Hello and welcome to this long lecture on
genome editing. This lecture is mostly
intended for university students
researchers clinicians and and the media
who are quite new to genome editing and
I want a good background in it. I cover
things like zinc finger nucleases and
TALENs
all the way through to CRISPR and the
latest CRISPR methods like prime editing.
So it's quite a long lecture. If you look
in the description, you'll see timestamps
for each section - which you can skip to
or play again. Also I've broken this
lecture down into smaller pieces if
you're just interested in a specific
thing like CRISPR or prime editing or
something like that. On the slides you'll
see numbers in circles in the bottom
right hand corner. Those are for the
references that I've used to create this
lecture so some of the graphics that you
see come from those papers and reviews.  I'm mostly giving you that list of
references as the best starting
point that I can see for you to continue
your own reading. In the
description you'll see all of those
references with links to those papers
and journal sites. Be aware that some of
those will not be open access, so you'll
need your institutional licenses to see
those articles unless you want to pay. I hope you enjoy it and let's get
started!
In this lecture I'm going to
cover the following sections. Each of
these sections is timestamped in the
description so you can jump straight to
those sections or you can go back and
watch a particular section again later
on if you want to just revise what
you've heard alternately to these
sections will get broken up into very
short videos later
if you've just got an interest in one
particular area so before I get into
details on the different kinds of
nucleases in the introduction i'll just
revise hopefully what you already know
about double strand DNA break repair and
we'll look at the kinds of genome
editing outcomes that you could achieve
using sequence specific nucleases then
I'll move on to describing zinc finger
nucleases and the pros and cons of those
and then I'll move on to talents and
then we'll move on to CRISPR and before
we get into how CRISPR is used for
genome editing we'll describe the
natural adaptive immune systems that
CRISPR belongs to because their debt
explains some of the issues of CRISPR
specificity then we'll get into what
CRISPR cash 9 is and how it's used and
then the longest section will be spent
on CRISPR specificity and the the
advanced methods that are out there to
overcome the limitations of CRISPR and
in the last section we'll talk about
precision editing where you want to make
very specific edits and you need a
absolute control over what you're doing
and there's two areas of that homology
directed repair and prime editing
genome editing describes the ability to
modify the DNA genomes of living
organisms in controllable ways so that's
modifying in living cells in living
species rather than taking a piece of
DNA out modifying it with some enzymes
growing it up in bacteria then put it
back again
rats limb modifying in living cells so
in the case of humans in human cells
genome editing is critical to the study
of genes and gene regulate regulatory
element functions so rather than to
understand the gene you take it out of a
cell and you study it somewhere else or
you're still studying a gene regulatory
element like a promoter or an enhancer
and you put that into some kind of
reporter assay with genome editing you
can study a gene or a gene regulatory
element in its natural context it's
chromosomal context so you can delete it
mutate it exchange it modify it however
you want to in its natural context this
is very powerful genome editing also
paves the way for the creation of models
for human disease so you can take the
specific mutations that occur in the
human population and put them into cell
lines or into animal models to try and
mimic the disease and study very
specific molecular aspects of that
disease and gain some insight maybe
develop some drugs and test them out
genome editing also obviously allows for
the advance correction of genetic
diseases so these are advanced gene
therapies so rather than when you've got
a patient who's got a defective gene
rather than overlaying that with a
normal gene and saw those cells in that
patient expressed both the normal and
the mutant gene you can now actually try
and correct the mutant gene and get rid
of the problems of that mutant gene
genome editing also allows for very
powerful improvements in the
biotechnology and food production
sectors so you could modify microbes or
plants to make better products for you
or be resistant to diseases or easier to
handle whatever it is that's limiting in
those industries and genome editing
editing can now be applied
should it be safe to do so prior to
genome editing
Amitha called gene targeting was used to
modify genomes this is what describes
the genetic replacement based on
cellular homologous recombination
machinery so this is obviously used in
yeast an awful lot it can be used in
mammalian and Brianna Steph cells and in
b-cells but in general it's very
inefficient in most cell types of
interest and for all those applications
list listed above gene targeting just is
just not useful in most situations so
genome editing involves a variety of
powerful synthetic biology strategies
that have been developed to create
sequence specific DNA endonucleases so
these are enzymes that cut a specific
DNA sequence of your choosing and you
you place those under nucleuses to
create the cut exactly where you want to
so if you're trying to knock out a gene
it's going to cut at your specific gene
this then stimulates cellular DNA repair
and DNA repair in fixing that break will
either make mistakes or you can persuade
it and trick it into making specific
changes so the combination of your
knowledge of DNA repair and your ability
to create these enzymes that will cut
where you want them to
you can then modify a target genetic
