Unexpected high rates of off-target CRISPR mutations? Um, probably not.

UPDATE, 3/30/18:
Nature Methods has retracted the paper.   Two of the authors formally agreed with the retraction, while the rest did not.

UPDATE, 3/26/18:
Schaefer et al. have now uploaded a manuscript to bioRxiv, stating that they have now sequenced additional CRISPR mice but did not find off-target mutations in these animals, in contrast to what they claimed in their Nature Methods paper.    My perception is that they (the authors) are not going to retract the first paper.

Original post:

FSSSSSSSSSSS……(sound of me being thawed out of cryogenic storage) I’m back to CRISPR blog after a long hiatus.  The reason for my revival is the recent fracas about the paper "Unexpected mutations after CRISPR–Cas9 editing in vivo" in Nature Methods, Schaefer et al. 5/30/17...    Like the first “human embryo CRISPR paper”, this one got a lot of press in the broader media, in part certainly because of the headline-grabbing title but mainly because it implies a counter-hype to the CRISPR hype. 

BOTTOM LINE:  The central conclusion of the paper is almost certainly wrong.  The variants reported were almost certainly pre-existing polymorphisms and were not caused by CRISPR.  This paper should be retracted. If I’m proven wrong, I’ll sheepishly publish my own retraction of this blog post!  

This paper got a lot of attention, and even temporarily eroded stock prices of CRISPR biotech companies.   Columbia University first pushed the story in a press release on May 30. It got picked up and the story was off and running.   

Cosmos: May 30. CRISPR gene editing causes hundreds of unintended, off-target mutations
The Conversation: May 31 by Ian Haydon. CRISPR Controversy Fuels Debate in the Science Community
Wired, June 4: Crispr’s Next Big Debate: How Messy Is Too Messy?
Seeker: June 6. CRISPR DNA editing can cause hundreds of off-target mutations.
The Scientist, June 7: Was a Drop in CRISPR Firms’ Stock Warranted?
New Atlas: June 12.  The CRISPR controversy: Scientists skeptical over recent critical study
USA Today, July 24: CRISPR gene editing tool: Are we ready to play God?
Forbes, July 1: CRISPR Gene Editing Controversy: Does It Really Cause Unexpected Mutations?

I will not present a very detailed breakdown of the paper here.   The serious problems in the paper have been addressed previously, and well, by others. Gatean Burgio has published a very good analysis on Medium.  A publicly available preprint also deals directly with the key data and concludes the authors were mistaken: Questioning unexpected CRISPR off-target mutations in vivo. Kim et al, bioRxiv, 6/30/17.

However, I can contribute a few pieces of key genetic-y information that have not been described in much detail.  And these really involve details, and they really do matter.  They have to do with what inbred mouse strains really are and how CRISPR mice are made.

Inbred does not mean identical.  First, there is a widespread assumption that separate mice from the same inbred strain are genetically identical.   For almost all practical purposes, this is a pretty useful assumption.  But technically, of course, it is wrong.  Each new mammalian embryo has on the order of 50-100 new mutations (mostly single base changes - “SNPs” -  but other types too).   So even siblings from (theoretically) genetically identical parents will not be perfectly identical.  You’ll have to look hard to find the differences though; the new mutations amount to fewer than 1 new base per million bases of genomic DNA.   That’s why hardly anyone needed to worry much about this problem before the advent of whole-genome sequencing, and its resulting power to actually find these variants.

Let’s pause briefly to think about inbred mouse strains.  By definition, these strains were initially derived by repeated brother-sister matings, carried out for usually 20 or more generations.  This is adequate to eliminate almost all of the initial heterozygosity present in the founding mating pair.  However, some new SNP mutations still arise in each new generation.  Each new mutation might be weeded out by chance thanks to the continued inbreeding.  Alternatively, it has a chance to rise in allele frequency or even become fixed. 

