Hello, and welcome to this
integrated DNA technologies
webinar As Cas12a 12 Ultra--
Precise Genome Editing
with a Mutant Protein
with the Reliability of Sp Cas9.
My name is Malcolm
McDougall, and I
will be serving as moderator
for today's presentation.
Today's presentation will
be given by Chris Vakulskas
and Bernice Thommandru.
Dr. Vakulskas is a
Senior Staff Scientist
in the molecular
genetics research
group at IDT Dr. Vakulskas
earned his PhD in microbiology
at the University of
Iowa, where he studied
genetic regulatory circuits and
pathogenic bacterial species.
After earning his PhD, he became
an NIH postdoctoral fellow
at the University
of Florida, where
he studied RNA binding proteins
and post transcriptional gene
regulation.
At IDT, Dr. Vakulskas has
managed contract research
projects, led process
development for CRISPR protein
purification, and develops
novel CRISPR proteins
including Alt-R S.p.
HiFi Cas9 Nuclease
3NLS and Alt-R A.s.
Cas12 CPF1 ultra.
Bernice Thommandru is
a research scientist
in the molecular genetics
research group at IDT.
She received her MS degree
in molecular physiology
and biophysics from
the University of Iowa,
studying transcriptional
regulation
of multidrug resistance and
pathogenic fungi of the lung.
At IDT, Bernice has
focused on optimizing
delivery strategies
for CRISPR reagents,
as well as methods for
increasing the rate
of homology directed repair.
The presentation should
last about 30 to 40 minutes,
and following the
presentation Chris and Bernice
will answer as many questions
as possible from attendees.
The question and
answer session will
be conducted by Molly
Schubert, research scientist
in the molecular genetics
research group at IDT.
As attendees, you
have been muted,
but we encourage you to ask
questions or make comments
at anytime during or
after the presentation
by entering your
question in the Q&A box.
Also, please note that you can
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In case you need to
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We will also post the recorded
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You will receive links to
these in a follow-up email.
And now, let me hand it over
to Chris for the presentation.
All right.
Thanks, Malcolm.
Good afternoon, everybody.
I'm excited to
share with you today
a project we've been working
on for the last few years
regarding a mutant form of A.s.
Cas12a known as the A.s.
Cas12a Ultra.
This was formerly known as
Cpf1, CRISPR editing enzyme.
And today, I'm going to run--
so this is the outline of what
I'll be speaking about.
I'm going over the
basics of genome editing
very quickly, touch
very quickly on the RNA,
and then finally settle in on
the topic of today's webinar,
the newly evolved mutant.
And then lastly,
Bernice is going
to go over some HDR
protocols and data
regarding this new enzyme.
So in general, the two most
popular genome editing enzymes
that people are using would
be Cas9 from strep pyogenes,
and then what is now known as
Cas12a from Acidaminococcus
or Lachnospiraceae.
And in general, what people
are using these enzymes to do
is to introduce
double-stranded breaks
into the genome of
living organisms
to do one of two things.
First, in the bottom
left-hand side,
to facilitate recombination
to introduce entirely
new DNA sequences through
homology-directed repair, HDR.
Or otherwise, on the right, to
allow error-prone native repair
pathways to allow insertion or
deletion of a few nucleotides,
otherwise known as an indel.
And Cas12a is a little different
than perhaps the more popular
SpCas9.
They're both RNA-guided
endonucleases.
The Cas12a variant that
IET promotes the use of
is from Acidaminococcus,
otherwise known as the A.s.
Variant.
There are, of course, a
few different proteins
from a few different
species out there.
And unlike Cas9, which
uses two separate RNAs
to guide the enzyme,
Cas12a is actually
natively guided by a single
RNA that's very short--
41 to 44 nucleotides
in length, depending
on how long your targeted
spacer sequence is.
Whereas Cas9 generates a
blunt-ended double-stranded
break, the Cas12a
enzyme actually
leaves a staggered-ended cut.
And then finally, and
probably most importantly,
the PAM site for Cas12a is
quite different than Cas9.
Cas9 has an NGG PAM sequence,
and the Cas12a enzyme
has a triple-TN or
triple-TV PAM site.
And we'll get into
a little bit more
of the specifics of
that in just a moment.
And at [? 9018, ?]
we recommend using
ribonucleoprotein or an
RNP-based delivery of CRISPR
reagents.
And one of the main
reasons we promote
that is because it's very,
very simple and easy.
It's associated with less
toxicity and less off-target
editing.
And this is just a quick cartoon
showing just how easy it is.
You can obtain purified Cas12a
protein, chemically synthesized
targeting RNA.
Add those together
in a test tube
on the benchtop for 10 to
20 minutes, and then those
can be delivered into
live cells by whatever
your favorite delivery
mechanism for whatever
your ultimate application--
electroporation, lipofection,
microinjection, what have you.
And so it's very,
very simple to use.
And what I'll do before I
get into this new enzyme
is give you an idea
of where we began
and why we decided to
evolve a new mutant.
We started with an
early version of Cas12a.
A couple of scientists
in the laboratory
used a little over
200 different RNAs
to look at different sites that
target different types of PAM
sites for Cas12a.
This is done in standard
human immortalized cell lines.
And we took the data
from this experiment--
on the y-axis, you could see
its total editing efficiency,
and then we rank-order the
performance on the x-axis.
And then we parsed everything
out by the particular PAM site.
And what we noticed is that the
triple-TA PAM sites facilitated
on average the highest
level of editing,
and the quadruple-T PAM sites,
in that sort of green or yellow
color--
those are by far the
worst performing sites.
And then the triple-TC NGs
were somewhere in the middle.
And when we sum this data
together, ultimately what
we wanted to do was
to use this system
and make it as
functional as Cas9 is.
