Reporter gene assays guide — Part 9: Troubleshooting
Even with the best design and the most carefully chosen controls, reporter assays fail. They fail in ways that are usually obvious in retrospect and almost never obvious at the time. The goal of this chapter is to short-circuit that pattern: to give you a checklist of failures that experienced researchers know to look for, organised by the symptom you actually observe at the bench.
"I have no signal"
This is the most common and most alarming failure. The first question to ask is whether the cells are expressing the construct at all.
The transfection failed. This is the most frequent cause. Verify transfection efficiency by including a GFP or mCherry control well in parallel, or by checking the Renilla signal in a dual-luciferase experiment. If the constitutive control is also at background, the transfection is the problem. Common causes: old or incorrectly-stored transfection reagent, wrong reagent for the cell type, DNA of poor quality, cells in poor condition, or cells at the wrong density. Each of these has a known fix: fresh reagent, switching to a transfection reagent validated for the cell type, re-prepping the DNA, thawing fresh cells, and adjusting density to 70 to 90% confluence at the time of transfection.
Double check whether you are using a cellular or a secreted reporter. If you reporter plasmid is cellular but you are assaying the media you wont get a signal!
The reporter gene has a problem. Confirm the sequence of the reporter. Silent mutations, frameshifts, and accidental truncations are surprisingly common. Sequencing the construct before troubleshooting further is time well spent.
The promoter is not active in your cell type. Some promoters are highly cell-type-specific. A neuron-specific promoter driving luciferase in HEK293 cells will give background, because the transcription factors required are absent. Verify by running a positive control (a known strong promoter driving the same reporter in the same cell line) in parallel. If the positive control works and your promoter does not, the promoter is the problem.
The cells are not receiving the substrate properly. For firefly luciferase, this usually means the lysis was incomplete or the substrate was cold. For fluorescent reporters, the substrate does not apply but excitation/emission settings can be wrong. For secreted reporters, the timing of substrate addition can be critical, particularly for flash-kinetics reporters like Gaussia luciferase.
The plate reader is malfunctioning. Less common than you would think. Check the instrument calibration, the gain setting, and the integration time. A gain that is set too low for a dim signal is a frequent cause of "no signal" that is actually "very little signal that the reader cannot see."
"I have signal but no induction"
This is more subtle than "no signal" because the assay is clearly functioning. The problem is that the experimental condition is not producing the expected effect. Common causes:
The positive control is also not working. A sign that the issue is not specific to your test condition. The pathway may not be inducible in your cell type, the time point may be wrong, or the stimulus may be inactive. Verify the stimulus with an independent method (e.g. Western blot for a downstream marker).
The reporter construct is not responding to the pathway. A common reason is that the binding sites in the synthetic promoter are insufficient, in the wrong context, or the wrong ones for your pathway. Verify the construct by testing it in a system where you know the pathway is robustly active (e.g. a cell line that constitutively activates the pathway).
The induction is happening at a different time point. Most signalling responses are transient. A reporter read at 24 hours will not see a response that peaks at 4 hours. A time course is the diagnostic: read at 0, 1, 2, 4, 8, 12, and 24 hours to find the peak.
The concentration of the stimulus is wrong. Too low, no response. Too high, toxic or off-target effects. A dose-response curve with at least 5 concentrations is the standard fix. The peak of the response is usually 3- to 10-fold lower than the toxic threshold.
The cells are not responsive. Cell lines lose responsiveness with passage number, particularly primary cells and lines that have drifted from the original stock. Verify by testing the cells with a known stimulus and reading out a different marker (e.g. qPCR of an endogenous target gene). If the cells are unresponsive to the stimulus, the problem is the cells, not the reporter.
"I have high background"
The reporter is leaky. All promoters have some baseline activity, and high-copy plasmids can give substantial signal even from "silent" promoters. Switch to a lower-copy vector, use a weaker promoter, or add a destabilisation tag to reduce accumulation. The promoterless control well tells you the magnitude of the problem.
The cells have endogenous activity. Some cells express endogenous alkaline phosphatase, β-galactosidase, or even luciferase-like activity. For alkaline phosphatase, heat-inactivation of the samples (65°C, 30 minutes) eliminates most of the endogenous activity. For β-galactosidase, switching to a luminescent reporter (e.g. β-galactosidase-driven Gaussia luciferase) avoids the background. For other reporters, run untransfected cells in parallel and subtract the background.
The substrate is contaminated or old. Old or improperly-stored substrate gives high background and erratic signals. D-luciferin is reasonably stable when stored dry at -20°C; coelenterazine is much less stable and must be protected from light. Furimazine (the NanoLuc substrate) is also light-sensitive.
