Assay Development for Monitoring DNA Damage

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Assays that monitor DNA damage play a pivotal role in drug discovery endeavors, providing critical insights into the genotoxic potential of candidate compounds. These assays assess a compound’s ability to induce various forms of DNA damage, including strand breaks and mutations. Their significance lies in their capacity to provide essential information about the safety profile of potential drugs, identify compounds selectively targeting DNA damage in cancer, and pinpoint inhibitors of DNA damaging agents or processes.


Our objective was to develop cell-based assays for DNA damage and cytotoxicity to screen novel genotoxic compounds and DNA damage inhibitors effectively. We established assays capable of directly and indirectly measuring DNA damage, leveraging high content imaging and flow cytometry techniques. These assays offer diverse approaches to screening genotoxic compounds and serve as primary and orthogonal assays in the discovery pipeline.

What We Did...

Measurement of DNA damage involves detecting exposed 3’OH groups in DNA strand breaks or assessing the accumulation of DNA repair proteins at damaged sites. We devised a screening cascade comprising assays designed to identify and evaluate compounds’ activity in modulating DNA repair enzymes/proteins. Ensuring the biological relevance and specificity of these assays to the target protein and signalling pathway was imperative. Moreover, the assays needed to be reproducible, robust, and capable of handling expected compound throughput at each stage of the screening cascade.

The screening cascade developed which aimed at identifying small molecule inhibitors of DNA damage response proteins is illustrated in Figure 1.

DNA Damage_Assay Cascade

Figure 1. Screening cascade based on target biology based on availability of commercial reagents and throughput formats, to achieve the timelines and milestones requirements of the project. 

NanoBRET™ Target Engagement Assay

The Promega NanoBRET™ target engagement assay facilitates the quantification of compound affinity for the target protein or binding to related off-target proteins within a cell-based system. This assay utilizes full-length target proteins, providing insights into compound cell permeability and residence time.

To execute the assay, cells undergo transient transfection with a cDNA construct, generating a fusion protein comprising the full-length target protein linked to NanoLuc® Luciferase. Upon addition of a cell-permeable fluorescent NanoBRET™ tracer, which is a fluorescently labeled compound capable of reversibly binding to the target protein, bioluminescence resonance energy transfer (BRET) occurs. This phenomenon transpires once the tracer binds and closely interacts with the NanoLuc® Luciferase.

The apparent affinity of test compounds is discerned through competitive displacement of the NanoBRET™ tracer, leading to a subsequent reduction in BRET signal.

DNA Damage_NanoBret diagram

Figure 2. Concept of the NanoBRET™ assay from Promega. In this format, a fusion construct of the target cDNA connected to the Nanoluc® cDNA is transiently expressed in cells. In the presence of substrate, the Nanoluc® enzyme generates photons. A fluorescence tagged tracer (a compound that binds to the target with high affinity) is added to the cells and binds to the target protein, bringing the fluorophore into close proximity with the Nanoluc® photon “engine” and activating the fluorophore. Compounds that bind to the target compete with the tracer and in so doing reduce the fluorescence in the presence of a consistent photon signature.

By utilizing a custom tracer compound, we successfully identified active compounds selectively binding to the target protein while minimizing off-target binding. An example of the results for one of the active compounds identified using this assay is shown in Figure 3.A. where potent binding and successful displacement of the NanoBRET™ tracer is observed for the target protein but only minimal binding is identified in the off-target protein (Figure 3.B.) counter-screen assay.

DNA Damage_Example results from NanoBret

Figure 3.A. A compound that competes for binding to the target protein decreases the fluorescence signal as it displaces the tracer from the binding site. Figure 3.B.  The same compound does not bind to the off-target protein and therefore is selective. Y-axis represents BRET ratio, which is the amount of specific fluorescence (tracer detected at 600+nm) divided by the amount of photons detected at 460nm.

Cell Toxicity

In drug discovery screening cascades, the inclusion of a cytotoxicity assay is considered essential due to the potential of toxic compounds to yield unforeseen outcomes in cell-based assays. This practice becomes particularly pertinent in our project, given that our downstream cellular assays involve counteracting the effects of a DNA damage inducing agent. The occurrence of compound-induced cytotoxicity could significantly compromise assay readouts. Thus, our objective is to screen compounds at concentrations below toxicity thresholds.

DNA Damage_Cell toxicity assay diagram

Figure 4. Principle of Promega Cell-Titre GLO assay. The substrate luciferin, in the presence of excess co-factors (provided in the assay buffer), utilises ATP from the lysed cells to generate oxyluciferin and photons of light. Since ATP is not stored inside cells but is manufactured to order, then decreases in ATP suggest perturbation of cell metabolism leading to cell death.

