This is a section containing a few Frequently Asked Questions (FAQs) for designs and application of our CRISPR genome engineering tools, as well as Trouble Shooting Tips when using our reagents and protocols as provided in this site. Please note all the reagents from our lab published in Cong et al., Science 2013 have  been deposited to Addgene (Feng Zhang Lab CRISPR Constructs). For any additional questions please feel free to post to our CRISPR Genome Engineering Discussion Group.



FAQs for Designing of CRISPR Genome Engineering Contructs and Targeting Experiments

A) Which version of backbone should I use for my genome engineering experiment, the chimeric (PX330) or the crRNA array (PX260)?
Answer: For targeting a single locus, we suggest using pX330 (chimeric RNA backbone). It should be more efficient than pX260 as we stated in our reagent page.

 

B) How should I design the oligos for simultaneous cleavage of two or more targets using the crRNA array backbone (PX260 or PX334) for multiplex genome engineering?
Answer: Let’s take a example for targeting two different genomic loci and design oligos for cloning into the crRNA array backbone. You should first define your two 30-bp protospacer targets with the proper ‘NGG’ PAM. Then order a pair of long oligos looks like:

5′ – AAACNNNNNNNNNNNNNNNNNNNNNNNNNNNNNN-DR-NNNNNNNNNNNNNNNNNNNNNNNNNNNNNNGT – 3′
and
 3′  NNNNNNNNNNNNNNNNNNNNNNNNNNNNNN-DR-NNNNNNNNNNNNNNNNNNNNNNNNNNNNNNCAAAAT – 5′

 

Where the DR sequence for S. pyogenes Cas9 we used is: GTTTTAGAGCTATGCTGTTTTGAATGGTCCCAAAAC. So the annealed product would be:

 

5′ – AAACNNNNNNNNNNNNNNNNNNNNNNNNNNNNNN-DR-NNNNNNNNNNNNNNNNNNNNNNNNNNNNNNGT – 3′
               3′ – NNNNNNNNNNNNNNNNNNNNNNNNNNNNNN-DR-NNNNNNNNNNNNNNNNNNNNNNNNNNNNNNCAAAAT – 5′

 

Which can be readily cloned into the target backbone vector PX260 or PX334. Note, for triple targets, you might need to order more than one pair of oligos, or maybe synthesize the insert with proper restriction enzyme sites for creation of the cloning 5′ overhang, then ligate into the digested backbone vector.

 

C) For plasmid pX260, 30bp target sequence is needed with a NGG PAM seq. Shall the NGG be exactly within the 3′ or immediately following the 3′ of this 30bp sequence?
Answer: Yes the NGG is located immediate next to the 3′ end of the 30bp sequence.

 

D) For plasmid pX330, 20bp target sequence is needed. In your illustration this 20bp sequence starts with a “G” and followed by a 19 nucleotides. Does it mean we shall always choose our target sequence starting with a G?
Answer: We recommend finding a 20bp genome target starting with the base ‘G’, due to the transcription initiation requirement of a ‘G’ base for human U6 promoter, as explained here in the reagent page.

If you have to use other bases at the starting position of your genome target, it might affect the transcription efficiency of the U6 promoter. However, in general we have been adding a ‘G’ to the target so effective forming a 21-bp target sequence with an additional ‘G’ base added to the 5′ initiation start position, and we have not observe any negative effects when using this design.

 

E) If I don’t find my desired targets in the USCS genome track provided in the Tools section, how could I design my own Cas9 target, especially if I would like to target a specific site within the genome of my favorite organism?
Answer: For application of Cas9 for site-specific genome editing in eukaryotic cells and organisms, we have computationally identified suitable target sites for the S. pyogenes Cas9. These sites are viewable as UCSC Genome Browser tracks for the human, mouse, rat, zebrafish, C. elegans, and D. melanogaster genomes.

