#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.

Move over, Cas9: Cpf1 may be your new #CRISPR competition.

This past weekend at the Pilgrimage music festival in Franklin, TN I had the pleasure of seeing Weezer rock out, then I walked a few hundred yards to another stage to watch Wilco do the same.  These bands each had their own stage, but in the CRISPR world, Cas9 is a superstar that now has to share the stage with a newcomer:  Cpf1.  (Yeah, I know that's a goofy setup but it really was a good festival and it was on my mind.  Now on to the science.)

Last Friday Feng Zhang’s group published a paper in Cell that immediately grabbed a lot of attention, and rightly so.  They reveal that the Cpf1 class of CRISPR effector proteins may be an attractive alternative to Cas9.   Although Cpf1 has many similarities to Cas9, it has some significant differences that are very interesting – and could lead to improved efficiencies for some types of gene editing.  

Cpf1 Is a Single RNA-Guided Endonuclease of a Class 2 CRISPR-Cas System.  Zetsche et al., 2015, Cell 163, 1–13October 22, 2015  (Avail. online Sept. 25 2015).

Here is my summary…First, they reviewed some background on CRISPR systems in bacteria to cover the basis of the study.   There are two major classes of CRISPR systems based mainly on the proteins involved; the cleavage effectors of class 1 are complexes of multiple proteins, while the class 2 effectors are single proteins like Cas9.  Within class 2 there are two subtypes of systems: those with Cas9, and another that has Cpf1.   Cpf1 means “CRISPR from Prevotella and Francisella 1”.  Some bacterial species carry both Cas9-CRISPR and Cpf1-CRISPR loci in their genomes.   Like Cas9, Cpf1 has RuvC-like DNA cleavage domains but it lacks some of the other domain and neighbor-gene features of Cas9, so it’s clearly distinct in its evolution.    It’s a largish protein of ~1300 amino acids, similar in size to Cas9.

They picked the Cpf1-CRISPR gene system of Francisella novicida strain U112 to study first since there were clear homologies of Cpf1-CRISPR spacer sequences to various prophage in this species – further suggesting that Cpf1 is important in bacterial immunity and so it’s well adapted to slice and dice target DNAs.   (Sidebar: what’s F. novicida? A pretty rare human pathogen, originally isolated from the Great Salt Lake in Utah.  It’s related to the better known bug F. tularensis which is one of the most infectious pathogens known.)     

By transferring the F. novicida Cpf1-CRISPR gene locus into E. coli they quickly established that it prefers a  “TTN” PAM motif that is located 5’ to its protospacer target – not 3’, as per Cas9.  So right away it’s distinct in having a PAM that isn’t G-rich and is on the opposite side of the protospacer. 

Like Cas9, Cpf1 binds a crRNA that carries the protospacer sequence for base-pairing the target.  But for me the biggest surprise in the paper is that unlike Cas9, Cpf1 does not require a separate tracrRNA – in fact, there’s no sign of a tracrRNA gene at the Cpf1-CRISPR locus.   Thus, Cpf1 merely needs a cRNA that is about 43 bases long –of which 24 nt is protospacer and 19 nt is the constitutive direct repeat sequence.   This is very different than Cas9 – even by fusing the crRNA and tracrRNA, the single RNA that Cas9 needs is still ~100 nt long.

Furthermore, the Cpf1 crRNA does not have the long stemloop structure that is typical of RNAseIII-mediated processing to cut it out of its primary transcript.   It has a much shorter stemloop that is required for Cpf1 activity, however.   But surprisingly, Cpf1 itself is apparently directly responsible for cleaving the 43-base cRNAs apart from the primary transcript in the first place! This isn’t conclusively proven yet, but is pretty likely based on their experiments.   

Next, two more surprises comes from the cleavage sites on the target DNA.  The cut sites are staggered by about 5 bases.  This should create “sticky overhangs” that might be exploitable to enable gene editing via NHEJ-mediated-ligation of DNA fragments with matching ends.   And, the cut sites are in the 3’ end of the protospacer, distal to the 5’ end where the PAM is.    The cut positions usually follow the 18^th base on the protospacer strand and the 23^rd base on the complementary strand (the one that pairs to the crRNA).