How many polymorphisms? It’s good to do a thought experiment about how many variants we might expect to see within a population of inbred mice.  Population genetics theory predicts that the number of existing polymorphisms in a fully inbred (brother-sister mating) mammalian population will be approximately 4N, where N is the mutation rate per individual per generation.  Let’s assume N is 50; 4N = 200.  So we’d expect about 200 polymorphisms to exist, all told, among the genomes of the brother and sister.   (e.g. one or both is heterozygous for the variant in question, or both are homozygous but for different alleles). 

But 200 polymorphisms is just a lower bound.  In theory this is what we’d expect for an inbred population of two animals, which is impractical in reality.  In practice, inbred strains have larger effective population sizes – JAX probably has hundreds of FVB mice in their actual stock colony at any moment, if not more.   Although they take pains to maintain as close inbreeding as possible with this colony, I believe it’s safe to assume that the number of SNP polymorphisms extant in a vendor’s FVB stock is in the thousands, when the whole population is considered.  Mind you, many or most of these variants may be at low allele frequencies - perhaps only in a single animal. But many will certainly exist in multiple animals.   (These individuals are still incredibly similar to one another as compared to, say, wild mice - any two of which almost certainly will have several million SNP differences between them.) 

Sibs have a lot in common.  Now we can do another thought experiment: if we take two true siblings from the inbred strain population, they will share more of these ~2000 SNPs in common with each other than either would share with another mouse that is not a full sibling but is from the same population.     This is one of the simplest concepts in genetics, but it’s still not something we usually consider when dealing with mice of the same colony of inbred mice. 

Let's make CRISPR mice.  But how?  Ok. Now back to CRISPR. How are CRISPR mice actually made?  The process begins with 1-cell zygotes, which must be physically injected with the CRISPR reagents.   The process is essentially the same as was perfected over thirty years ago for the production of transgenic mice; but instead of injecting a DNA transgene, we inject the zygotes with a mixture of CRISPR guide RNA, Cas9 protein or mRNA, and (optionally) a targeting DNA molecule.

To inject them we need to get the zygotes into a culture dish under a microscope.    But we’ll need to collect about 100 or more of them to be sure of getting several live CRISPR’d mice in the end.  Most, but not all of the zygotes will survive the initial injection process; but only some of these will implant and develop normally, and only some of these will be CRISPR-edited.   That’s why we need to start with a lot of zygotes. Mice usually ovulate and mate at night.     The next morning, fertilized zygotes can be dissected from the oviduct of the female mouse.  Now, even though we will have used hormone injections to induce super-ovulation (so the mouse will release more eggs than normal), we’ll only get about 15 or so embryos per FVB mouse.  And not all of those will be “injectable” – some may have abnormally cleaved too early or simply be unfertilized, and will be useless. So we’ll need to dissect zygotes from about 10 females to be reasonably confident we’ll have enough to do the experiment.  And oh yea, the females all need to be about 4-5 weeks old when mated.

Fine - so we need to have 10 females of the right age, freshly mated the night before the injections.  We’ll order these from a vendor like JAX.   The night before injections, we’ll set up each female mouse in a separate mating cage with a stud male of the same strain.  We keep these males on hand at all times; they are proven fertile males, usually somewhere between 3 and 12 months old. 

A batch of embryos.  On injection morning, we euthanize the female mice and quickly dissect the zygotes out of the oviducts into one culture dish.  Now, they are all in one batch, and at this stage nobody ever keeps track of which female produced which zygotes.  They are all supposed to be genetically identical, right? (sarcasm) 

Some are sibs.  So finally, we can do our final two thought experiments:  If we take any two of these embryos at random, what is the chance that they will be more closely related to each other than to a third, randomly chosen embryo from the same batch?  Obviously it is quite possible that the first two might be true siblings.  The batch of embryos is made of ten or so "families" of sibling groups.  But we can’t tell under the microscope which ones are true siblings or not.  The point is, any two that you pick might be siblings.   And it is certainly possible that the first two might simply be more closely related to each other than they are to the third one, even if none of them are siblings. 

And last, let’s assume that when we pick two embryos, that they are in fact true siblings and that the third one we pick is not a true sibling of the other two.   Now let’s sequence their genomes and see how many SNPs they all share.   If there are several thousand SNPs in the vendor’s mouse colony, we expect that full sibs will likely share a thousand or more SNPs in common that would not be shared with another randomly chosen mouse from the same colony.   This is based on the idea, as explained above, that any mouse is likely to have on the order of a few thousand polymorphisms.   Siblings, of course, will share about 50% of those between them.