The plate reader gain is too high. A signal that is at the top of the dynamic range saturates the detector and gives an apparent high background because the baseline is being amplified. Reduce the gain, dilute the sample, or use a smaller aliquot of lysate.
"My data is highly variable"
Well-to-well coefficients of variation above 20% are usually fixable. The first place to look is at the pipetting. Aqueous solutions of DNA and transfection reagent are viscous, and small variations in volume give large variations in transfection efficiency. Use a multichannel pipette for 96-well work, a positive-displacement pipette for viscous solutions, and a reagent reservoir rather than repeated aspiration from a tube.
Other sources of variability:
Cell density variation. Cells that are not evenly distributed across the well give different signals. Make sure the cell suspension is well-mixed before plating, and let the plate sit at room temperature for 10 to 15 minutes after plating to allow even settling before moving to the incubator.
Edge effects. The outer wells evaporate faster, experience temperature gradients, and have different gas exchange. Either fill the edge wells with media only (and exclude them from analysis) or use a plate with a moat.
Inconsistent lysis. Particularly for adherent cells, incomplete or variable lysis gives variable signals. The Promega passive lysis buffer is forgiving; other lysis buffers (Triton-based, RIPA) are less so. Use a consistent protocol with a defined incubation time.
Plasmid DNA quality. Preps of varying quality give different transfection efficiencies. Use a single high-quality prep for an entire experiment, or use a transfection-grade midi- or maxi-prep. If comparing activity of different plasmids, re quantify both of the plasmid concentrations.
Cell health. Cells that are stressed, contaminated, or at the wrong passage number give highly variable signals. Use healthy cells at a defined passage number, and verify by mycoplasma testing.
"My signal disappears when I scale up"
This is a common and frustrating problem when moving from a 24-well or 96-well format to a 384-well or 1536-well format. The causes are usually physical rather than biological:
Surface-to-volume ratio changes. Smaller wells have more evaporation per unit volume. The medium concentrates, the cells are exposed to higher effective drug concentrations, and the pH shifts. Use evaporation-resistant plates, reduced incubation times, or humidified incubators.
Cell density is wrong for the new format. Cells per well, not just percent confluence, matters. A 384-well plate with 5,000 cells per well is very different from a 96-well plate with 5,000 cells per well. Re-titrate the cell density for the new format.
Reagent volumes do not scale linearly. The volume of transfection reagent, lysis buffer, and substrate all need to be re-optimised for the new well size. Use the manufacturer's recommended starting points and titrate.
Plate reader settings change. The path length, the integration time, and the gain all need to be re-optimised for the new well format. Run a dilution series of a positive control to verify linearity in the new plate.
"My results do not reproduce"
This is the worst category of problem because it usually shows up after the experiment is finished, and the cause is often one of the ones above that was overlooked earlier. The most common specific causes:
Different cell passage. The cells used in experiment 1 were at passage 8; the cells used in experiment 2 were at passage 22. The cells have drifted. Use cells within a defined passage window and track this carefully.
Different plasmid preps. Different preps of the same construct can give different signals due to variations in supercoiling, endotoxin level, and DNA concentration. Use a single large-scale prep for an entire project, or normalise transfection efficiency with a co-transfected control.
Different reagents. New bottle of serum, new bottle of trypsin, new lot of substrate. Each one is a potential source of variation. Test new lots against the old and reserve sufficient quantities of a single lot for long projects.
Different protocol execution. Small variations in incubation time, temperature, pipetting technique, and reagent order can give measurable differences. Document the protocol in detail and follow it precisely. The details that seem unimportant (vortexing after substrate addition, the exact duration of the lysis incubation, the time from substrate addition to reading) often matter.
"My screen is full of hits that are not real"
This is a HTS-specific problem. A high hit rate (more than 1% of compounds activating the reporter) usually means the screen is detecting compound autofluorescence, luciferase inhibition, or non-specific effects. Common causes:
Luciferase inhibition. Compounds that inhibit firefly luciferase directly (and there are many in screening libraries) give an apparent repression of the reporter that is actually an assay artefact. The same compounds can give an apparent activation of a co-expressed Renilla control by suppressing the ratio, depending on the order of substrate addition. The fix is to use a dual-luciferase format and exclude compounds that affect both reporters.
Compound autofluorescence. Fluorescent compounds give signal in the reporter channel without any biological activity. This is particularly problematic for fluorescent protein reporters. The fix is to filter compounds by their spectral properties or to use a luminescent reporter that is not affected by compound fluorescence.
Cytotoxicity. Compounds that kill cells give apparent pathway activation or repression that is actually a stress response. The fix is to counter-screen for viability on every hit and exclude cytotoxic compounds.
Off-target pathway effects. Some compounds activate the pathway through an unexpected mechanism. The fix is to validate hits with a different reporter, a different cell line, and a different mechanism of action.