About using DNA or RNA for mouse embryo #CRISPR injections.

I got a question:

Isn't the disadvantage of injecting DNA the threat of integration and more frequent mosaicism than in the case of RNA as Cas is expressed quicker? Do you have some direct experience with that? Thanks! 

Um, well yes.  Yes.  Those are the disadvantages.  Also I will add that because the RNA should lead to quicker Cas9 expression,  mutagenesis efficiencies will likely be higher than with DNA vectors.

So why use DNA at all?  Well, the issues are mostly practical.  DNA vectors are easy to customize for CRISPR.  Although the issues of efficiency and mosaicism are potentially problematic, I have seen pretty consistent success* in generating simple indel mutations following injections of PX330-style CRISPR-Cas9 DNA plasmids.  That is, consistent double digit percentages of founders carrying mutations as assayed by PCR and/or sequencing.    In addition, in our core we have obtained HDR-mediated codon editing rates in the 10-20% range using PX330 vectors co-injected with appropriate "donor" oligos.  But this is dependent on cooperative CRISPR sites that have a high rate of baseline cleavage.

Another practical consideration is that not everyone can routinely synthesize high-quality RNAs in vitro with consistency.   Quality DNA is relatively easy to prepare and QC.   RNA is much less so - especially for the 4+ kilobase Cas9 mRNA.   OK, so some of you are saying "Come one, my lab makes RNAs all the time - no prob! " .    That's awesome, but the empirical observation is that it's not trivial to get proficient at making long mRNAs, and to keep on top of the key reagent issues (RNAses, enzymes going bad, etc.).

Also, CRISPR DNA plasmids are immediately useful for cell culture gene editing experiments.  Some labs will be making these anyway so they will have them on hand, ready to go.   

What I am also observing - which many others have reported - is that a fraction of CRISPR sites just don't cut very well, even when the sequence characteristics of the site seem OK.  (Like, somewhere on the order of 1/3 to 1/4 of CRISPR target sites?) Most of our injections to date have been using DNA plasmids.   It's possible that RNAs might save the day for some of these sites.    

The ability to do precise HDR-mediated editing/insertions, rather than simple indels, is very compelling and is the direction most of our CRISPR ideas are going in terms of new mouse models.  But coding modifications usually have extremely narrow CRISPR target choices that are imposed by the science;  if you want to change a codon, you'll probably need a target as close as possible - preferably overlapping the codon.  There won't be many to choose from.  So getting the highest efficiency cleavage rates may be critical for some of these projects - for these, Cas9 mRNA or protein may be needed.

Finally, these issues of target efficiency really call for pre-validation of sites.  This can be done by transfecting CRISPR plasmids into cooperative cell lines, e.g. NIH3T3 for mouse targets, followed by PCR and mismatch cleavage assays, which can then be quantified.  But then - if you go through the trouble to do that, you will have generated the DNA plasmids and thus have the DNA reagent ready for injection.   

Having said all that, although I really like the convenience of plasmids, the RNA problems are all about sourcing them.  A few vendors, such as Sigma-Aldrich can provide custom guide RNAs and Cas9 mRNA that work.  (FYI I do not receive any compensation from Sigma).  The RNA reagent expense is less than the cost of mouse embryo injections.  I suppose zebrafish researchers may balk at the cost, as they will have more capacity to inject fish eggs, in their own labs usually, and may be more willing to make RNAs in-house.  For mice, you'll be usually working with a transgenic core and spending thousands of bucks per experiment.  Vendor-supplied RNAs may be worth the money.    Thanks for the question!

*Actually, "consistent" may be misleading… To clarify, about 75% of the NHEJ projects I've been observing have had this level of success.   So - more success than not, but then again, not perfectly consistent.  

TIDE: an online tool for evaluating #CRISPR gene editing in sequence trace files.

TIDE is a neat new web tool that's designed for a specific problem that I've definitely been dealing with. Following a CRISPR experiment, either in cell lines or animals, it's not trivial to quantify how well editing/mutagenesis worked and what sort of mutations were generated.   This is well summarized in the introduction of this paper so I won't repeat that, but I have certainly had these situations:  first, staring at ABI chromatograms following sequencing of PCR products from founder mice, and second, trying to quantify cleavage in pools of transfected cells.     Of course, the target site PCRs are going to usually contain mixtures of molecules with different mutations, and likely some amount of wild-type allele (for sure in pooled cells, often in founder animals).   So direct sequencing is hard to interpret as the actual chromatogram data past the cleavage site is usually a jumble of overlapping staggered sequences.

