Journal:Laboratory diagnosis of COVID-19 in China: A review of challenging cases and analysis
|Full article title||Laboratory diagnosis of COVID-19 in China: A review of challenging cases and analysis|
|Journal||Journal of Microbiology, Immunology and Infection|
|Author(s)||Jing, Ran; Kudinha, Timothy; Zhou, Meng-Lan; Xiao, Meng; Wang, He; Yang, Wen-Hang; Xu, Ying-Chun; Hsueh, Po-Ren|
Chinese Academy of Medical Sciences, Beijing Key Laboratory for Mechanisms Research and Precision Diagnosis of|
Invasive Fungal Diseases, Charles Sturt University, NSW Health Pathology, National Taiwan University College of Medicine
|Primary contact||Email: tkudinha at yahoo dot com|
|Volume and issue||In Press|
|Distribution license||Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International|
Since the initial emergence of coronavirus disease 2019 (COVID-19) in Wuhan, Hubei province, China, a rapid spread of the disease occurred around the world, becoming an international global health concern at the pandemic level. In the face of this medical challenge threatening humans, the development of rapid and accurate methods for early screening and diagnosis of COVID-19 became crucial to containing the emerging public health threat, and preventing further spread within the population. Despite the large number of COVID-19 confirmed cases in China, some problematic cases with inconsistent laboratory testing results were reported. Specifically, a high false-negative rate of 41% on severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) detection by real-time reverse transcription polymerase chain reaction (qRT-PCR) assays was observed in China. Although serological testing has been applied worldwide as a complementary method to help identify SARS-CoV-2, several limitations on its use have been reported in China. Therefore, the separate use of qRT-PCR and serological testing in the diagnosis of COVID-19 in China and elsewhere presents considerable challenges, but when used in combination, these methods can be valuable tools in the fight against COVID-19. In this review, we give an overview of the advantages and disadvantages of different molecular techniques for SARS-CoV-2 detection that are currently used in several labs, including qRT-PCR, gene sequencing, loop-mediated isothermal amplification (LAMP), nucleic acid mass spectrometry (MS), and gene editing techniques based on the clustered regularly interspaced short palindromic repeats (CRISPR/Cas13) system. Then we mainly review and analyze some causes of false-negative qRT-PCR results, and how to resolve some of the diagnostic dilemmas.
Keywords: SARS-CoV-2, COVID-19, qRT-PCR, serology testing, challenging cases
Soon after coronavirus disease 2019 (COVID-19) fully emerged in China at the beginning of 2020, the Chinese government immediately implemented strong measures to contain the outbreak. With great efforts, the COVID-19 cases have stabilized in China as a whole to date, albeit a small number of imported cases that intermittently emerge. However, an epidemic began to rapidly spread around the world from April to date. As of August 21, 2020 (6:48pm CEST), there had been a total of 22,536,278 confirmed cases worldwide, with the largest cumulative number of COVID-19 confirmed cases (n = 5,477,305) in the United States of America (USA), followed by Brazil (n = 3,456,652), and India (n = 2.905,823).
Some challenging cases of COVID-19 diagnosis were encountered in China and elsewhere, involving inconsistent laboratory testing results, mainly caused by false-negative real-time reverse transcription-polymerase chain reaction (qRT-PCR) detection. In this review, we summarize and discuss some possible causes of false-negative results, including how to resolve the diagnostic dilemma. We also review and discuss the advantages and disadvantages of the different lab assays for diagnosing COVID-19, including different molecular techniques and serological assays, and the value of combining qRT-PCR assays with serological testing. In brief, it is crucial to select appropriate diagnostic methods according to the phase of infection, or to use a combination of different methods and other clinical parameters in confirming the infection status of individuals.
SARS-CoV-2 etiological characteristics and genome organization
There are four genera under the subfamily coronavirus (CoVs), including α, β, γ, and δ. The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) virus, responsible for COVID-19, belongs to the β CoV genus, the seventh member of the family of CoVs possessing a single-stranded, positive-sense RNA genome. The genome of the SARS-CoV-2 virus consists of about 29,000 bases. Studies show that there are at least 12 coding regions, including open reading frames (ORF) 1 ab, S, 3, E, M, 7, 8, 9, 10b, N, 13, and 14. Among them, ORF 1 ab is the region of the RdRp gene which codes for RNA polymerase and is responsible for viral nucleic acid replication.
The structural proteins include:
- a spike (S), crucially associated with virus transmission capacity, binding to angiotensin-converting enzyme 2 (ACE2) receptors on the cell surface to get into the host cell;
- an envelope protein (E), responsible for the formation of virus envelopes and virus particles;
- a membrane protein (M), responsible for membrane proteins encoded; and
- a nucleocapsid (N), having recognition with the host RNA of the virus genome.
These functional proteins play an essential role in genome maintenance and virus replication. Beyond these, several accessory proteins also help in virus replication, including ORF3, ORF6, ORF7a, ORF7b, ORF8, and ORF9b. The amplication fragments and loci of genes coding these proteins are shown in Fig. 1.