Final thoughts.   This is pretty exactly much what the authors found.   I see no reason to conclude the mutations were caused by CRISPR.    It certainly shines a bright light on the problem of distinguishing pre-existing variations from new mutations in gene editing experiments – which is exactly the problem that tripped up the authors of the first human embryo CRISPR paper.   The genome is big, and there are lots of variants to keep track of.  It’s actually a much tougher problem than doing CRISPR in the first place.

My recommendations for mixing and diluting #CRISPR gRNAs, mRNAs, and HDR oligo DNAs for mouse zygote injections.

It's super convenient to outsource CRISPR reagent construction to vendors (shout out to my friends at Sigma-Aldrich) and also to outsource generation of CRISPR mice to your friendly neighborhood transgenic core (shout out to the Vanderbilt TMESCSR).   However there still is that step where you may have to actually handle the RNAs/DNAs and mix them together, before handing them off to your friendly transgenic core staff.  I often get asked how I do this.  It's nothing tricky but it does involve RNA, so of course, be clean and careful.  Here are my current recommendations based on receiving RNAs shipped on dry ice from Sigma-Aldrich.

Recommendations for mixing and diluting CRISPR gRNAs , mRNAs, and HDR oligo DNAs for mouse zygote injections.  

Doug Mortlock Feb 2016

•A chart with the recommended final concentrations is found on the TMESCSR website ( https://labnodes.vanderbilt.edu/resource/view/id/11363 ) and is reproduced here.

•Sigma-Aldrich ships the RNAs at slightly higher concentrations than shown below.  Cas9 mRNA is provided at 500 ng/ul and gRNAs at 200 ng/ul.   

•I do not further clean up or process the RNAs in any way prior to dilution.   However, I do carefully re-precipitate the HDR oligos and resuspend them in sterile RNAse-free water.    This can remove some contaminating non-DNA material that is sometimes present in lyophilized oligos as shipped from the vendor.

1.     To prepare “N” injection days worth of injection mixture aliquotes, prepare N+1 tubes as follows.  Use clean, RNAse-free 1.5 ml microfuge tubes, sterile RNAse-free water, and RNAse-free arosol barrier tips.   Pre-rinse each tube by adding 1 ml of the water, vortex, and dump out all the water. Spin down the tubes briefly and remove any lingering dregs of water with a pipette tip. Place all the tubes on ice.

2.     Mix the RNAs, water (and DNA oligo as needed) to create a mix with the correct final concentrations of all reagents and volume that is equal to at least (N x 25) + 5 µl.  Mix briefly by flicking the tube gently (do NOT vortex). 

3.     Spin the mixture at full speed in microfuge for 1 minute. This is to pellet any small particulates – even if none are visible to the eye, before or after this step!

4.     Drawing from the top of the mixture volume, remove 25 µl and aliquot to one of the tubes.  Repeat for each aliquot.  There will be ~5 µl left over.  This hopefully has concentrated any particulate material (that might clog injection needles) and is discarded as a sacrifice to the CRISPR gods.   Although it is NOT usually visibly apparent that there are ANY particulates present, this step is added because it is TYPICAL for the injection technicians to have needle-clogging problems with CRISPR injections.  This reduces embryo survival.   While this pre-spin step may not always solve the problem it may help and is easy to do.

5.     Clearly label the aliquot tubes and bring them on ice to the transgenic core. They should be stored at -20˚ until injection date.   

The chart below was culled from our TMESCSR recommendations as of Feb 2016.  I know 'cause I wrote 'em.

Create the required mixture of components so that it has the final concentrations shown below.

The "min volume to bring to Core" is specific to the Vanderbilt's core's preferences.  So don't use this to argue with your core's staff about what is necessary.  They get to decide that for themselves!