The CellTiter-Glo® assay operates on the principle of ATP-dependent conversion of luciferin to oxyluciferin, resulting in luminescence. Cells serve as the source of ATP, and the luminescence signal is directly proportional to the concentration of viable cells. The impact of compounds on cell viability may arise from inhibiting cell growth, inducing cell death, or perturbing cell metabolism.

In this assay format, luminescence readouts are standardized against zero-hour controls and depicted as percent growth inhibition. The GIC50 (Growth Inhibition Curve) denotes the concentration at which 50% growth inhibition is achieved, while growth inhibition exceeding 100% indicates cytotoxicity, as the number of viable cells diminishes below the density observed at zero-hour. Zero-hour controls are established in wells where cells are seeded and allowed to adhere but remain untreated with compounds or DMSO. These controls provide a baseline for determining zero-hour luminescence, which reflects the number of viable cells and their metabolic activity at the initiation of compound treatment

DNA Damage_Cell toxicity assay_grapgh of results

Figure 5. Growth inhibition curve as a percent of the zero-hour control ATP level determined in control untreated wells just prior to compound addition. If the compound is non-toxic then the line will remain at 0% inhibition or less than 0%. If the compound causes the growth inhibition to plateau at 100% then the compound is cytostatic, the cells remain in the wells and are producing ATP but there are the same number of cells as time zero. If the compound is toxic, then the line will plateau at 200%, consistent with little or no ATP in the cells, in this case 24 hours post-treatment.

Quantification of DNA Damage by γH2AX Assay

Within cells, the protein H2AX plays a crucial role in orchestrating the repair of damaged DNA. When DNA damage is detected, H2AX undergoes phosphorylation at the residue Ser-139, thereby forming γH2AX. As a response to DNA damaging agents, γH2AX foci manifest in cell nuclei in a dose-dependent manner. This phenomenon enables the quantification of DNA damage, facilitating the screening of genotoxic compounds.

This process is applicable to agents that directly induce DNA damage, such as cisplatin or mitomycin C, as well as those that indirectly provoke DNA damage by impeding DNA repair mechanisms. The detection and quantification of foci are easily accomplished using selective γH2AX antibodies, resulting in a highly sensitive assay for monitoring DNA damage and its resolution.


Figure 5. DNA damage is reliably quantified by staining for γH2AX, which forms foci at affected DNA sites. In this instance HT115 cells were treated with a serial dilution of mitomycin C. γH2AX foci were detected with a 555 nm fluorescence labelled secondary antibody. Nuclei were stained with Hoechst. Images were captured using a ImageXpress® Confocal with a 40X water immersion objective.

Assay Development to monitor DNA Damage

Figure 6. DNA damage is quantified by detecting γH2AX foci formed in a dose-dependent manner in response to DNA damaging agents. Using mitomycin C as a High Control, the assay can achieve a window of >3 Signal to Background (S2B) and a Z’ Score of >0.5.

Quantification of DNA Single-Strand Breaks by TUNEL Assay

DNA strand breaks induced by genotoxic agents reveal free 3’OH groups (Figure 7). Terminal deoxynucleotide transferase (TdT) catalyses the addition of deoxynucleotides to these 3’OH groups. This can be quantified by Flow Cytometry via the addition of dUTP-FITC, which is integrated into DNA by TdT, allowing for an increase in fluorescence with DNA damage (Figure 7).

In this TUNEL assay, cells are treated with compounds and a tool genotoxic agent for 48 h, fixed, permeabilized and then incubated with the TdT enzyme and dUTP-FITC. Compounds are screened for their ability to inhibit the effect of the DNA Damaging Agent.

Figure 7. Reagents cause single strand breaks in the DNA of the cell. TdT hijacks the repair mechanism and adds dUTP-FITC to these repaired 3’OH ends. The degree of fluorescence (FITC), as detected by flow cytometry, is proportional to the level of DNA damage.

Treatment with the DNA damaging agent (depicted in Figure 8.A) results in the induction of single strand breaks in approximately 60% of cells within the UTP-FITC_FSC_+ve gate, contrasting with minimal induction observed in cells treated with DMSO. This consistent induction of UTP-FITC +ve cells by the DNA damaging agent across experiments contributes to the robustness of the screening assay, as illustrated in Figure 8.B.

In instances of additional cell stress, such as exposure to toxic concentrations of a compound, higher levels of UTP-FITC incorporation (as shown in Figure 8.A) occur due to apoptosis-induced formation of double stranded DNA breaks within the apoptotic_FSC gate. The gating of apoptotic cells facilitates the identification and unbiased exclusion of outliers resulting from cell death, ensuring the integrity of further analysis.

