild presentation of COVID-19, SARS-CoV-2 RNA levels in URT specimens begin to decrease but are still frequently detectable by NAAT. [14, 15, 16] However, some studies did not observe a sharp decrease after this first week. [17, 21] This phenomenon may be linked to the immune status of the patients in question; immunocompromised patients, as well as individuals with asymptomatic infections, have been noted to shed the virus for longer periods of time than immunocompetent or symptomatic patients, as measured by RNA detection. [22, 23, 24] Virus shedding decreases in subsequent weeks, but a low percentage of specimens may have detectable RNA for 6 weeks or longer after symptom onset. [25] Moreover, RNA in LRT specimens, such as sputum, may peak later and be detectable longer than URT-specimen RNA would. [19]  Likewise, in some patients, saliva has been shown to have higher viral loads and RNA that persists longer than do paired NP specimens. [26]  Saliva may provide a suitable alternative to NP specimens given the scarcity of swab supplies and does not necessarily need to be collected by HCP, which can reduce PPE usage.

RCT-PCR NAATs

The more than 200 NAATs with EUA approval from the FDA fall into two broad categories [7] : nucleic acid amplification based on reverse transcription polymerase chain reaction (RT-PCR) assays, and SARS-CoV-2 RNA detection through isothermal amplification.

With regard to RT-PCR assays, given that all coronaviruses have an RNA genome, it is necessary to synthesize complementary DNA (cDNA) from the RNA genome through reverse transcription, followed by PCR amplification of the cDNA with specific primers for the SARS-CoV-2 genes of interest. While all NAATs that utilize RT-PCR detect SARS-CoV-2 in this way, there are many variations that can be applied for the actual detection of the amplified genes. Most common is the use of a real-time RT-PCR, which employs fluorescence to detect the amount of amplified DNA in real time. A frequently utilized example of this is TaqMan hydrolysis. [27]  In real-time RT-PCR, the amount of gene target present in the sample typically determines the number of PCR cycles (known as the cycle threshold [Ct] value) needed before SARS-CoV-2 is detected.

All current SARS-CoV-2 RT-PCR assays with EUA approval are labeled only for the qualitative detection of gene targets specific to the virus and are not approved for quantitative measurement of the amount of virus present in the sample. While the Ct value can be reduced by increasing the amount of gene target in the sample, it can be influenced by many other factors as well, including the quality of the specimen collection technique, sample type (eg, NP sample vs saliva), gene target, and assay. There are no current recommendations for the use of Ct values in patient management, and more research on viral load kinetics is needed.

An alternative to fluorescent detection includes sequencing of the viral genome. While this has had limited applications in the detection of SARS-CoV-2, it can play an important role in determining whether a patient has become re-infected, as it may be possible to compare the sequence of the original isolate with that of a second one. [28]

Isothermal amplification NAATs

The main distinction between isothermal amplification and PCR is that in the former, amplification is achieved using a constant temperature, while in the latter, cycling of temperatures is required. [29] . Several methods of isothermal amplification are available, but the two most important ones for SARS-CoV-2 diagnostics are reverse transcription loop-mediated isothermal amplification (RT-LAMP) and transcript-mediated amplification (TMA). Both chemistries can generate more than 109 copies of a gene target in 1 hour [29] . Detection of the amplified target can be achieved with a nonspecific fluorescent dye, such as SYBR Green, that binds to double-stranded DNA, or a fluorescent probe that is specific to the target sequence. Some EUA assays also incorporate clustered, regularly interspaced, short palindromic repeats (CRISPR) for the detection of RNA amplified using RT-LAMP. [30, 31]

Throughput and turnaround times for NAATs

Many NAATs require off-board nucleic acid extraction prior to RT-PCR testing. The use of magnetic particle technologies that bind nucleic acid has allowed automation of this process, [32] and this in turn has led to the development of many high-throughput nucleic acid extraction devices that can accommodate a large number of patient samples at one time. NAATs that require nucleic acid extraction should be performed using the appropriate instrument(s) in the EUA submission and manufacturer’s IFU.