They tested if they could inactivate the DNA cleavage domains via homologous mutations in codons known to do this in Cas9.  However, the resulting Cpf1 mutants don’t have “nickase” activity – they can’t cut either strand.   So it’s not clear that Cpf1 nickases can be made and in fact the authors suggest that the cleavage might require some sort of dimerization.  I can’t visualize how that would work yet but I’m sure it will be figured out in the near future…

Base substitution experiments then showed that, as per Cas9 CRISPRs, there is a “seed” region close to the PAM in which single base substitutions completely prevent cleavage activity.   Therefore, unlike the Cas9 CRISPR target the cleavage sites and the seed region do not overlap.  This immediately suggests a potential improvement over Cas9 in mammalian HDR-mediated repair efficiency.    This is because any initial cleavage events that might lead to “simple” NHEJ indels might still be substrates for cleavage  - and thus allowing additional chances for HDR-mediated edting to occur.  With Cas9, an indel mutation will almost always disrupt the target seeds and then it’s game over for HDR.     

Finally, they did the important work to screen various Cpf1 proteins from different bacterial species to see if any would actually work in mammalian cells.  This is because that despite codon optimization and attachment of nuclear localization signals, most of these bacterial proteins just don’t work right when you put them inside human or mouse cells.  Therefore they tested 16 different Cpf1 proteins.  Of these, for seven proteins they could identify PAM signatures using their E. coli assay.  They all had similar T-rich PAMs. 

Of these seven proteins, only two worked well in human HEK293 cells – AbCpf1, from an Acidaminococcus, and LbCpf1, which is from a Lachnospiraceae; interestingly these are apparently both anaerobic bacteria sometimes found in mammalian intestines.  Anyway these can both generate indels at specific targets in human cells at a rate similar to Cas9 – typically, 10-20% Surveyor assay numbers were observed, when they tested HEK cells following simple transfections.

Bottom line: Cpf1 may be the real deal as a serious competitor for Cas9.  Is Cas9 suddenly obsolete?  Hardly.  First, we don't yet know if Cpf1 is as specific as Cas9 – though there is every reason to think it may be.   So the off-target effects need to be carefully measured. Second, we don’t know how widespread the targeting efficiencies will be across sites (although the initial tests seem very promising).  Third, although Cpf1 may be better than Cas9 for mediating insertions of DNA, it’s not yet been shown if that is true.   However it may have some nice advantages over Cas9, not the least of which is that its guide RNA is only 43 bases long.  It will thus be feasible to purchase directly synthesized guide RNAs for Cpf1, perhaps with chemical modifications to enhance stability.

Probably, Cpf1 and Cas9 will both be in the spotlight for a long while to come.  Look for more Cpf1 papers to start coming out very soon and for Cpf1 plasmids to appear in Addgene.  Happy CRISPRing, everyone.

Link to a timeline of recent discourse on #CRISPR human germline gene editing.

I've started a separate blog about human germline gene editing.  Hope you like it.  My first post is a list of links to recent news items on the topic.

Webtool link for getting Xu et al #CRISPR target scores for your sequence of interest.

A followup to yesterday's post about the Xu et al paper,  Sequence determinants of improved CRISPR sgRNA design:   They have also kindly made a public webtool for generating CRISPR scores with their model.   It's a cut-and-paste that accepts up to 10000 bases.   Simple and quick.    

Of course, their source code is available too in their supplemental material and here.   

#CRISPR target sequence preferences are being clarified. Xu et al Genome Research paper.

It's of huge value to be able to predict CRISPR target efficiency ahead of time.  Xu et al have published an analysis of multiple guide RNA data sets and extracted what they claim is an improved model for target cleavage efficiency prediction.   This data is all for the S.pyogenes native Cas9. 

Xu et al. Sequence determinants of improved CRISPR sgRNA design.
Genome Res. 2015 Aug;25(8):1147-57. doi: 10.1101/gr.191452.115. Epub 2015 Jun 10.

Their paper is important to me for several reasons.  First, they have examined two independently-published "large" guide RNA data sets that had mutagenesis-efficiency data, which allows more confidence that trends of sequence preferences are holding up across labs and platforms.   Second, they validated their predictive model on a small (in comparison to genome-wide, but still not bad) data set of new CRISPR targets and corresponding guide RNAs.  Third, they did "in silico validation" by turning their model loose on another target/indel data set, and showed improved performance of their predictive model over a previously published model.   See ROC curves in Fig. 4b.    This allows an ability to weed out "50-60% of the inefficient sgRNAs…at the cost of 10-20% of efficient sgRNAs misclassified."  That is, misclassified as inefficient.    

For those who are interested in genome-wide knockout screening experiments these sorts of models are very good for increasing efficiency of the screens.   Moreover, if you wish to knockout particular genes, it will allow you to test or use fewer targets per gene till you find one that works well.