locus precisely and quickly this is a
very very quick method so there are
in three generations of nucleosis that
have been created for the using gene of
for their use in genome editing the
first version involves ink finger
nucleases so I shall take you through
those then a better approach came along
which are the tale nucleus or talons but
in this most modern ear in the last few
years
CRISPR really has become the easy
easiest and most applicable method out
there so I'll take you through each of
these three things it's important to
consider zinc finger nucleases and tell
nucleuses and understand them because
those are the ones that have made it
through two clinical trials and a lot of
biotech applications so it's important
to know what they actually are on what
their value is so looking at double
strand break repair pathways and so I'm
mostly going to focus the talk on
mammalian systems but a lot of this is
true across eukaryotes the most dominant
pathway of DNA break repair is called
non-homologous end joining it's a very
fast set of enzymatic processes which
simply join breaks back together again
and they ignore the sequence at those
breaks so if you've got multiple breaks
present potentially different ends will
get joined to each other but also when
joining the correct ends together it can
do so in an error-prone way so on the
left is a diagram here of the typical
kinds of factors that recognize the
breaks signal the breaks and then start
to fill in any ends which aren't blunt
and then ligate them together this kind
of repair can be completed in minutes
it's extremely IRA prone because no
template copy is used
and canonical non-homologous end joining
can be accurate if the ends are our
blunt and phosphorylated and clean it
can join them back together the way they
came apart and it's fine but if you're
using one of these nucleases and genome
editing it's gonna cut that site again
because you've reinstated the target
sequence and what will happen eventually
is that non-homologous end joining will
insert an extra base to facilitate the
ligation of the ends and in which case
you've got one base insertion you've
changed the sequence that might
inactivated gene alternative non
homology an alternative enjoining
pathways typically reset the five prime
ends at the brakes so the two just that
one strand back so five prime to three
prime reception on what they're doing is
exposing a short sequence a single
standard sequence to look for local
micro homology so it might just be a
couple of a's or a couple of CS and it
will join those two a couple of T's or a
couple of g's on the on the other end
and it will ligate through that so those
will pair up and there may be bases that
will need to be removed and so often you
get these local deletions so
collectively you get insertions or
deletions from non-homologous end
joining we call them in Del's for short
and so the outcome of this kind of
repair pathway of seemingly random so if
you're trying to create a knock out of
your favorite gene by targeting a
nucleus at the beginning of the gene
then this dominant non-homologous end
joining pathways is excellent for
creating all these seemingly random
mutations and you'll get frameshift
mutations in there and you'll get a loss
of gene expression very very easily but
actually when people look at lots and
lots of targets and repeat genome
editing in lots of cells they find that
actually the kind of edits that occur
from non-homologous end joining are not
random and and to some degree that
they're based on the local sequence
context so you'll see at some sites you
always get +1 insertions at other sites
you always get deletions of a particular
length because of the local micro
homology a slower but accurate DNA break
repair pathway is called homology
directed repair and this is an enzymatic
process that uses a in normal biology
the sister chromatids template for
accurate repair so officer this is
normally associated with DNA replication
so if you've got a stalled replication
fork sometimes the system will will
actually cut that fork create a DNA
break and because you've got a local
template available you can then copy
that so a set of proteins that recognize
those breaks reset so a key part of
homology directed repair is a very
extensive 5 prime to 3 prime reception
at the break which exposes a long three
prime overhang this is typically many
kilobases in length this three prime
break is then protected with protease
like a replication protein a so they
protect their single-stranded DNA
binding proteins they protect these
three prime ends they then get exchanged
with route 51 this fantastic filament
protein and in combination with the
braket proteins these filaments are able
to then invade and search a sister copy
for a matching sequence and when it does
so it base pairs up and a strand
exchange occurs and so you get these
Holliday junctions where you've got a
crossover between one chromatid at
another and then these are resolved and
the gaps are filled in and so what you
end up with is a perfect copy from one
sister chromatid to another
so this process overall can take over an
hour depending on the sequence so it's
quite slow it's limited to a very
specific part of the cell cycle but
potentially we could use this and genome
editing to persuade the cell to make a
specific change for us but instead of
having a sister chromatid as the copy we
would deliver a donor DNA and that donor
could be a plasmid or a viral genome
that we've introduced into cells and we
do so in excess if we can and so the
exchange of the sequence were interested
in becomes the dominant event so a key
thing about considering what's going to
happen after you create a DNA break is
that you've got these error-prone
pathway and then you've got this
accurate pathway and they're active at
different times in the cell cycle so
non-homologous end joining is is
functional throughout the cell cycle
certainly it's very dominant in g1 and s
phase you know which is where most of