Compounds undergo assessment for their capacity to inhibit the DNA damaging agent-induced increase in UTP-FITC cells, with data normalization to controls yielding percent inhibition values (depicted in Figure 8.C). The utilization of flow cytometry for cell analysis has enabled precise replicates for compound concentration curves. Furthermore, a notable correlation exists between the potency of compounds observed in the Target Engagement assay and the TUNEL assay.

DNA Damage_TUNEL Assay Results

Figure 8. TUNEL DNA damage assay. Populations of cells can be differentiated into groups based on the FITC signal as they pass through the counting chamber. They can also be segmented and excluded based on cyto-toxicity and cell death, allowing the population with DNA damage alone to be identified and counted.

An Orthogonal Measurement of DNA Damage via a Cell Cycle Assay

DNA Damage_ On Target functional response diagram

Figure 9. Bromodeoxyuridine (BrdU) uptake is used to track the cell cycle stage of individual cells. Cells incorporate this synthetic thymidine analog while synthesizing new DNA during S phase. By providing BrdU for a brief period it is possible to mark a pool of cells that are in S phase while the BrdU is present. These cells are then tracked through the remainder of the cell cycle and into the next round of replication, permitting the duration of the cell cycle phases to be determined without the need to induce a potentially toxic cell cycle block. In this case a fluorescence tagged antibody is used to detect the degree of BrdU staining and 7AAD intercalates the DNA such that the amount of DNA can be quantitated.

In any screening cascade, confirming the results of active compounds using an orthogonal assay is crucial to mitigate the risk of false positives. To address this, we implemented a cell cycle assay to validate the activity of hit compounds.

During this assay, cells are subjected to treatment with compounds and a DNA damaging agent for 48 hours prior to pulse staining with BrdU, followed by fixation and permeabilization. Subsequently, cells undergo a limited DNase digest to expose BrdU-labeled epitopes before being stained with an anti-BrdU-FITC antibody and 7-AAD.

As cells progress through their cell cycle, the amount of DNA within each cell increases from G0/G1 phase to G2/M phase, with an intermediate amount of DNA present during S phase when DNA replication occurs. The binding of 7-AAD to DNA is stoichiometric, proportional to the amount of DNA in each cell, while BrdU is only incorporated into cells actively replicating their DNA during S phase.

By utilizing a combination of BrdU-FITC and 7-AAD staining, cells can be categorized into distinct phases of the cell cycle, including G0/G1, S, or G2/M phase, while apoptotic cells undergoing DNA cleavage are gated as SubG1 of the cell cycle.

Treatment with a DNA damaging agent (as depicted in Figure 10.A & .B) leads to G2/M arrest, a reduction in the percentage of cells in S phase, and a slight increase in the percentage of cells in SubG1. The impact of compound treatment on DNA damaging agent-induced G2/M arrest can be visualized for the entire cell cycle (as shown in Figure 10.C.) or as the variance from DMSO treated cells (illustrated in Figure 10.D.).

In both representations (Figure 10.C. & .D.), a dose-dependent inhibition of the DNA damaging agent’s effect is clearly evident. Additionally, percent inhibition values for the effect on S phase (Figure 10.E.) and G2/M phase cells (Figure 10.F.) can be plotted. Remarkably, complete compound inhibition curves are observed, along with a correlation in the IC50 values with those obtained in the Target Engagement and TUNEL assays.

DNA Damage_ On Target functional response results

Figure 10. Using the combination of BrdU-FITC and 7-AAD staining, cells can be gated (A) as being in G0/G1, S or G2/M phase, whilst apoptotic cells (undergoing DNA cleavage) are gated as SubG1. Treatment with a DNA Damaging Agent (A and B) induces G2/M arrest, a decrease in the % of cells in S phase and a small increase in the % of cells in SubG1. The effect of compound treatment DNA Damaging Agent-induced G2/M arrest can be displayed for the whole cell cycle (C) or as the difference to DMSO treated cells (D). In both displays (C and D) we can clearly observe a dose-dependent inhibition of the effect of the tool genotoxic agent. We can also plot % inhibition values (generated by normalizing the data to the controls) for the effect on S phase (E) and G2/M (F) phase cells. 



In summary, we have developed a comprehensive suite of assays for screening genotoxic compounds and DNA damage inhibitors. These assays serve as valuable tools in drug discovery, offering insights into compound safety, target engagement, and efficacy in modulating DNA damage response proteins.