Molecular diagnosis for COVID-19 confirmation
Real-time reverse transcription-polymerase chain reaction (qRT-PCR)
In many countries, the preferred testing method for COVID-19 confirmation is the qRT-PCR assay, which is regarded as the gold standard for virus infection confirmation. According to Diagnosis & Treatment Scheme for Coronavirus Disease 2019 (seventh edition, in Chinese), suspected COVID-19 cases are laboratory-confirmed for positive detection of SARS-CoV-2 RNA by qRT-PCR testing. This form of molecular testing offers several advantages in the diagnosis of COVID-19. Comparted to serology testing, qRT-PCR testing is much more valuable in the early phase of infection. Firstly, qRT-PCR results are generally available within a few hours, and the testing is easy to perform on a large scale, and with low cost per sample. However, high false-negative rates of SARS-CoV-2 detection have been reported in China (41%).
Common qRT-PCR amplification fragments and loci of SARS-CoV-2 are shown in the prior Fig. 1. Different countries have selected different targets and designed different primers for qRT-PCR assays. The available primer and probe sequences designed by different countries are summarized in Table 1 below, including COVID-19 infection confirmatory tests for different qRT-PCR assays.
Viral genome sequencing
According to the seventh edition of Diagnosis & Treatment Scheme for Coronavirus Disease 2019, a COVID-19 diagnosis can also be confirmed by detection of a partial or whole genome sequence of the virus, which is highly homologous with known SARS-CoV-2 strains. This is especially valuable in cases when only one SARS-CoV-2 gene target is detected for the known βCoVs by qRT-PCR. For example, Wang et al. have developed a nanopore target sequencing (NTS) method targeting 11 viral regions that is able to detect as few as 10 viral copies/mL within one hour of sequencing. In addition, next-generation sequencing (NGS) also played an important role in studying the origin of SARS-CoV-2 and was very valuable in the early stages of the COVID-19 outbreak in China. Based on phylogenetic analysis, SARS-CoV-2 is closely related (with 88% sequence identity) to bat-SL-CoVZC45 and bat-SL-CoVZXC21, and most closely related (with 96.3% of sequence similarity) to bat-CoV RaTG13, all detected in bats. However, it is not very closely related to SARS-CoV and MERS-CoV, with about 79% and 50% sequence similarity, respectively.
Molecular sequencing is also used to study the evolution of SARS-CoV-2 and monitor the virus' variability. For example, in China's Guangdong province, 53 genomes from COVID-19-confirmed cases were generated by using both meta-genomic sequencing and multiplex PCR amplification, followed by nanopore sequencing, to study the genetic diversity, evolution, and epidemiology of SARS-CoV-2 in China. The 53 genome sequences from Guangdong province, and some viral genome sequences from other cities in China and other countries, were scattered throughout the phylogenetic tree, suggesting that most of the 53 cases were imported from different regions rather than locally transmitted. Therefore, molecular sequencing can help investigators identify a native or imported species in order to evaluate if the large-scale surveillance and intervention measures implemented are effective.
Although NGS is used mostly for identification of new viral species, and understanding the impact of genetic variability to viral evolution, it can also be used to detect SARS-CoV-2 in samples with low viral load. Notably, studying the evolution and transmission patterns of SARS-CoV-2 after it emerges in a new population is crucial for implementing effective measures in infection control and prevention. However, NGS is currently impractical for routine use in diagnosing COVID-19 infection due to some limitations. The high cost and long testing cycles for NGS means that it is not suitable for clinical routines and thus is not available in most clinical labs. Besides, all sequence-based methods are susceptible to nucleotide substitution, which can affect the oligonucleotide hybridization efficiency and result in false-negative results.
Loop-mediated isothermal amplification (LAMP)
Loop-mediated isothermal amplification (LAMP) was developed as a rapid, accurate, and cheaper molecular technique to amplify the target sequence at a single reaction temperature instead of the sophisticated thermal cycling equipment required in qRT-PCR testing. The LAMP method has some advantages that make it useful for point-of-care (POC) testing. First, the amount of viral nucleic acid produced is much higher than in the qRT-PCR assay, and a negative or positive result can be visually differentiated by using a colorimetric change without requiring a machine to read the results. In addition, LAMP results are available in one hour, and there is no requirement for expensive reagents or specialized equipment, making it useful for POC diagnosis in remote clinical facilities without sufficient laboratory capacity. Moreover, some non-peer-reviewed studies have demonstrated that the LAMP assay has higher sensitivity and specificity compared to qRT-PCR assays as it utilizes six primers to identify multiple regions on the target in a single reaction. In Saudi Arabia, Kashir and Yaqinuddin demonstrated the effectiveness of LAMP in the detection of SARS-CoV-2 in samples with very low viral load. Additionally, cross-reactivity of RT-LAMP assays with other human coronaviruses was not demonstrated in a Korean research study. However, LAMP assays also have some limitations. Kashir and Yaqinuddin indicated that the complex primer design system of the LAMP assay may limit the choice of target sites and resolution or specificity. Another downside is that unlike the qRT-PCR technique, the LAMP technique is still in the developmental stage, so there is a lack of relevant literature on performance evaluation.
