Understanding Antibody Testing for COVID-19

Structure and Characteristics of Antibody Isotypes

Before discussing antibody tests, it is worth understanding how human antibodies are classified into 5 isotypes (IgM, IgD, IgG, IgA, and IgE) according to their H chains, which provide each isotype with distinct characteristics and roles.

Molecular Testing for Viral RNA

Molecular test methods are considered the only reliable means of diagnosing an active case of COVID-19, particularly early in the infection course, and are the only means of determining if a patient is contagious to others. By detecting the presence of viral genetic material in the nasal, oral, and respiratory tracts, one can determine if a patient is actively shedding virus which can be spread to others. These tests are perhaps of greatest utility early in the course of infection as they can confirm viral presence up to 2 days before the onset of symptoms. Given that antibodies may not be detectable until 6-7 days after symptom onset, molecular tests can accelerate the diagnostic window by up to 9 days. The duration of viral shedding can be highly variable and depend on severity of symptoms, length of illness, and patient-specific immune response. Generally speaking, viral shedding is undetectable 21 to 35 days after symptom onset or 3 to 5 days after a patient becomes asymptomatic. At the end of the disease course, viral load ultimately becomes undetectable and therefore a molecular test will not detect a prior infection, even one that has recently resolved. Although point-of-care molecular tests for SARS-CoV-2 are becoming more widely available with faster result times, all currently available tests require a laboratory analyzer platform which are generally in short supply and on back order for several months. There remains a backlog of samples for molecular testing at many laboratories and public health officials have put in place strict guidelines requiring severe symptoms to qualify to receive a molecular test.

Therefore, there is a clear role for antibody testing as an important tool in the diagnostic toolbox for COVID-19. Antibody testing can provide important insight to individuals about their functional immunity to the ongoing pandemic, giving peace of mind and assisting with decisions about return to community activities and the workplace. These tests also provide valuable information to public health officials about the spread of virus in different communities, especially in light of the high reported numbers of asymptomatic cases. Furthermore, although never a primary diagnostic tool, antibody status can be used to aid the clinical diagnosis of suspected noncritical cases that present 7 days or more after the start of symptoms. In these cases, the use of simple, cost-effective, point-of-care antibody cassette systems can offload the pressure on molecular testing throughput to make those resources more immediately available to the acutely ill and critical patients.

Principle of Antibody Assays

With the rapid acceleration of the COVID-19 pandemic, there has been a rush to develop tests that can detect the presence of antibodies produced by the body in response to exposure/infection with the SARS-CoV-2 virus. These testing methodologies rely on the antigen-antibody binding affinity described previously. The test principle relies on a recombinant antigen produced in the laboratory which is designed to mimic specific structures of the virus, causing any antibodies present in whole blood or serum with a binding affinity to attach to the antigen. The specificity of the test can be highly dependent on the target antigen that is chosen as some viral structures can be highly conserved across broad families of virus and others can be highly derived and specific to a given strain. Therefore, it is important to understand the cross reactivity to other viruses of the test being used so as to avoid misinterpretation of a false positive result that may be detecting an antigen from a similar virus, yet not confer immunity to the SARS-CoV-2. These tests fall into 2 broad categories: laboratory-based immunoassays which require a reader or analyzer to detect the reaction, and cassette-based systems which can be read at the point of care through a change in color in an indicator region visible to the naked eye.

Laboratory-based immunoassays have several technical advantages when compared with cassette-based systems. Because they are read by sensitive laboratory instrumentation and a controlled aliquot is delivered to the test system, they can be considered quantitative or semiquantitative tests. Therefore, they can determine how much of a given antibody (titer level) is present per unit volume of serum and, when operating at the margins of detection, their sensitivity and specificity can be higher than cassette-based systems. However, these systems have real disadvantages. Infrastructure costs, requirements for venous puncture, additional steps in sample processing, time to obtain results, and practical throughput challenges are problematic and substantially increase cost per test.

