Skip to content

Cycle Sequencing

After you’ve picked colonies and completed your minipreps, you’ve finished the fabrication phase. The next question: what did you actually make? This tutorial will guide you through how to determine if your plasmid contains the correct sequence.

What Is Cycle Sequencing?

Cycle sequencing, also often called Sanger sequencing, uses DNA polymerase and chain-terminating nucleotides (ddNTPs) to create truncated DNA fragments, each ending in a labeled base. These fragments are then separated by capillary electrophoresis to reveal the DNA sequence.

Diagram showing plasmid denaturation, primer binding, extension with ddNTPs, and detection by capillary electrophoresis.

Figure: Cycle sequencing overview. Fluorescently-labeled ddNTPs stop extension at every base type. The resulting DNA fragments are separated and analyzed.

Why Not Just Sequence the Whole Plasmid?

Full-plasmid sequencing services (e.g., Plasmidsaurus) cost ~$15/sample and return the entire sequence. This is useful in some cases, but often overkill. In the pP6 experiment, we only care about a small variable region between two restriction sites. A single cycle sequencing read (~$3.50) is sufficient.

Cycle Sequencing Details

In cycle sequencing, only one primer is used, which means the amplification is linear rather than exponential like in PCR. The read typically starts about 20–50 bp downstream of the primer, and you'll usually get around 400–1000 bp of usable sequence, depending on the quality of the reaction.

Each ddNTP is fluorescently labeled with a distinct color. After PCR-like cycling, the products are run through capillary electrophoresis.

Diagram of a polyacrylamide sequencing gel with bands in A, C, G, T lanes and the corresponding base calls listed beside the gel. Figure: Example of a traditional sequencing gel, where each lane contains fragments ending in a specific base. The band pattern reveals the sequence of the DNA strand. This image is adapted from Edvotek’s excellent DNA sequencing tutorial, which we recommend if you’d like a deeper dive into how the chemistry works.

From Data to Interpretation

From the sequencing facility, you get:

  • A .txt file containing the base calls (the 'read')
  • An .ab1 chromatogram file (shows raw fluorescent data), the 'trace'

You can view both in ApE or Benchling. ApE will also annotate known features if you hit ctrl-K with a feature database installed.

Step-by-Step: How to Sequence Your pP6 Clone

Choosing a Primer

The sequence you read will start ~20–50 bp downstream of your primer. So, you must choose a primer upstream of the region you want to check. For pP6, we use the standard primer G00101.

When to Use Other Sequencing Options

Method Cost/sample Output Use Case
Cycle sequencing ~$3.50 ~1 kb from a primer Quick check of a region
Full plasmid ~$15 Entire plasmid Final confirmation, mutation scan
Deep sequencing $750+ Millions of reads Libraries, high-throughput work

What to Do With Your pP6 Sequencing Result

Once your sequencing data is returned (usually in 1–2 days), you will:

1. Download your result

2. Check for the Expected pP6 Architecture

Before comparing your entire sequence to the model, begin by checking for key features within your read:

🔍 Step 1: Look for the BseRI → Promoter → BseRI Pattern

This is the core region you’re sequencing. You're looking for the structure:

BseRI → variable promoter region → BseRI

If this pattern is intact, your clone is a candidate hit. If not, it’s likely not usable, but it may still be worth investigating as an example of unexpected outcomes. Reads from this experiment sometimes contain odd duplications, deletions, or recombinations due to the N-rich promoter region.

🧬 Step 2: Use ApE to Confirm Key Features

Open your .seq file and pP6.seq in ApE. You can download the model file here: 📄 pP6.seq If your feature database is installed, hit ctrl-K to light up key landmarks such as:

  • BseRI restriction sites (you should see exactly 2)
  • consensus promoter pattern: NNNNTTGACANNNNNNNNNNNNNNNNNTATAATNNNNNNANNNN
  • T4 terminator

Use the feature list at the top of ApE to verify that all these landmarks are present, appear once, and are in the correct order.

🧪 Step 3: Align the Read to the Model

To verify flanking sequence accuracy: - Select your sequence in ApE - Go to Tools → Align with another sequence... - Choose pP6.seq - Look for 100% identity near and around the promoter region

A clean alignment confirms no point mutations or context disruption.

✅ Quick Checklist

  • [ ] Contains two BseRI sites
  • [ ] Includes the UBER promoter motif between them
  • [ ] Promoter is not duplicated, reversed, or truncated
  • [ ] Contains expected T4 terminator site
  • [ ] Alignment shows clean sequence on both sides

Use this structure to decide if your clone is usable or just interesting. Add your findings to the worksheet in the next step.

3. Confirm if your clone is a good read

  • Is the read clean, free of noise or ambiguous calls?
  • How long is the high-quality portion with no N's — 100 bp (low quality), 800 bp (good), 1000 bp (great)?

A high-quality read should give you several hundred bases of clean, mappable sequence. The more of the surrounding context you can confirm, the better.

4. Search for the target motif

  • Look for this key motif in your read: 
    GAGGAGTCCTGGGTTCNNNNTTGACANNNNNNNNNNNNNNNNNTATAATNNNNNNANNNNGTTAGTATTTCTCCTC
  • If it's found and the read is clean, mark your clone as usable.

5. Fill out the worksheet

Go to:
📝 pP6 Clones Worksheet

For each clone that has a clean and analyzable read, enter the following information:

  • clone_id — Your assigned clone label (e.g., 79A)
  • read_name — The filename of your .seq file (e.g., 62-pP6-14B_F08_054.seq)
  • date_sequenced — The name of the sequencing folder (e.g., 2022_04_24)
  • canonical — Mark "yes" if the read matches the model sequence (pP6.seq) across the entire promoter region with no mutations or rearrangements. Otherwise, "no".
  • usable — Mark "yes" if the expected UBER promoter motif is found and intact, even if the rest of the plasmid has issues. Otherwise, "no".
  • cassette — Paste the actual sequence you matched that corresponds to the expected promoter region.
  • Notes — Summarize what you observed.

🧪 Example Annotations

clone_id canonical usable cassette Notes
14A yes yes GAGGA...CTC Perfect match
14B no no Pcon region is shortened, no UBER present
14C no yes GAGGA...CTC Additional BseRI sites included, but Pcon site is fine
14D no no Contamination, matches pTP1
14E no yes GAGGA...CTC Extra BseRI and BsaI sites, but promoter is fine

Remember: usable means the promoter is intact and could be moved forward into our development pipeline. Canonical means it's a perfect match to what we designed.

You don’t need perfection to keep a clone — but you do need to understand it.

6. Close out the experiment with your supervisor

  • Discard cleanup DNA and used plates
  • Discard clones with bad reads
  • Clean and bleach culture block
  • Confirm image/data uploads
  • Move good clones ("hits") to TPcon6B box
  • Your pP6 work is complete when hits are logged and uploaded

🎉 That’s it! You’ve finished the pP6 experiment!

But there's one final question: we know the promoter works and we've seen that it's green — but how strong is it, exactly?

The pP6 hits you've found vary a lot in brightness. To move from a qualitative observation ("looks bright") to a quantitative measurement, your next tutorial — BestP — will walk you through how to assess promoter strength using fluorescence activity assays.

🧪 Quiz: Sequencing

1️⃣ Why Sequence?

Which of the following is not a reason to sequence your pP6 clone?





2️⃣ Usable Clones

What makes a clone “usable”?





3️⃣ Feature Identification

What ApE feature helps you quickly check for key sequences?





4️⃣ What to Look For

What are you primarily trying to verify in your sequencing read?