your genome editing is going to be
occurring HDR on the other hand is
linked to replication so it's only
active when DNA replication is occurring
and throughout S phase but accumulates
late in S phase and then in g2
particularly g2 is where a lot of these
stalled fox are getting resolved so if
your goal is to undertake destructive
gene editing like creating a gene
knockout then it's very easy because
non-homologous end joining is dominant
and you're going to get a lot of cells
were which are going to have mutations
which are useful to you in that the
create frame shifts are now inactivate
your gene if your goal is to undertake
an accurate DNA edit so you doing gene
correction or a specific mutation in a
gene regulatory element you creating a
specific model then somehow you need to
deal with all the unwanted
non-homologous end joining mutations and
you've got a sift
through all of this junk to find the
cells which have the specific edits that
you're interested in and this is a
hidden problem of genome editing that a
lot of people perhaps don't consider
when they first get started so in this
slide here I'm going to show you in very
simplified terms the possible genome
editing outcomes you can get just from
creating DNA breaks using sequence
specific nucleases so in these cartoons
we've got two stands representing DNA
and and then this this cut through here
just shows a DNA break that we have
introduced with our nucleus so if we
just cut in a place of interest so
that's the promoter early exons of a
gene then we get insertions and
deletions occur because of
non-homologous end joining and so we can
disrupt that genes function or that
regulatory elements function if we
provide a donor DNA at the same time so
that could be a linear piece of DNA or
it could be a plasmid which we are going
to cut at either end with our nucleases
at the same time as cutting Guard
genomic target then without any special
homology arms or special processes so
this is just using non-homologous end
joining you will get insertion of your
sequence of interest it's nearly always
a trans gene so I've colored it green
here in cases it could be a GFP trans
gene say and you will get insertion at
that target site but the key thing is
that you've got no control over the
orientation of that insertion it can go
in either way and also the ends either
side of the insertion will be will have
these in Dells on them so so you may
lose all gained sequences so if it's
important for you to for example in the
case of perhaps you wanted to
Tagg the c-terminus over of a gene and
created gfp fusion or a fusion of
another long tag in that case your
insertion needs to be the correct
orientation and it needs to be in the
correct open reading frame as your gene
and with this kind of approach very few
of your cells are going to achieve that
but potentially for example in the case
of a GFP fusion you can you can easily
select for those cells so if it's very
inefficient knocking and very
inefficient at creating the correct
knocking those cells are still bright
green so you'll be able to sort them out
through flow cytometry for example so it
depends on your application sometimes
you can just do this very simple quick
way alternatively you could make quite
large genomic rearrangements just by
cutting twice so let's say you've got a
gene locus which you want to invert for
some reason you can just cut either side
of it and the cell when repairing those
breaks may put that insert back in to
its normal location but do so the wrong
way around
and obviously alternatively probably
more dominantly
you will lose that fragment and you'll
end up with a deletion so that's just
simply cutting with two nucleuses at the
same time no other trips so small
percentage of cells will make these big
edits for you and we have done this
we've we've deleted out several genes
from a gene cluster and just by cutting
twice and not doing anything else and
you get a decent number of cells that
have made that edit for you and you can
do some experiments on them
alternatively you may want to persuade
the cell to use homology directed repair
for you so for example let's say we want
to knock in a gene at a specific
location so in this case we're going to
cut the target locus with our nucleus so
that's the target locus here and then
you'll have a donor fragment which
contains the sequence you want to in
but importantly that fragment has to be
flanked by homology arms which are the
same sequences as what's found at that
either side of the break at the genomic
target so this sequence here would be
placed here this sequence here would be
placed there and Stratton's strand
exchange would be used would occur using
these homology arms and in the case of
sort of traditional HDR where you're
knocking in large gene fragments you
need at least 750 bases in lens for that
homology arm and the longer the better
alternatively you may just want to make
precise edits so let's say you've got a
defective gene and you're wanting to
correct it with the real gene you would
cut as close as possible to the way you
want the change to occur and then you'd
have a donor fragment again like you've
seen on the left here with these
homology arms so if you wanted to
exchange a large gene sequence then you
do need these long homology arms but
let's say you're just making an edit of
1 2 or 3 bases as is typical for most
human genetic diseases then it's been
shown that your donor fragment can
actually be very short indeed so
actually no people use oligonucleotides
so very short single-stranded DNA
fragments you only need one strand which
will have the edit which may be 1 2 or 3
bases and then that can just be flanked
by 40 bases for zero basis of homology
arm and that's sufficient to get an
exchange to occur and so this has now
become was until very recently the main
way that you would make precise edits of
a gene
so in understanding zinc finger
nucleases were perhaps need to look
further back to what other enzymes cut