An important factor that may influence the choice of NAATs for patient testing is the speed with which the assay can detect SARS-CoV-2 RNA. Some assays can return results in as few as 5-10 minutes, although more commonly the turnaround time is an hour or more. Assays that require external nucleic acid extraction will be inherently slower. Conversely, assays that do not require prior nucleic acid extraction and can be performed with minimal hands-on time (ie, sample to answer) generally offer a much shorter time to results, though there can still be considerable variation between assays. Some of these NAATs can be performed at the point of care in patient care settings operating under the Clinical Laboratory Improvement Amendments of 1988 (CLIA) Certificate of Waiver, Certificate of Compliance, or Certificate of Accreditation. [33]  In some situations, when a specimen must be sent to a reference laboratory for testing, the turnaround time for results make take 2 days or more [34] .

Reagent shortages and specimen pooling

During the course of the pandemic, a number of laboratories have experienced shortages of one or more tests, consumables, or reagents necessary for SARS-CoV-2 testing. [35, 36] According to a survey, these shortages have limited testing in many laboratories to only about 43% of their maximum capacity, on average. [35] This reduction in capacity has impacted the ability of hospitals and labs to perform tests in accordance with IDSA guidelines regarding the repetition of initially negative tests for those patients with high clinical suspicion of COVID-19. [37]

Due to reagent shortages for SARS-CoV-2 RNA detection, many labs have considered pooling specimens for testing. Pooling of samples has been applied to surge testing and as part of epidemiologic surveillance for other respiratory viruses. [38] If the pool tests positive, the individual specimens that make up the pool must be tested individually. This strategy can be effective under two conditions. One is that there be a low community positivity rate or prevalence. If it is too high, every pool will contain a positive specimen, with reassessment needed to determine which one it is. This limits the potential reagent savings associated with pooling. The second condition is that the loss of sensitivity must not be too great with regard to the dilution of potentially positive specimens with negative specimens. If the loss of sensitivity is too high, the number of false negatives can impact patient care and management.

The number of specimens that can be pooled together will depend on the community prevalence and the loss of sensitivity of the assay. [39] Studies have shown that pooling of even a small number of individual specimens may result in 6-7% more false negatives. [40] In particular, pooling specimens appears to impact the sensitivity of detection in patients with low viral loads, as estimated by real-time RT-PCR. [40, 41]

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Viral Antigen Detection

As of October 26, 2020, the FDA had issued EUA certification for six commercially available SARS-CoV-2 antigen detection tests. [7] These are typically lateral-flow immunoassays intended for the qualitative detection of nucleocapsid protein antigen directly from NP and/or nasal swabs; the presence of such antigen implies current SARS-CoV-2 infection. Several of the antigen test systems require an instrument reader to determine the results, whereas others can be read visually by the operator.

In general, the workflow of the antigen detection test requires placement of the swab into a reagent solution that is mixed and then applied to a test cartridge. The sample is then allowed to migrate along the test strip, where, if the antigen is present, specific antibodies to SARS-CoV-2 antigen will react with the sample and produce a colorimetric or fluorescent signal within 10-15 minutes. All test systems require use of the swab provided in the kit, and in most cases it is applied directly without dilution in VTM. The tests include built-in internal controls that must be visualized by the operator or read by the instrument to ensure the validity of results. In all cases, the manufacturer’s IFU should be strictly followed.

Antigen detection tests with EUA from the FDA are authorized for diagnostic testing in symptomatic individuals within the first 5-12 days of symptom onset. It is best to perform this type of testing in the early stages of infection, when the viral load is generally highest, since antigen levels in specimens collected beyond 5-7 days post symptom onset may drop below the assay's detection limit. There is scant information on the utility of antigen testing for SARS-CoV-2 in asymptomatic persons. At this time, the CDC recommends that antigen tests not be used to make decisions about discontinuing isolation. [42]

From a regulatory standpoint, all of the EUA antigen detection tests can be employed by laboratories certified under CLIA to perform moderate-, high-, or waived-complexity tests and at the point of care in patient care settings operating under a CLIA Certificate of Waiver, Certificate of Compliance, or Certificate of Accreditation. In other words, the testing does not necessarily need to be performed in a laboratory by a trained staff of technologists but can be used in other settings (eg, physicians' offices) that have regulatory approval to provide this simple type of assessment. Currently, none of the antigen-based tests are approved for home use. Regardless of where they are performed, CLIA-certified laboratories or performing sites within the United States are required to report all positive results to the appropriate public health authorities. [42]

