Developmental Biology
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Generating CRISPR/Cas9 Mediated Monoallelic Deletions to Study Enhancer Function in Mouse Embryonic Stem Cells
Chapters
Summary April 2nd, 2016
Experimental validation of enhancer activity is best approached by loss-of-function analysis. Presented here is an efficient protocol that uses CRISPR/Cas9 mediated deletion to study allele-specific regulation of gene transcription in F1 ES cells which contain a hybrid genome (Mus musculus129 x Mus castaneus).
Transcript
The overall goal of this procedure is to use allele specific CRISPR/Cas9 mediated deletion to study regulation of gene transcription in mouse embryonic stem cells. The main advantage of this technique, which uses a hybrid genome to generate and analyze monoallelic deletions. Instead, it eliminates the need to analyze monoallelic enhanced deletions that may not be viable.
This method can help answer key questions in the functional genomics field. Such as, which gene does a specific enhancer regulate and how dependent is that gene on any one enhancer for its activity. Those this method can provide insight into stem cell pluripotency, it can also be applied to other systems.
For example, it can help us to understand how stem cells change their fate to become neurons, blood cells, or muscle. After designing, constructing, and confirming the sequence of the guide RNA, or gRNA, to be used, prepare to transfect the endotoxin-free plasmids into ES cells. Using a centrifuge, pellet one times ten to the sixth F1 embryonic stem cells generated by crossing Mus musculus 129 with Mus castaneus in a 1.5 milliliter tube at 300 times g'for five minutes.
After the spin, discard the supernatant and add 100 microliters of the resuspension buffer supplied by the kit manufacturer. For deletion of the target DNA region, add five micrograms each of P Cas9 GFP 5-prime and 3-prime gRNA plasmids. Using a pipette, gently mix.
Avoid introducing any bubbles. Next, use the electronic pipette tip of the microporator to aspirate 100 microliters of the electroporation mix, taking care to avoid introducing air into the tip. Then, plug the pipette into the transfection device.
Select the appropriate protocol and press start. While, the electroporation is running, observe the tip. With the correct set-up, tiny bubbles should emerge from the tube electrode.
A spark indicates the presence of an air bubble and will interfere with the transfection. If this occurs, the electroporation must be repeated with a new batch of cells. Following the electroporation, eject the transfected cells into a ten centimeter gelatin coated dish containing ten milliliters of ES cell medium.
Incubate the transfected cells at 37 degrees Celsius, 5%carbon dioxide. After 48 hours, use a flow cytometer to sort and isolate the cast nine GFP positive ES cells. Then, feed to 1 to 1.5 times 10 to the fourth GFP positive cells in a 10 centimeter, gelatin-coated dish containing 10 milliliters of ES cell medium.
Plating at this low density will facilitate picking up individual ES cell colonies. Four to five days later, use a microscope to identify individual ball-shaped, single ES cell colonies. Using a pipette transfer each colony to one well of a 96-well plate pretreated with gelatin and containing 30 microliters of tripsom.
Once all of the colonies have been picked and dissociated into medium, grow the cells at 37 degrees Celsius in the carbon dioxide incubator until most of the wells are over 70%confluent. This usually takes about two days. When the cells are ready for splitting, remove the medium and add 30 microliters of tripsom.
Incubate the cells for five minutes. Then, to each well add 180 microliters of ES cell medium to neutralize the tripsom. Pipette up and down to achieve a single cell suspension.
From each well, transfer 70 microliters to each of three gelatin-coated 96-well plates containing 130 microliters of ES cell medium in each well. Incubate at 37 degrees Celsius. When the plates reach 70 to 85%confluency, use them to perform genotyping and expression analysis of each clone, as described in the sections that follow.
Freeze the cell stocks as described in the accompanying document. Here, four sets of primers will be designed to screen the clones for the desired deletion. These include:inside primers, outside primers, and gRNA flanking primers for both 5-prime and 3-prime gRNA target sites.
Begin by visiting the website shown here to obtain the single nucleotide polymorphism or snip track corresponding to the 129 and Cas genotypes. This directs to the Mitchell Lab CRISPR webpage. There, click on the given link.
The site will redirect to the UCSC genome browser containing the track displaying snips between the 129 and Cas genomes in the MM-9 mouse genome assembly. Next, enter the coordinates of the region to be deleted. and click on go"Zoom in on a region of about 500 base pairs in the middle of the desired deletion containing more than three snips.
