Virus in a well – microplate-based assays for virology research

The COVID-19 pandemic has promoted global efforts to develop antiviral treatments and vaccines against SARS-CoV-2. The pandemic has put virology, a field of research that has been neglected for years, back on the map by Tobias Pusterla, BMG Labtech.

In fact, the increase in occurrence and types of virus outbreaks of recent years, such as SARS-CoV-1, Ebola, Zika, MERS and swine flu, illustrate the growing risks viruses pose to our health, our society and economy.

Virus research requires efforts to answer the fundamental questions that drive virological knowledge and future treatment development: how does a virus enter host cells? How does disease develop? How can hosts be protected? How can symptoms be relieved?

Modern medicine combats viral infections by vaccination and drug treatment. While vaccinations confer artificially acquired immunity to a specific virus, antiviral drug treatments typically target and reduce virus entry into cells and their replication. As viruses rely on cellular pathways and mechanisms to replicate, antiviral drugs may cause toxic side effects to the host. However, their selectivity and ability to interfere with viral replication, have been shown to be very effective.

The development, identification, repurposing and testing of antiviral drugs relies mainly on functional assays run mostly in microplates. The first microplates for laboratory use came into existence in 1951 due to the Asian flu pandemic. Hungarian doctor Gyula Takátsy, looking for a way to increase sample throughput in his laboratory, conceived the 96-well microplate format – today´s standard.

Since 1951, microplate-based assays have become an indispensable tool for modern laboratories, particularly now, as labs worldwide embark on the current virus challenge. Especially important are the microplate readers being used extensively in compound screens or for diagnostic tests.

As viruses encode a limited number of proteins, they must take over host cellular machineries and pathways to replicate and spread. Functional assays aim at understanding these host-virus interactions, clarifying the molecular mechanisms behind infection. Functional methods rely heavily on microplate-based assays and are employed in both basic life science research labs as well as in drug screening facilities. In fact, once the functioning mechanism of a virus is characterised and potential druggable target proteins are identified, functional assays need to be scaled up to screen for antiviral molecules that inhibit those very mechanisms.

Before moving to clinical studies, pharmaceutical regulatory agencies suggest the use of in vitroassays to determine cytopathic effect and mechanism of action of a virus, and to test resistance to novel antivirals ¹. The basic principles of functional assays, whether used for basic research or high-throughput screens are similar. The most relevant microplate-based assays utilised in virology are described below.

Infectivity and neutralisation assays

Infectivity assays determine and quantify the activity of a virus; its capability to enter the host cell and to use its molecular replication machinery. Methods to test the effect of antiviral molecules partially overlap with assays used to diagnose infections. Live cell-based microplate assays are generally employed for both low- and high- throughput assays.

Virus neutralisation is still the gold standard assay to assess the effectiveness of antiviral compounds or antibodies.This assay evaluates the ability of an antibody or compound to bind and neutralise the surface proteins a virus requires to enter host cells and replicate. The neutralisation assay combines incubation of the virus (or pseudo-virus) with potentially neutralising compounds or antibodies. Incubation is followed by a cell-based infection assay. This provides information whether the treatment can block viral infection of the host cells.

Traditionally, neutralisation assays have been performed with a virus solution mixed to potential neutralising samples at different concentrations. The use of native pathogens though exposed the operator to an elevated infection risk. Today, pseudo-viruses or pseudo-types are mainly used. These are chimeric viruses bearing the viral proteins of interest without the disease-causing genes. For a neutralisation assay, they are engineered to express fluorescent or luminescent reporter genes that are activated in the infected host cell upon viral replication (fig. 1). This way, infection and neutralisation can be easily detected in high-throughput and in real time by a microplate reader with gas control. ²For this purpose, atmospheric control units, able to regulate CO₂and O₂in the plate reader are essential.

Studying the cytopathic effect of a virus in presence or absence of a compound identifies whether the compound being tested can prevent infection. Screening assays to detect cytopathic effects are mainly based on the measurement of cell viability with an ATP-dependent luciferase, these assays are compatible with microplate formats up to 1536 wells. Reagents are added directly to cells after the appropriate incubation with virus and compound. The microplate is subsequently measured on a high-throughput plate reader where low signals correlate to a high cytopathic effect. In addition to infection-derived cell death, this assay also reports on compound-induced cell toxicity in control wells with compound and cells but no virus.

Microplate reader-based detection of cell viability upon infection is also used to determine the 50% Tissue Culture Infective Dose (TCID50). TCID50 quantifies the amount of virus required to produce cytopathic effect in 50% of inoculated cells. A titration of virus is applied to a constant number of host cells and given time to invade and replicate. TCID50 results are expressed as 50% infectious dose (ID50) per millilitre (ID50/ml). TCID50 is crucial in the development and testing of vaccines and antivirals, and for the production of viral antigens and viral recombinant proteins.

