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SARS-CoV-2 Detection Methods

Biosystems and Integrative Sciences Institute (BioISI), Faculty of Sciences, University of Lisboa, 1649-004 Lisbon, Portugal
REQUIMTE, Instituto Superior de Engenharia do Porto, 4200-072 Porto, Portugal
DNA & RNA Sensing Lab, University of Trás-os-Montes e Alto Douro (UTAD), 5000-901 Vila Real, Portugal
Authors to whom correspondence should be addressed.
Chemosensors 2022, 10(6), 221;
Received: 6 May 2022 / Revised: 1 June 2022 / Accepted: 9 June 2022 / Published: 11 June 2022
(This article belongs to the Special Issue State of the Art in Nucleic Acid Detection)


A fast and highly specific detection of COVID-19 infections is essential in managing the virus dissemination networks. The most relevant technologies developed for SARS-CoV-2 detection, along with their advantages and limitations, will be presented and fully explored. Additionally, some of the newest and emerging COVID-19 diagnosis tools, such as biosensing platforms, will also be introduced. Considering the extreme relevance that all these technologies assume in pandemic control, it is of the utmost relevance to have an intrinsic knowledge of the parameters that need to be taken into consideration before choosing the most adequate test for a particular situation. Moreover, the new variants of the virus and their potential impact on the detection method’s effectiveness will be discussed. In order to better manage the pandemic, it is essential to maintain continuous research into the SARS-CoV-2 genome and updated genomic surveillance at the global level. This will allow for timely detection of new mutations and viral variants, which may affect the performance of COVID-19 detection tests.

1. Introduction

The SARS-CoV-2 virus is responsible for the pulmonary disease COVID-19. It was first reported in December 2019, in Wuhan, China, but quickly spread around the world, being classified as a pandemic in March 2020. The management of this pandemic relies mostly on vaccination and preventive measures such as social distancing and the use of masks. Even though the number of new infections varies periodically, and we have seen the existence of several “waves” with an average separation of 4 months, it is still of extreme importance to maintain high levels of testing at all times. Due to the fact that COVID-19 is highly contagious and presents non-specific symptomatology, easily mistaken for the common flu, it is important to have readily available specific diagnostic tools. Currently, there are several detection methods available, but due to the worldwide need their availability was limited, especially in the first year of the pandemic. Most of these constraints were linked to the lack of equipment and reagents, defective test kits, and excessive waiting time before results, hence the emergency in developing fast, sensitive, and cost-effective tools [1]. Since it has been proven that early detection has a great impact in decreasing the number of cases, it is imperative to enhance the availability and specificity of tests, as well as diminish the time from sample collection to test result [2]. Most tests are usually performed in individuals who had close contact with positive cases and in individuals who present symptoms such as fever, fatigue, dry cough, and myalgia which is common in COVID-19 patients. However, with the ever-growing number of vaccinated people, the guidelines regarding diagnosis tests are expected to change. The last challenge regarding this pandemic is the rise of new viral variants. As is normal in RNA viruses, SARS-CoV-2 is associated with a relatively high mutational rate, which leads to the occurrence of mutations and recombination events that can lead to the emergence of viral variants associated with negative impacts, either in disease presentation, transmissibility, therapeutics, vaccination effectiveness or diagnostic test performance [3]. These variants usually have high prevalence due to their characteristics and are classified by the World Health Organization (WHO) as variants of concern (VOC) [4,5]. Until now, there have been five different VOCs: Alpha, Beta, Gamma, Delta, and Omicron [6].
Due to the great number of emerging techniques for SARS-CoV-2 detection, it is not feasible to present all the different methodologies described so far. Following, a review of some of the current employed and emerging detection techniques, and their advantages and disadvantages, will be presented. The impact of new variants in the currently available diagnostic tests will also be explored.