OK, now the sobering reality for nerds like me is that predictive models, even with great ROC curves, have false positive and false negative rates that will bite you in the behind on a regular basis if you are designing large projects around the function of single CRISPR targets.  I'm still facing this issue for precision knock-in projects, for which there are often not many targets to choose from.   And with transgenic mice we always want the efficiency as high as possible.   For cell lines, hey, that's not as much a problem if you can subclone the edited lines.

But let's get back to the CRISPR target sequence preferences.  The bottom line here is that the last three bases of the protospacer seem to have the most influence on cleavage efficiency, with a C preferred at the -3 position (relative to the PAM), and G's at -2 and -1.    Also, G's are helpful at the -17 to -14 region, while A's are good at the -12 to -9 region.  Finally, a C seems helpful at +1 following the PAM.

Looking back at the Wang et al paper, they also reported a preference for A's at around -10 to -8, and essentially a "GCRR" preference for bases -4 to -1.  This makes sense since Xu have based their model partly on the the Wang data.   However, Xu et al point out that the apparent G preference at the -20 position is probably an artifact of the Wang sgRNA library in that these may have had increased efficiency due to enhanced transcription, not activity per se.

General GC-richness in the protospacer is known to correlate with CRISPR mutagenesis.  Could that just be driven by the GC-rich preferences of the last few bases?   Otherwise, GC-richness doesn't clearly emerge from the Xu model, at least to me anyway.  I took a crack at this by looking at a data set from Gagnon et al, mostly because I could handle the size of their sgRNA list in an excel spreadsheet without exploding my own brain or my iMac.  My impression is that GC richness is still "good" even when the last 4 bases of the protospacer are similar.   Here's an example.  From Gagnon et al's list of 122 sgRNAs with indel numbers, I ranked them according to how well they matched the "GCRR" of the last four bases.  I based this on the Wang et al paper although I think it is very similar to that corresponding part of the Xu model.   My  "score" ranged from 0 to 7.  Then I examined the subset of 30 targets that all had a same "score" of 5.   So these targets are all controlled, at least kinda sorta, for their  variation in bases -4 to -1 in that they have similar strength of matching to the "GCRR" motif.  Finally, I graphed the indel frequencies versus the GC content of the first 16 bases of their protospacers.   Here is the data.  y axis= indel frequency (in a zebrafish model), x axis= # of GC base pairs in first 16 bases.

This ain't close to something I'd submit for peer review but I do see a trend.  GC richness in the first 16 bases of the protospacer correlates with cleavage efficiency, even within a group of targets for which the 3' ends are similar.   So for now - I will continue to prefer overall GC-rich targets that also have at least some matching to the "CGG", or "GCRR", motif at the very 3' end.  

Also, "CGG" matches the high-efficiency 3' end reported by Farboud and Meyer so there's another corroboration.

So, the answer to my previous post "Are there sequence preferences near the 3' end of the #CRISPR protospacer? …" is, yes.  And this holds up for S.pyogenes Cas9 when used across human, mouse, fish and C.elegans models.   

A final note - these data all refer to cleavage and/or knockout efficiencies.  CRISPRi and CRISPRa screens, which do not lead to or require DNA cleavage, have different sequence preferences which Xu et al also modeled in detail.   

Happy CRISPRing.

My review of recent Joung lab paper with new PAM specificities! for Cas9: will broaden choice of #CRISPR targets.

It was just a matter of time before someone mutagenized Cas9 to try to change its PAM preferences.  (What's a PAM? Look here if you aren't hip yet).   Although that "NGG" motif is pretty abundant in genomes - heck, it's only 2 bases - it sure would be nice to be able to target even more sequences with high specificity.  For example, the closer the CRISPR site is to the site where precise genome editing is required, the more efficient it will (probably) be;  having more PAM choices will only be helpful in this situation.  But, having more targets isn't very useful unless the properties of CRISPR specificity and sensitivity remain robust.  

So now the Joung lab has led the way with efforts to coax Cas9 into preferring new PAMs, as described in Kleinstiver et al's new paper in Nature.    I really like this paper.   It introduces new Cas9 variants with preferences for NGA, or NGCG PAMs. The new variants are not complicated to engineer, maintain high cleavage activity, work in vivo, and have low off-target effects similar to native Cas9.  