DNA sequences specific DNA sequences and
whether we could modify them or not
so obviously restriction enzymes exist
in lots of prokaryotes and we have a
large catalogue of these enzymes which
will cut a specific sequence and the
idea was to try and modify those enzymes
so by understanding them created crystal
structures of them doing lots of
mutagenesis on them potentially we can
take an enzyme that cuts sequence X and
persuade it to cut sequence Y
specifically well actually most
restriction enzymes cannot be readily
adapted to cleave new sequences in
general have got very short recognition
sequences which is not helpful that
they'll cut many many many times in the
genome and not cut to unique sequences
and when you persuade them to cut
another sequence they're cut cut them
very very weakly so really that wasn't a
route to develop a genome editing
however there's a class of enzymes the
type 2's restriction enzymes which are
quite interesting an example is the
enzyme Fock one fok one and these type
2's restriction enzymes have separate
cleavage and DNA recognition domains and
importantly the cleavage domain has no
specificity and can work on its own it
just needs to be recruited to the DNA so
for example here in the case of huaquan
which functions as a dimer that's an
important feature of Fock one is a
dimeric protein so two molecules of the
same protein in green and in blue here
and this domain is the DNA recognition
domain for combines this particular
sequence GG 80 g and then the cleavage
domain is here and a key feature would
tattooist restriction enzymes they tend
to bind in one place and cut a specific
distance away and the sequence that they
cut it doesn't matter it can cut up all
the sequences here so type 2s reduce
restriction enzymes are really useful in
synthetic biology for lots of different
techniques but in the case of genome
editing we can take this cleavage domain
which will only function as a dimer and
add it on to another DNA binding domain
so we've got the cleavage parts we need
the DNA binding part and the concept was
to fuse this catalytic domain of Fock
one onto a different DNA recognition
domain and when you look through the
genomes of of mammals the most common
DNA binding domain out there is the zinc
finger domain so for example there's
nearly 1500 human genes that have got
zinc finger motifs and there's a large
family of genes that contain a certain
kind of zinc finger called the sis to
hiss to offs or C to H to zinc finger
domain and these domains which I'll show
you on the next slide I've got a very
simple Beta Beta Alpha fold and follow a
very common amino acid motif so these
were discovered in 1985 and zinc finger
domains were heavily studied in the
1990s and from this very substantial
amount of work a DNA recognition comeup
code emerged from comparing the the
protein amino acid sequence of zinc
finger domains and the DNA but DNA
sequence that they specifically bound to
so if we look here i'm here's a crystal
structure of zinc finger proteins bound
to DNA and so they bind to normal b DNA
and the zinc finger domains
interdigitate or poke into the major
groove of DNA and all these linked
finger domains look like so you've got
an alpha helix
and to be two sheets here and it's all
held together with a molecule of zinc
hence the name zinc finger and then
there are these specific residues on the
Alpha helix and on this beta tone here
which are the ones which bind to DNA and
depending on on which amino acid in as
you can see in this table is in which
position in a triplet here will
determine which nucleotide is bound and
so you know you could actually look up
through this table and and create on
paper which ideal proteins would bind to
your sequence of interest and indeed
this is possible so you can then create
zinc finger nucleases so zinc finger
nucleases are where you've got an array
of zinc finger proteins usually three or
four each zinc finger binds to three
bases so in the case of four zinc
fingers is by mister twelve bases now as
I said before fuckwad only functions as
a dimer so you need to make two of these
proteins so you've got to assemble two
proteins so R for zinc fingers with a
fuc one cleavage domain at the
c-terminus and Fuquan needs six bases of
space to bind to and cleave so your
target sites are these twelve bases
there's a gap and then these twelve
bases and there are tools to help you
design these so key features of zinc
finger nucleases is that the cleavage
domain has no sequence specificity so
you can cut what whatever you want to
you do need to make two zfn proteins but
this does increase sequence specificity
so you've now got 24 bases of
specificity which is pretty good there
are mutants of the Fock one domain that
would
votes which are obligate heterodimers so
there's a left fuckwad mutant and
there's a right fuck well mutant and
what this means is that if one of these
let's say this left ZF Xenophon protein
bound at an off target site through its
12 bases it wouldn't be able to
homodimer eyes with itself and therefore
cut that off target site so by creating
obligate heterodimers you you're really
restricting the possibility of
Xenophon's cutting off target you do
still get some off target active
activity but it's greatly reduced the
four times in finger domains is about as
good as it gets longer race generally
don't work so you're looking at 24 bases
of specificity at best assembling zinc
finger arrays now is quite
straightforward because there are
wonderful synthetic biology tools out
there to to join fragments of DNA
together quickly in cells the real
problem with zinc finger nucleases is
the vast majority of zinc finger domains
do not function very well when assembled
together in these arrays but for reasons
that are not fully understood so the
best approach is to take a sort of mass
action approach and so the company
sangamore based in california and and
you can buy products license from sigma
they had found zinc finger arrays that