We sum this data together
and compare it to Cas9.
In blue-- this is, again,
the same type of format.
Y-axis is total
editing efficiency.
X-axis is rank-ordered for
each of the different sites--
in blue is the first
version of our Cas12a
when we allowed the use of all
of those different PAM sites,
including those very
poor quadruple-T sites.
And in orange is what
we had seen for Cas9.
So there's quite a
bit of difference.
The Cas9s seem to be much
more reliable in general.
And then we were able to
make Cas12a more reliable
simply by telling people,
OK, don't use those very
bad quadruple-T sites.
You can see that
that blue set of data
gets shifted far
over to the left,
resulting in the gray
series of data, when you
don't use quadruple-T sites.
So this is what we saw with
one of our first offerings.
We decided to take a little
deeper dive into this
and ask, why is this system
not performing as well?
One of the problems is
simply that the enzyme is not
delivered as efficiently
as it could be,
so we're getting poor
nuclear delivery.
And secondarily, we noticed
even in a cell-free context
or in bacteria, this
enzyme has inherently lower
enzymatic activity.
So it's just not
as potent as Cas9.
So we wanted to
take these problems
and use protein engineering
techniques to try and solve
the challenges.
The first of which is
by using simple things
like changing protein linker
sequences, the things that
are outside of the Cas12a
open-reading frame itself.
So we played around with
a lot of different nuclear
localization signals, put those
on either end of the protein,
put multiple different ones,
different identities-- changed
all those things, and
ultimately settled
on what worked best for us.
And then, second,
to solve the problem
of lower intrinsic
enzymatic activity,
we embarked on a directed
evolution screen of bacteria
to screen for mutants.
So these are mutations
actually within
the Cas12a
open-reading frame that
generated an enzyme with higher
intrinsic enzymatic function.
Starting with the
first, easy strategy--
so on the left-hand
side, this is
data we found for a really
nice academic publication,
just demonstrating what
happens when you do or do not
have a nuclear localization
signal on a non-human protein.
So on the left-hand
side, you can
see all the GFP is
cytoplasmic-bound
when you don't have an NLS
appended to the protein.
And on the right-hand
side, when you
add the right NLS in
the right context,
the vast majority of
the GFP fusion protein
ends up where it's supposed
to be in the nucleus.
And then this
illustrates what we
did between our V1, or
our version 1 offering,
and our V3 Cas12a offering.
In the data set on
the right is-- again,
this is the same y-axis gives
you the editing efficiency,
and x-axis gives you the
ranked-ordered performance--
showing you the difference
between, in blue,
our first offering, which had
poor nuclear delivery, and then
our current website,
offering up the wild-type end
in the V3 enzyme, in this
yellow or green color.
So the data set has shifted
remarkably to the left,
showing you that just
making the nuclear delivery
better makes the overall
performance better.
So that's a little
bit of data that
summarizes a heck of
a lot of work finding
the best form of Cas12a, or best
form of the wild-type Cas12a.
So then we moved on
and asked, what can we
do to the actual Cas12a
protein sequence?
Here, what we did is we
used a similar strategy
that we used to generate
our HiFi Cas9 enzyme,
where we set up a cleavage
reporter system in E. coli.
There's a cartoon here
of how this works,
and I'll try to
briefly summarize it.
In general, we have a
high-copy plasmid in E. coli
that has two very
important things on it.
One, it has a toxin
that, when induced,
will kill the bacteria.
And the second part
is, it has a target
site for either a good
PAM site for Cas12a,
triple-TC, or a bad
PAM site, quadruple-T.
Basically, what
happens is if you
get survival in this
system, that results
from successful Cas12a cleavage.
And the mechanism
behind that is,
bacteria do not like,
in general, linear DNA
to be hanging around.
So when a plasmid encounters
a double-stranded break
and is linearized, the
bacteria have exonucleases that
rapidly degrade those plasmids.
And we're asking Cas12a
to linearize the plasma,
to degrade it before the
toxin can be induced.
So if we look on the bottom
left-hand side with a good PAM,
we put this into bacteria.
The minus toxin is
simply a control
that shows you everything
that could have survived,
and the plus toxin
represents the number
of colonies that represent from
successful Cas12a cleavage.
So with the wild-type enzyme,
with a good triple-TC PAM,
we get a high percentage of
survival in the plus toxin
case.
Now, moving over to the
right, with a poor PAM site,
a quadruple-T PAM--
the minus toxin,
again, a control
that shows us everything
that could have survived.
And then, with
plus toxin, you can
see a vast majority of those
controls do not survive,
because CPF1 struggles
to cleave this site.
And this is what we
used for the screen.
So the basic strategy was
to generate a mutant library
of Cas12a using
low-fidelity PCR,
and then put it through this
series of bacterial strains
to select for mutants that could
survive with a poor PAM site.
And we did several different
rounds of screening.
We actually selected
for surviving colonies,
isolated the DNA, put
those back into bacteria--
we did this five times, having
coupled all of the results
to NGS.
And we plotted the
redacted data here
on this chart, where the higher
the percentage of mutation rate
on the y-axis, the more enriched
that particular mutant is.
So the better those mutants are
cleaning that quadruple-T PAM
site.
These are the three mutant
hot spots that we found.
The mutant 1 and 2 seem
to be the overall best.
And these are color
coded by round,
so these are the ones that were
enriched over the fifth round
of mutagenesis.
Through a lot of different
tinkering with this,
and combining, mixing, and
matching different mutations,
we ultimately
settled on something
that gave us the highest
efficiency without compromising
other properties of the
enzyme, for which we're calling
the AsCas12a Ultra protein.
And for the remainder
of this talk,
I'm going to go over
some of the performance
properties of this mutant.