RNA dependent RNA Polymerase assays

Viral RNA-dependent RNA Polymerase (RdRP) is a specific target for RNA viruses such as SARS-CoV-2 and influenza. Viruses localised in the cytoplasm of a cell cannot access the host RNA polymerase to encode their own polymerase to promote replication. In these viruses, the RdRP enzyme transcribes RNA from a viral RNA template, in contrast to DNA-dependent RNA polymerase. Understandably, the analysis of RdRP activity and its inhibition are important factors.

Traditionally, the measurement of incorporation of radioactive nucleotides during RNA transcription has been used to determine RdRP activity in vitro. Today the use of fluorophores and microplate readers enables a more efficient and cost-effective quantification.³RdRP is mixed with a biotin-labelled primer and fluorescently labelled nucleoside triphosphates (NTPs) to initialise the reaction. If RdRP is active, it elongates the primer strand incorporating fluorescent NTPs. The complex of primer and newly synthesised RNA is subsequently immobilised on a streptavidin coated microplate, excessive NTPs can be washed away. The amount of immobilised NTPs depends on the activity of RdRP and can be quantified by the plate reader. Alternatively, fluorescent dyes that increase their intensity emission in the presence of RNA can be used. The increase in RNA catalysed by RdRP is then measured in a fluorescence microplate reader.

Fluorescence polarisation can also be used to determine RdRP activity by its complementation of a template RNA strand. Fluorescently labelled single strand RNA displays a low polarisation value. By RdRP-dependent synthesis of the complementary strand, the RNA molecule increases its size, leading to an increase in fluorescence polarisation (fig. 2).⁴ This method can efficiently be run on a plate reader with fluorescence polarisation detection.

Neuraminidase assay

Neuraminidase is a key glycoprotein found on the surface of influenza viruses. It cleaves sialic acid groups from glycoproteins on the host cell surface, leading to virion release from infected cells. It is crucial for the spread in the respiratory tract. Neuraminidase is a target of new therapies such as inhibitors zanamivir and oseltamivir.^{5, 6}Neuraminidase activity and inhibition can be efficiently measured on plate readers using a synthetic substrate which, when processed by the enzyme, releases a fluorophore. In this assay, the virus is mixed with the substrate and fluorescence intensity is detected and directly related to neuraminidase activity. ⁷Alternatively, a chemiluminescence substrate may be used, purportedly providing higher sensitivity than fluorescence.⁸

Interaction assays for compound screens 

In the field of antiviral drug development, interaction assays play a fundamental role. They are used to screen for molecules that interact with and inhibit structures essential for the lifecycle of a virus. Common targets include host cell receptors used by viruses to enter the cell, proteases cleaving viral precursor or cellular proteins, and polymerases, integrases and methyltransferases involved in genome replication.

Interaction assays rely mainly on proximity detection methods such as resonance energy transfer (FRET, BRET and TR-FRET) and AlphaScreen. Alternatively, fluorescence polarisation is used to track changes in molecular size induced by binding events.

Proximity methods are mainly based on Förster´s resonance energy transfer taking place between two fluorophores (FRET), a luciferase and a fluorophore (BRET), or between a fluorophore with long emission half-life (lanthanide) and a fluorophore (TR-FRET). High-throughput screens rely on simple mix-and-read, homogeneous assays with low background signals, an inherent feature of resonance transfer assays.

Transfer of energy (photons) takes place from a donor to an acceptor chromophore, if found in proximity. The two molecules whose interaction being investigated are labelled, one with the donor, the other with the acceptor chromophore. The acceptor only emits a signal if binding between the two molecules occurs, bringing donor and acceptor in close proximity.

Screening for interacting molecules generally uses a competitive setup: a target bearing the donor and a known target binder bearing the acceptor lead to acceptor signal emission. If a compound displaces the acceptor-labelled molecule, acceptor signal decreases due to separation, signalising binding inhibition.

FRET-based methods are commonly ratiometric: data are internally normalised, eliminating the influence of the number of molecules present in a sample and minimising interferences from variable assay conditions. This represents an obvious advantage for screening campaigns.

Low background noise is specifically observed in assays that use long lifetime fluorophores (TR-FRET) or lumiphores (BRET) as donors. For instance, a TR-FRET-based assay was used to screen for inhibitors of the interaction between the HIV-1 integrase and its main co-factor. Using this method, a strong inhibitor being regarded as a promising candidate for HIV therapy was found. ⁹

Although based on a different physico-chemical principle, AlphaScreen is as well often used for interaction assays. This method has been used in 1536 well plates to establish a high-throughput assay to investigate inhibitors of the interaction between the SARS-CoV-2 spike protein and the ACE2 receptor. Engagement of the ACE2 receptor by the spike protein allows entry into host cells and an inhibitor of this interaction is a promising therapeutic. The assay provided an excellent assay quality with Z’ > 0.7 and identified 25 compounds out of 3400 which inhibited the spike protein – ACE2 interaction. ¹⁰