function well and I think these are
pairs of zinc fingers and they've got
pretty much every possible combination
of pairs and they found that these pairs
are happy to be joined to each other so
they can create sets of 4 zinc finger
domains for most sequences very quickly
on a robotic platform and therefore
create functional a defense quite
quickly so a lot of those function well
but for ordinary groups assembling them
themselves and labs it
really was quite a painful process
because the majority of that offends you
made just didn't function anywhere near
as well as this should have done on
paper however if you've got a sink thing
in you clears that binds well and is
specific it's a it's a superb tool for
genome editing there's nothing
fundamentally wrong with zinc finger
nucleases it's just that they're
difficult to make if you don't have one
of these huge assembly platforms
available to you a key advantage of zinc
finger nucleases over all the methods
that follow which I describe in this
lecture is that the fans are quite small
so they're much easier to deliver into
cells and tissues than some of the more
advanced tools that are used later on so
they certainly do have their place in
genome editing although you'll find in
the modern era not many people use them
so cell phones were first successfully
used for genome editing in 2003 they've
been used in many species for a very
wide variety vadik editing applications
and a lot of the genome editing
strategies and approaches that that we
know news today were developed using
zetas fans at first so some of the
foundation papers that are out there all
use their defenses so another reason for
knowing about them there are over 20 zfn
based and genetic therapies that are
going through different stages of
clinical trials so they have their place
but they were slow to make there was a
low chance of them functioning well
particularly when created in in in most
standard research laboratories and a big
breakthrough came in 2009 when the DNA
recognition code of a different protein
domain family was solved and that leads
us on to learning about talyn's in the
next section
transcription activator like effector
proteins or tail proteins I find in
plant pathogenic bacteria of the genus
Xanthomonas and there are class of DNA
binding proteins which have a
predictable DNA specificity and these
different effectively transcription
factor genes that their role in
Xanthomonas is to activate specific host
plant genes to support the virulence of
Xanthomonas and a key reason for these
domains evolving is that they have a
very simple code and they're very easy
to mutate and adapt so if a plant
changes the the sequence of the promoter
of these important host genes to try and
stop Xanthomonas functioning and
infecting them then the Xanthomonas can
change the sequence of their tail
proteins very quickly so what do these
proteins look like well they have these
large repeats in them called repeat
variable domains and they all have the
very same sequence protein sequence with
the exception of these two residues at
position 12 and 13 out of this 34 amino
acid repeat and the structure of the
repeats looks like this which is
helix-turn-helix motif and these two
residues are what specify which
individual base each domain binds to so
the zinc finger domains were good
because one domain just binds to three
bases but the tails are even better
because one domain binds to one base so
you only need a library of four proteins
and then you can assemble them in any
order you want to to bind to any
sequence you want to so all the natural
tail proteins have very long arrays of
these are V DS
and all these are V DS are very happy
very good neighbors with each other so
they're very happy to be assembled and
in to longer race to create proteins
with longer more specific DNA binding
sequences so here's the RVD code so
depending on what these two amino acids
are here in the turn and I will specify
bind to an a base h DC base and so on
and so on we've also got a couple here
which are nonspecific or bind to G or na
so these are incredibly powerful tools
so in the lab you just got fragments of
DNA that encode each one of these four
domains or these other specific domains
here and you just need to assemble them
together into long arrays and then put
them into a longer protein there's very
nice crystal structures of them which
just show how they bind in that they
just again interdigitate into the major
groove of DNA they don't modify the DNA
sequence at all the DNA structure at all
so you've got very nice B DNA very
straight when you look down the end of
it and you just sort of get like this
propeller of Procope of protein and
sticking out from it so very very
elegant protein and you can use tail
proteins much like you do with zinc
fingers and but instead of having is
that a fan now you have something called
a tail nucleus or Talon and again like
before you have left and right proteins
and you create this dimer so fuck one or
will only cleave as a dimer and the
difference in the architecture of
talyn's is that more space is required
between the DNA binding domains for the
fuck want to work so you have a gap of
between 14 and 20 bases unfuck one cuts
in the middle
you've just got to assemble two proteins
just like before but a key feature now
is that you've got much more specificity
in the complex you've got typically
thirty to forty basis of specificity and
the majority of tail nucleus proteins
that you make do function very well I
would say about 75% of the talents that
you make will cleave very well and
they're quite straightforward to
assemble in the northern area lab Anna
and I have done sort of myself the kinds
of breaks it creates both zinc finger
nucleases and talons create breaks with
the four base five prime overhang and
that's just what fuck Wan does it
doesn't create a blunt end it creates a
five-prime overhang and you could use
those overhang sequences if you want to
