No Amplification in Known Positive Samples — A Practical Troubleshooting Checklist

If your known positive sample is flat-lining, the experiment is broken at a fundamental level and nothing else on that plate is interpretable. Stop analyzing your unknowns and fix this first. The positive control exists precisely to catch these failures — it's doing its job. Now you need to do yours.

The most common culprabb? In roughly this order: reagent problem (master mix or primers), template problem (degraded, too dilute, or inhibited), or instrument/setup error (wrong protocol, no activation step, optical misconfiguration). Less common but worth checking: wrong well mapping, sealed plate punctured during loading, or you simply forgot to add a component. I know that last one sounds insulting, but I've seen it from first-year students and senior PIs alike. No shame — just re-make the reaction.

Rule Out the Obvious First

Before you start swapping reagents and re-extracting RNA, spend five minutes on the things that take five minutes to check:

1. Did you actually add template? Check your tube — is the volume where you'd expect it, or is it still full? If you were loading a 96-well plate, it's surprisingly easy to skip a well or pipette into the wrong one.
2. Did you add master mix? Look at the wells. Reactions without master mix will look visibly different in volume (10 µL vs. 20 µL is noticeable). Some people aliquot primers and template first, then add master mix — if you got interrupted, maybe you never finished.
3. Check your run file. Open the protocol on your QuantStudio, CFX96, or LightCycler 480 and confirm: Did the hot-start activation step actually run? PowerUp SYBR needs 2 min at 50°C then 2 min at 95°C. Luna Universal needs 1 min at 95°C. TaqMan Fast Advanced needs 20 sec at 95°C. If you loaded someone else's protocol or the wrong saved file, the polymerase may never have activated.
4. Check your detection settings. On the CFX96, make sure the correct fluorophore channel is selected (SYBR/FAM). On the QuantStudio, confirm the experiment type matches your chemistry. I once spent an afternoon troubleshooting a "failed" TaqMan assay that had been set up as a SYBR experiment — the software was reading the wrong channel.
5. Check the plate seal. A loose or punctured optical seal causes evaporation, which concentrates inhibitors, creates bubbles in the light path, and generally ruins everything. If you see condensation patterns on the seal or salt crust in wells after the run, this is your problem.

If all of the above checks out, move on to the reagents.

Reagent Troubleshooting

Primers first. Primers are the most failure-prone reagent in a qPCR setup because they're handled constantly, diluted by everyone in the lab, and degrade if someone left the working stock at room temp over a long weekend.

• Make a fresh 10 µM working stock from your 100 µM freezer stock. Use nuclease-free water, not TE — EDTA at working concentrations can chelate enough Mg²⁺ to reduce efficiency, and at higher concentrations it'll kill the reaction entirely.
• Confirm the final primer concentration in your reaction. Most SYBR assays work well at 200–400 nM each (forward and reverse). If you're using 10 µM stocks in a 20 µL reaction, that's 0.4–0.8 µL per primer. A pipetting error that delivers 0.04 µL instead of 0.4 µL means you have essentially no primers in the tube.
• If you suspect primer degradation, run them on a 3% agarose gel or check on a Bioanalyzer. But honestly, just ordering new primers is faster and cheaper than diagnosing degraded ones.

Master mix. If other people in the lab are getting amplification with the same master mix lot, it's probably fine. If nobody is, or if you're the only user, consider:

• Has this aliquot been through many freeze-thaw cycles? Most commercial mixes (PowerUp SYBR, Luna Universal, SsoAdvanced) tolerate quite a few, but eventually activity drops.
• Did someone accidentally add the wrong thing to the tube? Contamination with bleach, ethanol, or detergent will kill the polymerase. If the master mix looks unusual (precipitate, color change), toss it.
• Try a fresh aliquot from a sealed tube, or borrow a colleague's known-working mix.

Probe (TaqMan assays only). Probes are more fragile than primers. They're light-sensitive, and the fluorophore-quencher pair degrades over time, especially with repeated freeze-thaw.