Previously, Anders et al had published a Nature paper describing Cas9-DNA structural interaction.  (Senior author of this paper was Martin Jinek, who was first author of the seminal 2012 Dounda/Charpentier Science paper).  An interesting nugget in that paper was a first attempt to alter Cas9 specificity by mutating the two amino acids (Arg1333 and Arg 1335), which apparently interact directly with the two guanine bases of the PAM motif, to glutamine.  This was inspired by Cas9 variants from non-S.pyogenes species which prefer A-rich PAMs and have glutamines in the homologous positions.   However these changes alone couldn't make S.pyogenes' Cas9 prefer NAA instead of NGG.

Enter Kleinstiver et al.  They began a systematic attempt to engineer new PAM specificities into S.pyogenes Cas9.  First, they used a clever bacterial assay to measure PAM preference in which the CRISPR target is within a toxic gene.  Cas9 variants with different codon changes were then introduced.  In this setup, target cleavage disrupts the gene and allows the bugs to grow, allowing one to sequence the survivors to figure out which Cas9 variants worked.  In this manner they identified combinations of codon changes that allowed Cas9 to recognize a NGA PAM.  2 variant combos, "VQR" and "EQR" , emerged as being best at now preferring NGA over NGG.

Then, they used a different assay to measure preference for all the possible different PAMs for selected Cas9 mutants.  See Figs. 1e, 1f for these data nicely visualized.  For example you can clearly see how wild type Cas9 greatly prefers NGG, but has a little ability to use NAG as had been previously reported by many - in fact many off-target analyses consider NAG PAMs as well as NGGs.   Then, they tested the VQR and EQR variants, which revealed that these now are sensitive to the fourth base in the PAM.  Specifically, Cas9-VQR "likes" NGAG, NGAA, NGAT, NGCG the best.    Interestingly, Cas9-EQR preferred NGAG almost exclusively.  The authors concluded that the T1337R variant is a gain-of-function allowing sensitivity to the fourth base, which is then specified by other codon variants.  Cool.

Next, they found that the "VRER" combination allowed specific preference for a NGCG PAM.  Note that this GCG motif is much less common in mammalian DNA than the other PAMs - after all, it's got a CG in it - but that also means it's off-target potential is lower.   Since I know lots of genes with GC-rich regions I'm betting this PAM will come in handy.

A few more points from the paper:   The new variants work in zebrafish in human cells, and have good activity and low off-target effects.  Additionally, they noticed that the D1135E variant actually increased PAM specificity for the wild-type NGG PAM relative to NAG - see Fig 3a, and furthermore it reduced off-target cleavage on other off-target sites that have mismatches in protospacer but (presumably) the NGG PAM.  In other words D1135E reduces off-target cleavage in general (at least somewhat) without reducing on-target cleavage.   Sounds good to me!

Finally, they examined two Cas9 genes from other bacterial species and showed they could carefully measure their normal spectra of PAM preferences (which are different from the NGG of S.pyogenes).  In other words they are poised to do the same mutagenesis work on these other Cas9 proteins, which will add even more PAM choices to the toolkit.   

i was about to write "these Cas9 variants should be widely available soon", then I thought Hmm, better check Addgene.   Sure enough:   VQR, EQR, and VRER expression plasmids are already available!  Kudos to Keith Joung and his lab for making these available to the world.  Happy CRISPRing with new PAMs!

Photoactivatable #CRISPR-Cas9 systems!

There are a few recently developed light-activated CRISPR-Cas9 tools that have been reported lately.  I'm motivated to post this based on the most recent one, which demonstrated gene editing using modified "split" Cas9 protein halves that were conjugated to newly developed light-inducible dimerization domains named "Magnets".   This was a paper just published by Nihongaki et al. in Nature Biotechnology.   (PDF is as of the post only published online at this link). This system is nice in that it just requires expression of normal gRNA plus the two modified coding portions of Cas9, plus, blue light to activate dimerization and Cas9 targeting function and cleavage.   It's reversible too - removing the light stimulation lets the complex fall apart.  Neat!

Other groups in parallel have created very similar tools that allow light-inducible activation of Cas9 to allow targeting.  In a related paper Nihongaki and colleagues showed they could use this to activate transcription at CRISPR target genes using light, and Polstein and Gersbach have made very similar tools.  Both groups used the CRY2 and CIB1 light-induced dimerization domains from Arabidopsis.

Using a different strategy, Hemphill et al used a caged amino acid strategy to encode a light-activatable codon into Cas9.  This system is a bit more complex to set up, as it requires engineering a pyrrolysl tRNA synthetase into the cells being targeted - basically, re-engineering the genetic code to get a light-activated lysine into the guts of Cas9.  This seems very different mechanistically than the dimerization approach and so it maybe a good alternative for some applications, as it probably has some distinct wavelength and kinetic properties.  Always a good thing to have different tools in the toolkit.