in some strategies but that does mean
that those breaks are a bit more
mutagenic because non-homologous end
joining in general will get rid of that
five prime overhang and so you'll end up
with deletions
so talons were first used for genome
editing back in 2011
they've become widely used in many
species for a whole variety of editing
applications that can do everything that
seda fans can do it takes one to two
weeks to make them in a standard
molecular biology lab using the tools
that are out there I've seen here half
of talons I've got good cleavage
activity I think it's more than that's
about 75% there's a wide the wild
application of talents has led to many
improvements to reduce off target
activity so there's been new RV d--'s
have been developed which we've got
better based specificity
there's mutations and the fuc one domain
to make it function as an obligate
heterodimer as I've mentioned there are
other mutations that improve the
cleavage activity of Fock one and
there's been lots of changes to the
architecture of talents too to make sure
that it will only cleave when the space
is a certain length which again improves
its specificity because on off target
where two proteins might bind by chance
the the the spacings not the same so
it's a very powerful robust technology
for genome editing again there's really
nothing wrong with talyn's whatsoever
they definitely have their place and
talents are being pursued to final
applications and a lot of different
strategies however like was there
defense you've still got to make these
engineered proteins there's a certain
commitment to to making these engineered
proteins testing them and getting them
working before you do any genome editing
and this is the reason why they're not
used so much these days because there's
an easier approach out there
so in summary I think the way you could
consider zinc finger nucleases and
talons is that they represent versions 1
& 2 of genome editing these technologies
are very well established they're proven
we know what the limits of use are these
enter nucleus is these engineered
nucleases can be very active and very
specific and the quite a small payload
when you're considering how to deliver
them to cells and tissues but there's a
sort of barrier to entry for a lot of
people which is that that initial
protein engineering can be
time-consuming and and expensive you
need to make two proteins per target and
only a fraction of what you make will
will perform well and and perhaps a
another issues that if you're wanting to
create more than one break or edit at
the same time so let's say you're trying
to mutate two genes at once or you
trying to delete a genomic region by
cutting twice in those situations for 2
cups you'll need to make 4 proteins and
deliver four proteins to the settle at
the same time 3 cups you up to 6
proteins it becomes very difficult very
quickly so what if there was an approach
out there where you could avoid having
to make engineered proteins and that's
what we'll find out about when we look
at CRISPR
clustered regularly interspaced short
palindromic repeats is a very long
acronym that is abbreviated to CRISPR
which is certainly a CRISPR way of
talking about the adaptive immune system
found in about 40% of all bacteria and
90% of archaea bacteria and it was
discovered from the sequencing of lots
of prokaryotic genomes and we you kept
seeing these repetitive regions so in
black here are all of these different
repeats and in front of the repeats were
all these open reading frames and so
groups I'm guarding to studying what
these CRISPR loss I were what were these
clustered repeats all about and what it
is is a system with which to detect
bacterial bacteria phage genomes detect
and cut them and also save fragments of
those phage genomes into these
repetitive arrays as a record of
previous infection and so you can target
them again so there's a bit of a chicken
and egg situation goes on here but if
the CRISPR casts system has identified a
viral genome and cleaved it and
therefore inactivated that phage
infection and cut it up into small
pieces some of those small pieces which
are called spacers gets inserted into
this CRISPR array so all these different
colors here are fragments of phage
genomes that this particular bacterium
as-as-as cleaved before in its evolution
this repeat array this CRISPR array is
then transcribed as one long pre CRISPR
transcript the polymerase 2 transcript
which has got all these repeats in it
those repeats well each one of those
repeats binds this other tracer RNA to
form a double-stranded DNA sequence here
and then RNA s3 then binds to this
duplex of RNAs and Cleaves them so each
one of these spacer and repeat regions
called a crisper RNA gets separated from
this pre crispr RNA transcript so you
end up with lots and lots of crispr RNA
tracer RNA fragments these are then
bound by caste 9 and caste 9 is an RNA
guided DNA and a nuclease so these
double stranded RNAs bind to caste 9 and
program it they activate it for search
in the genome cast 9 searches the genome
for short DNA sequences called pro to
prote of space at adjacent motifs or Pam
sequences so each one of these spaces
that we talked about before these phage
sequences these spaces when present in
the crisper RNA is now called a proto
spacer and cast line needs to find a
sequence in the genome called the pam
which centers it it allows it to sit on
to DNA cast and then unwinds the strands
and if there's a match between the proto
space so it has in its loaded crispr RNA
if that matches the genomic target then
the unwinding continues on and cast iron
will then cleave that target
so we can look at this in more detail
again where we're looking at the the
natural CRISPR system and this is a
class two CRISPR system so we have cast
nine and it loads up this duplex of RNAs
of the crispr RNA which contains the
proto spacer which is twenty bases which
what it is supposed to match a phage
genome and then that's paired up with