But this is essentially
how we made it.
So moving right
along, what we'll do
is we'll compare this
new mutant enzyme
to the V3 wild-type version that
we're selling on the website
as well.
So the wild-type
data are in blue,
and then the new Cas12a
Ultra mutant are in orange.
And we've done this
in two different ways.
We've shown when we allow
for all possible PAM
sequences, including those
very poor quadruple-T PAMs,
triple-TN.
And now we've also
shown the data
when we subtract those out.
So you could do it every way.
What we find is the
best overall reliability
and potency we
achieve with Cas12a
is when we use that Cas12a
Ultra enzyme, in orange,
and when we subtract out
all the quadruple-T PAMs.
So using triple-TV and using
the Cas12a Ultra enzyme
gives you a vast majority of
sites that get nearly 100%
indel rate by NGL.
This is a phenomenal change that
we find is better than anything
we've ever seen
for CPF1 or Cas12a.
We further parse that data out
into the individual PAM sites.
So this is the same data
set, just showing you
the performance
difference that's
actually seen as a function
of the particular type of PAM
site.
And you can see that for
every type of PAM site,
moving left to right,
triple-TA, there's
a huge performance boost
going from wild type to Ultra.
And the key thing when
analyzing this box plot
is to look at the
averages, which
is the line in the
middle of the box plot.
So you can see it's
around 65%, maybe,
for triple-TA and
wild type, that
gets boosted to nearly
100% of the Ultra mutant.
And that trend
continues down the line.
And even the quadruple-T
PAMs in orange--
you can see they do not
perform very well at all
with the wild-type mutant.
You can see that number goes
up quite a bit with Ultra.
But nevertheless, on average,
even with the Ultra mutant,
quadruple-T PAMs
are inconsistent.
So we're still recommending that
people select triple-TV target
sites if possible.
And ultimately,
what we wanted to do
with this enzyme was
produce something that was
as reliable and potent as Cas9.
That was the idea we came
up with before ever doing
any screening.
So ultimately, we wanted to
actually do that experiment
with whatever we found.
And since Cas9 and Cas12a
use entirely different target
sites-- the PAMs couldn't
be more different--
we obviously couldn't
target the exact same site.
But the approach we used was to
take a large number of sites,
and then pick target sites for
each enzyme that were as close
together in the same
genomic locus as possible,
and then look at a large
amount of data in the aggregate
and ask, how useful
is this enzyme,
relative not only to our prior
offering, but now to Cas9.
And we parse that data out in
terms of triple-TN on the left,
and then triple-TV on the right.
And you can see, if you
look at our first offering,
the V3 offering in blue-- this
is just the wild-type enzyme
with optimized NLS--
it is not nearly as
reliable as SpCas9 in grey.
Because then the average goes
from somewhere around 15%
to up to 90% for Cas9.
But if we now look at the
Cas12a Ultra mutant in orange,
the average is very, very
similar, and maybe just
a fraction above,
what we see with Cas9.
And then if we move to
the right of the panel,
you can see when using
triple-TV sites-- again,
the wild-type V3
Cas12a enzyme in blue--
not nearly as good or
reliable as Cas9 in gray.
But now, our Cas12a
mutant in orange--
the average is
significantly higher
than even the Cas9,
indicating that we actually
did what we set out to do.
We set out to make
something that
was as reliable
and potent as Cas9,
and I believe we achieved that.
And most of this data has
been in either HEK293 cells
or Jurkat.
We've expanded that to a
series of different cell lines,
and although we
can't show the data,
we have sent this to
some collaborators, who
are working on plans.
And they've seen some very, very
positive, encouraging results.
And along those lines,
I had previously
mentioned that there are other
sources of Cas12a out there
that come from different
bacterial species,
and why would you
pick one over another.
Well, the reason for
that is that when
working in ectothermic
organisms like plants, things
that you need delivery to occur
in temperatures lower than 37,
it was previously determined
that the Lachnospiraceae,
or L.b.
Cas12a variant, actually works
better at lower temperatures.
And we actually find
that in-house as well.
But since our Ultra
mutant-- which,
again, is an A.s.,
Acidaminococcus derivative--
since it has overall
higher enzymatic activity,
we asked if it
could possibly serve
as a universal
solution for Cas12a,
even at lower
temperature, and perhaps
work better than the wild type.
And that's, in
fact, what we see.
So if you look on the
left-hand side, at 30 degrees
you can see, consistent with
what others have reported,
the L.b.
Cas12a variant is superior
to the wild-type Cas12a.
However, the Ultra
mutant is actually
superior to both
wild-type forms,
regardless of whether
it's from L.b.
or A.s.
So since the guides are slightly
different between the Cas12as
from those two
species, we had hoped
this would be a
universal solution,
so you'd only have
to order the A.s.
RNA.
And that's in fact what we see.
And then on the
right, at 37 degrees,
is the same story
we've shown before.
The two various wild-type
variants are very similar,
but the Ultra
mutant wins the day.
So we believe that the
Cas12a Ultra protein
is a universal solution.
Whether you're
working in organisms
that require
low-temperature delivery
or whether you're looking at
37, no need to switch RNAs.
We've also analyzed some of
the biochemical properties.
I'm just going to
summarize this data.
What we found with
the Ultra mutant is,
the change is actually
not in its specificity.
We haven't broadened
the PAM space.
All we've really done
is change the affinity
for the target site.
And what this shows is that the
Cas12a Ultra mutant chemical is
on the y-axis.
The wild type is on the x-axis.
You can see this
trend of data trends
towards the Ultra
mutant, indicating
that the DNA binding
affinity for the target site
is, in fact, higher.
And then, probably more
importantly, as I mentioned,
we've also looked
at specificity.