Experiment type	Final conc. Cas9 mRNA	Final conc. all gRNAs	Final conc. all ssDNA donor oligos	Buffer	Min. volume to bring to Core
Knockout experiment, RNA reagents only	100 ng/µl	50 ng/µl	-	Sterile, RNAse-free water	25 µl PER INJECTION DAY
Knock-in experiment, RNA + ssDNA oligo(s)	100 ng/µl	50 ng/µl	200 ng/µl	Sterile, RNAse-free water	25 µl PER INJECTION DAY

Requested deviations from these concentrations will require consultation and approval from the core manager.   The core staff may need to dilute the reagents more if injection problems arise (e.g. clogging).

REQUIREMENTS FOR PURIFICATION:

• We strongly suggest ssDNA oligos be re-precipitated before use to remove potential contaminants from the vendor.    See TMESCSR website for this protocol.

For #CRISPR HDR, use donor oligos that are complementary to the "gRNA strand". A new paper shows why; see my blog post.

(ERRATUM:  I made on correction to this post on March 11 2016.  In the original version of the post I stated that the paper implied that the donor oligo should have "additional length of homology on the PAM-distal side as compared to the PAM-proximal side".  That was a mistake - it turns out the opposite was true.   The authors found that additional length of homology on the PAM-proximal side was favorable.  I got confused because the PAM in figure 3 is on the "bottom" strand, not the top, so the PAM-proximal side is to the left of the cut site in their oligo schematics.  Figure 3 has an "upside down" Cas9 icon in keeping with this fact.  Thank you Scot for letting me know about the error!).


After a long break from blogging... here's a nice nugget of insight about CRISPR-mediated homologous recombination.    Back in late 2014 I blogged about 
Optimal design of ssODNs (donor oligos) for #CRISPR - length and strandedness data? . In that post I pointed out a curious observation that HDR oligos work "better" when they are designed from the strand that is complementary to the protospacer/gRNA sequence.   This was somewhat counterintuitive to me, as one might think that in a complementary HDR oligo would tend to anneal to the gRNA, reducing its availability or kinetics somewhat and generally interfering with Cas9's job.   But empirically, this was not the case; complementary oligos work better.

Now, Richardson et al seem to have found an explanation. (Richardson et al, Enhancing homology-directed genome editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA. Nature Biotechnology (2016, Published online 20 January 2016)). Turns out that following double-stranded DNA cleavage by Cas9, the first component of the DNA molecule that it releases is the 3' end of the DNA strand that corresponds to the protospacer/gRNA.  Quote: Hence, although Cas9 globally dissociates from duplex DNA in a symmetric fashion (Fig. 1c), it appears that the enzyme locally releases the PAM-distal nontarget strand after cleavage but before dissociation."  And here is a nice diagram from the supplemental material of the paper that shows how this strand "breathes" after cleavage.  I thank the senior author, Jacob Corn, for graciously allowing me to reproduce the image here:




(OK - before moving forward, let's get clarity on the terms here; when we're comparing the two strands of the DNA containing the CRISPR target, the "target" strand is the DNA strand that directly will anneal to the gRNA.  Thus, the "target strand" is complementary to the gRNA.  The non-target DNA strand encodes the protospacer and the "NGG" PAM sequence.   Got it? )   

Read the above quote again.  The first bit of DNA that is released by Cas9 is the single-stranded 3' end of the non-target strand.  This immediately suggests 2 things:

1.  Cas9 releases the non-target strand before the target strand.  Thus the non-target strand is available sooner than the target strand to potentially engage with a complementary donor molecule and jump-start homologous recombination.      

2.  The 3' end of the non-target strand, which is "PAM-distal", is released first.  This also suggests that the design considerations for homology might be different for the PAM-distal and PAM-proximal sides of the cleavage location.

For this second point, the practical consideration is that commercially available ssDNA oligos are usually limited to 200 bases or less (depending on the vendor's capabilities; 120 base oligos can work well too).  So, we are limited in the length of homology we can actually apply to each side of the cleavage site.   Instead of centering the oligo (e.g. for a 120 base HDR oligo = ~60 bases of homology on each side) it may be better to skew the oligo design to have more homology on one side of the cut site versus the other.  That is what Richardson et al found - at least for one target they investigated in detail  (See Fig. 3c, d, e.)