Clustered regularly interspaced short palindromic repeats (CRISPR-Cas)
Clustered regularly interspaced short palindromic repeats (CRISPR-Cas)-based nucleic acid detection technology can be used for site-specific modifications and gene editing in microorganisms. A research group from China developed the CRISPR/Cas13 system, using two guide RNAs (gRNAs) to identify the S and ORF1ab genes of the SARS-CoV-2 genome. If SARS-CoV-2 is present in the sample, each of the two gRNAs will recognize its associated S and ORF1ab gene, and then guide Cas13 to cleave the two targets. Finally, bands from the cleaved SARS-CoV-2 RNA can be visualized. If the visualized bands are available, it means the presence of specific targets in the sample, thus achieving the purpose of detecting SARS-CoV-2. This method has been shown to consistently detect SARS-CoV-2 RNA of between 10 and 100 copies per μL of input, and as Hou et al. have demonstrated, can be completed within 40 minutes by visually reading the detection result from a lateral flow dipstick. In non-peer-reviewed research, Hou et al. have also evaluated the diagnostic performance of "CRISPR-nCoV" for SARS-CoV-2 RNA detection and reported a 100% sensitivity in 52 samples. Given the rapidity, simplicity, and higher sensitivity and specificity of CRISPR-nCoV compared to PCR-based methods, the prospect of CRISPR/Cas-based SARS-CoV-2 detection looks very promising. However, this technique is still in the exploratory and research stage and needs to be further evaluated by more tests.
Nucleic acid mass spectrometry (MS)
A powerful new method for rapid identification of emerging diseases has been recently described, based on polymerase chain reaction (PCR) to amplify nucleic acid targets from large groupings of organisms using electrospray ionization mass spectrometry (ESI-MS) for precise mass measurements of PCR products and characterization of base composition to identify organisms in a sample. During the last decade, MS has successfully been used for molecular diagnosis of viral infections. Sampath et al. demonstrated that this method could identify and differentiate between SARS and other known CoVs, including the human CoV 229E and OC43. The method has the high-throughput capabilities of automated analysis of more than 1,500 PCR reactions per day, with a detection sensitivity of 1 PFU/mL. This makes it useful in the surveillance of viral infections, and boosts rapid identification of known or emerging pathogens. Darui Biotech Company in China has developed a nucleic acid MS method with a capacity of simultaneously detecting more than 20 pathogens (including SARS-COV-2), but it requires professionally trained personnel to perform the method.
Analysis of challenging cases inconsistent with clinical testing
Despite the significant increase in the number of laboratory-confirmed cases, and the identification of common clinical characteristics in the diagnosis of COVID-19, some rather odd or difficult cases have been reported in China and elsewhere, with inconsistent clinical laboratory testing results and/or clinical symptoms. These problematic or odd cases mainly have involved some asymptomatic or clinically mild cases, with no typical COVID-19 radiological indications or defined clinical symptoms, but with positive detection of SARS-CoV-2 RNA. Conversely, some suspect cases, with the typical viral pneumonia radiological features of COVID-19 but with negative detection of SARS-CoV-2 RNA, were also reported in China.
Some studies have reported that asymptomatic cases are common in younger and middle-aged populations without underlying diseases. Additionally, some studies have shown that a large number of those asymptomatic cases involved medical staff. Thus qRT-PCR testing plays a crucial role in high-risk population screening, close contact tracing, and longitudinal surveillance for better controlling and reducing the effects of this epidemic.
On the contrary, there have also been some odd cases in which qRT-PCR detection for COVID-19 is negative, but with highly suspicious clinical symptoms and radiologic findings consistent with the disease. Although, detection of viral nucleic acid is regarded as the gold standard for virus infection confirmation, a negative result cannot exclude the possibility of COVID-19 due to possible false-negative results. To date, many cases of suspected false-negative detection of SARS-CoV-2 RNA have been reported in several hospitals both in China and elsewhere. These false-negative cases present challenges for prevention and control of the COVID-19 pandemic, especially when the test result plays a crucial role in determining whether the patient receives continual medical care and isolation, or is discharged. Given the high infectious potential of COVID-19, it would be ideal to treat these false-negative cases as positive, but due to limited space in hospitals, this might present another challenge.
Causes of false-negative molecular diagnosis of COVID-19
Many possible causes for false-negative COVID-19 results have been proposed.
First, the level of virus shedding differs in different parts of the body as the infection progresses. As such, low viral load levels in different samples and time periods of illness could result in false-negative detection of SARS-CoV-2 RNA, especially for discharged patients. SARS-CoV-2 RNA has been detected in oral cavity-associated specimens during early infection, and in anal swabs during late infection. In a study by Wu et al. involving 74 patients, viral shedding in the throat (throat swabs) was detected at a mean of 16.7 days, in comparison to a later appearance of viral RNA in fecal samples with a prolonged viral clearance for a mean of 27.9 days. Wang et al. also found a longer duration of viral shedding in throat/nasal swabs for over 72 days after onset of illness. In addition, a study from Germany showed that shedding of viral RNA in the sputum could outlast the end of symptoms (over three weeks) in six of nine patients. As for nasopharyngeal swabs, a study showed that in about 53% of cases, viral clearance was achieved 21 days after onset of symptoms. In short, if the sampling time is out of sync with the viral shedding dynamics at different anatomic sites, or the viral load is below the qRT-PCR detectable limit during the viral shedding, this will increase the possibility of false-negative results of qRT-PCR tests in the samples.