Although antigen detection tests are simple, easy to perform, and fairly inexpensive, one of the major concerns associated with their use is the lack of analytic and clinical sensitivity compared with RNA detection tests. A meta-analysis of four commercially available antigen tests available outside of the United States showed that the average sensitivity was 56.2%. [43] In a study using HCP-collected nasal swabs in two EUA antigen tests, overall agreement between them was 98.1%, while, compared with RT-PCR, one of the tests displayed a positive percent agreement (PPA) of 82.4% and a negative percent agreement (NPA) of 99.5%, in adult patients with onset of symptoms less than 7 days prior to collection. [44]  

Due to these limitations, it may be necessary to confirm a negative antigen test with NAAT when the pretest probability is relatively high, especially if the patient is symptomatic or has a known exposure to a confirmed COVID-19 case. A negative antigen test does not necessarily need to be confirmed by NAAT if the pretest probability is low, as in cases in which the individual is asymptomatic or has no known exposures. [42] Generally speaking, interpretation of the results from antigen detection tests will depend on the local prevalence of COVID-19, the assay performance characteristics (ie, sensitivity/specificity), and the patient’s clinical signs, symptoms, and history.

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Antibody Detection

Tests used to detect anti–SARS-CoV-2 antibodies in blood and saliva specimens are employed to identify patients currently or previously infected with SARS-CoV-2. In general, detectable levels of antibodies can take several days to weeks to develop; therefore, antibody detection tests have limited utility in the diagnosis of acute infection, with testing by NAAT being preferred for this. [5]  However, the use of antibody tests is viable as an aid to the diagnosis of COVID-19, even if the tests should not represent the sole basis for the determination of acute SARS-CoV-2 infection. [45, 46]  

Crucial to the interpretation of an antibody detection test is knowledge of the nature, dynamics, and timing of the antibody response to SARS-CoV2 infection. Several studies have shown that in most patients, seroconversion occurs by 2 weeks after the onset of symptoms; almost all patients have detectable levels of anti–SARS-CoV2 antibodies by day 28 post symptom onset. [47, 48] Studies involving hospitalized patients with SARS-CoV-2 infection confirmed by NAAT showed the presence of all isotypes of anti-SARS-CoV-2–specific antibodies, including immunoglobulin M (IgM), IgA, and IgG. [49] Detectable titers of IgM and IgA appeared 7-14 days post symptom onset (and in rare cases, as early as day one after symptom onset). The timing of peak IgG levels varies. In some cases, IgG can be detected simultaneously with IgM, but in the majority of patients it is delayed by a few days and plateaus between 15 and 21 days. [47] In most individuals, the antibody titer correlates with disease severity; however, some patients appear to have undetectable levels of antibodies. [45, 50]

Types of antibody detection tests

Two SARS-CoV-2 antigens have been used as antigenic targets for the development of antibody detection assays: spike (S) and nucleocapsid (N) proteins. S protein binds to the cell surface angiotensin-converting enzyme 2 (ACE2) receptor, whereas the N protein plays crucial roles in viral replication and assembly, is highly conserved, and induces antibodies sooner than S protein during infection. [51] However, studies suggest that a host’s neutralizing antibodies are predominantly those directed against the S protein. [52] .

Available immunoassays are capable of detecting all isotypes of antibodies: IgA, IgM, and IgG. Total antibody detection assays are designed to determine levels of all isotypes combined. All currently authorized tests provide either qualitative or semi-quantitative results.

Commercial SARS-CoV-2 serologic assays include the following:

  • Automated direct chemiluminescence immunoassay (CIA)
  • Enzyme-linked immunosorbent assay (ELISA)
  • Rapid lateral-flow assay (LFA)
  • Bioassays such as those employing plaque reduction and microneutralization

Automated direct chemiluminescence immunoassay

In CIA, quantification is provided by luminescence detection, with this method using a combination of recombinant antigens coated onto magnetic beads. Data show that the CIA technique for detection of SARS-CoV-2 antibodies offers excellent sensitivity and specificity for determination of the presence of total antibodies or selected isotypes. It also provides an automation option, allowing high-throughput sample testing. [53]