• Check that the probe concentration is correct (typically 100–250 nM final).
• If you're multiplexing, confirm that the probe isn't quenched by spectral bleed-through from another channel — though this would cause high background, not no signal. No signal from a probe usually means no probe in the well or completely degraded probe.

Template Problems

Your "known positive" is only as good as its last quality check. Templates fail for a few specific reasons:

Degraded RNA/cDNA. If your positive control is cDNA that's been sitting at –20°C for six months, it may have degraded. cDNA is more stable than RNA, but it's not immortal — especially in dilute solutions (< 1 ng/µL) where adsorption to tube walls becomes significant. Run a test with a freshly reverse-transcribed batch from the same RNA, or use a gDNA positive control if your primers allow it.

Inhibitors carried over from extraction. This is the classic problem with tissue samples, plant material, blood, and stool. Common inhibitors include:

• Ethanol from wash steps (most common — the "I can smell ethanol in my eluate" situation)
• Phenol from TRIzol extractions
• Heparin from blood samples
• Humic acids from soil/environmental samples
• Melanin from skin/hair follicle samples

The test for inhibition is simple: spike a known quantity of an exogenous control (like the SPUD assay, described by Nolan et al. 2006) into your sample and into water, and run both. If the sample's Ct is delayed by >1 cycle compared to the water-only control, you have inhibition. Alternatively, run your positive control at 1:5 and 1:10 dilutions — if the diluted samples actually amplify (or amplify with better efficiency), inhibition is your problem.

Template too dilute. If your positive control is at the edge of detection — say, Ct 35+ on a previous run — any small loss of template (adsorption, degradation, pipetting error) can push it to no amplification. Use a positive control that gives a Ct of 20–25 so you have headroom.

Instrument and Thermal Cycler Issues

Less common, but when it's the instrument, it affects everything on the plate — so if all wells (including your positive) are flat, consider this:

• Block/rotor temperature calibration. If the block isn't reaching 95°C, denaturation is incomplete and you get no amplification. Most instruments have a diagnostic self-test. Run it, or contact your core facility.
• Optical system failure. A burned-out excitation LED or a misaligned detector will give you flat lines across the board. Check by running a plate of just master mix (no template) — you should see a flat baseline with some fluorescence. If the raw fluorescence values are at zero or near-zero across all wells, the optics are the issue.
• On the Rotor-Gene Q specifically, make sure the locking ring is properly seated. If the rotor wobbles, optical reads become erratic and the software can't call a Ct.

A Systematic Approach When You're Stuck

If you've checked the common causes and you're still staring at flat lines, here's how to isolate the variable:

1. New master mix + old primers + old template → tests master mix
2. Old master mix + new primers + old template → tests primers
3. Old master mix + old primers + new template (or gDNA) → tests template
4. All new reagents → if this works, one of your original reagents was the problem; swap them back one at a time

Run these on a small scale — 3–4 reactions total, not a full plate. Use a well-characterized housekeeping gene assay (GAPDH, ACTB, or 18S) as your readout, since you know it should amplify from any intact cDNA or gDNA. If even 18S at 200 pg of human gDNA doesn't give you a Ct around 10–12, you've either got an instrument problem or a master mix problem.

When It's Not Really "No Amplification"

One more thing. Check your baseline and threshold settings before concluding there's no amplification. On the QuantStudio software, auto-baseline sometimes sets the baseline window incorrectly, especially for late-amplifying targets, and the amplification curve gets subtracted into oblivion. Look at the raw fluorescence (multicomponent plot or amplification data in raw Rn mode). If you see a sigmoid curve in the raw data but not in the ΔRn plot, your baseline settings are wrong, not your biology.

On the CFX96, the equivalent is checking "RFU" vs. "Baseline Subtracted" in the data analysis tab. I've seen people report "no amplification" when the target was clearly there at Ct 33 but got buried by an aggressive baseline correction.

If you're tired of manually stepping through these checks every time something goes wrong, VoilaPCR flags failed positive controls, baseline anomalies, and outlier replicates automatically when you upload your run file — so you can spend your time fixing the actual problem instead of hunting for it.