Fig. 2 shows a general relationship between viral load kinetics of SARS-CoV-2 from the upper respiratory tract (URT) and the course of COVID-19 infection. He et al. suggested that viral shedding might begin two to three days in the URT before onset of symptoms (Fig. 2). Then viral load (in throat swabs) peaks during the first week of illness and gradually decreases in the second week (Fig. 2), with researchers suspecting that infectiousness peaks on or before symptom onset, as per data obtained from 23 patients. However, a research study in Germany indicated that viral shedding in pharyngeal swabs reached a peak in the first week of symptomatic presentation. Feng et al. reported on a case from China with fever and patchy ground-glass opacity on chest CT on admission, but with four negative sequential qRT-PCR results on the pharyngeal swabs. It was not until the fifth day of admission that the fifth qRT-PCR test was positive. This case indicates that the first four negative qRT-PCR testing results were possibly false-negatives. One possible reason is that although the virus had already started shedding in the patient's pharyngeal site before or after admission, it was not detected until the fifth day due to the low viral load below the detectable limit of the qRT-PCR assay. In Korea, a similar case was reported in a patient with a fever who had SARS-CoV-2 detected from a mixed specimen of nasopharyngeal and oropharyngeal swabs on the second day of symptom onset. However, the viral load started to decline from the seventh day, and viral RNA was undetectable by qRT-PCR for two successive days from day 15 in spite of the ongoing infection, suggesting that viral load kinetics, sampling time, and duration of the illness can have an influence on qRT-PCR results.
Multiple COVID-19 cases, which were SARS-CoV-2-positive by qRT-PCR assays in the respiratory tract swabs after patients had been discharged from hospital, have become highly controversial in China. Zhou et al. reported a case where the patient met the criteria for hospital discharge but tested positive for SARS-CoV2 again 10 days after discharge. Thus a longer observation period should be considered for discharged patients.
On the other hand, some patients tested positive for SARS-CoV-2 RNA in their fecal samples for nearly five weeks after hospital discharge, but with consecutive respiratory samples being negative, possibly due to extended duration of viral shedding in faeces. A study by Wu et al. reported on two cases with detection of viral RNA in the fecal samples for 33 continuous days after testing negative in respiratory tract samples, and with positive SARS-CoV-2 RNA in their fecal samples for 47 days after first onset of symptoms. Notably, live virus isolation from fecal samples has rarely been successful in mild cases, mainly due to low viral load. Therefore, despite the presence of SARS-CoV-2 RNA in the fecal samples, further research is needed to determine the infectivity potential of these patients. In summary, it is suggested that follow-up testing be done on discharged patients with prolonged viral shedding, using fresh fecal samples at specific time points, and to extend the follow-up period for discharged patients through testing of respiratory tract swabs, to minimize potential transmission of COVID-19. Additionally, collectings samples from multiple sites at different time points can minimize the incidence of false-negative detection of SARS-CoV-2 RNA due to viral shedding dynamics.
Second, the quality of samples at different phases of infection also plays a role in the detection of SARS-CoV-2 nucleic acid, and hence in the incidence of false negative detection of COVID-19. For example, two highly suspected cases were reported in China where there was no viral RNA detected in the URT specimens, but results were positive in bronchoalveolar lavage fluid (BALF). Furthermore, a patient from Switzerland was reported to have had a two-day history of dyspnea and a six-day history of fever (39 °C) with suspect chest imaging features, but with two false-negative results of nasopharyngeal and oral swabs by qRT-PCR assays. The patient was finally confirmed COVID-19-positive by SARS-CoV-2 RNA detection in a BALF sample. In Thailand, a patient with persistent fever tested continually negative for SARS-CoV-2 RNA in nasopharyngeal and oropharyngeal samples up to the fifth day. On the eighth day, a BALF sample tested positive for SARS-CoV-2 RNA by the qRT-PCR assay.
It was unclear why these patients’ URT specimens tested consecutively negative for SARS-CoV-2 RNA. Some possible causes include improper collection or handling of specimens, and low viral load due to diminished viral shedding in URT specimens. Another possible explanation is the relatively lower sensitivity of nasopharyngeal and oral swab qRT-PCR assays for SARS-CoV-2 RNA, ranging from 56% to 83%, in comparison to lower respiratory tract (LRT) specimens. Although BALF specimens increase the detection rates of COVID-19, their collection requires a suction device and a skilled operator, and is also painful for patients, so they are not convenient for routine laboratory diagnosis of SARS-CoV-2 RNA.
Yang et al. revealed that save for BALF, sputum was the best specimen for laboratory diagnosis of COVID-19, followed by nasal swabs—which were most recommended—with detection rates ranging from 74.4% to 88.9% and 53.6% to 73.3%, respectively, for both severe and mild cases during the first 14 days after onset of illness. However, not all COVID-19 patients always present a dry cough, with only 28% able to produce sputum for diagnostic evaluation. In most studies of respiratory virus infections, nasopharyngeal or throat swabs are normally used for viral load monitoring. However, the collection of nasopharyngeal swabs is an invasive procedure; it is uncomfortable for the patient and poses a risk of transmission of the virus to the healthcare workers from coughing and sneezing. Previous studies have also demonstrated a relatively low SARS-CoV-2 RNA detection rate in throat swabs (collected ≥ eight days), especially in samples from mild cases, and thus throat swabs are not recommended to limit the incidence of false-negative results. Compared with nasopharyngeal swabs, saliva is much more acceptable to patients and is safer for healthcare workers to collect. A previous study has shown that saliva has a high and consistent coronavirus detection rate of >90% with nasopharyngeal specimens. Hence, if the clinical, laboratory, and radiological features are highly suspicious for COVID-19, but with negative qRT-PCR tests on URT specimens, performing qRT-PCR assays on LRT specimens might improve the detection rate of SARS-CoV-2 in specimens such as sputum and BALF. Thus for challenging COVID-19 cases, different types of samples are recommended from a patient for combination testing to reduce the incidence of false-negative results.