What TIDE does is actually to quantify the underlying non-wild-type sequence signal in the chromatogram data 3' to the expected cleavage site, then it quantifies the apparent contribution of specific, underlying mutant alleles, based on the pretty good assumption that most of the mutations generated by CRISPR will be short indels.  This seems to be a extension of PolyPeakParser, which I blogged about previously, but it's able to deal with multiple mutant alleles.  

I had a recent data set of sequence files from a mouse CRISPR experiment, so I thought I'd compare our independent analysis of the founder mice to TIDE's interpretation.  The gene is anonymized but I can state that it was a straightforward attempt to create indel mutations in a gene of interest.    Here's what we did:   About 25% of pups were positive for new mutations as revealed by Surveyor assays.  We then sequenced PCR products on 8 founder littermates, of which 6 were known Surveyor-positive and 2 were of unknown status.    

The last 2 (#19, #20) mice had normal, wild-type sequencing data.   The other 6 mice had very jumbled sequences past the cleavage site.   After some serious staring at the chromatograms - which took a while - I made some guesses that some of them had specific indel mutations.  However some of them were just too complex for me to figure out.      

Then I analyzed all of them with TIDE, using the sequence file from wild-type mouse #20 as the control file (which TIDE requires).   Here's the results:

Pup #	Pre-TIDE manual interpretation	TIDE result
2	WT allele and at least 2 different mutant alleles present. Could not interpret mutations at all.	No significant results, but the sequence quality was rather poor to begin with.
6	WT allele and a 1-bp deletion allele. Germline transmission confirmed.	66.5% WT, 24.8 % 1-bp deletion.
10	No WT allele; one 3-bp deletion; plus a complex (discontiguous)  4-bp deletion.  Germline transmission confirmed of both alleles at essentially mendelian rates.	10.9% WT, 44.9% 3-bp deletion, 33.7% 4-bp deletion.
16	WT allele and 2 different mutant alleles present. Could not interpret mutations.	22.5% WT, 58.5 % 1-bp deletion, 9.7% 5-bp insertion.
22	WT allele and a 1-bp deletion allele.  Germline transmission confirmed.	60% WT, 30.2% 1-bp deletion.
24	No WT allele, but multiple (>3) mutant alleles.	At least 4 different deletions of -2, -12, -28, -29 bp, each at low levels.
19	WT allele predominates.	75% WT;  7.4% 2-bp insertion; 8% 8-bp deletion.
20	WT allele predominates.	(Used #19 as control) 82.8% WT, 10.7% 4-bp deletion.

I was fairly impressed by the TIDE results.  First, it agreed with my specific interpretations for #6, 10 and 22, which were actually confirmed by germline transmission.   Second, it was able to correctly call 2 mutations at the same time in mouse #10.   Third, it made interpretations that made sense for founders #16 and 24, which I had given up on.    

Finally, I didn't really give the algorithm the optimal control sequence.  Instead I used the file for an apparently wild-type founder animal (#20).  However - when the files from #19 and #20 were used as controls to analyze each other, low levels of mutant alleles were detected.  And yes, if you go back to the chromatograms you can see a little underlying signal that may be a bit more than "usual" past the cleavage site - but it's very easy to miss.   This result is actually consistent with the experiment, since the embryos were injected with a PX330 plasmid, which may persist past the 1-cell stage and thus lead to low levels of mosaicism.      

Based on the imperfect controls I used, I would not take the TIDE quantitation of allele fractions literally.   However the qualitative results were pretty good and I wasn't able to find anything manually that TIDE didn't.  Also, this is a very fast analysis if you are performing sequencing on the PCRs anyway.   Moreover, it's easy to apply this analysis to PCRs on transfected pools of cells to measure CRISPR mutagenesis.  I'm looking forward to trying TIDE in this context as well.

Easy quantitative assessment of genome editing by sequence trace decomposition.  Eva K. Brinkman, Tao Chen, Mario Amendola and Bas van Steensel.    Nucleic Acids Research, 2014, Vol. 42, No. 22 e168

Correction to my post "Protocol for cloning protospacer adapters...PX330-family #CRISPR plasmids".

There was an error - now corrected - in my previous protocol post  "Protocol for cloning protospacer adapters into Zhang lab PX330-family #CRISPR plasmids.":  I omitted by mistake the instruction to add the ligase enzyme to the ligation reaction. (line II - 2 - e in the protocol).  Duh!    I apologize profusely.

