New Accurate Diagnostic Technology Uses Sugars To Detect SARS-COV-2 Coronavirus


Scientist from the department of chemistry at the University of Warwick-UK along with experts from Iceni Diagnostics Ltd-UK and University Hospitals Coventry and Warwickshire (UHCW) NHS Trust-UK have developed and demonstrated a new diagnostic technology using sugars instead of antibodies to detect the SARS-CoV-2 coronavirus.

The study team developed a prototype flow-through device (related, but distinct to LFDs), utilizing N-acetyl neuraminic acid-functionalized, polymer-coated, gold nanoparticles as the detection/capture unit for SARS-COV-2, by targeting the sialic acid-binding site of the spike protein.
The prototype device can give rapid results, with higher viral loads being faster than lower viral loads. The prototype’s effectiveness is demonstrated using spike protein, lentiviral models, and a panel of heat-inactivated primary patient nasal swabs.

The study findings showed that the device was able to retain detection capability toward recombinant spike proteins from several variants (mutants) of concern.

These findings provide the proof of principle that glyco-lateral-flow devices could be developed to be used in the tracking monitoring of infectious agents, to complement, or as alternatives to antibody-based systems.

The study findings were published in the peer reviewed journal: ACS Sensors

The COVID-19 pandemic has led to >171 million confirmed cases and ∼3.7 million deaths worldwide, reported to WHO, as of the 4th of June 2021. (1) COVID-19 is caused by the coronavirus SARS-COV-2, first reported in Wuhan (China). (2) Despite global efforts, there are still a limited number of effective therapeutics. Vaccines have now been approved for use, but with limited supplies; a major mechanism for controlling disease spread remains testing, identification, and patient isolation.

The testing system deployed by more economically developed countries (MEDCs) and less economically developed countries (LEDCs) has been based primarily on molecular (genetic) approaches such as real-time reverse-transcription polymerase chain reaction (rRT-PCR). (3−6)

However, RT-PCR-based approaches require dedicated laboratory facilities and trained personnel, meaning early in the pandemic CT scans, which are not recommended for routine use, were initially employed. (7) Due to the infrastructure needs of RT-PCR and long processing times, RT-PCR does not typically provide a rapid turnaround, especially in a high volume laboratory setting, although it is considered the gold standard for COVID-19 diagnosis.

In July 2020 during the early stages of the COVID-19 pandemic, in the United States, the average wait time for an RT-PCR test result was 4 days with 37% of people receiving the results within 2 days. (8) The availability of RT-PCR testing also varies significantly between countries; per 1000 people (31/7/2020) (9) the United Kingdom (2.27) and the United States (2.91) have significantly out-tested LEDCs such as Zimbabwe (0.07) or Myanmar (0.01). (9)

In Iran, for example, CT scanners are more abundant (10) than RT-PCR machines. (11,12) Faster RT-PCR devices, such as those based on DNAnudge, have been developed and allow for decentralized testing outside of hospital or lab environments but do have capacity requirements of one machine to one test. (13) Other molecular genetic techniques have also been developed, which similarly do not require centralized testing infrastructure.

For example, loop-mediated isothermal amplification (LAMP) (14) can return a diagnosis in just over 90 min (LamPORE device). Although faster than conventional RT-PCR, neither of these offer rapid results at a capacity that would facilitate mass screening or at a cost per device that would allow point-of-care testing in the home or in low-resource environments. (15,16)

Lateral-flow devices (LFD) are established tools for rapid diagnosis, giving results often in under 30 min and therefore can rapidly identify infected individuals. LFDs, such as the home-pregnancy test, (17) use antibodies as detection units in both the stationary phase (test line bound to nitrocellulose) and as a coating for the mobile phase (on the surface of a gold nanoparticle). Upon binding the target analyte, the stationary and mobile phase form a “sandwich” with the analyte in the middle.

The results are visible by the eye as a red or blue line depending on the precise gold formulation, although other nanomaterials, such as fluorescent particles, can be used. (18) LFDs are typically cheap (compared to molecular methods), require little to no training or clinical infrastructure to use, and can be scaled up to enable large population testing. LFDs tend to have lower sensitivity (some false negatives) but high selectivity (few false positives).

The cost-effectiveness and clinical usefulness of LFDs have been demonstrated by malaria rapid diagnostic tests, (19,20) in the diagnosis of cutaneous leishmaniasis (21) and in comparisons with more expensive RT-PCR approaches for Ebola diagnosis. (22) Consequently, the appeal of LFDs in the COVID-19 pandemic is that their low cost and rapid turnaround time may enable mass testing of large populations. (23) This could find asymptomatic individuals spreading the virus, who would not be identified by symptomatic RT-PCR testing only, (24−26) currently the preferred option in most healthcare systems.