Thirdly, false-negative detection of SARS-CoV-2 by qRT-PCR is possibly associated with difficulty in detecting residual virus resident in pulmonary tissues. A patient reported by Yao et al. was initially confirmed as SARS-CoV-2 positive by qRT-PCR testing on nasopharyngeal swabs. Later on, it was demonstrated histopathologically that residual SARS-CoV-2 virus was present in pulmonary tissues, but with three consecutive negative results by qRT-PCR of nasopharyngeal swabs in the following days. Unfortunately, the patient died in the end. This case raised the possibility that non-detection of SARS-CoV-2 in the nasopharyngeal swabs might not be fully indicative of the virus status in lung tissue. Thus detection of SARS-CoV-2 RNA in BALF and extension of quarantine or hospital discharge periods are recommended, especially for elderly patients with underlying diseases.
Fourth, co-infection with other viruses may have an impact on qRT-PCR detection accuracy. Influenza A virus was one of the most common viral pathogens causing co-infection among patients with SARS-CoV-2 infection in China. Lai et al. reported on two COVID-19 cases co-infected with influenza A virus but which yielded false-negative results for SARS-CoV-2. Zhao et al. reported on a COVID-19 patient with HIV-1 and HCV coinfection, who showed continuously negative SARS-CoV-2 RNA tests by qRT-PCR but with a delayed antibody response against SARS-CoV-2 in the plasma. Therefore, co-detection of SARS-CoV-2 with another virus present creates additional challenges in the diagnosis of COVID-19. Further research is needed to verify the influence of other viral infections on SARS-CoV-2 detection in viral co-infected patients.
Kit sensitivity and extraction methodology
Fifth, false-negative results are possibly associated with in vitro viral nucleic acid diagnostic kits with unstable sensitivity, and some methods of RNA extraction. Many countries have designed different SARS-CoV-2 diagnostic kits with different targets and primers for qRT-PCR assays (summarized in the previous Table 1). Although it is commonly accepted, as per data from many studies, that E-gene based qRT-PCR assays have a higher diagnostic sensitivity than other targets, the specificity of the RdRp and N genes have been shown to be higher. Actually, during the early stages of COVID-19 outbreak in China, there were a series of false-negative detection of SARS-CoV-2 RNA in some samples caused by some poor sensitivity of diagnostic kits developed in an emergency (no data available). However, the sensitivity of currently available diagnostic kits from different manufacturers has improved significantly. In the last few months, very few reported false-negative cases were related to low or unstable sensitivity of the kits.
For highly suspected or already confirmed cases, if only one target is used for COVID-19 confirmation or follow-up diagnosis, it is important to improve the accuracy rate of qRT-PCR tests by comparing with different diagnostic kits. Some researchers from Beijing Centre for Disease Prevention and Control found that thermal inactivation might reduce the detectable amount of SARS-CoV-2 in qRT-PCR assays, thereby resulting in false-negative results; this is particularly common in the early phase of infection with low viral load in samples. Although thermal treatment of samples before RNA extraction is not recommended by WHO, thermal inactivation of samples under 56 °C for 30 minutes is required to ensure biosafety for laboratory personnel based on Chinese guidelines.
Causes from other countries
Sixth, some other causes of false-negative qRT-PCR results have been analyzed in other countries. Tahamtan and Ardebili from Iran indicated that mutations in the primer and probe target regions in the SARS-CoV-2 genome could result in false-negative qRT-PCR results. They indicated that it was possibly caused by genetic variability of SARS-CoV-2 resulting in mismatches among the primers, probes, and the target sequences. In fact, since the first SARS-CoV-2 genomic sequence became available, several studies have reported on a rapid genetic evolution of SARS-CoV-2 through a phylogenetic tree analysis. Both natural mutation and active viral recombination are able to weaken the efficiency of oligonucleotide annealing, declining the sensitivity and specificity of qRT-PCR detection. In order to avoid false-negative results due to unknown mutation, continuous monitoring of genetic variability is necessary, and targeting multiple regions in the viral genome is crucial to SARS-CoV-2 detection.
Sample collection and storage
Last but not least, proper management of sample collection and storage is essential for reducing the incidence of false negative qRT-PCR detection of COVID-19. For example, if samples are collected too early or too late during an infection, this may have an effect on viral load. Furthermore, improper storage and/or transportation of specimens can result in RNA degradation, leading to a false-negative result. Additionally, whether a standardized clinical laboratory is adequately equipped and has well trained laboratory personnel for virus detection is also an important factor. Therefore, strengthening the professional training of laboratory operators and improving the laboratory's quality management system can also reduce the incidence of false-negative results.
Supplementary serological testing
To resolve the limitations of qRT-PCR testing and difficult COVID-19 suspected cases, serological testing (IgM/IgG antibody detection) is suggested as a complementary identification assay. The clinical significance of false-negative qRT-PCR results (related to the course of infection) combined with serological testing is summarized in Table 2. Specific IgM and IgG antibodies can be used in determining whether the patient has a recent or previous viral infection, and also in quantification of SARS-CoV-2-positive cases, including asymptomatic and recovered cases. For example, SARS-CoV-2 antibodies were detected in 10.1% (28/276) of asymptomatic medical staff at one hospital in China, and five of them were in close contact with confirmed COVID-19 patients, but they were qRT-PCR negative. Another study detected IgM and IgG antibodies in 84.21% and 94.74% of 19 patients with negative SARS-CoV-2 detection by qRT-PCR assays but with typical clinical symptoms, respectively. This strongly suggests that serological testing can significantly reduce the risk of misdiagnosis and play a crucial role in timely diagnosis, treatment, and prevention of COVID-19.