2015 Gruber Prize in Genetics awarded to Charpentier and Doudna for #CRISPR.

The 2015 Gruber Prize in Genetics is being awarded to Emmanuelle Charpentier and Jennifer Doudna for their pioneering work on CRISPR biology.   This prize is presented annually at the American Society in Human Genetics annual meeting, which will be held in Baltimore this year in October.

If you're not familiar with the Gruber Prizes, they are awarded in several disciplines including genetics and they include a $500,000 cash prize, so it's quite an award. Congratulations once again to Drs. Charpentier and Doudna!

@LluisMontoliu guest post! about low off-target #CRISPR rates in embryos.

Lluis Montoliu is very well known to the transgenic mouse community and and expert on all things related to mouse genetic engineering.  Therefore I was very happy when he sent a message to the ISTT mailing list describing the recent in-depth confirmation that yes, CRISPR can have extremely low off-target cleavage rates in mouse zygotes, as alluded to in one of my previous posts (and probably true for human embryos too despite a recent report).   

He has kindly agreed to let me re-post his message on this blog.  Thanks Lluis!  You can also follow @LluisMontoliu on Twitter, and check out his own CRISPR information web site and also his lab's web page.  

Subject: [ISTT_list] Off-target mutations are rare in CRISPR-Cas9-edited animals

Dear colleagues,

Anyone who has already carefully analyzed mice edited by CRISPR-Cas9 will have confirmed the almost absence of off-target mutations, in contrast to what was initially predicted and announced. Off-target mutations appear to be very rare in genome-edited animals, if present at all. We and other have usually taken a shortcut and have opted to analyze a limited number of off-target sites in our genome-edited mice, selecting a few off-target sites (those with higher score, higher probability to be modified) and cloned and sequenced these DNA pieces from all founder animals generated, just to find that none of them appear to be modified.
http://www.ncbi.nlm.nih.gov/pubmed/25897126

Now, Bill Skarnes and collaborators (Sanger Inst., Hinxton, UK) have done the proper experiment, the experiment we and other would have liked to do, namely: whole deep genome sequencing on CRISPR-Cas9-edited mice. And they found the same result. Even if you don't select for sites and you review the entire genome there appear to be no off-target sites that are modified by the CRISPR-Cas9 reagents.
Off-target mutations are rare in Cas9-modified mice Vivek Iyer, Bin Shen, Wensheng Zhang, Alex Hodgkins, Thomas Keane, Xingxu Huang & William C Skarnes Nature Methods 12, 479 (2015) doi:10.1038/nmeth.3408 http://www.nature.com/nmeth/journal/v12/n6/full/nmeth.3408.htmlhttp://www.ncbi.nlm.nih.gov/pubmed/26020497

Hence, these amazing tools are far more precise and accurate than initially considered, particularly when these are injected as RNA (orprotein) into zygotes (into fertilized oocytes). Of course, this does not mean that you should not aim to obtain and analyze at least two independent mutant/edited animals to confirm the robustness of the associated phenotype, as you would be doing with any other genome alteration you would be producing.  And, bear in mind, the whole picture might be different in cells, particularly if they are transfected with DNA plasmids transcribing Cas9 constantly and in high amounts,  and hence providing lots of opportunities (and time) for this endonuclease to cut elsewhere, other than the expected targeted sequence. In contrast to what happens in zygotes, where a limited amount of Cas9 RNA (or protein) is used, does the job and vanishes away.

Further enjoy your genome-edited animals!

Lluis
--
Dr. Lluis Montoliu
Investigador Cientifico - Research Scientist CSIC Centro Nacional de Biotecnologia (CNB-CSIC) Campus de Cantoblanco C/ Darwin, 3
28049 Madrid (Spain)

More confirmation that SCR7 increases #CRISPR insertion rates by inhibiting NHEJ.

I'm kicking myself for not finding this paper two months ago when it came out - I've been waiting for this sort of data!   Maruyama et al have published a more complete description of SCR7 tests in CRISPR modifications.   

Increasing the efficiency of precise genome editing with CRISPR-Cas9 by inhibition of nonhomologous end joining.  

Takeshi Maruyama

,

Stephanie K Dougan,

Matthias C Truttmann,

Angelina M Bilate,

Jessica R Ingram

& Hidde L Ploegh.  