First, in the biochemical
sense, looking
at three different sites,
comparing wild type on top
to Ultra on bottom, we
find that both specificity
within the spacer as well
as within the PAM space
is essentially identical.
So unlike some of the
literature-reported mutants,
which have both higher
intrinsic enzymatic activity
and broadened PAM
space, our mutant
seems to strictly have higher
intrinsic enzymatic activity
without the concern of reduced
target site specificity.
And I think that matters,
because this enzyme has
been sold as a good
option for genome editing
because it has intrinsically
higher fidelity than Cas9.
And that was the last thing
we wanted to change with it.
We wanted to keep that
good property of it,
but at the same time make
it more useful for people.
And I think as soon as you
start making the PAM site less
predictable, the
higher the likelihood
for unintended off-target
sites to occur.
So one other thing
I wanted to mention
is, for the wild-type
version of Cas12a,
much like for Cas9, we have an
enzyme-specific electroporation
enhancer, a Cas12a
electroporation enhancer.
So for people using
that delivery method,
that particular product
is pretty essential
to get the maximum levels
of editing you can see.
We wanted to know if that was
true for this new Ultra mutant
as well, and in fact that
does seem to be the case.
So you can see that performance
is dramatically higher
when you have the Cas12a
electroporation enhancer.
Now, this is the same
product, whether you're
using the wild type
or the Ultra mutant,
and it's already on
the website to order.
So very, very
important for customers
using electroporation delivery.
And then, finally, for
my portion of the talk,
I'll just summarize
what we found.
When we had originally set
out to use Cas12a in-house,
we found that it had
poor nuclear delivery
for a particular clone.
The enzyme had lower
intrinsic activity,
both in a cell-free
context and in bacteria.
We set out to
improve those things.
We used optimized nuclear
localization signals
and linkers to make
delivery better.
And then we had a directed
evolution scheme in bacteria
that we used to isolate what
we are now calling the A.s.
Cas12a Ultra protein.
And finally, when testing
this enzyme and using it,
if you're using
electroporation, we still
recommend that you use the
Alt-R Cas12a electroporation
enhancer.
So I'll be available
to take questions
about this section of
the talk at the very end,
but for the time being,
I will turn this over
to Bernice to talk about HDR.
Thanks, Chris.
Hi, everyone.
My name is Bernice.
I've been a researcher
at IDT for three years,
and at least two of
those years my research
has focused on improving
HDR rates with Cas12a.
So I'm really excited
to have the chance
to show everyone what
we've discovered here.
So let's jump into it.
I'll go over some basic
guidelines of HDR with Cas12a.
I'll start with the benefit of
using a high-activity enzyme
to mediate cleavage, and also
talk about how best to use
a potent enzyme like that.
And since most of what
I'm presenting today
is HDR using
single-stranded donors,
I'll talk a bit about strand
preference for the donor
sequence, as well as how
best to position your insert
within that donor molecule.
And finally, I'll discuss
some additional enhancements
that can help boost your
HDR rate even further.
So with that outline,
let's look at some data.
We heard Chris talk a lot
about the improvements
that Cas12a Ultra offers,
specifically the fact
that the on-target
rate of editing
is greatly improved over
the wild-type enzyme.
One thing that's common
to all types of HDR,
even beyond the
Cas12a context, is
that maximizing
editing efficiency
is key to maximizing HDR.
In this figure, there's
a direct comparison
of the wild-type
and Ultra enzymes,
and this is 110 sites
tested in Jurkat cells using
short single-stranded
DNA donors.
The top panel shows
editing efficiency,
and the bottom panel shows the
corresponding perfect HDR rate.
And these are all
measured by NGS,
for each of these 110
sites divided by PAM.
In the top panel, with the
wild-type enzyme in blue
and the Ultra enzyme
in orange, it's
very obvious to see that
the average rate of editing
is significantly improved
when the Ultra mutant is used
for every site, including TTTT.
As Chris mentioned before,
generally we recommend TTTV,
but the TTTTs are
also available.
In the bottom panel,
showing HDR rate
with the wild type in
dark green and the Ultra
mutant in lime green, we can
see the HDR rate improve simply
by using the Ultra
enzyme and improving
the overall rate of editing.
And the main takeaways
here are that not
only is it important
to use a potent enzyme,
it's also important
to pick a guide
RNA with a high rate of
cleavage that can help maximize
your potential for HDR.
And also, a quick
note I wanted to make
is that we know it's been
reported in recent literature
that Cas12a has some
non-specific single-stranded
DNase activity.
But we've seen that if
you're using a potent enzyme,
like Cas12a Ultra, you
can titrate back your RNP
dose to where you're still
achieving very high editing,
but your single-stranded
DNA donor
isn't at risk for degradation.
So next, I'll talk about
strand preference for the donor
sequence.
In the cartoon at
the top left, you'll
see that what I'll refer to
as the targeted strand is
the strand to which the
guide RNA is complementary.
And then the non-targeted strand
is just the opposite strand
of the target strand.
So that would not be
complementary to the guide
or any sequence.
When designing an HDR
donor, you can use sequence
from either the targeted
or non-targeted strand
for the homology arms
of your donor molecule.
In the graph below,
we've directly
compared using either the
targeted or non-targeted
strand sequence for HDR at eight
human sites in Jurkat cells,
where the bars represent
perfect HDR by NGS
and the dots represent the
total editing efficiency.
So I'll direct your eyes
to the orange dots first,
to show that when using the
targeted sequence as your HDR
donor, this causes
the total editing
to decrease, in comparison to
when the non-targeted strand is
used, which is the
blue dots up above.
As we see in the bars below,
the corresponding HDR rates
also decrease when the
targeted strand is being used.