However, serological testing also has some limitations, mainly the slow antibody response to SARS-CoV-2 virus means that they cannot be helpful in the early stages of infection. Seroconversion is usually detectable between five and seven days and 14 days after onset of symptoms. Based on findings from the U.S. Food and Drug Administration, the non-specific IgM antibodies to SARS-CoV-2 are detectable just a few days after initial infection, but IgM levels throughout the course of infection can rapidly decline and finally become undetectable. However, IgG antibodies remain detectable for a longer period, or even when SARS-CoV-2 RNA is undetectable, as described in the prior Fig. 2. In China, Zhang et al. reported on 15 patients with relatively low or undetectable IgM and IgG titers on day 0 (the day of first sampling). However, increasing antibody titers were demonstrated on the patients on the fifth day, and this was interpreted as a transition from the early to the later phase of viral infection, with dynamic changes of viral presence. On the contrary, there was a relatively low positive detection rate by qRT-PCR assays during the same period. Thus, serological testing alone cannot confirm or exclude COVID-19 infection. For example, a negative result cannot rule out the infection because the patient may not be infected at the sampling time, as the individual may be in the "window period" (delay in the production of antibodies), especially for those who have a history of close contact with confirmed cases. Moreover, false-positive detection of IgM and IgG antibodies have been described, mainly associated with cut-off values of the kit. A weak positive result near the cut-off value is likely to be a false positive.
Another reason is that some existing interfering substances in plasma samples (including interferon, rheumatoid factors [RF] and non-specific antibodies) can lead to false-positive results. Jia et al. demonstrated differing detection results of IgM/IgG antibodies in serum samples with different RF concentrations. In a total of nine serum samples with different RF concentrations, detection of IgM-specific antibodies was observed at an RF concentration of >331 IU/mL, and both IgM and IgG test results were positive in samples with an RF concentration of 981.2 IU/mL. Additionally, potential cross-reactivity of SARS-COV-2 antibodies with antibodies generated by other coronaviruses probably also results in false-positive results. For example, Lv et al. found a high frequency of cross-reactivity between the S protein of SARS-CoV-2 and SARS-CoV among plasma samples from 15 COVID-19 patients.
In short, although serological testing alone is not enough to confirm COVID-19 infection, combining both serological testing and molecular techniques can improve the identification rate of COVID-19. Serological testing is valuable in evaluating the overall immune response in large scale population surveillance.
In summary, to resolve the COVID-19 challenging cases, more comprehensive analysis and/or further evaluation of different diagnostic methods is needed. Improving the identification rates of SARS-CoV-2, including reducing the incidence of false-negative/false-positive results, still remains a considerable challenge in the laboratory diagnosis of COVID-19 in China, requiring further research. At present, vigilance is still required in China, as there remains a risk that SARS-CoV-2 transmission may reignite, with an increasing number of COVID-19 imported cases being reported.
The authors of this paper contributed equally to this work.
This work was supported by Chinese Academy of Medical Sciences Innovation Fund for Medical Sciences (Grant No. 2016-I2M-1-014) and Beijing Nova Program (Z201100006820127).
Declaration of competing interest
The authors declare that they have no conflicts of interest.
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- ↑ 9.0 9.1 Corman, V.M.; Landt, O.; Kaiser, M. et al. (2020). "Detection of 2019 novel coronavirus (2019-nCoV) by real-time RT-PCR". Euro Surveillance 25 (3): 2000045. doi:10.2807/1560-7917.ES.2020.25.3.2000045. PMC PMC6988269. PMID 31992387. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC6988269.
- ↑ 10.0 10.1 World Health Organization (24 January 2020). "Molecular assays to diagnose COVID-19: Summary table of available protocols". World Health Organization. https://www.who.int/publications/m/item/molecular-assays-to-diagnose-covid-19-summary-table-of-available-protocols.
- ↑ U.S. Department of Health and Human Services (29 May 2020). "2019-Novel Coronavirus (2019-nCoV) Real-time rRT-PCR Panel Primers and Probes" (PDF). Centers for Disease Control and Prevention. https://www.cdc.gov/coronavirus/2019-ncov/downloads/rt-pcr-panel-primer-probes.pdf.
- ↑ University of Hong Kong, School of Public Health (2020). "Detection of 2019 novel coronavirus (2019-nCoV) in suspected human cases by RT-PCR" (PDF). World Health Organization. https://www.who.int/docs/default-source/coronaviruse/peiris-protocol-16-1-20.pdf.
- ↑ National Health Commission of the People’s Republic of China (4 March 2020). "Diagnosis & Treatment Scheme for Coronavirus Disease 2019, 7th Edition". http://www.nhc.gov.cn/yzygj/s7653p/202003/46c9294a7dfe4cef80dc7f5912eb1989.shtml.
- ↑ Wang, M.; Fu, A.; Hu, B. et al. (2020). "Nanopore Targeted Sequencing for the Accurate and Comprehensive Detection of SARS-CoV-2 and Other Respiratory Viruses". Small 16 (32): e2002169. doi:10.1002/smll.202002169. PMC PMC7361204. PMID 32578378. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7361204.