2. SARS-CoV-2 Detection Techniques

2.1. Clinically Useful Methods

2.1.1. Antibody Tests

These are serological tests that analyze blood or serum samples to detect specific SARS-CoV-2 antibodies. These tests will inform if one person has been infected with COVID-19 by detecting the presence of either immunoglobulin G (IgG) or immunoglobulin M (IgM) [7]. These antibodies are directed against the Spike (S) and Nucleocapside (N) proteins and have their highest levels 2 weeks after symptoms onset in the case of IgM and 3 weeks after symptom onset for IgG, the latter also providing long-term immunity. The most used techniques to detect SARS-CoV-2 antibodies are the Enzyme-linked Immunosorbent Assays (ELISA) and Lateral Flow Immunoassays (LFIA).
ELISA is an immunological laboratory-based procedure which uses plates coated with viral proteins, usually the N or S protein to detect specific antibodies (Figure 1B). After adding the biological sample, the binding of any antibodies to the viral proteins occurs. In the case of a positive sample, the presence of the antibody–protein complex will be detected by color change or fluorescence after the addition of a marked antibody. In order to obtain a more accurate result, the detection of both IgM and IgG is recommended, as well as a waiting time of at least 14 days after symptom onset [8]. This test is faster than RT-qPCR and requires minimal equipment; however, there is the risk of cross-reactivity to antibodies from other coronaviruses [9]. It also does not allow for early detection since it takes several days for the human immune system to make a detectable antibody response, hence the inconsistency of these tests in the first 15 days after infection [10]. Moreover, this diagnostic is usually based on the detection of only one protein. These limitations make these tests more prone to inaccurate results when considering the high mutation rate of the virus. Although they are indeed limited for diagnosis, these tests are useful for estimating the number of individuals who had been in contact with SARS-CoV-2, regardless of having exhibited symptoms or not [11]. Another type of immunoassay available for the detection of IgG and IgM is the Chemiluminescence Immunoassay (CLIA), which is similar to ELISA as it uses chemiluminescent labels for antibody detection [12].
LFIA also uses N and S proteins to detect a SARS-CoV-2 immune response. After being loaded into the cassette, the sample will flow through the conjugation pad by a capillary effect. In the conjugation pad there are viral antigens and control antibodies, both marked with a reporter molecule which is usually gold nanoparticles. In the case of a positive sample, the antibodies present (IgG or IgM) will bind to the marked antigens. All of the constituents will continue to flow until they reach the test zone, where immobilized anti-human IgG or IgM are placed. When present, the marked antigen–antibody (IgG/IgM) complex will bind and give rise to a colorimetric reaction. The rest of the sample, which includes the control marked antibodies, will continue to flow until the control zone and bind to the anti-control antibodies that are immobilized in that zone, also causing a colorimetric reaction. The biological samples are usually whole blood, serum, or plasma. For more accuracy, both IgG and IgM are recommended to be detected in the same LFIA test. Therefore, these kits will present a three-lined positive result: one for the control, one for IgG and another for IgM [13]. Additionally, when there is only one line in the zone test, an inference can be made in relation to the individual’s infection time. That is, assuming that the control line is positive, the appearance of only the IgM line will indicate a shorter time of infection, while only IgG will mean that the exposure to SARS-CoV-2 was long ago, since the IgM antibodies are no longer present in the bloodstream [14]. Antibody LFIAs are faster, cheaper, portable, and more user-friendly when compared to ELISA; however, the specificity and sensitivity is lower. Nevertheless, the accuracy of antibody LFIA is correlated to time of infection, so its use can be optimized according to the suspected time of infection. In the fourth week after infection, the accuracy of the test is expected to be very high, as the levels of antibodies present in the bloodstream are at their peak. Therefore, these tests are not adequate for control of the population in early stages of infection. Moreover, as with ELISA assays, it usually only tests one SARS-CoV-2 related protein per test, which is highly limiting. The ideal scenario for this type of test is, for example, large epidemiological population studies where the main objective is not to diagnose but to investigate antibody prevalence [15]. However, with the current high levels of vaccinated individuals, the use of these antibody tests is more recommended as a tool to assess immune response to vaccination since it would be difficult to differentiate a positive test due to previous COVID-19 infection from a positive result due to vaccination [16].

2.1.2. Antigen Tests

As in antibody detection, the most used techniques for antigen detection are ELISA and LFIA. When applied for antigen detection, the ELISA test will use SARS-CoV-2 specific antibodies or other affinity ligands, such as aptamers or synthetic peptides. These will recognize and bind to the viral proteins, usually S or N, if present in the sample. The use of the N protein as a target is recommended since it is highly abundant during viral replication and has low cross-reactivity with other coronaviruses (CoVs) [17]. However, these types of tests are limited by the availability of antibodies and epitope preservation [18].
Regarding LFIA, they usually detect viral antigens in a paper-like membrane with immobilized antibodies at two different sites, leading to the appearance of color zones after interaction with the virus [11]. As with the antibody LFIA, these tests have a conjugation pad, but this time it is loaded only with marked SARS-CoV-2 specific antibodies. There is also a test zone with immobilized antibodies, specific to the SARS-CoV-2 antigens that, when present in the sample, will bind after a linkage reaction with marked antibodies in the conjugation pad. A control zone with immobilized secondary antibodies is also present where the excess marked antibodies present in the conjugation pad will bind. The first line gives the test result, while the second serves as a control. These types of colorimetric bioassays rely on the use of gold nanoparticles conjugated with antibodies [19]. Usually, these tests only require a self-performed nasal swab or saliva sample, which is next mixed with a buffer (available in the kit). The technology used in LFIA allows a better manipulation of fluids, which will be reflected by a smaller sample volume and an efficient mixing of the reagents, giving a faster result read-out. These types of tests are very fast (<20 min) and inexpensive, and therefore suitable in the POC context. However, sensitivity and specificity are relatively low. There are reports of 56.2% sensitivity and 99.5% specificity, with a 27.9% percentage of false negatives [20,21]. Nonetheless, these values can vary depending on the specific LFIA used. Another limitation is the fact that these tests are only qualitative, giving a positive or negative type of diagnosis, which does not allow for quantifying patient viral load. Moreover, as in the previously described antibody tests, it also relies on the detection of only one SARS-CoV-2 protein, which can lead to false negatives.
Comparing the two assays, ELISA has a higher specificity and sensitivity. However, the waiting time is considerably higher (1–5 h), the amount of sample needed is larger, and there are several manual steps that require a laboratory setting and technicians, which limit its use as a POC test. These antigen tests are particularly recommended to confirm suspected cases of infection. In the case of a negative result, the individual is then indicated to perform a more sensitive RT-PCR test to confirm the result, since the result was inconsistent with the clinical context [22]. As these tests are prone to generating false negative results, they can give a false sense of security. This can lead to unexpected outbreaks in a given group.