Nature Biotechnology 

33, 

538–542

 

(2015)

They confirm what Singh et al previously reported in a small but exciting data morsel last fall, which is that substantially higher rates of HDR-mediated insertion can be achieved in mouse zygotes by treating them with the NHEJ inhibitor, SCR7, during the injection process.   They actually mixed SCR7 (final conc. 1mM) directly into the injection cocktail of gRNA + Cas9mRNA + donor ssDNA oligo.

After some preliminary tests in cell lines, they moved to zygotes.  Using a donor oligo to insert a short peptide tag and validated CRISPR targets/gRNAs, they did tests with and without SCR7.    Bottom line:  HDR-mediated insertion rates increased by several fold for the two genes they tested.   Although that may not sound like a breakthrough to some of you, many of you will know that in the world of mouse engineering it's key, because it will probably often mean the difference between getting zero versus a few correctly engineered pups out of an injection series.  

Some other highlights are:

1.  The embryos seem to tolerate SCR7 application under these conditions with no problem; no toxicity or increased death was noted.  Various other studies seem to support that transient inhibition of NHEJ is well tolerated.   Note that the SCR7 target, ligase IV, is critical for embryonic development so it can't be globally knocked out.  

2. No increase in off-target effects.   Cool.

Technical notes:

1.  Yesterday's google searching quickly turned up 3 companies selling SCR7.  Yay.

2.   SCR7 must be dissolved in DMSO.  I think making a stock solution of 100 mM SCR7 in DMSO is reasonable.  So the final injection mix, with 100-fold SCR7 dilution from the stock, will have 1 mM DMSO and also 1% DMSO.   I couldn't dig out the SCR7 stock solution details from the paper but it's probably close to these parameters.

3. SCR7 very strongly inhibits the recovery of NHEJ-style mutations from the CRISPR targets.  

4.  The zygote injections were all done cytoplasmic, not pronuclear, although they were done at the pronculear stage.  Thus it is clear that HDR edits with ssDNA oligos can be efficiently done by cytoplasmic injections.   This is great because it results in higher rates of pup survival than pronuclear injection.

Still lingering questions for me:

1. Although the authors showed they could increase the insertion rate of a "large" cassette - a GFP-style reporter ORF - in cell culture, they did not repeat this experiment in embryos.  Or at least they didn't show the data.  Was there negative data to report?   Or just not enough live pups yet for them to feel comfortable with publishing a negative result?  Or have they not tried it yet?   The routine insertion of kilobase-sized cassettes in embryos is now my next CRISPR mountain to climb!

2.  I would kinda like to know if there may be an increased rate, or change, in the genome-wide mutation rate by SCR7 treatment. After all we are mucking around with the DNA repair pathway here.  Since each mammal embryo probably has on the order of 50-100 new mutations anyway, it would have to be a pretty substantial change in mutation rate to scare me off.   I'll bet there is no detectable effect.  Besides, NHEJ usually results in new mutations anyway - so I would imagine that we'd observe cell or embryo death following SCR7 treatment, long before we could observe a change in mutation rates or spectrum in surviving embryos.

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.  

The reported off-target effects in the recent Liang et al human embryo #CRISPR paper are partly incorrect.

As widely reported last week, a group in China has published results of CRISPR editing experiments in human triponuclear embryos (Liang et al, Protein & Cell 2015).   The news blurb in Nature is worth a read to get the context of the paper, which follows on the heels of a previous statement published in Science by leaders in the CRISPR field and others, in which they discourage CRISPR experiments in human embryos at this time pending further discussion of the implications of such research.    

In this post I won’t get into the ethical implications of the paper (which is more than I can deal with in one post anyway!).   Here I’ll discuss the technical results.   The Liang et al paper does not actually present much data that is very surprising - it is not unexpected that CRISPR can induce targeted mutations in humans, since it works in basically every species in which it’s been tried.    Their target gene was HBB (beta-globin) and they attempted HDR using the familiar approach of coinjecting Cas9 mRNA, guide RNA, and donor oligo ssDNA.

Here’s their 4 main points, paraphrased from the abstract:   

1. Efficiency of HDR was low.
2. Edited embryos were mosaic.   
3. Off-target mutations were evident.
4. A separate, highly homologous gene (HBD) could serve as donor template for repair, thus introducing sequences inadvertently from the other gene into the target gene.

Of these, points #1 and 2 were not surprising to those who have injected CRISPR reagents into mouse embryos, and not all that different.   The reported HDR efficiency was 14%, which is in line with at least some published mouse experiments (e.g. Singh et al 2014).   I will state that in our mouse core we are apparently seeing HDR results around 10-20% efficiency across several experiments.    Mosaicism has also been previously reported in CRISPR mice (Yen et al 2014).  Point #4 was kind of novel, but in retrospect not completely weird, since the HBD gene (delta-globin) is over 90% identical to HBB.