- ↑ 15.0 15.1 15.2 15.3 15.4 Lu, J.; du Plessis, L.; Liu, Z. et al. (2020). "Genomic Epidemiology of SARS-CoV-2 in Guangdong Province, China". Cell 181 (5): 997-1003.e9. doi:10.1016/j.cell.2020.04.023. PMC PMC7192124. PMID 32359424. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7192124.
- ↑ Paraskevis, D.; Kostaki, E.G.; Magiorkinis, G. et al. (2020). "Full-genome evolutionary analysis of the novel corona virus (2019-nCoV) rejects the hypothesis of emergence as a result of a recent recombination event". Infection, Genetics and Evolution 79: 104212. doi:10.1016/j.meegid.2020.104212. PMC PMC7106301. PMID 32004758. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7106301.
- ↑ 17.0 17.1 Álvarez-Díaz, D.A.; Franco-Muñoz, C.; Laiton-Donato, K. et al. (2020). "Molecular analysis of several in-house rRT-PCR protocols for SARS-CoV-2 detection in the context of genetic variability of the virus in Colombia". Infection, Genetics and Evolution 84: 104390. doi:10.1016/j.meegid.2020.104390. PMC PMC7272177. PMID 32505692. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7272177.
- ↑ Wang, X.; Fang, J.; Zhu, Y. et al. (2020). "Clinical characteristics of non-critically ill patients with novel coronavirus infection (COVID-19) in a Fangcang Hospital". Clinical Microbiology and Infection 26 (8): 1063-1068. doi:10.1016/j.cmi.2020.03.032. PMC PMC7195539. PMID 32251842. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7195539.
- ↑ Park, G.-S.; Ku, K.; Baek, S.-H. et al. (2020). "Development of Reverse Transcription Loop-Mediated Isothermal Amplification Assays Targeting Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2)". Journal of Molecular Diagnostics 22 (6): 729-735. doi:10.1016/j.jmoldx.2020.03.006. PMC PMC7144851. PMID 32276051. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7144851.
- ↑ 20.0 20.1 Kashir, J.; Yaqinuddin, A. (2020). "Loop mediated isothermal amplification (LAMP) assays as a rapid diagnostic for COVID-19". Medical Hypotheses 141: 109786. doi:10.1016/j.mehy.2020.109786. PMC PMC7182526. PMID 32361529. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7182526.
- ↑ Yang, W.; Dang, X.; Wang, Q. et al. (2020). "Rapid Detection of SARS-CoV-2 Using Reverse transcription RT-LAMP method". medRxiv. doi:10.1101/2020.03.02.20030130.
- ↑ Zhang, Y.; Odiwuor, N.; Xiong, J. et al. (2020). "Rapid Molecular Detection of SARS-CoV-2 (COVID-19) Virus RNA Using Colorimetric LAMP". medRxiv. doi:10.1101/2020.02.26.20028373.
- ↑ 23.0 23.1 23.2 23.3 23.4 Qiu, F.; Wang, H.; Zhang, Z. et al. (2020). "Laboratory testing techniques for SARS-CoV-2". Journal of Southern Medical University 40 (2): 164–7. doi:10.12122/j.issn.1673-4254.2020.02.04. PMC PMC7086127. PMID 32376546. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7086127.
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- ↑ 29.0 29.1 Gao, Z.; Xu, Y.; Sun, C. et al. (2020). "A Systematic Review of Asymptomatic Infections with COVID-19". Journal of Microbiology, Immunology, and Infection In Press. doi:10.1016/j.jmii.2020.05.001. PMC PMC7227597. PMID 32425996. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7227597.
- ↑ 30.0 30.1 Lai, C.-C.; Liu, Y.H.; Wang, C.-Y. et al. (2020). "Asymptomatic carrier state, acute respiratory disease, and pneumonia due to severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2): Facts and myths". Journal of Microbiology, Immunology, and Infection 53 (3): 404–12. doi:10.1016/j.jmii.2020.02.012. PMC PMC7128959. PMID 32173241. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7128959.
- ↑ 31.0 31.1 Li, Q.; Guan, X.; Wu, P. et al. (2020). "Early Transmission Dynamics in Wuhan, China, of Novel Coronavirus-Infected Pneumonia". New England Journal of Medicine 382 (13): 1199–1207. doi:10.1056/NEJMoa2001316. PMC PMC7121484. PMID 31995857. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7121484.
- ↑ 32.0 32.1 32.2 Zhang, W.; Du, R.-H.; Li, B. et al. (2020). "Molecular and serological investigation of 2019-nCoV infected patients: Implication of multiple shedding routes". Emergining Microbes and Infections 9 (1): 386-389. doi:10.1080/22221751.2020.1729071. PMC PMC7048229. PMID 32065057. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7048229.
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- ↑ Yong, S.E.F.; Anderson. D.E.; Weu. W.E. et al. (2020). "Connecting clusters of COVID-19: An epidemiological and serological investigation". The Lancet Infectious Diseases 20 (7): 809–15. doi:10.1016/S1473-3099(20)30273-5. PMC PMC7173813. PMID 32330439. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7173813.
- ↑ 37.0 37.1 He, X.; Lau, E.H.Y.; Wu, P. et al. (2020). "Temporal dynamics in viral shedding and transmissibility of COVID-19". Nature Medicine 26 (5): 672-675. doi:10.1038/s41591-020-0869-5. PMID 32296168.