Cassette-based systems rely on a color change that is visible to the human eye and is more appropriate for use at the point of care with almost immediate results available to health care workers and patients. Cassette systems use the principle of lateral flow immunoassay or immunochromatography. The cassette contains a shallow well into which approximately 10 to 15 μl (eg, 2 small drops from a finger prick) of whole blood, serum, or plasma are placed along with a small quantity of buffer specific to the test kit. The blood and buffer are absorbed into a porous test strip which is impregnated with recombinant viral antigens doped with an indicator (eg, colloidal gold, latex particles, europium, or quantum dots). Antibodies from the serum bind to antigens in the test strip and are wicked laterally along the length of the test strip. In the indicator regions of the test kit, anti-human antibodies which are immobilized in the test strip will bind to the antigen-antibody complex and hold them in the indicator region. The colloidal gold, or other colorant, accumulates in the indicator region leading to a visible change in color along a narrow band of the wicking substrate. There may be one or more indicator regions with anti-human antibodies that are specific to IgM, IgG, or other immunoglobulins. All tests should also contain a “control” indicator line to confirm that the test sample has appropriately wicked along the length of the assay and antigens in the test kit remain viable, therefore confirming a valid test procedure (Fig. 1 ). This entire process, from finger-stick to result, occurs in less than 15 minutes.

These point-of-care test cassettes are intended to be read as a binary outcome (presence/absence) for each antibody indicator region. Although the degree of color change may be an indicator of the quantity of antibody present in a sample, variability between antibody concentrations in whole blood, plasma, and serum along with a buffer that is added in “drops” make it unreliable to attempt interpretation of color change intensity with most currently available technology. Kits that use fluorescent indicators can provide semiquantitative results with a hand-held or tabletop reader and appropriate sample preparation techniques but cannot be read by eye. The advantages of lateral flow immunoassay kits are that they can be produced at low cost and in large quantities, they can be used point of care with finger-stick whole blood samples, require low human and facility resource utilization, and can provide very rapid test results—while a patient remains on site. This makes lateral flow immunoassay cassettes an ideal choice for population level sampling and workforce sampling for small and large employers, as discussed further.

Interpreting Point-of-Care Antibody Tests

Because most test kits contain indicator lines for both IgM and IgG antibodies, there are several different combinations and corresponding interpretations of results that may occur. Results themselves should always be interpreted in accordance with the package insert and instructions for use with the specific test kit being utilized, but the following discussion is helpful in linking the binary results of the typical antibody test to clinical meaning and making informed recommendations for patients and the workforce. As discussed earlier, IgM antibodies are produced in the short term after infection, whereas IgG are produced in a more delayed timescale. The use of both is advantageous [[18], [19], [20]]. In a recent study, of 535 plasma samples taken from 173 patients with SARS-CoV-2, the median seroconversion time for total antibody (Ab), IgM, and then IgG were day 11, day 12, and day 14, separately. The presence of antibodies was <40% among patients within 1-week since onset, and rapidly increased to 100.0% (Ab), 94.3% (IgM), and 79.8% (IgG) 15 days after onset. By contrast, RNA detectability decreased from 66.7% in samples collected before day 7 to 45.5% during day 15-39. Combining RNA and antibody detections significantly improved the sensitivity of pathogenic diagnosis for COVID-19 (P < .001), even in early phase of 1 week since onset (P = .007). Moreover, a higher titer of Ab was independently associated with a worse clinical classification (P = .006) consistent with the understanding that more symptomatic patients are likely to have higher antibody responses, and higher inflammatory markers, on average. Using the SARS coronavirus as an example, IgM antibodies generally rise above the detectable threshold for these point-of-care tests in approximately 5 to 7 days after the initial onset of symptoms (assuming a patient does develop symptoms after infection). The IgM then remains above the detection threshold for 14 to 21 days from symptom onset. About midway through the rise and fall of IgM production, around day 14 after symptom onset, IgG will rise above detection levels. IgG production will generally continue to rise for 28 to 35 days after symptom onset, peaking around or after clinical recovery. IgG typically has a long half-life and will remain above detectable thresholds for months or even years after the resolution of infection.