What we're observing here
is that the donor molecule
that contains the guide
complementary sequence--
when using the
targeted strand, this
is interfering with RNP binding,
and therefore dragging down
the overall rate of editing.
As I mentioned in
the last slide,
editing and HDR are
very closely linked.
If you're dragging down editing,
you'll also drag down HDR.
And the strand
preference is something
that we've observed as
being unique to Cas12a.
We've not seen such a strong
strand preference for Cas9.
And also, towards
the right-hand side
of the graph, where
editing is most robust,
there's not as much of a
strand preference, here.
But if you're unsure
of which strand to use,
or you are using more of
a mid-range editing site,
we do just recommend to use
the non-targeting strand
for your donor sequence, just to
sidestep this problem entirely.
So now, I'll talk about how to
place your insert in that donor
sequence.
In this graph, the same guide
RNA is used with 15 different
types of single-stranded
donors, where each donor has
the insert--
in this case, an EcoR1
restriction site--
each donor has the insert
placed a different distance
from the PAM.
The x-axis below is distance
from the PAM and base pairs.
In the direct
middle of the graph,
the insert is placed 18
bases from the PAM, which
is generally
considered the position
of the five-prime cut of the
staggered cut that CPF1 makes.
Directly left of that, inserting
16 bases from the PAM--
so just two bases shifted over
towards the five-prime end--
there's a dramatic
improvement in HDR.
And most importantly,
what we see
here is that there's an optimal
window of HDR insert, which
is about 8 to 16 bases from
the PAM, where HDR is highest.
If you compare that optimal
window to the sequence
above, showing both the PAM
and the surrounding sequence,
we can see that that optimal
window, 8 to 16 bases
from the PAM, is right in
the middle of the protospacer
sequence.
And
Again, because of Cas12a's
tight binding affinity
and specificity, we find
that using your insert
to fully disrupt that
protospacer sequence can
prevent repeated
rebinding and recutting
by the Cas12a RNP over time
as these edits are being made.
This can allow for
higher HDR rates
simply by moving your insert
just two or four bases away
from that five-prime cut.
Now we've looked at
the insertion profile
for a number of sites
in different cell lines
and different cell
types, human and mouse.
What we seem to find is that the
insertion profile differs site
to site.
That means that the
optimal window of insert
may be different for your
project-specific guide RNAs.
There may be multiple
plans available to you
in the space around
the HDR base change
you want to make so we recommend
testing these different guide
earnings in combination
with your insights
and different
in-store positioning
to find what would give
you the best HDR rate
and what would maximize
your potential for HDR.
Lastly, I want to talk about
a couple other factors that
can help boost your
HDR rate even further.
The first of that is IDT's
Alt-R HDR Enhancer product.
This is a small molecule
enhancer and solution
that when added to
cells post-transfection
helps bias the repair
pathway towards HDR,
therefore boosting
your HDR rate.
And we know a number of
these so-called HDR enhancers
have been reported
in the literature.
And the main
difference between all
of those in our HDR
enhancer product
is that our product
is very effective.
So on the graph on the
left, you can see here
that the untreated and DMSO
controls in this example
give about 10% HDR.
Using our Alt-R HDR enhancer
at our recommended dose,
that rate goes all
the way up to 44%.
So that's a
four-fold improvement
just using our HDR enhancer.
And the other
compounds we've tested
in this experiment, SCR7,
Brefeldin A, and a few others
from the literature
that we found just
didn't deliver on the
promise of improving HDR.
And some even were
detrimental to the HDR rate.
And our Alt-R HDR
Enhancer product
we found to be effective
for both Cas9 and Cas12 HDR
with multiple donor
types, whether that's
single-stranded
or double-stranded
with long inserts like megamers
and in multiple cell lines.
And this graph on
the right shows
some of that data where
we've tested our HDR
enhancer in four of our
common human cell lines--
Jurkat, HEK293, HeLa,
and K562, and found
that our HDR enhancer
really helped boost
HDR in all of those cell lines.
So this last figure
encompasses everything
I've talked about how to
get good HDR with Cas12a.
I've used Cas12a Ultra
at a moderate dose
for the high cleavage rate.
The donors were designed to
have the EcoR1 insert placed
16 basis from the
PAM to fully disrupt
that protospacer sequence
for each of the 20 sites
tested here in this graph.
And the non-targeting
strand sequence
was use for the donor designs.
And the last element that
I wanted to mention here
is that these
single-stranded donors that
were chemically
modified did contain
two phosphorothioate
linkages at either end.
And this is important to protect
the single-stranded donor
from any excellent
nucleus activities that's
present in the cell.
This is especially important
for high nucleus cell
types like Jurkat
and IPSEs where
can be very difficult
to mediate HDR.
And here in the graph we
can see through the middle
and towards the right-hand
side where editing
is most robust, we're seeing HDR
rates of 40%, 50%, almost 60%
with no sorting, no fuss, just
using all of the guidelines
that I've laid out here today.
So that's my
portion of the talk.
I wanted to sum up some
conclusions for you.
For doing Cas12a HDR,
we recommend following
these simple guidelines.
Number one, use a Cas12a enzyme
with a high rate of cleavage
like A.s Cas12a Ultra.
Use the non-targeted strand
as the donor sequence
and position your insert such
that the protospacer sequence
is disrupted.
And lastly, you can use modified
donors and Alt-R HDR Enhancer
to further boost your
HDR rate even further.
And for more info on any of
our products and to see some
performance data for
the Alt-R HDR Enhancer,
you can visit our website
idtdna.com/CRISPR.
And some overall
conclusions we'd
like you to take
away today, A.s.
Cas12a Ultra is a new
genome editing solution
that is as potent and
reliable as S.p Cas9.
