Sanger sequencing, also known as chain-termination sequencing, is a method used to determine the exact order of nucleotides in a DNA molecule. Developed by Frederick Sanger in 1977, it remains the gold standard for validating DNA sequences and detecting mutations.

How Sanger Sequencing Works

1. Setting Up the Reaction

The DNA to be sequenced is mixed with a DNA primer, DNA polymerase, normal nucleotides (dNTPs), and a small amount of fluorescently labeled dideoxynucleotides (ddNTPs). Each of the four ddNTPs is labeled with a different fluorescent dye.

2. Chain Termination

The reaction starts with the primer binding to the template DNA. The DNA polymerase extends the primer by adding dNTPs. Whenever a ddNTP is incorporated instead of a dNTP, the chain stops growing because ddNTPs lack the 3’-hydroxyl group needed for further extension.

3. Fragment Generation

This process produces a mixture of DNA fragments of varying lengths, each ending at a specific nucleotide position. The fluorescent label on the terminating ddNTP identifies which base is at the end of each fragment.

4. Capillary Electrophoresis

The mixture is loaded into a capillary electrophoresis instrument, where size-based separation is performed by capillary gel electrophoresis. The fragments migrate through a polymer matrix within the capillary and are detected by laser-induced fluorescence as they pass the detection window. A laser excites the fluorescent labels, and a detector records which color passes by at each time point.

5. Chromatogram Analysis

The instrument produces a chromatogram—a series of colored peaks representing the sequence of nucleotides. The sequence is read from the chromatogram, typically from the shortest fragment to the longest, giving the complete DNA sequence.

Practical Sanger Sequencing Workflow

Prepare the template DNA by purifying the PCR product or plasmid using a column cleanup kit. For plasmid sequencing, use 200–500 ng of purified plasmid; for PCR products, use 1–10 ng per 100 bp of amplicon. Set up the cycle sequencing reaction in a 10 µL volume: 2 µL of BigDye Terminator v3.1 Ready Reaction Mix, 2 µL of 5× sequencing buffer, 1 µL of primer (3.2 pmol/µL), template DNA, and water to 10 µL. Use the following thermal cycling program: 96°C for 1 minute, then 25 cycles of 96°C for 10 seconds, 50°C for 5 seconds, and 60°C for 4 minutes. Purify the extension products by ethanol precipitation or a magnetic bead cleanup to remove unincorporated dye terminators. Load the purified sample onto a capillary electrophoresis instrument (e.g., Applied Biosystems 3730). The run separates fragments by size, and a laser excites the fluorescent dyes as fragments pass the detection window. Examine the resulting chromatogram using software such as FinchTV, SnapGene, or Chromas. Good-quality data typically extends 600–900 bases from the primer. Check for clean, evenly spaced peaks with baseline resolution. Single nucleotide polymorphisms (SNPs) appear as two overlapping peaks at the same position. Poor quality at the beginning (first 20–40 bases) and end (after 800 bases) is normal — trim these regions before analysis. Troubleshoot failed reactions by increasing template, redesigning primers with Tm of 60–65°C, or adding DMSO (2–5%) for GC-rich templates.

Real-World Application

To confirm a CRISPR-edited gene in mouse embryos, the target region is PCR-amplified and Sanger sequenced. The chromatogram shows mixed peaks near the cut site, indicating indels. ICE analysis decomposes the trace and reports 72% editing efficiency with a 1 bp insertion as the most common allele. Sanger sequencing remains the gold standard for validating genome edits due to its accuracy and cost-effectiveness.