Here's another thought. The authors note that Cas9 actually stays on the DNA for quite a while after it cleaves both strands - about 5 hours.    This might have something to do with why DNA repair takes significantly longer on Cas9-cleaved breaks than on breaks induced with radiation - Cas9 may just sitting there, sterically hindering the DNA repair proteins from accessing the free DNA ends at the break.  Perhaps, CRISPR mutagenesis efficiencies could be further enhanced by increasing Cas9's intrinsic off-rate?  On this note, another recent paper by Kleinstiver et al (Keith Joung lab) suggests that directed mutations can destabilize Cas9's non-specific interaction with DNA.   The authors note that this reduces off-target cleavage significantly while preserving on-target cleavage .   While they did not see dramatic increases in on-target efficiency, perhaps in some experimental contexts there might be?    Hmm.    

#CRISPR editing mouse embryos by direct zygote electroporation - no microinjection needed.

I’ve been intending to blog on this for a while now and finally got around to it.   This paper comes from a group at the Jackson labs; the first author is Wenning Qin, and the senior author is Haoyi Yang who also holds a primary appointment at an institute in Beijing.

Qin et al.  Efficient CRISPR/Cas9-mediated genome editing in mice by zygote electroporation of nuclease.  Genetics, 2015 v.200 (2) pp. 423-430.

In this paper they show that CRISPR-Cas9 reagents can be electroporated directly into mouse zygotes to generate gene-edited animals.  It’s not quite as efficient as direct injection into zygotes – but it’s not too shabby, considering the ease of the electroporation step.

To remind those who are unfamiliar, the main method to deliver DNAs or RNAs into mouse zygotes is through direct pronuclear or cytoplasmic injection, using ultrafine capillary needles.  It requires micromanipulators for the needles, carefully controlled pressure to deliver the injected material, and a quality inverted microscope with high-contrast optics.  Plus a steady hand and skill at injections.  In mammals, zygotic DNA transgenesis has generally required direct injection of DNA into the pronuclei (unless a retrovirus is used, which has its own drawbacks).  This was shown in papers from the early days of transgenic mouse research.  

Historically, electroporation has not been used for engineering mouse zygotes.   There are some good reasons for this.  First, zygotes have a zona pellucida surrounding the zygote itself.  This can be dissolved away rather easily with brief acid treatment – but without the zona, the embryos are sticky and much more difficult to handle.  Second, electroporation doesn't immediately transfer material into the zygote nucleus, and the chance of DNA integrating into the genome is very low.  In fact, even direct injection of DNA into the zygote cytoplasm does not yield transgenic mice efficiently – you’ve got to inject it into the pronuclei.    

Now, there are research applications for zygote injection apart than transgenesis.  You might just want to transiently express an mRNA in a zygote, for example.  Embryologists who work with zebrafish and xenopus will be totally familiar with this idea.  It’s not done frequently in mice but can be done.  I think the labs that ever do this with mouse embryos are hardcore enough they have access to microinjection equipment, and presumably haven’t bothered to try electroporation much - why would you, if you are all set up to perform the established method.

However…what if you either (1) want to try transgenic manipulation, but don’t have access to a microinjection apparatus, or (2) you just want to really streamline the labor involved?  Then electroporation might be useful…  Enter CRISPR, in which we actually do want to transiently express the reagents in zygotes.  At Jax they fall into the (2) category.

To develop this method, Qin et al. first confirmed previous reports that brief incubation in acid can be carried out to weaken the zona pellucida without completely dissolving it, while not affecting embryo viability.  Next, they tested electroporation parameters to optimize both the media/TE mixtures compatible with embryo survival and the maximum voltages the embryos could tolerate and still live.   Finally, they mixed acid-treated embryos with Cas9 mRNA plus guide RNAs for known pre-validated targets in the Tet1 or Tet2 genes and did electroporations.  Surviving embryos were either genotyped after in vitro culture, or transferred into recipient females and analyzed after birth.  