And we offer this in
500-microgram and 100-microgram
quantities.
We can do custom protein orders
if you house a specific yield
or formulation that you'd like.
And you can just contact us at
CRISPR@idtdna.com via email,
and we can get that
process started.
Our Alt-R Cas12a crRNAs and
Cas12a Electroporation Enhancer
are your best bet for
maximizing editing efficiency.
Our crRNAs are available
in either single tubes
or in 96 well plate formats.
They come in two nanomole
or 10 nanomole quantities
on the website.
But again, we can offer
custom crRNA orders
if you have a specific yield
or modification pattern that's
interesting to you.
You can again contact
us at the CRISPR email,
CRISPR@idtdna.com.
And lastly, Cas12a
is great at HDR
and HDR rates can
be maximized simply
with proper donor design and
using modified donors using
Cas12a Ultra and lastly using
the Alt-R HDR Enhancer really
boost that HDR rate.
And for more info, please
visit our website site
at idtdna.com/CRISPR.
There's lots of
great performance
data for all of our
products and more info
right on that website.
So that's all the data
we have to show today.
Chris and I wanted to
say thank you to everyone
for joining us today.
And we'd be happy to
take any questions.
Great.
And thank you Chris
and Bernice for
your informative presentation.
If you have a question and
have not done so already,
please type it into the Q&A box.
OK, there are a few
questions for you two.
Here's the first one.
With regards to
your HDR Enhancer
and the electroporation
enhancer,
are they ever used together?
Absolutely.
We see them as two different
ways to boost your HDR rate.
The Electroporation enhancer
helps boost editing.
And the addition of the Alt-R
HDR Enhancer helps boost HDR.
So we do use them in
tandem all the time.
Great.
Another question here.
Chris, this one might
be great for you.
What is the size of Cas12a
Ultra compared to Cas9?
And when would you want
to use one or the other?
A follow-up to that,
have you tested
the Cas12a Ultra in vivo?
Yeah.
So they're about--
they're very similar size.
And importantly,
in terms of what
we have available
commercially, it's
about the same concentration.
Cas9's about 61 micromolar,
the Cas12a's about 62,
so approximately the same, both
at 10 mg per ml quantities.
And we've certainly used
both in living cells
and show great performance
with both wild type and then,
of course, accompanying
improvement with the Ultra
variant.
And I think there's
a second part of that
question I might have missed?
Have you tested in vivo, I
think you've answered that.
Yeah, yeah.
Yep.
OK.
Yeah.
So one more question here
back to the enhancers.
Are these designed
for mammalian cells,
or have they've been tested
with other organisms like plants
or fungi?
We don't do plant and fungi
testing here in-house.
We just don't have
the bandwidth for it.
But we are always
looking for beta testers
to test our products
in their cell types.
I don't know that we have
any data ready to share
for the HDR Enhancer in fungi.
OK.
Have you compared the
efficiency of this Cas12a Ultra
with other published
mutants in terms of activity
or namely MAD7 from Inscripta
says this question here?
Yeah, so MAD7's an
interesting one.
We have never looked
at that internally,
so we don't have
data to compare MAD7.
We have evaluated some
other variants or mutants
of Cas12a that have come
out throughout the years.
And it's a tough
thing because there
are a lot of
different things that
go into summing the
performance of an enzyme.
Part of it is how well
it gets in the nucleus.
The other part of it is the
inherent enzymatic activity.
And just to summarize all
the data we've produced
is we have never
found anything that
is as potent as this
particular mutant in terms
of its combination of NLSs
and then on top of that
the mutations that
we've introduced.
It's simply the best
that we've seen.
Great, thank you.
I've gotten a few
questions here actually
that Bernice you might
be able to answer
about the optimal length for a
donor DNA and the hemology arms
that you're using,
what length those are.
So a majority of the
data that I showed today
is about 100 total basis for--
or nucleotides for the
single-stranded donor.
We usually choose to
use around 40 base pairs
for the hemology arms, and then
the insert varies from there,
so usually around 100 for the
total length of the oligo.
Great.
And a follow-up to that,
this is a two-fold question
from this researcher.
One, choosing the non-targeted
strand for an HDR template
design is only an option when
using a single strand of DNA
template.
Is that correct?
Yes, I'd say so, yep.
And then the second part of
this is, have you ever tried
adding three prime
overhangs such as UAU
to the crRNA to increase
on-target activity?
Yeah, that's an interesting one.
So I believe there
have been, I think,
four or five
different things that
have come out about adding
different extensions
to either end of
the crRNA, things
as short as a couple of
U's, as was mentioned.
But there have also been
some things about appending
an entire tRNA at the end.
And we've looked at, again,
a wide variety of these.
And what we found is that in
the context of an unmodified
RNA, so something that has
no two primal methyl bases,
has no N blockers, in the
context of a completely naked
RNA, some of those
modifications are
superior to just
the standard crRNA,
but it's very guide specific.
And what I'd finish
on is that what
we have commercially
available is an end block,
a proprietary end-blocked RNA.
And we haven't found any
of these end extensions
that are superior to
the end-blocked RNA
that we have available
on our website.
Great, thank you.
This is a question about
the actual delivery.
They're wondering if you
could elaborate a bit further
on the two micromolar
RNP composition.
For example, how much crRNA
and how much Cas12a or Cpf1
is individually delivered?
So generally we mix our
crRNA and the Cas12a enzyme
at a one-to-one ratio.
And we deliver that at
a final concentration
of two micromolar.
So, most of our
experiments are using
the [? MAX ?] nuclear factor.
And those are very
small reaction volumes.
So we'll mix our RNP in a
small, five microlayer volume.