2.1.3. RT-qPCR

Real-time reverse transcription quantitative polymerase chain reaction (RT-qPCR) is the most used detection method for COVID-19, it is considered the “gold diagnostic technique” due to its reliability. Firstly, a biological sample for viral RNA extraction is collect-ed. By recommendation of the U.S. Centers for Disease Control and Prevention (CDC), the samples can be from both the upper respiratory tract (URT) (nasopharyngeal or oropharyngeal swab) or the lower respiratory tract (LRT) [23]. After RNA isolation, the analysis consists of a two-step process: a reverse transcription, which converts viral RNA into cDNA, followed by the amplification of certain viral regions using specific primers and probes while the levels of amplification are measured by a fluorescence signal (Figure 1C).
Primer and probe design are fundamental due to the high mutation rates of SARS-CoV-2; therefore, a relatively conserved target region must be chosen in order to avoid false negatives. The primers and probes recommended by the WHO include the N, E, S, ORF1ab, and RdRP genes [24]. RT-PCR usually targets multiple regions in order to avoid false results due to the possibility of mutations in one region or other unforeseen errors. This is quite important. The RT-PCR result is considered the most robust, precisely because it relies on two/three targets. It also usually targets human RNase P, as an internal control, to ensure that the RNA extraction has been well performed and there were no malfunctions with the equipment/reagents [25].
The critical step in this procedure is RNA isolation since RNA molecules are very prone to degradation. This step must be handled very carefully with optimized protocols and under strict laboratory conditions in order to avoid any contaminations and minimize RNA degradation, thus avoiding false results [26]. Usually, the entire process, from RNA extraction to results, lasts from 4 to 8 h depending on the protocols used and on the number of samples and labels that the platform (Realtime PCR thermocycler) can comport. The limit of detection of RT-qPCR kits varies from 0.3 copies/μL to 100 copies/μL [27], which is one of its major advantages since it allows detection at an early stage of infection with high specificity [28]. However, there is a possibility of false negatives due to technical errors, sample collection and storage, low viral load, and other laboratory conditions. The occurrence of false positives can also happen since this method requires several steps (e.g., sample collection, transport, DNA extraction, PCR reaction), and it is possible for cross-contamination of samples to occur if all security measures are not followed. Even the smallest contamination from a positive sample can originate a false result in a negative one due to the very high sensitivity of this technique [27]. Additionally, the implementation of this diagnosis methodology requires qualified technicians, specific and expensive equipment, general optimization of the methods, and several preparation steps [29]. Therefore, and even though it is the most robust and reliable technique for SARS-CoV-2 detection, it has time, cost, and need for specialized personnel, laboratory settings, and equipment as major limitations for a wide application, particularly in third world countries. In order to maximize the accuracy of this test, it is best to perform it at least 4 days after exposure, to minimize the risk of false negatives due to low viral loads [30].
It is also relevant to highlight that this detection method also provides a quantitative result, known as cycle threshold (Ct), that allows for the inference of the original viral concentration and consequent approximate patient viral load. The Ct value is inversely proportional to the initial concentration of target DNA; therefore, if the Ct value is 40 or higher, it indicates that there was a low concentration of viral target near the assay’s limit of detection. However, the Ct values can be influenced by several variables, such as quality of the sample, quality of the extraction, and primer choice. Therefore, it cannot be used as a direct way to assume a completely realistic and accurate patient viral load [31].
As a complementary approach, aiming to reduce costs and time, the use of multiplex RT-qPCR targeting different viruses, e.g., SARS-CoV-2 and influenza, is also possible. This allows for a more efficient diagnosis, since both COVID-19 and flu mostly share symptomatology [32]. Other PCR based techniques, such as digital PCR (dPCR) [33], and technologies using microfluidic devices, such as the IFC chip and the “lab-on-a-disc”, are also being developed and offer several advantages, such as faster result turn-out, higher sensibility, and shorter processing time, often not needing to proceed with RNA extraction [34,35].
The techniques described in this section are the most commonly used and have proven clinical usefulness. However, the global impact of COVID-19 led to an exceptional common effort and investment from several organizations in order to develop better alternatives for the diagnosis of this disease in terms of sensitivity, reliability, and delivering quick results. Next, some of the developed technologies will be described, even though the majority of them are not yet fully available for the general public. Nevertheless, they still represent potentially useful diagnostic tools.