Ok, so regarding point #3 - off-target mutations...I was very interested in this because the authors reported four distinct off-target (OT) mutations in human embryos associated with the single CRISPR guide RNA they used, and this has already been described in the media as being substantially higher than OT rates in animal embryos.  Meaning, mouse embryos.      

One of these OTs was particularly surprising, as it looked like a very poor match to the target protospacer indeed - although the 3’-most 11 bases (the “seed” region”) matched the target, the 5’ bases only matched 1 out of 9 bases, for a total of 8 mismatches.    Frankly, this really scared me, because if true it means that the current methods used to predict OTs are not nearly broad enough.   But this level of OT mismatch was much greater than any OT I had seen before.

Bottom line: After looking at their data, I now firmly believe that only one of the four OT mutations were actually new mutations caused by CRISPR.  The other three were simply polymorphisms, already present in the  germline, that the authors mistakenly classified as OTs.

Here is how they did their OT analysis and my interpretation of the data.   Tripronuclear human embryos were obtained from a fertility clinic; you can identify these microscopically at the 1-cell zygote stage.  Fertilized by 2 sperm by accident, they are effectively triploid, and are absolutely unable to survive to term as normal pregnancies - but they can survive well enough during short-term CRISPR experiments, in which the embryos are only kept alive for a few days in vitro.   Briefly, 86 tripronuclear embryos were injected; 71 survived the injection; 56 of these were GFP-positive (used to as a reporter to show expression of injected reagents) and used for DNA analyses of on- and/or off-target effects.   28 of these embryos had on-target indel mutations and/or the desired HDR edit and were used for OT analysis.   

Note that they had originally chosen this particular CRISPR target from 3 potential targets they looked at in their gene;  one of these didn’t cut well and was not used further . For each of the other two, 7 sites were identified as the “top” potential OTs by using the MIT tool.  Of the two targets, one was found to have no OT mutations at the 7 potential OT sites when it was tested in 293T cells.  So they decided to work with this CRISPR target.  

2 of the 7 OT’s were found to have mutations in the injected embryos.  These were named G1-OT4 and G1-OT5 and are the first two of the four total OT mutations they claimed to identify (Figure 3A).  T7 mismatch assays were used for this analysis.   293T cell transfections with the guide RNA had already shown a lack of mutations across the 7 OT sites, that is, they were negative by T7 assays.  I’ll come back to these later.

They then did whole-exome sequencing on six of the embryos to identify potentially even more mutated OTs.  From this data, they first called indels and SNVs (single nucleotide variants) and then searched for protospacer similarity “allowing for ≤6 mismatches or perfect match of the last 10 nt 3′ of the gRNA” anywhere within 100 bp of the indels.  (Not sure if they did anything more with the SNVs.)   This identified two apparently new OTs, in the 3’ UTRs of  the C1QC and TTR genes, each found in one embryo (Figure 3B).  These were confirmed by T7 assays.

So - what does the T7 mismatch assay really indicate?  It reveals heterozygosity within the PCR product.   Of course, new mutations can cause this.  But so can plain old polymorphisms.   This is a drawback of using mismatch assays when applied to polymorphic samples.     

The next question is, simply, are there common human polymorphisms in the PCR products used in the OT analysis?  It’s easy to check this using the UCSC genome browser and the 1000 genomes site.   

For OT #1, a.k.a. “G1-OT4”, (Fig. 1C and 3A; PCR, hg19, chr11:132761837-132762356; intron of OPCML) there are no known common polymorphisms within the PCR that are close to the OT.    The closest SNP, rs79549129, has a minor allele frequency (MAF) of 1.2% but zero in asian populations.   There are no other annotated variants near the OT with a significant MAF.  The closest “common” SNP is rs2659601 but it’s about 50 bp from one end of the PCR product.  I don’t think that could produce the band sizes seen in the Fig. S3 T7 assays. From what I can tell from their Fig. S3 & S4, their T7 assays are compatible with new mutations that have been induced by cleavage close to the CRISPR OT site.   Thus, these look like “real” CRISPR OT effects at this site.    6/20 of on-target embryos, or 30%, had mutations at this OT.  So this looks like a real OT effect that replicates across embryos, but not in 293T cells.   

But then, polymorphisms become apparent in the other OTs...

