How to Set Up a No-Template Control in PCR and qPCR

A no-template control (NTC) is a reaction mixture containing all PCR or qPCR components except the DNA or cDNA template, replaced by an equivalent volume of nuclease-free water or the buffer used to dilute samples. The NTC serves as the primary contamination control in every PCR-based assay, detecting whether any component of the master mix—water, primers, probes, nucleotides, polymerase, or buffers—has been contaminated with nucleic acids from previous amplifications or environmental sources. You should include at least one NTC in every PCR run, and ideally duplicate NTCs when running high-throughput or diagnostic assays, because a positive NTC signal invalidates all results from that run and requires systematic decontamination before repeating the experiment.

At a Glance

Scientific Principle of the No-Template Control

The NTC operates on a simple but essential principle: if any component of the PCR master mix contains contaminating nucleic acids, those contaminants will be amplified alongside your intended targets, producing false-positive signals. The NTC contains every reagent that goes into your sample reactions—polymerase, dNTPs, primers, probes, buffer, and water—but substitutes the template volume with a clean, nuclease-free liquid that should contain no amplifiable DNA or RNA.

The sensitivity of PCR and especially qPCR makes this control indispensable. A single contaminating molecule can be amplified to detectable levels within 35–40 cycles, and qPCR can reliably detect as few as 1–10 copies of target DNA. This means that even trace contamination from previous experiments, aerosolized amplicons, or contaminated pipette tips will produce a signal in the NTC. The NTC thus functions as the earliest warning system for contamination, allowing you to identify problems before they compromise your experimental data.

The underlying risk of contamination is well recognized in molecular biology laboratories. The CDC and NIH, in the Biosafety in Microbiological and Biomedical Laboratories (BMBL), 6th Edition, emphasize that standard molecular biology procedures involving nucleic acid amplification require careful attention to contamination control, particularly when working with recombinant or synthetic nucleic acids [1]. The NIH Guidelines for Research Involving Recombinant or Synthetic Nucleic Acid Molecules further establish that investigators must implement appropriate containment practices, which include the routine use of negative controls such as NTCs to monitor for unintended amplification [2].

Materials and Instrumentation Choices

Water and Diluents

The choice of water for your NTC is critical. Use only molecular biology-grade nuclease-free water that has been certified DNase/RNase-free and tested for absence of amplifiable nucleic acids. Do not use autoclaved distilled water, deionized water from laboratory stills, or water that has been stored in shared containers, as these frequently contain trace nucleic acids from environmental sources. Commercial nuclease-free water from reputable suppliers is the standard choice, and you should test each new lot before using it in critical assays.

For some applications, particularly when your samples are diluted in a specific buffer (e.g., TE buffer, low-EDTA TE, or sample dilution buffer), you should use that same buffer for the NTC. This ensures that any contaminants present in the buffer itself will be detected. However, nuclease-free water remains the most common and widely accepted NTC diluent.

Master Mix Components

The master mix for your NTC should be prepared from the same stock reagents used for your sample reactions. Do not set aside a separate "clean" master mix for NTCs, as this defeats the purpose—you need to test the exact reagents that will contact your samples. Prepare a single master mix containing all common components (polymerase, dNTPs, primers, probes, buffer, and any additives), then aliquot this master mix into tubes for samples and NTCs.

When using commercial master mixes, note that some pre-formulated mixes contain stabilizers, glycerol, or other additives that can occasionally harbor contaminants. Always include an NTC when opening a new tube of master mix for the first time, as this establishes a baseline for that lot.

Consumables

Use only sterile, DNase/RNase-free pipette tips with aerosol barriers (filter tips) for all PCR setup steps. Standard non-filter tips can allow aerosolized contaminants to enter the pipette barrel and be transferred to subsequent reactions. Similarly, use only sterile, DNase/RNase-free PCR tubes or plates that have been certified free of nucleic acid contamination.

Instrumentation

The choice of thermal cycler or qPCR instrument does not affect NTC setup, but the instrument's detection method influences how you interpret NTC results. In conventional PCR with gel electrophoresis, the NTC should show no visible band after staining. In qPCR using fluorescent probes or SYBR Green, the NTC should produce no amplification curve that crosses the threshold, or if it does, the Cq value should be clearly distinguishable from positive samples (typically >5 cycles later than the weakest positive sample).

Conceptual Workflow for NTC Setup

Step 1: Prepare Your Workspace

Before beginning any PCR setup, clean your work area with 10% bleach (sodium hypochlorite) followed by 70% ethanol. Bleach degrades nucleic acids, while ethanol removes residual bleach and provides a clean surface. Use dedicated PCR setup areas that are physically separated from areas where post-amplification products are handled. The BMBL recommends establishing clean and dirty zones in molecular biology laboratories, with unidirectional workflow from pre-amplification to post-amplification areas [1].