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- ↑ 39.0 39.1 Kim, J.Y.; Ko, J.H.; Kim, Y. et al. (2020). "Viral Load Kinetics of SARS-CoV-2 Infection in First Two Patients in Korea". Journal of Korean Medical Science 35 (7): e86. doi:10.3346/jkms.2020.35.e86. PMC PMC7036338. PMID 32080991. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7036338.
- ↑ 40.0 40.1 40.2 40.3 Yao, X.-H.; He, Z.-C.; Li, T.-Y. et al. (2020). "Pathological evidence for residual SARS-CoV-2 in pulmonary tissues of a ready-for-discharge patient". Cell Research 30 (6): 541-543. doi:10.1038/s41422-020-0318-5. PMC PMC7186763. PMID 32346074. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7186763.
- ↑ Zhou, M.; Li, Q.; Cao, L. et al. (2020). "Re-emergence of SARS-CoV2 in a discharged COVID-19 case". Journal of Microbiology, Immunology, and Infection 53 (3): 501-502. doi:10.1016/j.jmii.2020.03.031. PMC PMC7194673. PMID 32303482. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7194673.
- ↑ 42.0 42.1 42.2 42.3 Yang, Y.; Yang, M.; Shen, C. et al. (2020). "Evaluating the accuracy of different respiratory specimens in the laboratory diagnosis and monitoring the viral shedding of 2019-nCoV infections". medRxiv. doi:10.1101/2020.02.11.20021493.
- ↑ 43.0 43.1 43.2 Marando, M.; Tamburello, A.; Gianella, P. (2020). "False-Negative Nasopharyngeal Swab RT-PCR Assays in Typical COVID-19: Role of Ultra-low-dose Chest CT and Bronchoscopy in Diagnosis". European Journal of Case Reports in Internal Medicine 7 (7): 001680. doi:10.12890/2020_001680. PMC PMC7357997. PMID 32670990. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7357997.
- ↑ Winichakoon, P.; Chaiwarith, R.; Liwsrisakun, C. et al. (2020). "Negative Nasopharyngeal and Oropharyngeal Swabs Do Not Rule Out COVID-19". Journal of Clinical Microbiology 58 (5): e00297-20. doi:10.1128/JCM.00297-20. PMC PMC7180262. PMID 32102856. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7180262.
- ↑ 45.0 45.1 45.2 To, K.K.-W.; Tsang, O.T.-Y.; Yip, C.C.-Y. et al. (2020). "Consistent Detection of 2019 Novel Coronavirus in Saliva". Clinical Infectious Diseases 71 (15): 841-843. doi:10.1093/cid/ciaa149. PMC PMC7108139. PMID 32047895. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7108139.
- ↑ Chen, F.; Liu, Z.S.; Zhang, F.R. et al. (2020). "First case of severe childhood novel coronavirus pneumonia in China". Chinese Journal of Pediatrics 58 (0): E005. doi:10.3760/cma.j.issn.0578-1310.2020.0005. PMID 32045966.
- ↑ 47.0 47.1 Lai, C.-C.; Wang, C.-Y.; Hsueh, P.-R. (2020). "Co-infections among patients with COVID-19: The need for combination therapy with non-anti-SARS-CoV-2 agents?". Journal of Microbiology, Immunology, and Infection 53 (4): 505–12. doi:10.1016/j.jmii.2020.05.013. PMC PMC7245213. PMID 32482366. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7245213.
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- ↑ Pan, Y.; Long, L.; Zhang, D. et al. (2020). "Potential False-Negative Nucleic Acid Testing Results for Severe Acute Respiratory Syndrome Coronavirus 2 from Thermal Inactivation of Samples with Low Viral Loads". Clinical Chemistry 66 (6): 794–801. doi:10.1093/clinchem/hvaa091. PMC PMC7184485. PMID 32246822. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7184485.
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- ↑ 52.0 52.1 Peñarrubia, L.; Ruiz, M.; Porco, R. et al. (2020). "Multiple assays in a real-time RT-PCR SARS-CoV-2 panel can mitigate the risk of loss of sensitivity by new genomic variants during the COVID-19 outbreak". International Journal of Infectious Diseases 97: 225-229. doi:10.1016/j.ijid.2020.06.027. PMC PMC7289722. PMID 32535302. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7289722.
- ↑ Zhao, J.; Yuan, Q.; Wang, H. et al. (2020). "Antibody responses to SARS-CoV-2 in patients of novel coronavirus disease 2019". Clinical Infectious Diseases In Press: ciaa344. doi:10.1093/cid/ciaa344. PMC PMC7184337. PMID 32221519. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7184337.
- ↑ Zhao, R.; Li, M.; Song, H. et al. (2020). "Early detection of SARS-CoV-2 antibodies in COVID-19 patients as a serologic marker of infection". Clinical Infectious Diseases In Press: ciaa523. doi:10.1093/cid/ciaa523. PMC PMC7197602. PMID 32357209. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7197602.
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- ↑ Lauer, S.A.; Grantz, K.H.; Bi, Q. et al. (2020). "The Incubation Period of Coronavirus Disease 2019 (COVID-19) From Publicly Reported Confirmed Cases: Estimation and Application". Annals of Internal Medicine 172 (9): 577-582. doi:10.7326/M20-0504. PMC PMC7081172. PMID 32150748. http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pmcentrez&artid=PMC7081172.
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