2.2. Potentially Useful Methods

2.2.1. RT-LAMP

As in RT-PCR, Reverse Transcription Loop-Mediated Isothermal Amplification (RT-LAMP) relies on the amplification of specific cDNA sequences to detect SARS-CoV-2 (Figure 1D). This technique uses four to six primers, and the reaction happens all at the same temperature; therefore, there is no need for a thermocycler or any expensive and bulky equipment. The results can be obtained through gel electrophoresis, UV-light illumination, or through colorimetric readouts based on reaction by-products [36]. The latter is the most used on-field since it allows a visual result within a few minutes. A usual colorimetric RT-LAMP reaction only requires a stable heat source, reverse-transcriptase, DNA polymerase, and a pH indicator to provide the color change. The time reaction is around 40 min from sample collection to result and has high specificity and sensitivity [37]. The samples are usually nasopharyngeal swabs or saliva samples and may not require previous RNA extraction, depending on the method used. The primers usually target regions in the N, S, E or ORF1ab gene; however, only one target can be used in each assay which limits its accuracy and increases the probability of false negative occurrence [38]. A multiplex RT-LAMP assay has also been developed which allows for more accurate detection since it targets two SARS-CoV-2 regions [39]. There are also tests that employ both CRISPR–Cas and RT-LAMP technology at the same time in order to circumvent some of their individual limitations [40,41].

2.2.2. CRISPR–Cas

Currently, there is great interest in CRISPR–Cas based detection methods which rely on the use of guide RNAs and a ribonuclease to detect specific RNA targets. Two examples are Specific High-sensitivity Enzymatic Reporter unlocking (SHERLOCK) and DNA Endonuclease-Targeted CRISPR Trans Reporter (DETECTR), which use Cas13 and Cas12, respectively [42,43]. Both of these methods rely on the detection of specific SARS-CoV-2 sequences by the guide RNA, which then will trigger the cleavage of the reporter molecules by either Cas12 or Cas13 (Figure 1E). This will provoke a consequent detectable fluorescence signal. This assay takes from 30 to 40 min, plus RNA extraction, and the results can be observed either by fluorescence detection or by using a lateral flow strip which is able to capture labeled molecules.
The advantages of these methods are: its speed, bulky equipment or reverse transcriptase reaction is not needed, low cost and easy result interpretation. However, the target sequence choice requires optimization. It also requires specialized personnel for the RNA extraction step and is not as sensitive as RT-PCR. Moreover, the generated results are only qualitative, and each reaction only reports the result for one SARS-CoV-2 target sequence [43].
Even though some of the newly developed technologies have already been tested in clinical samples and approved for commercialization, their use in the general population is not common. This is probably due to the existence of antigen rapid tests and RT-PCR that, even though having some inherent disadvantages, have already been accepted and normalized by the population. Moreover, both these methods can cover different necessities, the antigen tests being fast and cheap making them ideal for point of care and regular use, while RT-PCR provides a more complete and reliable result while being more time consuming and costly. These factors make it difficult and not completely necessary to introduce other diagnostic methods. However, the existence of alternative methods is always benefic, more so because it is possible to improve the conditions of the considered “standard test” which, as seen in the previous section, can be slightly flawed.

2.2.3. Biosensors

A biosensor is a device that combines a biological component to detect an analyte, and a transducer to detect a physicochemical reaction in order to generate a measurable signal. Biosensors are comprised of three components: a bioreceptor, a transducer, and a signal processor. The bioreceptor is a biological element, for example a nucleic acid probe, a viral protein, or an antibody, which will recognize a specific analyte present in the sample (Figure 2). The binding of the analyte to the bioreceptor will cause a type of alteration that will be detected by the transducer. This alteration will be transformed into a measurable signal, and the signal processor will then be responsible for its display on an electronic device [44]. Depending on the type of transducer, biosensors can be mainly classified as electrochemical, thermal, optical, or piezoelectric.
One of the techniques used to enhance sensitivity and lower the limit of detection of biosensors is the addition of nanoparticles. Nanoparticles are particles with a dimension of 1–100 nm which also have special physicochemical properties. They can be synthesized from materials such as gold, silver, carbon, or silica. Depending on the type of material, they can exhibit photoluminescence, magnetic capability, low toxicity, high stability, or good biocompatibility and conductance [45]. Another advantage is the possibility to chemically modify them in order to make conjugates with either nucleic acid probes, viral proteins, antibodies, or other ligands. There are numerous nanoparticle-based biosensors being developed for COVID-19 detection; nevertheless, their advantages are the same: fast, low-cost, portable, user-friendly, and highly sensitive and specific. However, the use of nanoparticles usually appears hand in hand with the need to optimize these systems due to their highly unexplored potential.
Several biosensors have already been developed or adapted to the detection of SARS-CoV-2, although its use is not regular since most of them are still in the process of optimization and/or validation and general commercialization is still limited.
Below, several biosensors with possible different types of bioreceptors targeting either SARS-CoV-2 nucleic acid, viral proteins, or antibodies will be explored (Table 1).