Step 2: Prepare the NTC First

When preparing your master mix, add the NTC water or buffer to the master mix tube before adding any sample templates. This order ensures that if your pipette tip or water contains contaminants, they will be distributed throughout the master mix and detected in the NTC. Some laboratories prepare the NTC as the very last reaction to be loaded, but this practice risks contaminating the NTC with amplicons from nearby sample tubes. The safer approach is to prepare the NTC master mix first, then aliquot it into the NTC tube or well, and only then proceed to add templates to the remaining master mix.

Step 3: Use the Correct Volume

Replace the template volume exactly. If your protocol calls for 2 µL of template DNA, add 2 µL of nuclease-free water to the NTC. Using a different volume can alter the final concentration of master mix components, potentially affecting amplification efficiency and making the NTC non-comparable to sample reactions.

Step 4: Cap or Seal Immediately

After adding the NTC water, immediately cap the tube or seal the plate well. This prevents aerosolized contaminants from entering the NTC during subsequent pipetting steps. If using a plate, consider sealing the NTC wells with optical adhesive film before adding samples to other wells.

Step 5: Place the NTC Strategically

In qPCR plates, place NTC wells in positions that are not adjacent to high-concentration positive controls or strong samples. Edge effects and well-to-well contamination are more likely when NTCs are placed next to wells with high template concentrations. A common strategy is to place NTCs in the first column (e.g., A1, B1) and positive controls in the last column, with samples in between.

Quality Checks Before and During the Run

Pre-Run Checks

• Verify that your nuclease-free water has been tested for nucleic acid contamination. Many commercial suppliers provide certificates of analysis; review these before use.
• Check that all pipettes used for PCR setup are dedicated to pre-amplification work and have been calibrated within the last 6–12 months.
• Confirm that filter tips are being used and that the tip boxes have not been contaminated by opening them in post-amplification areas.
• Ensure that your master mix has not been previously used for reactions containing template. Always use freshly prepared master mix or master mix that has been stored in single-use aliquots.

During the Run

• Monitor the NTC amplification curve in real-time if using qPCR. A sudden upward curve in the NTC during the run indicates contamination and should prompt you to consider whether the run should be aborted.
• For conventional PCR, note that you will not know the NTC result until after gel electrophoresis. This makes pre-run quality checks especially important.

Result Interpretation

Negative NTC (Expected Result)

A properly functioning NTC shows no amplification. In qPCR, this means no fluorescence signal that crosses the threshold within the number of cycles run. In conventional PCR, this means no visible band on the gel at the expected product size. Some qPCR instruments may show a slight increase in fluorescence in later cycles (e.g., after cycle 38–40) due to primer-dimer formation or non-specific amplification, particularly with SYBR Green chemistry. This is acceptable if the signal does not cross the threshold and if the melting curve (if performed) does not match the target amplicon.

Positive NTC (Contamination Detected)

If the NTC shows amplification, you must assume that one or more components of your master mix are contaminated. Do not use results from that run for any quantitative or qualitative analysis. The appropriate response is:

1. Record the NTC result in your laboratory notebook or electronic records.
2. Discard all results from that run.
3. Identify and eliminate the contamination source before repeating the experiment.

Interpreting Weak or Late NTC Signals

A weak NTC signal (e.g., Cq > 35 in a 40-cycle qPCR run) is still a positive result and indicates contamination. The contamination may be at very low levels, but it is still present and could affect samples with low template concentrations. Some researchers mistakenly dismiss late NTC signals as "background," but this is incorrect—any detectable amplification in the NTC means that contaminating nucleic acids are present in your reagents.

Troubleshooting

Limitations and Edge Cases

NTC Does Not Detect All Contamination Sources

The NTC only detects contamination present in the master mix components. It does not detect contamination introduced during sample collection, nucleic acid extraction, or reverse transcription (for RT-PCR). To monitor for contamination during these earlier steps, you need additional controls such as extraction blanks (processed alongside samples but containing no biological material) and no-reverse transcriptase controls (for RT-qPCR).

NTC in Multiplex Reactions

When running multiplex PCR or qPCR with multiple primer-probe sets, the NTC should show no amplification for any target. If one target shows amplification in the NTC but others do not, the contamination is likely specific to that primer-probe set or its target. This can occur if primers or probes for that target have been contaminated with amplicons from previous experiments. Replace the contaminated primer-probe set and retest.

NTC with Degenerate or Universal Primers

Assays using degenerate primers or universal primers (e.g., 16S rRNA gene amplification) are particularly susceptible to contamination because these primers can amplify DNA from a wide range of organisms, including environmental bacteria present in reagents. Commercial Taq polymerase preparations have been known to contain trace bacterial DNA, leading to positive NTC signals even with careful technique. In such cases, you may need to use specialized "clean" DNA polymerases that have been treated to remove contaminating DNA, or include enzymatic digestion steps (e.g., with restriction enzymes) to eliminate contaminating templates.