The clinical level of functional immunity imparted by SARS-CoV-2 IgG antibodies, and their critical concentration cutoffs, have not been determined for this virus. Therefore, recommendations must be based on similar viruses and knowledge of the immune system. Ultimately, this information will be available and quantitative tests will be valuable in determining immunity titers months and years after COVID-19 infection or vaccination, once vaccines exist. Based on experience with other viruses, if reinfection does occur with the same pathogen one would expect symptom severity and duration to be greatly reduced because of the inherent memory of the immune system. Furthermore, the hyperimmune response that leads to acute respiratory distress syndrome and which is associated with the high mortality rate of this virus would expect to be mitigated by the presence of a prior immune response.

For the interpretation of results in an effort to make meaningful recommendations on community behavior to test recipients, there are several factors that must be considered:

• 1.

  Is the subject symptomatic or asymptomatic at the time of antibody testing?

• 2.

  If symptomatic, how long since symptom onset?

• 3.

  IgM positive or negative?

• 4.

  IgG positive or negative?

Interpreting Results When the Subject is Asymptomatic at the Time of Testing

Interpreting Results When Symptomatic at the Time of Testing

Long-Term Immunity—Does It Exist With COVID-19?

A study in the wake of SARS, the similar coronavirus that triggered an epidemic in 2003, showed that survivors maintained neutralizing antibodies for 2 years on average, with the number of antibodies declining thereafter. Other coronaviruses in circulation in the human species also lead to at least partial immunity for some period. The immunity question has implications for whether COVID-19 follows an annual cycle like seasonal influenza or becomes dormant for multiple years and then erupts again, according to a recent publication in the journal Science. The authors noted that 2 other coronaviruses in circulation (OC43 and HKU1), which cause common colds, result in about 45 weeks of immunity on average. If the new virus follows that pattern, it would probably create annual outbreaks, they found [21]. However, generally speaking, the sicker you become due to an infection and the robust immune system response to it, the more robust the immunity provided. Therefore, one might assume that those people who get the most ill have the longest and most effective immunity after a re-exposure to the same virus. In a study by Callow et al. [22], people volunteered to have coronavirus 229E, which also causes common cold symptoms, inoculated up their nares. Ten became infected, and 8 developed cold symptoms. One year later, all but one of them returned to be reinfected again. Most reinfected, but those who had been ill before did not develop cold symptoms. Moreover, the period during which the patients shed the virus, and were potentially contagious, was shorter. The new virus, SARS-CoV-2, is genetically similar to the first SARS virus—hence the “2”—but it affects people differently. It is not as lethal as the original SARS but is more easily spread (R_O is higher) [23]. Many people who are infected do not develop symptoms at all and yet can potentially transmit the virus to others.

A very recent report from China that has not yet been peer-reviewed found a wide range of antibodies among people with mild cases of the virus. Most strikingly, younger people had fewer antibodies in the wake of the disease—and 30 percent of those sampled had low levels. Some individuals had no trace of antibodies. That has raised the question of whether a person with a mild or asymptomatic infection, but confirmed by the sensitive polymerase chain reaction test, might still be susceptible to a second infection. The neutralizing antibody titers were positively correlated with plasma CRP levels but negatively correlated with the lymphocyte counts of patients at the time of admission, indicating an association between degree of humoral response and cellular immune response. However, those with mild disease who do not develop high levels of antibodies may be at increased risk of reinfection, but are not likely to get significant symptoms, although they may be contagious. Therefore, those without antibody production should likely be treated as if they never had the disease from a social distancing perspective, even if they were proven to be positive with molecular testing.