Sites are selected such that the seed sequence for each SpCas9 target site, 5’-NNNNNNNNNNNN-NGG-3’, is specific to the relevant genome. because there are too many possible sites we only listed a few of all the sites we find using our algorithm. Essentially you will see from our website that we picked ngg PAM within a gene and BLAST the seed region of the targets to make sure it’s unique along the genome to guarantee specificity.

 

F) Besides the NGG and the general uniqueness of the target sequence, are there any special concerns we shall care about regarding the design of the targeting sequences?
Answer: We haven’t found any specific requirement for targeting sequence selection yet, but as you could see from our website, it’s advisory to blast the seed sequence within your target to ensure uniqueness if desired, see our Tools page here for more explanation.

 

G) Do you have a good sequencing primer for detecting whether the target sequence has been cloned in to backbone vectors, pX260/330, etc.?
Answer: Please use the U6 promoter primer as listed here. Human U6 Seq F_Insert: ACTATCATATGCTTACCGTAAC.

 

H) Is there a quick / easier way to check if the colonies on my plate (my clones) have the correct insert other than miniprep followed by Sanger sequencing?
Answer: Yes. There are several ways to check cloning success prior to sequencing verification. A very good way to check which of your colonies carry the correct insert (gRNA) in pX330/pX335 is to perform a BbsI/AgeI double digestion. As a successful insertion of oligos will destroy the BbsI sites, a double digest should discriminate between positive and negative clones. Clones with insertion will show only linearized plasmid of ~8.5 kb (only AgeI will be able to cut). Clones without insertion will show a ~1kb and ~7.5kb fragment (both BbsI and AgeI will be able to cut).

If you would like to perform direct colony digestion, one of the protocol we have is to use a kit called Colony FastScreen (restriction screen) from Epicentre, here is the link to the product.

G) If I want to introduce two cuts within the same genome to either introduce a deletion, or make sure there is a cleavage of target genomic locus, would you recommend using PX260 or PX330 (i.e., using the processing crRNA design or the chimeric RNA design)?
Answer: Please consider using PX330 for this type of ‘double-cut’ application through co-delivery of two PX330-derived targeting plasmids, as we have observed that this would be more efficient than using a single PX260-derived construct. Please refer to the reagents page for more information.

 

 



Trouble Shooting Tips for Cloning of CRISPR Genome Engineering Constructs

A) What should I do if I have no colonies on my plate following transformation?
Answer:
1. Please double check your oligo design as described in detail here: REAGENTS.
2. Make sure to use fresh (newly-ordered) BbsI enzyme, and also we have found that FastDigest BbsI(BpiI) from Fermentas seem to be very effective in supporting cleavage of backbone vector.
3. Please be sure to gel purified the digested vector carefully prior to ligation.
4. Please note the antibiotic resistance of the vector matches that of your plate.
5. Please check transformation efficiency of your competent cell using a positive control plasmid like pUC19.

 

B) I have a lot of colonies on the negative control plate.
Answer: Please first refer to previous answer to question A. Here are additional tips for this particular case:
1. Make sure the ligation was properly set-up with diluted annealed oligos, and the ligase kit you are using is as we recommended in the protocol, some people use overnight T4 ligation and sometimes this could lead to excessive amount of vector self-ligation, leading to higher background colony counts on the negative control plate.
2. Did you perform plasmid safe treatment?
3. Sometimes you might get a lot colonies on control but you still have very good success with the ligation plate, maybe try sequencing 4-6 colonies from your plate to check efficiency.

 

C) I don’t have or cannot set up a ramp-down protocol on my PCR machine, can I still anneal the oligos properly?
Answer: Yes, you could use the same set-up for Phosphorylate and then anneal by performing an incubation of the entire mixture at 95 degree for 5 minutes, then let the mixture gradually cool down under room temperature on the bench. This annealing protocol should work similarly to the one using a PCR machine. Another good way is to put your tube in boiled water (~ 100 degree Celsius) for 3-5 min, and then let the tubes cool down in the water bath under room temperature.

 

 

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