The first LFDs for the COVID-19 pandemic were designed to detect antibodies in patient blood samples produced in response to SARS-COV-2 infections. (27−29) These were intended to report if a patient has previously been infected; not to indicate active infection, so could not effectively be used in screening/triage settings or mass testing for active infections.

Antigen LFDs, in contrast, are designed to diagnose the presence of the virus i.e., an active infection. Several antigen lateral-flow tests, by late 2020, had passed Phase 3 testing in the United Kingdom, (30) gained WHO “Emergency Use Listing” approval, (31) or had emergency approval granted by The United States Food & Drug Administration. (32,33)

These devices all utilize antibodies as detection/capture units. To the best of our knowledge, these devices all use antibodies to target the nucleocapsid protein of SARS-COV-2. A university-based validation testing between LFDs and PCR confirmed that LFDs cannot detect lower viral loads but were estimated to be capable of identifying up to 85% of infections in the cohort trialed (26) showing their potential for frequent, low-cost testing when deployed appropriately.

It is important to note that antibodies are not essential components in LFDs, and other recognition moieties could be used, including nucleic acids, (34) glycans, and lectins. (35) Glycan-based LFDs could offer advantages over other recognition moieties.

For example, glycans are the site of pathogen adhesion during many viral infections (36,37) especially respiratory viruses such as influenzas, (38) and glycans can be chemically synthesized at scale. Glycan binding can also explore the “state” of a pathogen; for example, LecA/B are upregulated by Pseudomonas aeruginosa during infection. (39,40)

Furthermore, glycans are often more thermally robust than proteins (41) making them ideal candidates for low-resource environments. Glycosylated gold nanoparticles (the mobile phase) are well established having been used in colorimetric/aggregation-based diagnostics, surface enhanced-Raman, and other bioassays. (42−45) Despite this, glycans as capture units have not been widely applied in lateral flow (46) and to the best of our knowledge have not been shown to function using clinical samples, only models.

We have previously reported that the S1 domain of the SARS-COV-2 spike protein can bind α,N-acetyl neuraminic acid (Neu5NAc), a sialic acid, (47) and similar binding has been observed for other zoonotic coronaviruses toward sialic acids (48−50) (e.g., MERS).

The exact biological role of sialic acid binding is not yet understood for SARS-COV-2, with clear differences in its role in cell entry compared to MERS. (51) Microarray, ELISA and STD NMR have been used to further demonstrate that sialic acids are receptors for the SARS-COV-2 spike protein. (52−54) It has also emerged that sulfated glycosaminoglycans (including heparin sulfates) bind SARS-COV-2 spike protein, and can inhibit viral entry. (55−57) Glycans (including those carrying terminal sialic acids) have been shown to participate in the angiotensin-converting enzyme 2 (ACE2) receptor binding during SARS-COV-2 cell adhesion/entry. (58)

Incorporation of α,N-acetyl neuraminic acid onto a polymer-stabilized glyconanoparticle platform enabled detection of (purified) spike protein in an LFD (5 μg·mL–1) and also detection of a pseudotyped lentivirus presenting the SARS-COV-2 spike protein at 1.5 × 104 transduction units·mL–1 in a dipstick test. (47)

Herein, we demonstrate that glycan-based flow-through devices can detect SARS-COV-2 in heat-inactivated primary patient swabs and validate these initial results against RT-PCR. Compared to an LFD format, no test line was used, rather the sample is directly absorbed onto the nitrocellulose strip. Device optimization was achieved using a lentivirus and recombinant SARS-COV-2 spike protein showing that heat (or chemical) inactivation did not prevent usage.

The prototype devices were then used with a panel of primary heat-inactivated swabs, demonstrating the principle that flow-through glyco-assays can be used to detect viral infection and hence that glyco-LFDs are feasible if suitable test lines can be developed.

Furthermore, the devices were shown to detect recombinant mutant spike proteins showing that glycan-based detection may still detect variants of concern. This conceptual approach could be broadly applied to other pathogens/disease states and provide redundancy in testing regimes compared to using antibodies alone.

Figure 1. Nanoparticle synthesis and flow-through devices. (A) Neu5NAc-terminated polymer coating; (B) TEM micrograph of polymer-coated AuNPs; (C) C 1s portion of the XPS spectrum of polymer-coated AuNPs; and (D) flow-through device layout and assay procedure (top to bottom).


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