the tracer RNA calf nine then searches
the genome for short pam sequences these
are often just three years old bases
long and it then unwinds the strands at
these pam sequences and starts to see
whether there's a match between the
crispr RNA and the genomic target if
there is this unwinding continues this
bubble gets bigger and when it reaches a
full length and you've got this long our
loop forms or an hour loop describes
this RNA DNA duplex here and cast lands
holding the the other DNA stand
separately when it's got far enough then
the cleavage domains of cast nine are
activated and the cast iron systems used
in most laboratories create a double
strand break which is blunt ended and it
cuts three bases away from that pam
sequence so very predictable cleavage
so in 2012 Jennifer dude in this lab and
in one world shop NCA showed that cast
nine from the bacterium streptococcus
pyogenes is a programmable RNA guided
DNA and a nucleus very quickly the big
synthetic biology labs out there jumped
on this and modified it in quite a
simple but powerful way to turn it into
a tool to allow anybody out there to use
short guide RNAs to edit their genome of
interest so in the tool that we used in
labs the CRISPR class 9 system now only
has one RNA that's called a single guide
RNA and it combines one end of a crispr
RNA with the other end of a tracer RNA
so these RNAs are typically 98 to 100
bases long at the five prime end the 20
bases are the pro 2 spacer that Maps
your genomic target of interest and then
you have the structural RNA that binds
to and programs cast 9 and that's all
you need you need cast iron protein and
this short RNA and so you can very
quickly cut your target of interest
there is nothing more involved it's easy
to multiplex Plex this so you could cut
to many targets all at once
or you could create a large library of
guide RNAs so the only difficulty is
expressing and delivering those guide
RNAs into cells at the same time as cast
9 just to home in on this a little bit
more there are two nucleus domains in
cast 9 once called the H&H domain which
cuts the target strand or the non pam
strand and then there's a rough sea
domain that cuts the pam strand or you
can call it the
non-target stand because it's that's not
the one bound by the guy Darren E and
they cut pretty much the same time the
rough seiderman does indeed cut first
but what you end up with is a blunt
double strand break
that's three bases away from your pan so
to perform a crisper experiments very
straightforward and you've got lots of
options about how you would deliver
crisper into cells and the choice of
these is very much based on how easy it
is to get started the amount of money it
might cost you to get going but then
also what are the limitations of the
cell type use that you are using or what
specific experiment are you trying to
complete so the most simple way is to
use plasmids that Express cast 9 and the
guide RNA and you just deliver 2
plasmids into your cells of interest
sometimes you can put the guide RNA
expression on the same plasma this cast
9 and you just delivering one plasmid
that's the simplest way to get going but
you need a way of delivering plasmids
into cells efficiently and for a lot of
primary cells that is not so
straightforward another approach is to
make cast 9 messenger RNA in cell in the
lab in the tube and make the guide RNA
in the lab as well so through in vitro
transcription and then you deliver those
to RNAs into cells and that's got the
advantage of these RNAs getting to work
very quickly and in some situations
they're easier to deliver than plasmids
another approach is to use recombinant
cast 9 protein that's been made in a
laboratory and you've guided RNA that's
been synthesized or transcribed in the
lab you mix the RNA in the protein
together and
tube and then you deliver that Rabb or
nuclear protein into cells and this is
probably the most potent way of doing
CRISPR it's the most active way and
finally you could also use lentiviruses
to deliver a cast nine expression vector
and RNA expression vectors and the
reason for using lentivirus is that you
could have a library of different lenta
viral genomes that encode lots and lots
of different guide RNAs so you could
have a whole genome a whole gene library
or you could have a library of guides
against every kinase that sit there in
the genome and then you could do a mass
CRISPR experiment and select for cells
with specific properties of interest and
then those cells that you selected you
then sequence the Lenti viral genome to
see which guide RNAs were present after
your experiment and so now you know
which genes might be important in your
in your particular process that you're
interested in so very powerful tool for
for performing screens and an important
thing to consider here is the level of
castellón expression and how long it
takes to become expressed and how long
it lasts is very different depending on
the different ways that you deliver your
CRISPR into cells so if you use the sort
of standard approach of using plasmids
or let's say a viral vector to deliver a
CRISPR expression vector and a guide
expression vector into cells then it
takes very many hours before that DNA is
made its way into the nucleus it's been
transcribed the messenger RNA forecast
line has been exported to endoplasmic
reticulum translated cast line is folded
imported into nucleus and then
accumulates that takes many hours so
it's often 16 to 24 hours before you've
got maximal
last nine levels and then that plasmid
does often go into cells and quite high
copy number and sticks around for quite
a while even though the cells are
dividing they're still enough plasmid
air to to make casts nine and so the
CRISPR may go on for several days if you
use messenger RNA it gets to work quite
quickly so it can get translated and
cassiejenkins can get to work in just a
few hours another advantage of messenger
RNA is you can really sort of titrate
how much of it is you you're using in