NTC in Digital PCR

In digital PCR, the NTC should show no positive partitions. However, due to the extreme sensitivity of digital PCR, occasional false-positive partitions may occur. A common threshold is that the NTC should show fewer than 3 positive partitions out of 20,000 total partitions. If the NTC exceeds this threshold, contamination is present.

Documentation Requirements

Proper documentation of NTC results is essential for data integrity and reproducibility. For each PCR or qPCR run, record:

• Date and time of the run
• Operator name
• Master mix lot number and expiration date
• Water or buffer lot number
• Primer and probe lot numbers
• Thermal cycler or qPCR instrument used
• NTC result (Cq value for qPCR, band presence/absence for conventional PCR)
• Any corrective actions taken if NTC was positive

This documentation allows you to track contamination trends over time and identify recurring issues with specific reagents or instruments. The NIH Guidelines emphasize the importance of maintaining accurate records for all experiments involving recombinant or synthetic nucleic acids, including control results [2].

Biosafety Considerations

While NTC setup for PCR and qPCR typically falls within Biosafety Level 1 (BSL-1) practices, the BMBL provides important guidance that applies to all molecular biology work [1]:

• Use standard microbiological practices, including hand washing after handling potentially contaminated materials.
• Decontaminate work surfaces daily and after any spill of potentially contaminated material.
• Use mechanical pipetting devices; never pipette by mouth.
• Minimize the generation of aerosols, which can spread amplicon contamination.
• Segregate pre- and post-amplification areas to prevent cross-contamination.

For work involving recombinant or synthetic nucleic acids, the NIH Guidelines require that institutional biosafety committees review and approve the research, and that investigators follow appropriate containment levels [2]. Even at BSL-1, the use of PCR to amplify recombinant DNA sequences requires adherence to these guidelines.

Frequently Asked Questions

1. Can I use the same water for NTC that I use to dilute my samples?

Yes, and you should. Using the same water ensures that any contaminants present in your sample diluent will be detected in the NTC. However, if you use different water sources for sample dilution and NTC preparation, you may miss contamination in the sample diluent. Always use the same lot of nuclease-free water for both purposes.

2. How many NTCs should I include per plate?

At minimum, include one NTC per PCR run. For high-stakes assays (clinical diagnostics, GMO testing, forensic analysis), include duplicate NTCs placed in different locations on the plate. For multiplex assays with many targets, consider including one NTC per primer-probe set or at least two NTCs per plate. The more NTCs you include, the more confident you can be in detecting sporadic contamination.

3. My NTC shows amplification at cycle 37 in a 40-cycle qPCR run. Can I still use my sample data?

No. Any amplification in the NTC, regardless of how late it occurs, indicates that contaminating nucleic acids are present in your reagents. This contamination could affect samples with low template concentrations, producing false-positive results. You must discard all data from that run, identify the contamination source, and repeat the experiment with fresh reagents.

4. What is the difference between an NTC and an extraction blank?

An NTC monitors for contamination in the PCR master mix only. An extraction blank (also called a no-template extraction control) is a sample that goes through the entire nucleic acid extraction process alongside your biological samples but contains no biological material. The extraction blank detects contamination introduced during extraction, including from extraction reagents, columns, or laboratory surfaces. For comprehensive contamination monitoring, include both an NTC (for PCR reagents) and an extraction blank (for extraction reagents and procedures).

References and Further Reading

1. CDC and NIH. Biosafety in Microbiological and Biomedical Laboratories (BMBL), 6th Edition. U.S. Department of Health and Human Services, 2020. Available at: https://www.cdc.gov/labs/bmbl/index.html. This authoritative reference provides principles for risk assessment, containment, decontamination, and microbiological laboratory practice relevant to contamination control in molecular biology.

2. National Institutes of Health. NIH Guidelines for Research Involving Recombinant or Synthetic Nucleic Acid Molecules. NIH Office of Science Policy. Available at: https://osp.od.nih.gov/policies/biosafety-and-biosecurity-policy/nih-guidelines-for-research-involving-recombinant-or-synthetic-nucleic-acid-molecules/. This document establishes the institutional and biosafety framework for research involving recombinant and synthetic nucleic acids, including requirements for containment and documentation.

3. National Center for Biotechnology Information. NCBI Bookshelf: Molecular Biology and Laboratory Methods. Available at: https://www.ncbi.nlm.nih.gov/books/. This searchable collection provides authoritative biomedical books and methods references that support the molecular biology principles underlying PCR and qPCR controls.

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