The good news is that PCR failures follow patterns. The specific pattern of what you see on the gel, combined with the results of your controls, nearly always points to a small number of likely causes. A systematic approach to troubleshooting, starting with the controls and working through the most common causes, resolves the majority of PCR problems without guesswork.
This page gives you that systematic approach. It covers the most common patterns of PCR failure, their causes, and the specific steps that fix them. It covers primer design issues, template quality problems, cycle optimisation, PCR inhibitors, and qPCR-specific troubleshooting. At every step the focus is on what is actually happening in the reaction tube, not just what to try next.
Check Your Controls First, Every Time
Before any troubleshooting of the sample reactions begins, the controls tell you where the problem lies.
The positive control is a sample known to contain the target sequence. If the positive control shows the correct band at the expected size, the PCR reaction itself is working. The problem is with the test samples, not the assay. This directs troubleshooting toward sample quality, template DNA, inhibitors, or a technical issue specific to how those samples were set up.
If the positive control fails (no band, wrong size band, or faint band), the problem is with the assay itself: the primers, the polymerase, the buffer, the cycling conditions, or the template used as the positive control. Do not proceed to interpret test sample results from a run where the positive control failed.
The no-template control (NTC) contains all reaction components except template DNA. It should produce no visible band. A band in the NTC indicates contamination of the reagents with template DNA or PCR product from previous runs. This is a serious result that invalidates the entire run and requires decontamination of the workspace and reagents before repeating.
No Bands at All
No bands in either positive control or test samples: the reaction has completely failed. Possible causes in order of likelihood:
Forgotten component: Check the reaction setup. The most commonly forgotten component is the polymerase. Review the protocol and your pipetting record to confirm every component was added. In a multi-tube setup, it is easy to miss one tube when adding polymerase.
Reagent failure: Polymerase is the most temperature-sensitive component. If the enzyme was left at room temperature, thawed and refrozen multiple times, or stored incorrectly, it may have lost activity. Try a fresh tube of polymerase. Similarly, dNTPs degrade with repeated freeze-thaw cycles. Use a fresh aliquot.
Template failure: If the positive control uses the same template as the test samples, both would fail together if the template is degraded, absent, or too dilute. Quantify the template DNA on a NanoDrop or by gel to confirm it is present at adequate concentration (typically 1 to 100 ng per reaction for genomic DNA).
Cycling program error: Confirm the cycling program was set up correctly on the thermocycler: correct denaturation temperature (94 to 98 degrees Celsius depending on polymerase), correct annealing temperature, correct extension temperature (72 degrees Celsius for Taq), correct extension time (at minimum 1 minute per kilobase for Taq, 30 seconds per kilobase for high-fidelity polymerases), and correct number of cycles (25 to 35 for most applications).
Primer failure: Primers that have degraded, been stored at incorrect concentrations, or were synthesised incorrectly will not work. Verify the primer stock concentration. If primers have been in use for a long time or have been subjected to many freeze-thaw cycles, order fresh primer resynthesis.
Faint or No Bands in Test Samples but Positive Control Works
The assay works (positive control is positive) but the test samples are not amplifying. This pattern points to a problem with the test samples, not the assay.
Insufficient template: The most common explanation. Too little DNA was added. Quantify the template with NanoDrop or a fluorometric method (PicoGreen is more accurate for low-concentration samples). Increase the amount of template added to the reaction.
Template inhibitors: Clinical specimens and environmental samples contain substances that inhibit Taq polymerase and other PCR components. Common inhibitors include heme (from blood), humic acids (from soil and stool), SDS (from DNA extraction buffers), calcium, bile salts, and many others. Inhibitors can cause complete PCR failure or just reduce amplification efficiency. Detection: add a small amount of a known template (a spike) to the inhibited reaction alongside the experimental template. If the spike also fails to amplify, inhibitors are confirmed. Solutions: dilute the template (diluting 1:5 to 1:20 often resolves inhibition), use inhibitor-tolerant polymerase formulations (hot-start Taq with inhibitor-resistant buffer), or re-extract the template using a purification column to remove the inhibitor.
Degraded template: DNA or RNA that has been partially degraded by nucleases produces fragments shorter than the PCR target size. If the target is 500 bp but the template is fragmented to 200 bp average, you will not amplify the full-length product. Run an agarose gel of the template to check for degradation (intact genomic DNA should show a high-molecular-weight smear on gel). Store templates correctly (DNA at -20 degrees Celsius, RNA at -80 degrees Celsius in appropriate buffers).
Non-Specific Bands
Multiple bands of the wrong size suggest the primers are annealing at non-target sites in the genome and amplifying unintended products.
Annealing temperature too low: The most common cause of non-specific bands. Reducing the annealing temperature allows more mismatches between primer and template, causing primers to bind at multiple sites. Raise the annealing temperature in 1 to 2 degree increments using gradient PCR (running the same reaction simultaneously at multiple temperatures in a gradient thermocycler block). Start at Tm - 5 degrees Celsius and test up to Tm + 2 degrees Celsius.
Primer design problems: Primers that have internal secondary structure (hairpin loops) or that can form primer-primer duplexes (primer dimers) produce non-specific products. Check primer secondary structure and dimer propensity using online tools (such as OligoAnalyzer from IDT). If problems are found, redesign the primers.
Too many PCR cycles: Running too many cycles amplifies trace amounts of non-specific products to visible levels. Reduce cycle number to 25 to 30 and see whether non-specific bands disappear.
