I transfected Cas9 expressing cells with two sgRNAs targeting two different genes (separately, not co-transfection). Both proteins show significant Knockdown in Western Blot, indicating that the guides are working. I extracted genomic DNA from these cells and PCR amplified the target region. However Sanger Sequencing of the PCR amplicons yielded a clean trace, with no aberrant signal observed near the target site. So, TIDE is unable to predict the efficiency of my sgRNAs. Has anyone come across similar issue? What could be the reason for getting a clean chromatogram from a mixture of sequences?
1. Try Dhrmacon tool for sgRNA designing. it works fine for me.
2. Also, might be the sgRNA you have used is not very efficient. I believe one should always start with 2-3 sgRNA/gene to achieve the highest knocked-out clone among the 2-3 guides used.
3. Lastly, sometimes the knock-out effect is observed in western due to low protein translation happening in the cell but still on the gene level, sanger is unable to show that editing in a pool v=where diverse population exists. Try NGS and you have the clear picture.
4. You can also try serial dilution of the edited pools to achieve unique population of the edited cell which will give better sanger analysis compared to pools.
Hope this helps. All the best
Dear Sruthi and John,
Thank you for the answers. I'm certain it is not an issue of the efficiency of sgRNAs or their ability to get into the nucleus. The KD is around 70% for both, indicating they are both highly efficient sgRNAs. With one of the sgRNAs, I have already performed clonal isolation and the percentage of KO clones is quite high (based on Western Blot).
@John, yes the PCR should in theory amplify both wild type and non-WT DNA, meaning you should expect a mixture of sequences near the target site, which results in aberrant signal in Sanger Sequence. This is the principle on which TIDE (Tracking of Indels by DEcomposition) works, it uses the aberrant signal in the chromatogram to compute an indel spectrum and estimate sgRNA efficiency. However, the problem seems to be that I cannot observe the aberrant signal, even though every evidence points that the underlying population is a mixture.
Dear Ayon, maybe I have found your question too late and you have solved the problem long ago. But I'd like to express a guess, that maybe the absence of the abbreant signal after Sanger sequencing is caused by your editing yelding long deletion with the loss of primer annealing site? Thus, PCR yelds only wild-type products. TIDE in absence of abberant signal, obvious on sequencing chromatogram will not find any mutations. I usually look through the chromatogramm files to see if there are a clear signal for alignment and an abberant signal for decomposition before feeding the files to TIDE.
Please, write, if you have found the solution.
TIDE (Tracking of Indels by DEcomposition) is a computational method developed to analyze the efficiency and specificity of CRISPR/Cas9-induced mutations, such as insertions and deletions (indels), within a targeted genomic region. It is particularly useful for assessing the outcome of gene editing experiments in a simple and cost-effective manner, using standard Sanger sequencing. Here’s how you can use TIDE for your CRISPR/Cas9 experiments:
1. Perform CRISPR/Cas9 Experiment and Collect Samples
- Design and deliver CRISPR/Cas9 components (guide RNA and Cas9) to your cells of interest to induce targeted mutations.
- After a suitable period allowing for gene editing, collect samples from your edited cell population.
2. PCR Amplification and Sanger Sequencing
- Extract genomic DNA from your edited cell population.
- Use PCR to amplify the region surrounding your CRISPR target site.
- Purify the PCR product and subject it to Sanger sequencing. It's advisable to sequence both untreated (control) and treated samples for comparison.
3. Use TIDE to Analyze Sequencing Data
- Access the TIDE web tool (available at the TIDE website). The tool is user-friendly and does not require advanced computational skills.
- Upload the Sanger sequencing trace files (usually in .ab1 or .scf format) for both your control (non-edited) and edited samples.
- Input the details of your CRISPR target site, including the genomic location and the sequence of the guide RNA (gRNA).
- The TIDE algorithm will decompose the sequencing traces into a mixture of indels present in the population and estimate the frequencies of each mutation type.
4. Interpret Results
- TIDE provides a detailed report, including:The efficiency of indel generation (i.e., the percentage of sequences with mutations). The types of indels introduced (insertions or deletions) and their distribution. A graphical representation of the mutation spectrum.
- These results help in assessing the editing efficiency of your CRISPR experiment and understanding the mutation landscape induced by the CRISPR/Cas9 system.
5. Advantages and Limitations
- Advantages: TIDE is a straightforward, cost-effective method that doesn’t require deep sequencing. It’s suitable for evaluating the efficiency of CRISPR-mediated editing and the pattern of induced mutations.
- Limitations: TIDE is best suited for analyzing clonal cell populations and might not accurately represent the mutation spectrum in highly heterogeneous samples. It also primarily detects indels but not other types of mutations like large deletions or base substitutions.
Conclusion
TIDE is a powerful tool for researchers conducting CRISPR/Cas9 gene editing, offering insights into the efficiency and outcomes of their editing efforts. By following these steps to perform your experiment and analyze your data with TIDE, you can obtain valuable information about the mutations introduced in your target gene, helping to guide further experiments and applications of CRISPR technology.
l Check out this protocol list; it might provide additional insights for resolving the issue.
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