For OT #2, a. k. a. “G1-OT5”, (Fig. 1C and 3A; PCR, hg19 chr22:31000551-31001000; intron of TULP4), there are two common SNPs on either side of the OT:  rs616358 (G/C) and rs628203 (T/C).   Haplotype derivations in Southern Han Chinese suggest haplotype population frequencies of ~56% GT, 35% GC, 9% CT, and zero % CC.  So we would expect to see plenty of heterozygosity in this PCR - easily observable by T7 assays, in the range of 50% or so being positive in a population based sample from this geographic location  - no CRISPR required.  This OT was a false positive.  

UCSC screen grab showing SNPs close to the OT (black bar in middle)

This leaves the two additional OTs they discovered by whole-exome sequencing.  Remember their workflow: they called indels in their data sets, then looked for nearby partial matches to the CRISPR target.   However, they apparently did not filter out known polymorphisms first.

OT #3 was in the 3’ UTR of the TTR gene.   Inspection of the OT sequence location (given in Fig. S6) on the UCSC browser clearly shows that rs143948820 is a known 9-base indel contained completely inside the OT.       Turns out that it’s uncommon outside of Asia but it has a MAF of ~2% in Southern Han Chinese.  With a heterozygosity of ~4% in normal diploids, it’s totally possible that 1 out of 6 triploid embryos would carry this variant.  This OT was very likely a false positive. 

UCSC screen grab; OT is black bar, indel variant is long red bar.

Finally, OT #4 was in the 3’ UTR of the C1QC gene.   And similar to the case above, this OT overlaps with a known 17-base indel, rs142916975, that has a MAF of 38% in Southern Han Chinese.   Heterozygosity should be close to 50%.   In fact, I’m surprised they got a false positive in only 1 of their 6 samples.   This is almost certainly a false positive.

UCSC screen grab; OT in black, indel variant in blue.

In summary, only 1 of the OTs holds up to scrutiny.    Importantly, neither OT found by exome sequencing holds up.  This flips their conclusion on its head: “Our whole-exome sequencing result only covered a fraction of the genome and likely underestimated the off- target effects in human 3PN zygotes.”.   While it’s certainly possible that some more OTs could be found by whole genome sequencing, the exome data was essentially totally negative.   Note that they chose a CRISPR to work with because it had a low apparent OT rate in 293T cells.   In retrospect, it was just by luck that G1-OT5 has a negative T7 assay in 293T cells.  It could have been heterozygous, but it's apparently not.

To their credit, the authors rightly restate that it’s going to be critical moving forward to carefully analyze off-target effects in any human applications of CRISPR.  This is widely agreed upon (see below).  However this paper made some technical mistakes in this regard.  While underestimating off-target effects could certainly have serious negative consequences for future CRISPR-based clinical treatments for genetic disease - and nobody wants that - overestimating them could generate an excess of hesitation to research the feasibility of such treatments within the broader scientific community.  

As stated by Baltimore et al in their Science commentary:

“It is critical to implement appropriate and standardized benchmarking methods to determine the frequency of off-target effects and to assess the physiology of cells and tissues that have undergone genome editing.”

I don’t think I’m blowing smoke here that we all need to get this right, as the media quickly reported on the Liang et al conclusions:  

From Wired:  “But—and this is a big but—using the technique without proper guidance could result in unforeseen consequences. The Chinese researchers, for example, found mutations in many of the embryos in genes other than the ones they’d targeted with CRISPR/Cas9.”

From Time, quoting Carl Zimmer from National Geographic:   “The experiment “came out poorly,” Zimmer says; in some cases, DNA was placed in the wrong spot and “off-target” mutations were discovered in the DNA.”

From the Washington Post:   “And in some of the embryos, the gene editing caused unintended mutations in other genes.”

From USA today (emphasis is mine): “The team also found that the complex used in the procedure was also acting on other parts of the genome, leading to other bits of it mutating. That happened much more than in previous experiments on adult human cells and animal embryos — and could happen yet more if the whole genome were used, as it would be if the embryo were to be implanted.”

            (OK, note from this last article the specific comparison to the very observations that I have blogged about in more detail than most people probably ever wanted to hear about...My point is that, due to the technical problems in the Liang paper, I don’t think we can yet say the off-target effects were “much more than in previous experiments on adult human cells and animal embryos”. )

One final note - my analysis of this paper should not be interpreted to mean that I fully endorse CRISPR experimentation or applications in human embryos.   I also applaud the authors' cautionary tone that the incomplete efficiency of CRISPR editing in humans is a problem that any therapeutic applications need to address.

Whew, this was the longest post yet.