Biosensors Targeting Nucleic Acids

Regarding choice of bioreceptor, nucleic acid probes targeting a specific SARS-CoV-2 sequence are usually the most sensitive and specific. This type of detection relies on hybridization of complementary strands. Nevertheless, the same challenges regarding choice of the target remains, as it must be sufficiently conserved in order to avoid any potential mutations, and therefore false negatives, but still be SARS-CoV-2 specific to avoid cross-reactivity [46].
Table 1. Potentially useful biosensing methods targeting SARS-CoV-2 nucleic acid (FET—field-effect transistor, PMO—phosphorodiamidate morpholino oligos, CD—carbon dot).
Table 1. Potentially useful biosensing methods targeting SARS-CoV-2 nucleic acid (FET—field-effect transistor, PMO—phosphorodiamidate morpholino oligos, CD—carbon dot).
OligonucleotidesRdRP, E, Orf1aSynthetic targets0.22 pM-[47]
FETPMORdRPThroat swab/Serum2.29 fM/3.99 fM2 min[48]
ElectrochemicalOligonucleotidesRdRP-0.30 pM25 min[49]
ElectrochemicalOligonucleotidesNNasopharyngeal swab/Saliva6.9 copies/μL5 min[50]
ColorimetriccDNA/AuNPsNOropharyngeal swab0.18 ng/μL10 min[51]
ElectrochemiluminescentThiol-modified oligonucleotideORF1abSpiked human serum514 aM-[52]
ElectrochemicalBiotinylated probeORF1abSpiked samples807 fM-[53]
VoltammetriccDNA-Au@CD bioconjugatesRdRPSputum0.15 pM75 min[54]
ElectrochemiluminescentY-DNA probeRdRPPharyngeal swab59 am180 min[55]

Biosensors Targeting Viral Proteins

Another used biorecognition element is antibodies which will detect specific viral antigens, such as the S and N proteins. However, this kind of biosensor will only be able to detect actively replicating viruses, as in the case of acute infections. Opposite to antibody application, Molecularly Imprinted Peptides (MIP) can also be designed to detect specific analytes, increasing the biosensor’s specificity [56]. Even though the previously described antigen LFIAs are the only biosensors targeting viral proteins that can be considered clinically useful and are widely used, there are several others with potentially better sensitivity and specificity currently being developed. Some of them are described in Table 2.

Biosensors Targeting Antibodies

Another possibility is the use of viral proteins to detect antibodies. However, these biosensors have the same disadvantages as the commonly used serological tests: they can only detect past infections since the immune response only starts several days after symptoms onset [68]. As mentioned before, they can also be used to study immune response due to the vaccination plan in some countries [16].
Table 1, Table 2 and Table 3 exemplify the vast number of biosensors in development and how the technologies used can vary so greatly. This represents many potential alternatives for the detection of not only SARS-CoV-2 but probably other viruses, because many of the developed systems are easily adaptable. Even though they have many differences in the speed and limits of detection, in general biosensors seem to represent a fast and very sensitive diagnosis tool. However, the novelty and range of materials and technologies used in this area makes it a complex task to compare the available biosensors. Moreover, sometimes it is difficult to compare the sensibility among different biosensors since there is a lack of uniformity in the units used to define detection limits. This also happens in regard to the type of samples in which those values were obtained, which often are not clinical samples but synthetic targets and which are sometimes not even being disclosed.

2.2.4. Sequencing

Sequencing techniques, specifically whole-genome sequencing, are the most complete and comprehensible approaches for the identification of viral RNA. Besides an accurate identification, it also allows the scientific community to keep track of the viruses’ genome evolution by detecting new mutations. Since the beginning of the pandemic, more than 10 million genome sequences have been shared on online platforms worldwide [79]. This global collaboration has allowed scientists to keep track of epidemiological outbreaks and new variants so as to increase vaccine efficiency and implement local preventive measures [80].
However, since sequencing is an expensive and slow approach, it is not recommended for large-scale testing in an emergency situation. Nevertheless, sequencing, in particular next-generation sequencing (NGS), was crucial in the first identification of SARS-CoV-2 and consequent characterization of its molecular structure. This, along with viral culture and electron microscopy, has allowed a deeper understanding of the virus structure and its mechanisms. The employment of these techniques has permitted the development of all the diagnostic methods and therapeutic strategies. This serves to highlight the importance of these unorthodox detection methods, that despite not being suitable for large-scale testing continue to be irreplaceable in the research context [14].
In Table 4, comparison between the advantages and disadvantages of the different SARS-CoV-2 detection tools is summed.