And then the total volume
will be 20-some microliters
where the final concentration at
the time of the electroporation
is two micromolar.
Thank you.
I think I just want to add
one little thing under that.
I think that on
the subject of dose
and all that kind of stuff,
we have recommendations
that I think are the best
idea we can put out there
based on all the available
information we have on hand.
But we can't test
everybody's application.
And I think, in general,
the best approach
is to do a dose response
to make sure that you're
trying everything you can
with your specific application
that we may not have
looked at internally.
We don't have all the
nuances to every experiment
that people might have
in their own labs.
So I think a dose response is
always going to be prudent.
And I would encourage
anybody who finds--
and it maybe in a system
we haven't tested--
that finds a specific dose
requirement or delivery
requirement to reach
out to us because we'd
love to hear about it if you
find some specific dose that
works best in a
particular application
that we may not have tested.
Very good points.
Do you recommend to have
an incubation period--
typically overnight at
a lower temperature,
lower than 37 degrees
post electroporation?
Can you take that?
I'm curious, is that a
question about HDR or just
getting indels?
It came in pretty early in the
webinar before the HDR section.
So I think this is in regards to
the total editing at the lower
temperature data
that you showed.
Yeah.
So the data that we showed,
we maintain the cells
before and after
electroporation at 30 degrees.
So we were able to obtain
those efficiencies, editing
efficiencies, just by keeping
the cells at 30 degrees.
I would point out
that, as I mentioned,
we also have sent
this out to people
that are working in plants.
And they've gone as low
as 27, 30, 32 degrees.
And they've seen great editing
rates with the Ultra mutant
at those temperatures.
So I think--
I think you could
certainly expect
it to work well for the vast
majority of cases at lower
temperatures by maintaining
throughout the entire delivery
experiment.
So this question
is about targeting
multiple different
genes in parallel.
They're asking if you
should prepare separate RNPs
and then electroporate
them together,
or would you prepare a single
RNP with several crRNAs?
Yeah.
So we've done a bit of testing
with multiplexing RNPs.
And we did generally
recommend to mix your RNPs
and then combine them
right before delivery.
OK.
So this is a question about
the PAM, non-canonical PAMs.
They're wondering how
the Ultra mutant performs
on, for example, a
TYCV and TATV PAMs,
the non-canonical
PAMs for A.s Cpf1.
Yeah.
So as I mentioned, some
of the literature variants
that have come out of
the last couple of years
have shown expanded PAM
space on top of higher
intrinsic enzymatic activity.
And we had wondered if the same
would be true of our mutants.
We certainly didn't screen--
excuse me-- screen for mutants
with expanded PAM space,
but we wanted to look
at it because that's
what others had seen.
And we found no indication that
we had expanded the PAM space
outside of enabling more of the
quadruple tPAM sites to work.
We haven't looked at an
exhausted list of those PAMs.
We have looked in a in
vitro biochemical sense
to ask what types of PAM
sites are targetable,
and we did not find
expanded target space
whereas we did with some of
these other literature mutants.
Great, thank you.
So back to HDR,
someone is wondering
what is the largest piece
of DNA that has been
inserted with Cas12a by HDR?
Good question.
That's a great question.
What's the largest
insert we've tested?
This is a new space for us
is testing larger inserts.
I don't know that I have
any one number to give
of what's the last insert.
We've done some
megamers with it maybe?
That data is in the
process right now being
tested with larger inserts.
And hopefully we'll have an
answer for you on that soon.
Yeah, it's funny, having
this enzyme be as efficient
as it is for indels alone was
as surprising enough to us.
And we had-- this lends
into another question
I think might come up.
We had assumed that one of
the reasons that we previously
saw poor HDR with Cas12a
was because of this reported
inherent nuclease activity.
And we thought that
maybe the donors were
being degraded in the cell.
But when we isolated this
mutant and started testing it
for HDR, which is a fairly
recent thing for us,
we found that the HDR
rates seem to go way up,
suggesting that all that
we were really limited by
was the inherent
nuclease activity
of the enzyme for standard
targeting and not--
we weren't limited by
degradation of the donor.
So a lot of this is
pretty new to us.
And we're hopeful
that we'll come out
with some of that data
in the near future.
Yeah.
These two questions are related.
So we're talking about
off-target effects.
So have we tested for
off-target effects
with the new Ultra mutant?
And then does the
Ultra mutant have
the same fidelity as our
HiFi mutant that we have?
Yeah, so we've looked with
the Guide-seq procedure,
and we find pretty
much the same.
Looking in, I think,
four different sites,
we found pretty much
the same profile.
We found higher
on-target editing.
And we saw the same site.
We didn't see expanded sites.
And we saw about the
same level of editing
with a few modest increases
in off-target editing.
So, in general, we believe
that this mutant maintains
the same specificity profile.
But we've gone back and
looked into cell-free context.
And again, we find that it's
still very, very specific.
And one thing we did for the
high fidelity Cas9 was to use
Guide-seq as an identification
procedure to tell us what
the off-target sites were,
and then used our Ramp-seq
multiplex [? ample ?]
[? common ?] GS procedure
to give much better our
inaccurate quantitation
of the level of editing at
all those different sites.
And that's something
we're in the process
of doing with this mutant.
But based on the
cell-free data and based
on the Guide-seq procedure,
you can expect those mutants
to be still highly specific.
And I think those comparisons
to the high HiFi Cas9
will be coming in
the next few months.
And it's something we'll
try to get on the website.
So back to multiplex
editing, is there
any reason why Cas9
and Cas12a should not
be combined for multiplex
editing in the same time?
I think it's just something that
we've never really looked at.
I know that Editas Medicine had
shown that you can multiplex
Cas9 and Cpf1 together.
And in some cases they showed
reduced translocations.