Primer Dimers
A very bright, low-molecular-weight band, typically below 100 base pairs, with no or faint specific band is the primer dimer pattern. Primer dimers form when the two primers in the reaction anneal to each other (or to themselves) rather than to the template and are extended by the polymerase.
Solutions: Increase the annealing temperature. Use hot-start polymerase, which is inactive below around 55 degrees Celsius, preventing primer extension during the setup and initial temperature equilibration. Reduce primer concentration (typical working concentration is 200 to 400 nM; reducing to 100 to 200 nM can reduce dimer formation). If the problem persists, redesign the primers to avoid complementary 3-prime ends.
Optimising Magnesium Concentration
Mg2+ is an essential cofactor for Taq polymerase. The standard concentration in most commercial PCR buffers is 1.5 mM free Mg2+, but many primers and templates perform better at slightly higher or lower concentrations. If all other troubleshooting has failed, try testing Mg2+ concentrations from 1.0 to 3.5 mM in 0.5 mM increments. Excess Mg2+ increases non-specific amplification. Insufficient Mg2+ reduces polymerase activity and can cause failed amplification.
qPCR-Specific Troubleshooting
High Ct values (late amplification): Template is too dilute, inhibitors are present (dilute the template), or the primer/probe design is poor. Efficiency below 90 per cent in the standard curve indicates a problem with the assay.
Inconsistent duplicate Ct values (high standard deviation): Pipetting error is the most common cause. Use calibrated pipettes and mix the master mix thoroughly before aliquoting. Template may also be inadequately mixed, especially if it is a viscous extraction product.
Signal in NTC: Contamination of reagents with template or amplicon. Decontaminate all surfaces with 10 per cent bleach followed by UV light exposure. Prepare new master mix using fresh aliquots of all reagents.
High baseline fluorescence: Probe or primer concentration too high. Reduce to recommended levels. Insufficient denaturation during set up: check that the polymerase activation step (for hot-start) reached the correct temperature.
Frequently Asked Questions
Why does my PCR not work even though I followed the protocol?
PCR has many possible failure points: reagent quality, template quality, primer design, cycling conditions, and inhibitors. The first step is always to check the controls: did the positive control amplify? If not, the problem is with the assay components. If yes, the problem is with the test samples. Work through the most likely causes systematically rather than changing multiple things at once.
What is a no-template control (NTC)?
The NTC is a negative control in which all PCR components are included except the template DNA. It should produce no amplification signal. A positive result in the NTC indicates contamination of the reagents or workspace with template DNA or amplified PCR product from previous runs, which invalidates the entire run.
What are PCR inhibitors?
PCR inhibitors are substances present in a sample that reduce or completely block the activity of the DNA polymerase or otherwise prevent amplification. Common inhibitors include heme from blood, humic acids from stool and environmental samples, bile salts from stool, calcium, EDTA (if present in excess in the extraction buffer), and SDS from lysis buffers. Inhibition can be confirmed by spiking the reaction with a known template alongside the test sample.
What is the optimal annealing temperature for PCR?
The optimal annealing temperature is typically 3 to 5 degrees Celsius below the melting temperature (Tm) of the primers. If the two primers have different Tms, use a temperature based on the lower Tm primer, or use a buffer system designed to compensate for differences. The optimal temperature for a given primer pair is best determined empirically using gradient PCR.
What is hot-start PCR?
Hot-start PCR uses a modified polymerase that is inactive at room temperature and is only activated by the initial high-temperature denaturation step in the thermocycler. This prevents non-specific extension of primers during reaction setup (when the mixture is at room temperature and primers can anneal non-specifically), and dramatically reduces primer dimer formation and non-specific amplification.
What is gradient PCR?
Gradient PCR runs the same reaction simultaneously across a range of annealing temperatures (typically spanning 10 degrees Celsius) in a thermocycler with a gradient block. The temperature at which the specific product appears with the best yield and the cleanest gel result is the optimal annealing temperature for that primer pair and template combination.
What does a smear on a PCR gel mean?
A smear, where the bands merge into a continuous blur rather than discrete bands, usually indicates DNA degradation (the template is broken into many different sizes and the PCR produces fragments of many sizes) or gross non-specific amplification producing many products of different sizes. Check template quality by running the template on a gel before amplification. If the template shows a smear, it is degraded and needs to be replaced.
What is the extension time and how is it calculated?
The extension time in PCR is calculated based on the expected product size and the extension rate of the polymerase being used. Taq polymerase extends at approximately 1 kilobase per minute, so a 2 kb product requires at least 2 minutes of extension time. High-fidelity polymerases such as Phusion or Q5 extend at approximately 1 kb per 30 seconds and require shorter extension times. Using insufficient extension time causes truncated products or failed amplification of larger targets.
Why do I need to optimise Mg2+ concentration?
Mg2+ is the essential divalent cation cofactor for DNA polymerase. Too little Mg2+ reduces polymerase activity and decreases amplification efficiency. Too much Mg2+ stabilises non-specific primer-template interactions and increases non-specific amplification. The standard concentration of 1.5 mM works well for most reactions, but some primer-template combinations perform better at slightly different concentrations, particularly with complex secondary structure templates or challenging primer pairs.
What is DMSO and when is it used in PCR?
DMSO (dimethyl sulphoxide) at 2 to 5 per cent final concentration is added to PCR reactions involving GC-rich templates or target sequences with strong secondary structure. High GC content causes template strands to re-anneal more readily during denaturation (because GC base pairs have three hydrogen bonds rather than two), preventing primer access. DMSO destabilises these secondary structures, improving denaturation and primer annealing to difficult templates. It should not be added routinely as it can reduce polymerase activity when unnecessary.