3. Test Selection

The numerous available diagnostic tools are a proof of the work and investment made by several institutions worldwide during this pandemic. However, the many options available also bring indecision and confusion regarding which method is best for each individual situation. Only in Europe, more than 365 tests have been approved and are available for commercialization [14]. The difference in test performance regarding specificity and sensitivity, the equipment needed for each one, the cost, and the information that each result provides are all variables that need to be taken into consideration when choosing the best option. Each test has a more or less specific window of time in which its results have higher accuracy, and sometimes the test chosen is simply not the most adequate for a given situation, which can lead to false results. Symptom presentation, time of infection, and type and quality of the sample are some of the variables that can greatly influence test results and their efficacy. Therefore, the main objective is to be able to perform “the right test, on the right sample, at the right time” [14]. Depending on the type of sample, test performance may vary. SARS-CoV-2 can be detected in samples from the upper respiratory tract (URT) (which includes the most commonly used nasopharyngeal swab, along with oropharyngeal swab, nasal mid-turbinate swab, anterior nares swab, nasopharyngeal/nasal wash and saliva), samples from the lower respiratory tract (LRT)(such as sputum and bronchoalveolar lavage fluid (BALF)), and lastly, samples from serum, urine, and feces. For example, regarding molecular tests, samples from either the URT or LRT are expected to yield better results, especially in the first two weeks after infection, with a peak in the first three days after symptom onset because the virus is actively replicating. Among these samples, BALF and sputum usually have higher viral loads [81]. After this point, the viral load in the respiratory tract is expected to diminish, as is the accuracy of the tests using these samples. In cases of prolonged infection, the best way to obtain a positive result is by analyzing urine and stool, as the presence of the virus has been reported one month after infection in these samples [82]. Regarding serological testing, antibody detection is recommended only at least two weeks after infection. Among the antibodies detected, IgM is expected to start decreasing six weeks after infection, while IgG remains in the organism for a longer period of time [14,83]. Besides choice of adequate sample type, it is also necessary to follow the adequate procedures regarding collection, transport, and storage which are provided by the CDC [84]. Figure 3 illustrates, in a simplified way, the relationship between the type of sample, test, and time after infection and which is the best choice combination to detect a specific analyte, either viral RNA, antigens, or antibodies.

4. New Variants

Although there are multiple options regarding COVID-19 diagnosis, the emergence of new variants can impact the performance of most of them. SARS-CoV-2 genetic variability may compromise some of the available diagnostic methods, especially with the recurrent emergence of new variants with high prevalence, e.g., variants of interest (VOI) and variants of concern (VOC). Regarding molecular tests, the most common event is that a mutation in the primer/probe binding region will no longer be recognized or will reduce amplification efficiency, and thus generate false-negative results. In order to avoid these problems, test manufacturers are recommended to perform routine sequence-alignment analysis to detect the mutations as soon as possible and redesign their assays. This work is facilitated by the large number of available and up-to-date genomes that are deposited daily on online public platforms and may lead to the optimization of some oligonucleotides [85]. Nonetheless, it is necessary to consider the timespan between the new variant emerging, the detection that the region used for the test is no longer adequate, and the removal of the kits from the market and the end-user. This timespan needs to be as short as possible, otherwise a local outbreak may no longer be identified before it becomes widespread. Therefore, the importance of using multiple targets as the probability of several sequences being mutated simultaneously is diminished.
Until now, there have been several reports that mention mutations influencing test performance, the majority of them with low impact worldwide (Table 5). For example, the Alpha, and more recently, the Omicron variant are associated with failure detecting a target within the S gene in the TaqPath RT-PCR assay, due to a deletion [86,87]. Failures in RT-PCR assays that target regions in the E and N gene have also been detected [88,89]. Another reported example consisted of a mutation in the primer used by the China CDC (named N-CHINA-F), present at the 5′ end of the N gene in a region where a substitution of 3 nucleotides (GGG to AAC) took place, being present in some SARS-CoV-2 variants, namely Alpha, Gamma, and Lambda [90]. Another study using 27 different RT-PCR assays found that 7 of them had mismatches in primer/probe binding zones. Even though the presence of a mismatch in the binding region does not necessarily mean that the primer/probe will not bind, it is better to update the assay in order to improve its accuracy and prevent eventual false negatives [91]. The position and number of mismatches both influence primer performance, the most detrimental being mismatches in the 3′ end of the primers [92]. Overall, studies have found that tests targeting the N gene are more prone to failure (Table 5). This is due to a high probability of mismatches in that zone, based on probe/primer and genome sequence information, namely high mutation ratio in this particular region [93].
These reports are particularly upsetting since they report errors on the test that is considered to be the most accurate and reliable for SARS-CoV-2 detection. However, it is necessary to consider that RT-PCR assays use more than one target in order to provide a more accurate result, as opposed to the POC tests that target only one region. A strategy that is also being used to minimize the impact of mutations is the design of degenerate nucleotides in specific SNP positions, but this is not an easy task [91].
Regarding antigen test performance, it can be compromised when the mutations present in the new variants change the viral protein structure, especially the N and S proteins which are the most commonly used targets in antigen testing. Since the S protein is a known hotspot for non-synonymous mutations, it is probable that eventually the accuracy of some commercialized tests will be compromised. Until now, there has been little reported evidence of failures in antigen tests, but this does not mean they did not fail. The Quidel Sofia SARS Antigen FIA test, targeting the N protein, has been reported to have some problems linked to the D399N mutation in the N protein generating false negative results [97]. However, the impact of this mutation is considered to be low since it is only present in less than 0.01% of all genomes present in worldwide databases [97].
Tests using antibodies have similar concerns. Since there are some VOCs that have reported changes in human immune response, the proteins used in LFIA may not recognize the slightly different antibodies produced by the organism. However, until now there have been no reports describing failures in antibody tests due to the existence of mutations.
Lastly, the best way to mitigate the influence of new mutations in all of these tests, besides keeping them regularly updated, is to use multiple targets. Most RT-PCR tests target two viral regions plus one positive control. If one of the targeted regions has a mutation and fails to be detected, the test result would be considered inconclusive and the next step would be to use one of the following strategies: repeat the test with another sample; perform a complementary test; or, as a last resort, sequence the sample.