Enzyme-linked immunosorbent assay

ELISA can detect total antibody content or IgA, IgM, or IgG selectively. ELISAs are often utilized to study the timeline/seroconversion of antibody production in patients. The basis of the ELISA format is antibody-antigen reactivity; the analyte in a patient sample is detected using an enzyme conjugate that converts specific substrate into a measurable signal that is used as a readout. The advantages of ELISA include high-throughput patient testing. Disadvantages include limited reproducibility due to lack of standardization, variable detection limits, and use of variable antigens. Therefore, sensitivity and specificity vary widely across assays and even within assays validated by different users. [53]

Rapid lateral-flow assay,

LFA, similar to ELISA, employs SARS-CoV-2 antigen as a capture agent, but in a lateral-flow strip format. [54] The most appealing advantages of the lateral-flow format are fast turnaround time (10-30 minutes) when compared with classic ELISA (several hours), along with minimal sample processing. Most LFAs provide qualitative, visual results that are subjectively interpreted by the operator, enabling near-patient testing in low-resource settings. However, the current cost of LFA is higher than that of high-throughput ELISA. Moreover, although LFA has the potential to expand COVID-19 diagnostic capacity, these tests are still being evaluated, and a large study found alarming inconsistencies among 10 commercially available LFAs. [55] Therefore, LFA results should be interpreted with caution, and follow-up testing is recommended.

Bioassays

Bioassays such as those employing plaque reduction and microneutralization provide essential data for the validation of candidate diagnostic tests. However, they require specialized expertise and are offered by a limited number of highly specialized laboratories.

Other considerations in antibody testing

The FDA has issued EUAs for SARS-CoV-2 antibody tests using serum, plasma, or whole blood; [7]  it has also required commercial manufacturers who develop COVID-19 antibody tests to submit EUA requests together with an assay’s validation data. An updated list of serology tests that received EUA, along with their diagnostic specifications/test performance (including assay specificity, sensitivity, positive predictive value [PPV], and NPV), is available on the FDA website. [7] The FDA is actively involved in evaluation of the accuracy of serology tests.

The CDC has published interim guidelines for COVID-19 antibody testing, with recommendations that include preferential use of antibody assays that were granted EUA by the FDA. [46] Specific recommendations have been made with regard to maximizing the PPV of antibody detection tests; guidelines include performing serology testing on individuals with a high pretest probability of prior infection and employing antibody assays with high specificity. Moreover, the CDC recommends implementing a two-step/orthogonal testing algorithm involving two independent tests, with an initial positive result being confirmed by a different antibody assay. [46] The CDC strongly recommends interpretation of antibody test results in correlation with a patient's clinical history and that the results be interpreted and reported with caution.

Similar to the CDC, the American Association for Clinical Chemistry (AACC) recommends the use of assays that have received EUA by the FDA or laboratory-developed tests (LDTs) that have been developed and validated by high-complexity, CLIA-certified laboratories. At-home serology testing is not supported by the AACC at this time. [56]

The overall utility of antibody testing allows for the following:

  • Diagnosis of current infection in symptomatic patients, which can improve overall sensitivity of diagnosis in conjunction with NAAT
  • Monitoring of disease course, definition of time of seroconversion, and correlation of antibody titers with clinical presentation and disease severity
  • Screening or surveillance of asymptomatic patients to help determine population prevalence of COVID-19
  • Identification of convalescent serum donors - Convalescent plasma therapy could offer “passive immunity” and has been approved by the FDA as an emergency investigational new drug or provided with EUA, for early therapy during public health emergencies. Identification of optimal donors has been challenging due to a lack of established correlation between antibody titers and clinical efficacy. The FDA, however, has published guidelines on collection, antibody titer testing, and use of convalescent plasma. [57] Guidance from August 2020 recommends testing a donor’s plasma for anti–SARS-CoV-2 antibodies using an Ortho-Clinical Diagnostics VITROS anti–SARS-CoV-2 IgG assay, in which a signal-to-cutoff ratio of 12 or greater reveals high-titer convalescent plasma. In addition, the FDA recommends testing of neutralizing antibody titers if available. [57]
  • Assessment of vaccine immunogenicity and identification of individuals with protective immune status

There is an urgent need for research providing insight into immunity against SARS-CoV-2. Appropriate, validated, and well-performing serology assays play an essential role in the public health response to the COVID-19 pandemic.