Bottom line: they could generate mutant animals at double digit percentages.   Not surprisingly, efficiency increased with higher RNA concentrations.  The final standard conditions involve at least 30 to 50 embryos per electroporation, in a total volume of 20 µl buffer/media with final concentrations of 600 ng/µl Cas9 mRNA and 300 ng/µl guide RNA.  Note that this requires a total of 12 µg Cas9 mRNA and 6 µg gRNA per batch of 50 embryos.   They also showed HDR is possible by co-electroporation with donor oligo DNAs.

So what is the efficiency?  Table 2 is a very nice comparison of microinjection vs. electroporation across ten genes.  Kudos to them for a nice big data set!  Good news: electroporation generated mutants for 5/10 genes tested.  However, 8/10 microinjections were successful for the same genes/reagents.   The overall rate of mutants was lower in the electroporation set as well; the average efficiency in the 5 successful electroporations was 20%, while with microinjections it was 42%.   Interestingly, the overall rate of embryo survival to birth was higher for electroporations – about double that of microinjections.   So in the end, the lower efficiency of mutant generation via electroporation might be pretty well balanced by the higher birth rate per embryo.

I’ll still note that zygote manipulation may not be for the faint of heart even if you don’t have a microscope/microinjection setup handy.   Remember that to make engineered live mice you have to transfer the manipulated zygotes back into a pseudopregnant recipient female mouse.  This requires microsurgery, people.     

However – maybe even this last step can be improved upon.   Takahashi et al have now reported that embryos can be electroporated – and CRISPR-mutagenized – without taking them out of the oviduct.    (Takahashi et al, Scientific Reports, June 22 2015 (5:11406).)  This still requires injection of the CRISPR RNA solution into the oviduct of a live mouse, but it’s probably easier than transferring embryos back in to the oviduct.   Hmm… maybe one doesn't need to pre-weaken the zone pellucida?  Their method requires rather high RNA concentrations, but bears promise.   

#CRISPR / gene editing featured at Festival of Genomics Nov. 3-5 in San Mateo. #genomicsfest

I will be participating in a CRISPR mouse editing workshop next week (Nov. 3) at The Festival of Genomics, in San Mateo, CA.  The full workshop title is "CRISPR/Cas9 Genome Editing Pipelines for Mice and Rats" and I'll be joined by Thom Saunders (University of Michigan) and Kevin Peterson (The Jackson Laboratory).  

The Festival of Genomics is a fairly new, recurring conference that has a lower registration fee than most traditional scientific-society meetings.  The speaker lineup for the meeting next week is pretty strong, with a strong biotech presence not surprising given the locale, but also strong plenaries from academics as well (Jennifer Doudna, of course; Carlos Bustamante, Manolis Kellis etc).  

Looks like Tues. Nov. 3 is mostly workshops, and Weds & Thurs Nov. 4-5 will feature plenary talks in the morning and then four concurrent tracks of sessions: Genome Editing, Genome Analysis, Genomic Medicine, and Data Analysis.    If you're in the SF bay area, check it out.

Updated guidelines for mouse #CRISPR injections in the Vanderbilttransgenic core.

I have revised the CRISPR guidelines on our Vanderbilt transgenic core's website.

 https://labnodes.vanderbilt.edu/resource/view/id/5265/collection_id/14/community_id/8

Basically it describes the basic information about what to do, and more importantly what to know, to get a CRISPR mouse project started through our core.   This may be most useful for my Vanderbilt peeps but others may find it interesting, as it gives some insight into how our core facility is communicating these sorts of guidelines to our users.  

These guidelines are not heavy on the up-front, nitty-gritty CRISPR design aspects as there are other places to find that stuff - for example, in the archives of this blog, as well as through the links I provide on the right side of this blog page to some helpful tools.  

UK researchers ask you to submit your opinions about gene editing - link to web survey. #CRISPR

Dr. Lara Marks and Dr. Silvia Camporesi would like you to tell them your opinions about gene editing technology via their online survey.   What do you think? Let them know.  

Dr. Marks edits the hosting website, What is Biotechnology.   Dr. Camporesi is a bioethicist at Kings' College London.