5. Conclusions

The pandemic caused by SARS-CoV-2 has highlighted the importance of effective detection methods. Although there are several techniques available, COVID-19 diagnosis mostly relies on two techniques: RT-PCR and rapid antigen tests, e.g., LFIA. However, the constant research and development of other technologies, namely biosensors, allows a great range of potential options, not only for SARS-CoV-2 detection, but also for other viruses. Nevertheless, there are parameters that must be considered before choosing the most adequate test for each situation. Presentation of symptoms, infection time, and sample type all influence the outcome of a diagnostic tool. Other factors that can limit test choice are cost, time of response, availability of infrastructure, equipment, and specialized personnel. Moreover, the rise of new viral variants comes as a challenge, as it may affect the effectiveness of currently commercialized diagnostic tests. Hence, there is a necessity to maintain an updated genomic surveillance of the SARS-CoV-2 virus at the global level, as it is the only way to anticipate the possible failure of COVID-19 tests and proceed to the substitution and update of affected tests.

Author Contributions

A.L., M.A.C., P.M.-L. and H.M.R.G. conceived the project and contributed to writing the manuscript. All authors have read and agreed to the published version of the manuscript.


This work was funded by National Funds through FCT—Foundation for Science and Technology under Cdots Biosensing COVID19, nº 041_596518523, by RESEARCH COVID-19 and TEST3_Upscaling SARS-CoV-2, nº NORTE-01-0145-FEDER-072537. Was also funded by FEDER funds through the POCI and by National Funds through FCT under the NORTE-01-0145-FEDER-030858 and by the European Union (FEDER funds through Compete) UID/QUI/50006/2019. H.M.R. Gonçalves work was supported through the project UIDB/50006/2020, funded by FCT/MCTES through the project PTDC/BTM-MAT/30858/2017. M.A. Cardoso acknowledges the Post-doctoral grant (REQUIMTE 2021-55) subsidized by National Funds (FCT/MCTES) through Project PTDC/BAA-DIG/1079/2020.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.