Current thoughts on targeting reporter cassettes in mice using #CRISPR.

 As an advocate of the PX330 plasmid for implementing CRISPR in mouse zygotes, I am trying to figure out optimal conditions for enabling homology-dependent repair (HDR) with this system.  Genome editing requires HDR, and a donor DNA molecule must be supplied along with the CRISPR/Cas tools such as PX330.  

For introducing "smallish" edits or insertions - like, under 70 bases or so - one can order a single-stranded super-long oligo such as IDT's ultramers.  Larger insertions are going to require double stranded DNA fragments.  Most molecular biologist are well used to purifying these, once they are designed.   Here's a few considerations:

1.  If the CRISPR target is also present in the homology arms of a double-stranded donor, it's probably gonna cut the donor before it gets a chance to donate anything.

2.  What should the optimal ratio of donor/CRISPR/Cas9 be?    I am assuming roughly a 1:1 mass ratio.   My only guidance so far is from the Yang et al 2013 Cell paper.  They were coinjecting donor DNAs with RNAs for guide RNA & Cas9.   Here I will focus on their data for fluorescent reporter fragment insertions, which were injected as circular dsDNA plasmids.   Although this group mainly does cytoplasmic injections, they tested pronuclear injections too.

For pronuclear injections of these reagents. Yang et al  reported 9% and 18% targeting rates, using two different donors and targets.    The concentrations were:  10 ng/µl donor plasmid, 2.5 ng/µl guide RNA, 5 ng/µl Cas9 mRNA.    Thus the total nucleic acid burden of the pronuclear injection material was ~17 ng/µl.   Those with actual experience with mouse embryo injections will note this is rather high;  most DNA transgenes are injected at 1-5 ng/µl, no more.  Why no more?   It can cause toxicity, but even more often and more frustrating, it leads to frequent needle clogging at the injection microscope.   Injected material at these concentrations will need very careful preparation beforehand.  I suggest that a 0.22 micron spin filter be used before each injection over a total 5 ng/µl limit.  Anyway, a ratio of ~1:1 or maybe ~3:2 mass ratio of donor DNA vs. CRISPR reagents seems reasonable.    For PX330 I am starting off by suggesting 4 ng/µl PX330 + 6 ng/µl donor plasmid - although I have no data yet showing whether this is optimal.

Protocol for cloning protospacer adapters into Zhang lab PX330-family #CRISPR plasmids.

This is also publicly accessible at the Vanderbilt labnodes website.

Cloning protospacer adapters into PX330/PX335/PX458/PX459 (Zhang lab, MIT) via BbsI site.

I.               Anneal top and bottom oligos of adapter

1.     Resuspend oligos in water to 1 µg/µl concentration.

2.     Combine the following in a 200 µl PCR tube:

a.     5 µl “top” oligo

b.     5 µl “bottom” oligo

c.     10 µl of 10x annealing buffer

d.     80 µl water

(10X Annealing Buffer: 1 M NaCl / 10 mM EDTA pH 8.0 / 100 mM Tris pH 7.5)

3.     Mix well.  Run the mix in a thermal cycler with an annealing program, such as:  Step (1) 94˚ for 3 minutes; Step (2) cool from 94 ˚ to 25˚ slowly, such as over a 30 minute period.  Alternatively to a thermal cyler:  Heat a beaker of water to boiling. Float the tube in the boiling water bath for 5’. Remove the beaker from the heat and let it cool off naturally on a benchtop, till it is room temperature.

4.     Transfer the annealed adapter to a 1.5 ml tube.  Add 900 µl water.  The final concentration of annealed adapter is now ~ 10 ng/µl.

II.              Ligate to BbsI-cut vector.

1.     Before you start:  You will need to have the vector DNA previously cut with BbsI and gel-purifed, and in 10 mM Tris or Lo-TE at a concentration of at least 10 ng/µl.

2.     I use the NEB Quick Ligation kit, and I reduce the volumes by half to save reagents.  It works great.  Combine the following in this order:

a.     2.5 µl of 10 ng/µl BbsI-cut vector

b.     1 µl of 10 ng/µl annealed adapter

c.     1.5 µl of water

d.     5 µl of 2x Quick Ligase buffer

e.   0.5 µl of Quick Ligase enzyme

3.     Mix briefly, and incubate at room temp. for 5’.  Use immediately for transformation.

III.            Transform DH5a cells.

1.     You will need a 42˚ water bath and LB+AMP plates.  You will also need to get a tube of competent DH5a cells from the 9^th floor core in Light Hall.  Thaw the cells on ice.