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Testing For Patients with Multisystem Inflammatory Syndrome

COVID-19 in children often has a relatively mild presentation. However, starting in April 2020, reports from the United Kingdom and other countries described a Kawasaki-like disease in children and adolescents linked to SARS-CoV-2 infection. [58] The condition has been recognized as multisystem inflammatory syndrome (MIS) in children (MIS-C). MIS is considered a rare, yet serious, complication of SARS-CoV-2 infection. The exact pathophysiology and incidence of MIS remain to be determined.

The CDC and the World Health Organization (WHO) have each provided criteria for an MIS-C case definition. The definitions require patients to be less than age 21 years (CDC criteria) or less than age 19 years (WHO criteria) and to have evidence of recent or current SARS-CoV-2 infection or exposure (CDC only), the presence of documented fever, elevated markers of inflammation, at least two signs of multisystem involvement, and lack of an alternative diagnosis (eg, bacterial sepsis, toxic shock syndrome). The CDC also includes the presence of severe illness requiring hospitalization in the case definition. As more published data emerge, diagnostic criteria may be altered.

It is believed that immune dysregulation underlying MIS-C is related to past rather than acute infection with SARS-CoV-2. Serology testing for the presence of anti–SARS-CoV-2 antibodies plays an essential role in MIS-C diagnosis, since the majority of pediatric patients have a negative SARS-CoV-2 PCR result at the time of presentation with the syndrome. [58]

Laboratory tests recommended for the initial diagnosis and monitoring of disease progression include complete blood count (CBC), kidney and liver function markers, cardiac function biomarkers, and coagulation parameters. Cell counts with CBC differential, with special attention paid to platelet, lymphocyte, and neutrophil counts, allow for control of lymphopenia, neutrophilia, mild anemia, and thrombocytopenia in patients with moderate or severe symptoms. Serum electrolytes and renal function have been reported as abnormal even in patients with mild symptoms.

Although not fully understood, myocardial injury is one of the main clinical manifestations of MIS-C. [59] Therefore, close monitoring of the levels of markers of myocardial function such as troponin and brain natriuretic peptides (BNPs) are an essential part of the MIS-C workup. Inflammatory markers whose levels have been shown to correlate with disease severity include C-reactive protein (CRP), which has been reported as critically elevated in the majority of patients; erythrocyte sedimentation rate (ESR); and interleukin 6 (IL-6). Coagulopathy resulting in high fibrinogen, D-dimer, partial thromboplastin time (PTT), prothrombin time (PT), and factor VIII levels has been observed in MIS-C patients experiencing moderate to severe symptoms. Ferritin is also elevated in MIS-C, being higher in 55-76% of patients.

With the rapid spread of SARS-CoV-2, reports have emerged of MIS in adult patients (MIS-A) as well. [60]  Since there have been only a limited number of reported cases, however, the epidemiology, comorbidities, and incidence of MIS-A remain largely unknown. Similar to MIS-C, clinical presentation includes cardiovascular, gastrointestinal, and dermatologic symptoms in patients with confirmed SARS-CoV-2 infection.

Serology testing for the presence of anti–SARS-CoV-2 antibodies is crucial in the diagnosis of MIS-A. Published reports have shown that up to 30% of patients had a negative NAAT but were positive for the presence of SARS-CoV-2 antibodies. [60] The working case definition of MIS-A includes severe illness requiring hospitalization in patients aged 21 years or older, along with current or previous infection with SARS-CoV-2. Patients are evaluated for the presence of severe dysfunction of extra-pulmonary organs, including cardiac abnormalities, toxic shock, acute liver injury, and coagulopathy in the absence of severe respiratory illness. The laboratory-confirmed presence of severe inflammation is important for the diagnosis of MIS-A. The same markers are evaluated in MIS-A as in MIS-C, with CRP, ferritin, D-dimer, and IL-6 being greatly elevated. [61]

Additional, carefully conducted research is needed to understand the pathophysiology and long-term effects of MIS in children and adults. Currently, the differential diagnosis of MIS-C and MIS-A is broad, and clinicians and health departments should consider multidisciplinary care for patients who meet the criteria for these conditions.

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