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Figure 1. Schematic representation of the different detection technologies available and the sample under analysis: (A) Rapid Diagnostic Tests, (B) ELISA, (C) PCR, (D) LAMP, (E) CRISPR–Cas.
Figure 1. Schematic representation of the different detection technologies available and the sample under analysis: (A) Rapid Diagnostic Tests, (B) ELISA, (C) PCR, (D) LAMP, (E) CRISPR–Cas.
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Figure 2. Schematic representation of biosensor design.
Figure 2. Schematic representation of biosensor design.
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Figure 3. Scheme displaying the most appropriate options regarding type of sample, diagnostic test, and time after infection in order to detect each analyte: RNA, antigens, or antibodies (IgM and IgG).
Figure 3. Scheme displaying the most appropriate options regarding type of sample, diagnostic test, and time after infection in order to detect each analyte: RNA, antigens, or antibodies (IgM and IgG).
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Table 2. Potentially useful biosensing methods targeting SARS-CoV-2 viral proteins (NeuNAc—N-acetyl neuraminic acid).
Table 2. Potentially useful biosensing methods targeting SARS-CoV-2 viral proteins (NeuNAc—N-acetyl neuraminic acid).
Field-Transistor Effect (FET)AntibodiesS proteinNasopharyngeal swab242 copies/mL-[57]
ElectrochemicalACE2S proteinNasopharyngeal swab/Saliva229 fg/mL6 min[58]
ElectrochemicalAntibodiesS proteinEngineered Vero cells1 fg/mL3 min[59]
ThermalMIPS proteinNasal/Throat Swab<10 fg/mL15 min[60]
AntibodiesS proteinNasopharyngeal swab/Saliva1fg/mL10 min[61]
Surface Plasmon Resonance (SPR)AgNP/Antibody nanoconjugateS proteinSerum12 fg/mL-[62]
Plasmonic biosensorAuNP/Antibody nanoconjugateS proteinSolution containing S proteins4.2 fmol80 min[63]
FETAntibodiesN proteinSolution containing N proteins10 ag/mL4 min[64]
PlasmonicNeuNAcS proteinNasal swab--[65]
Plasmonic Fiberoptic AbsorbanceAnti-SARS-CoV-2 N-protein IgG1N proteinSolution containing N proteins2.5 ng/mL10 min[66]
Biotinylated antibodyN proteinPBS-diluted saliva6 pg/mL-[67]
Table 3. Potentially useful biosensing methods targeting SARS-CoV-2 antibodies (SPR—Surface plasmon resonance; GO—graphene oxide).
Table 3. Potentially useful biosensing methods targeting SARS-CoV-2 antibodies (SPR—Surface plasmon resonance; GO—graphene oxide).
SPRN proteinAntibodiesSerum30 nM15 min[69]
ElectrochemicalS proteinAntibodiesAntibodies in synthetic media9.3 ag/mL20 min[70]
ElectrochemicalGO-Au-antigen nanocompositeAntibodiesNasopharyngeal swab/Serum1 fg/mL-[71]
Square Wave Voltammetry (SWV)S protein + gold clustersS protein antibodyOropharyngeal swab/saliva0.03 fg/mL30 min[72]
Electrochemical ImpedanceRecombinant S proteinAntibodiesSerum1.99 nM-[73]
Cyclic VoltammetryRecombinant S proteinAntibodiesSerum2.53 nM-[73]
FluorescentB-cell epitopesAntibodiesSerum100 pM5 min[74]
SWVS protein epitopeIgGSerum-22 min[75]
SPRHistidine-tagged S proteinAntibodiesSerum0.057 μg/mL-[76]
SPRRBDAntibodiesSerum/Whole Blood-30 min[77]
ElectrochemicalN proteinAntibodySolution with antibodies13 fM<1 min[78]
Table 4. Principal advantages and disadvantages of the different types of SARS-CoV-2 detection tools.
Table 4. Principal advantages and disadvantages of the different types of SARS-CoV-2 detection tools.
RT-PCRHigh specificity and sensitivity, detects multiple targetsTime consuming, high cost, needs a laboratory setting and technicians
ELISAHigh sensitivity, faster and cheaper than RT-PCROnly detects 1 target, risk of cross-reactivity, needs a laboratory setting and technicians
LFIAVery fast, portable, and cheapLow specificity and sensitivity
CRISPR-BasedFast, high sensitivity and specificityOnly detects 1 target, needs a laboratory setting and technicians
RT-LAMPFast, high specificity and sensitivity, no bulky equipmentDifficulty in primer design
BiosensorsFast, portable, cheap, high specificity and sensitivityNeeds optimization
SequencingMost complete, detects all mutationsTime consuming and very high cost, needs a laboratory setting and technicians
Table 5. Mutations associated with failures in COVID-19 diagnostic tests and their prevalence in VOCs and VOIs.
Table 5. Mutations associated with failures in COVID-19 diagnostic tests and their prevalence in VOCs and VOIs.
ΔH69/V70SThermoFisher TaqPath COVID-19Alpha, Omicron[86,87]
C26340TECobas SARS-CoV-2-[89]
C28887TNChina CDCBeta, Eta, Mu[88]
C28977TNChina CDCAlpha[88]
C29200ANCepheid Xpert Xpress SARS-CoV-2-[96]
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Lino, A.; Cardoso, M.A.; Gonçalves, H.M.R.; Martins-Lopes, P. SARS-CoV-2 Detection Methods. Chemosensors 2022, 10, 221.

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Lino A, Cardoso MA, Gonçalves HMR, Martins-Lopes P. SARS-CoV-2 Detection Methods. Chemosensors. 2022; 10(6):221.

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Lino, Alexandra, Marita A. Cardoso, Helena M. R. Gonçalves, and Paula Martins-Lopes. 2022. "SARS-CoV-2 Detection Methods" Chemosensors 10, no. 6: 221.

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