But a lot of that stuff we
simply haven't looked at yet.
And it's a really
interesting question
and something we should take
a peek at in the future.
Great.
So we talked a bit
about modifications
to the crRNAs for Cas12a.
I have a few
questions about that.
One, is there an XT version for
the crRNAs that is available?
And then the second was--
I think you kind of
addressed this-- but do
you see any
improvements in cutting
with changes in the
guide RNA structure
or design for the Cas12a?
Yeah.
So much like with
the Cas9 guides,
we had embarked on a very large
chemical modification testing
program for the Cas12a guides.
And we did try two
primal methyl bases.
We tried phosphorothioate
bonds, walking completely
through the constant
and spacer sequence.
And through a lot of work
and testing, what we'd found
is that we couldn't find
anything superior to simple end
blocks on those RNAs.
And that comprises what we
ultimately have on our website
is the Alt-R Cas12a crRNAs.
But, again, I would
reach out and say
we don't test these things in
every possible application.
So if somebody has something
interesting that they think
may be more stable in
certain applications,
we'd love to hear about it.
And feel free to reach
out CRISPR@idtdna.com
Thanks, Chris.
So we have a couple questions
about our HDR Enhancer,
specifically if it works in
other species such as zebrafish
or mouse.
If you're using mouse embryos,
for example, were the embryos
cultured post-injection?
Do you have any comments
on that Bernice?
So, we here don't do
any mouse embryo work.
We have tested the HDR Enhancer
in a few mouse cell lines
and found it to be effective
in those mouse tissue culture
cell lines.
Was the first part of the
question, did I answer her?
Yeah, just if we've
tested in other organisms,
zebrafish or mouse,
and then how you
might go about using it for
mouse embryo engineering.
Sure.
Our general recommendation
is to test a couple of doses
that you think are safe.
We have a range of
doses that we use,
which is maybe around 10
micromolar to 30 micromolar
if you are worried
about toxicity
from these small molecules.
So, we generally
recommend that you
test a few concentrations
that may work for you
in your specific cell type.
Great, thank you.
So I have a few questions here
about guide design for Cas12a.
For example, if you're
expressing this in a vector,
what sequence do you
add to the target?
And then also, can we
recommend any tools
for designing guide
RNAs, for example,
predicting on-target
cleavage and identifying
off-target effects in
particular for uncommon genomes
where you can input a
custom genome, does anyone
have any comments on how to
design the crRNA or Cas12a?
That's a really great question.
Unfortunately right
now, we do not
have a Cas12a guide RNA design
tool available on our website,
nor do I have a good
recommendation for one to use.
But it is a very active
area of research for IDT.
And it's something
that we do hope
to have available in the future.
But what I would
say about this is
one of the benefits
of using the Cas12a
Ultra mutant is I think
we made it as reliable
or more so the Cas9.
So the vast majority of
guides that you select to use
are going to work.
And I think one thing
you can do to maximize
your chance of finding a guide
that gives editing rates that
are desirable to you would be to
consider choosing ones that are
not quadruple t because there
are still some of those sites
that even the Ultra
mutant does struggle with.
So unfortunately
for the time being,
the best recommendation I have
is to use the Ultra mutant in
avoid quadruple tPAMs.
Thanks, Chris.
This is kind of a
technical question,
but where can I find
the conditions of use?
Is this for research purposes
only, patent information,
et cetera?
Can you comment on that
or point this scientist
in the right direction?
Yeah.
So in terms of
conditions for use,
I assume that means just
scientific information
on how to use it.
That should be available on
the user guide on the website.
This particular set of
mutations has been submitted
for patent protection.
So that at some
point in the future
should be a published
patent application.
Otherwise for more specific
questions along those lines,
I would reach out to
CRISPR@idtdna.com,
and we can address that
more specifically offline.
Great.
And I think we have time
for one final question.
Let's just go with this one.
So when would you
want to use Cas12a,
and when would you want to use
Cas9 for an HDR experiment,
for example?
For an HDR experiment,
I think that depends
on the organism
you're working in,
what PAMs are available to you.
Since the improvements that
were made with Cas12a Ultra
have brought up the
on-target editing
to arrange that's
as good as Cas9,
I feel like for
HDR purposes, they
can be used
interchangeably, whatever
works best for your organism.
If you're working in
an AT-rich genome,
you might look at Cas12a.
If you have only an NGG
PAM available to you,
then you could use Cas9.
Yeah, I think sometimes you
just don't get a choice.
If you're trying to correct
a snip that you don't have
Cas9 sites around for,
Cas12a would be the enzyme
to use and vise versa.
And I think previously that
was a losing proposition
because it was a
bit of a crap shoot
as to whether you'd
find a Cas12a guide that
would work very well.
And now that I think
as Bernice said it,
that we've made this
enzyme as reliable
as Cas9, if both
sites are available,
you can choose at will.
But you're not limited by which
enzyme works and which doesn't.
And I think in the
past, Cas12a has maybe
been presented as only
an alternative when
you couldn't use Cas9.
And we just don't think
of it that way anymore.
We find that you can use either
for what suits your needs.
All right, that's all the
time we have for questions.
I want to thank all of you for
attending today's presentation.
I would also like to
thank Chris and Bernice
for the presentation
as well as Molly
for conducting the question
and answer session.
This is one of a
series of webinars
we will be presenting on
CRISPR as well as other topics.
We will email you about
these future webinars
as they are scheduled.
Also as a reminder, a recording
of this webinar will be posted
shortly on our website and
at www.youtube.com/idtdnabio.
There you will find several
other educational webinars
on such topics as
next-generation sequencing,
genotyping, qPCR, and
general molecular biology.
Thank you again for
attending, and we wish you
success in your research.