Allele-specific primer design can be complicated if snips are scarce. If it's not possible to design absolute allele-specific primers, moderate allele-specific primers may be used. Just remember to compare the CT values between the deleted and non-deleted clones during analysis of the qPCR results.
Now, click on View"in the top panel. From the drop-down menu, select DNA. The Get DNA in Window"page will appear.
In the sequence formatting options, choose all uppercase, then click on Get DNA"to display the target sequence in all uppercase format. Copy this sequence and past it into a Word document file. Then, compare the sequence with the 129 cast snip track and highlight the snip positions and mark the 129 snips with the lowercase.
When you are finished with the 129 sequence, copy it to create two fast day formatted sequences. One for 129 and one for Cas. In the cast sequence, substitute the correct base at each snip position, marking the snip with a lowercase.
Next, label the sequence with its corresponding genome, 129 or Cas. Then, in the browser, type in the address for Primer3Plus. In the appropriate area, paste the snip substituted 129 sequence.
Use the default settings to design the primers. To design the inside allele-specific primers, click on the drop-down menu to the right of Task"and select Primer List"the click on Pick Primers. A new page will open displaying a list of forward and reverse primers.
Check the select box to choose a forward or reverse primer that has snip marked by a lowercase either at the 3-prime end or within four bases of the 3-prime end. Note the primers that have a snip at the 3-prime end display increased allele specificity during qPCR. Click Send to Primer3 Manager"at the bottom of the screen.
A new page will open displaying the selected primer. To select the second primer, return to the primer list page, and click the back button to return to the main page. In the task drop-down menu, select Detection.
The main page will refresh and a left and right primer sequence box will appear at the bottom. Paste the first primer sequence in the appropriate box. Next, navigate to the General Settings tab located next to the Main tab and change the setting for the product size range to 80 to 200 base pairs.
Click Pick Primers"A page will open displaying five possible primer pairs for the first primer sequence. Choose a primer set from the list of primer pairs by checking the boxes to the left of the primers and click on Send to Primer3 Manager"at the bottom of the screen. These will be the 129 inside allele-specific primers.
Repeat these steps to design primers for the cast allele. Then, apply this technique to the design of outside allele-specific primers and non-allele-specific primers. Finally, test the primers for allele specificity using qPCR as described in the accompanying document.
Extract the genomic DNA from the 96-well plate prepared earlier. Centrifuge the plate briefly to settle any condensation to the bottom of the well. This DNA will serve as the template for deletion screening.
Next, on a 384-well plate, preform qPCR and duplicate for each clone. Use a two step PCR followed by melt curve analysis with detection. Following the run, first check each allele for amplification with the inside allele-specific primers.
No amplification of one allele or CT value differences of more than five cycles between alleles, suggests these clones carry a heterozygous deletion. No amplification of both alleles suggests that they carry a homozygous deletion. Next, check each allele for amplification with the outside allele-specific primers.
If the target deletion is larger than one KB, amplification with outside primers will only occur if a deletion is present. A CT value of 22 to 28 confirms the deletion. For target deletions smaller than one KB, confirm the amplicon size by eletrophoresis.
A 96-well plate of deletion clones was screened by qPCR using allele-specific inside primers. Gray bars represent from Cas-specific primers and black bars represent amplification from 129-specific primers. A low CT value or no amplification with inside primers indicates target deletion.
Clones with a deletion on one or both alleles are then screened with the outside primers. With outside primers, amplification confirms the deletion. Clones with a monoallelic deletion are screened using primers flanking the gRNA target regions to confirm the absence of insertions or deletions, called indels, on the non-deleted allele.
Only monoallelic clones without large indels at the target sites on the non-deleted allele are used in the subsequent expression analysis. Shown here are representative results from 129 or Cas SCR deleted clones. The SCR is a recently described Sox2 specific enhancer in ES cells.
Red bars represent expression of the Sox2 Cas allele, and blue bars represent amplification of the Sox2 129 allele. As can be seen in this figure, deletion of the SCR on the 129 or Cas allele reduced expression of Sox2 in CIS. After watching this video, you should have a good understanding of how to generate a large deletion using CRISPR technology and use allele-specific primers to genotype your deleted clones.
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