Fluorescence polarisation-based virus assays

Fluorescence polarisation (FP) is a cost effective and high-throughput compatible method to screen for molecular interactions. Polarisation of fluorescent emission light increases with a growth in molecular size of the complex formed with the fluorophore. Accordingly, FP-based interaction assays rely on binding or dissociation -dependent gain or loss of molecular weight of the complex formed with the fluorophore, respectively. FP is used to monitor binding events between nucleic acids or proteins and virus components required for replication, host proteins, or inhibitors. In particular, FP is ideal for studying interactions between the viral genome and replication proteins, as fluorescent labelling of oligonucleotides representing specific regions of their viral genome is easy and inexpensive. A FAM-labelled RNA template from the Zika virus was for instance used to analyse binding of RdRp to RNA. The results helped to identify a methyltransferase as a potential Zika virus therapeutic target. ¹¹

Currently, FP is experiencing somewhat of a revival in biological research. This is mainly caused by the increased performance and sensitivity of modern microplate readers that can deliver robust results with minimal variability and larger assay windows. This also applies to FP assays using red fluorophores, the low photon yield of which has previously been a problem. The use of red fluorophores in FP can be advantageous as it minimises unspecific background fluorescence which is usually in the blue-green spectral range.

Microplate-baseddiagnostic virus assays

Besides the need for a profound understanding of SARS-CoV-2 biology and for the swift development of treatments, the COVID-19 pandemic focussed significant efforts in the development of diagnostic methods to identify infected individuals. Identification of infection is necessary to control and limit virus propagation via an effective disease management. ¹²Precise identification of past or enduring infections helps to prevent transmission and supports appropriate therapies and treatment monitoring. ¹³

Diagnostic assays vary in efficacy, approach and cost. Besides accuracy, precision and specificity, ease of use, throughput and turn-around time (the interval time between sample registration and result reporting) are important factors. ¹⁴

Traditionally, identification methods of viral infections focussed mainly on visual detection and counting of virions by microscopy techniques. These methods require highly trained personnel, adequate facilities, a relatively large volume of patient sample and are subject to long turn-around times. ^{13, 15, 16}Accordingly, such techniques are now more often adopted for research rather than for diagnostic purposes.

Today, more rapid methods allowing qualitative testing for screening and surveillance, as well as confirmation of diagnosis are used. These rely mainly on the detection of the viral genome present in patient samples or on serological tests.

Nucleic acid amplification methods

Detection of viral genetic material in patients diagnoses infections early and with sensitivity, speed and reliability. ¹⁷As nucleic acid amplification techniquesdetect and amplify specific regions of the viral genome, they oftentimes can confirm infection even before symptoms are evident, or antigens and/or antibodies appear in the bloodstream.Common methods include Polymerase Chain Reaction (PCR) and quantitative PCR (qPCR).

A new approach that is increasing in popularity is the Loop-mediated Amplification (LAMP) assay. LAMPis a colorimetric or fluorescent assay based on an isothermal DNA amplification reaction run at 65ºC, and has already been successfully applied to SARS-CoV-2 testing. ¹⁸Unlike PCR, it can be easily measured in real-time in a microplate reader with periodic shaking, as it does not rely on temperature cycles. In addition, it bears cost and speed advantages over PCR and is highly automatable, making it a perfect assay for rapid virus diagnosis.

Serological tests 

A shortcoming of genetic amplification techniquesis that they cannot report on acquired immunity or past infections. To get these answers, serological tests are required.

Enzyme-Linked Immunosorbent assays (ELISA) measure the presence and concentration of antibodies against a virus or viral antigens in serological samples. A positive test is not an indicator of active infection though, as antibodies may persist in the blood for years after infection. ELISAs rather determine patient immunity upon exposure, reinfection, or reactivation and are invaluable for studying the epidemiology of disease. ELISAs produce a colorimetric, fluorescent or chemiluminescent output that is detected by a microplate reader. The signal is proportional to the amount of specific antigen or antibody in the sample.

A more sensitive but usually more expensive alternative to classic ELISAs are homogeneous immunoassays. Unlike ELISAs, washing steps to remove the unbound components from the well and reduce the background are not required. This minimises both handling steps and operation time, and makes them particularly suited for automation-supported screening (fig. 3). Homogeneous assays for virus detection include fluorescence polarisation immunoassays, and different TRF, TR-FRET and AlphaScreen-based assays. Besides recognition of antibodies, immunoassays are also helpful to understand the immunology behind viral infection, detecting and quantifying for instance different cytokines.

Microplate-based assays are fundamental tools for virologists to analyse the molecular and cellular mechanisms of infection objectively and quantitatively. Recent developments in detection technology broadened the range of methods for virus detection, analysis of interactions and biological responses. These approaches are in part complementary, in part alternative to previously established methods in virology. Modern microplate readers further enhance these approaches, increasing sensitivity and speed, as well as providing the capability of real-time monitoring in living cells for prolonged periods.

The combination of microplate-based assays and readers is currently improving and will further improve data quality and throughput, possibly enabling assays and analyses that have been previously problematic.

Volume 22, Issue 1 – Winter 2020/21

About the author

Tobias Pusterla has a degree in biotechnology and a PhD in cellular and molecular biology. He is international marketing manager with BMG LABTECH.

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