2.     Transfer 100 µl cells to a 1.5 ml tube.

3.     Add 5 µl of the ligation reaction. Mix well (do not vortex).

4.     Incubate 45’ on ice.

5.     Heat-shock the cells at 42˚ for 2’.

6.     Transfer cells back to ice.

7.     Add 900 µl of LB media to the cells. Transfer the cells to a 15 ml tube.

8.     Recover the cells by incubating in a 37˚ shaker incubator for 30’ at 250 rpm.

9.     Plate out 100 µl of the cells on an LB+AMP plate. Incubate at 37˚ overnight.  You should get at least a few dozen colonies (e.g. 10-100 is typical). Most of them will contain the correctly cloned product.   When I have done negative controls with no adapter, I get zero or just a couple of colonies.

IV.           Screen colonies for correct insertion of adapter.

1.     Inoculate 2 colonies separately into 15 ml tubes with 2 mls of LB+100 µg/ml AMP.  Shake overnight at 37˚.

2.     Perform Qiagen minipreps on 1.5 ml of each plasmid culture.  Save the remaining ~0.5 ml for making a glycerol stock to store at -80˚. (see below).  The culture sample can be stored at 4˚ for a few days before you make the glycerol stock.

3.     Spec the DNA.  Usually the concentration is about 40-100 ng/ul.    Prepare a sample for direct Sanger sequencing using this primer:

U6F1:              TACGATACAAGGCTGTTAGAGAG

This sequencing will read-through the region of the BbsI site and verify that the adapter has inserted properly.

V.             Make glycerol stocks of the correct clones.   To the remaining ~0.5 ml of the miniprep culture in step IV above, add 0.5 ml of sterile 30% glycerol.   (Glycerol solutions should be sterilized by filtration, never autoclaving.) Mix and store the glycerol stock at -80˚.

Doug Mortlock 7/2/14

DNA or RNA concentrations for mouse embryo #CRISPR injections: some considerations.

I was asked a few questions recently about this subject.  Here's my responses (edited for clarity):

Q: What is the recommended concentration of ssDNA to use for HDR in mouse embryo injections?

This depends if you are doing cytoplasmic (RNA + DNA) or pronuclear (DNA only) injections.  (More on the differences between those injections in the next question.)   Let's consider cytoplasmic injections of guide RNA, cas9 mRNA, and ssDNA as the HDR donor.   Yang et al. (Cell 2013 v154(6)pp.1370-1379) - the first group to publish HDR in mouse embryos -  reported using 50, 100 and 200 ng/ul for these components, respectively, for cytoplasmic injections.     Other groups have apparently reported using lower concentrations for these components with good success.  However, in general the concentrations used for cytoplasmic RNA/DNA injections will be higher than DNA concentrations used for pronuclear injections.   The cytoplasm can tolerate a higher burden of injected nucleic acid than the pronuclei can.

Q:  What are the main differences between cytoplasmic and pronuclear injections?

Cytoplasmic injection is technically slightly easier and (I believe) can result in higher embryo survival and implantation rates, since the risk of damaging the pronucleus is lower.    But it is only appropriate for RNA injections, such as Cas9 mRNA with guide RNAs, with or without HDR DNA donor molecules.     Although Cas9 mRNA injections can be very efficient, they have the disadvantage that they depend critically on the mRNA quality, which can vary depending on who made it, which kit was used, how it was stored and for how long, etc.   I am aware of l difficulties that several groups have encountered with this - some of this is anecdotal but some is from my direct experience. 

Because of this, I prefer injection of DNA plasmids (e.g. PX300) to transiently express Cas9 and guide RNAs.    Although this is slightly less efficient than RNA injections (when the RNA is good), it is very consistent, and it's easy for basically any lab that does cloning to make decent mini prep plasmid DNA suitable for injection.   This is not so straightforward for in vitro mRNA synthesis.       

So, how much DNA can you inject into a pronucleus?   It seems that up to about 10 ng/µl are OK as a maximum concentration, without incurring too much toxicity.    This is a total DNA burden.  A typical HDR experiment will then have at least one guide RNA/cas9 plasmid co-injected with an HDR ssDNA oligo or ds DNA fragment.   How much of each is optimal is still being worked out.    I suggest a 1:1 mass ratio of plasmid concentration to donor DNA concentration.  (note mass ratio, not a molar ratio; e.g. similar ng/ul of plasmid and donor.)