cells so you can prevent over expression
of casts 9 which is a problem for off
target activity of cast 9 and its
expression at last for a shorter period
than than what you will get with past
mood DNA and then if you deliver protein
into cells and obviously it gets to work
straight away and it's the
shortest-lived method of cast 9 delivery
so that cast 9 protein cannot be
expressed again because there's no
messenger RNA and that protein will just
degrade and dilute as the cells divide
so depending on these sorts of four main
approaches of delivering CRISPR into
cells you've got sort of different pros
and cons which you can sort of read
through later but in principle if you
deliver plasmids into cells this is also
the cheapest and simplest ways of doing
CRISPR and it's quite straightforward to
express and deliver for guide RNAs into
cells at the same time but depending on
the cell you're using the transaction
might be quite difficult in all these
situation plasmid situations the cast 9
expression will persist for quite a long
period of time and off targets
mutagenesis by cast 9 will be at its
highest with this kind of approach if I
just jump down to the protein approach
so using Cassadine protein which and
typically you would be purchasing from a
company so that will cost you money so
you've got to buy that Cassadine protein
and also you've got to prepare your
guide RNAs through transcription in the
lab or you're going to spend money
buying those but the advantages of the
RNP approach is that the transactions
very efficient it's suitable for doing
small screens and you've got the lowest
level of off-target activity because it
cost - is short-lived
and you can titrate in exactly how much
RMP you need just to get your target to
be cleaved and don't use anymore cast 9
than that
and then the lentiviral approach
obviously is suitable for these large
screens and and it can be lentiviruses
can be a better way of delivering their
CRISPR system into certain kinds of
cells particularly non-dividing cells so
just to point out that when you want to
express guide RNAs and cells from a DNA
template the way to do this is to use
RNA polymerase 3 promoters and these
promoters are very useful because
they're quite short they've got very
high activity and importantly they start
at a defined base so for example the u6
and 7sk promoters started to G base the
h1 promoter starts at an a base this
base here will be the first base of your
pro 2 spacer so you can only express
guide RNAs that begin with a G or an A
and then another key feature of
polymerase 3 promoters is that they are
terminated very easily with a row of T's
so five T's or more terminates the
transcription so you can easily get
these short transcripts that start at a
defined base and are a defined length
and you can express them in high levels
so how do we measure mutagenesis after
performing CRISPR how do we know that
the CRISPR has worked so all
applications generally involve
performing a genomic PCR of your target
so you got oligonucleotide primers
either side of your genomic target and
you amplify that from genomic DNA from
cells your parental cells before you
perform CRISPR and then from your
crisper treated cells and then you've
got different ways of seeing whether
mutations have occurred following
Cassadine cleavage and DNA repair so one
way is the surveyor assay and the
surveyor I say involves performing those
genomic PCRs of your parental cells or
wild-type cells and then of your crisper
treated cells and the key trick in this
assay is that these PCR products are
melted so they heat it up and then
cooled down very quickly and so the all
those different DNA strands don't
necessarily pair up with the Strand it
came with the the sister strand they
came from so if you've got a mixture of
wild-type and mutant sequences then
you'll end up with mismatches between
the annealed strands you'll get these
little bubbles in the DNA and then you
can cut those bubbles with an enzyme
like t7 and a nucleus one which cuts
mismatches within a double stranded DNA
sequence so it'll cut any mismatched
fragments into into pieces and then you
can run these digestion products out on
an agarose gel and through looking at
the digestion pattern if you've got
fragments of fallouts then that's
because CRISPR has has occurred cation
is bound cleaved and
every Pro and repair has happened and
the sequence has changed so this is a
very easy universal assay you don't need
to have a specific asset designed for a
specific target but I think one thing
you can often find when performing these
surveyor assays is that you can get
nonspecific cutting of your PCR products
dependent on how clean they are how you
prepare things in the lab and one thing
certainly about the surveyor I say is
that it does underreport mutagenesis
when you compare it to other more
sensitive methods but it is quite a
quick way of screening whether the
CRISPR is working on whether one guide
is better than another for example so
it's a commonly used assay another
approach which is far more sensitive as
the RFLPs a so the same as a but as I've
described before so you're doing a
genomic PCR of your parental cells and
of your CRISPR treated cells but in this
case you're going to digest your PCR
fragments and you're not doing this
melting reeling thing you just taking
the PCR fragments and digesting them
with the restriction enzyme which cuts
your CRISPR target so when you're
designing your guide RNAs you are
looking at what restriction sites
overlap that site that's three bases
away from the Pam for that particular
guide that restriction enzyme will
obviously cut the wild-type PCR fragment
but if CRISPR has occurred and it's
changed that sequence in any way then
that restriction enzyme site will then
be lost and so that fragment won't occur
and so by comparing the digestion
pattern of your parental cells and your
CRISPR treated cells you'll be able to
see whether any restriction sites have
been lost therefore whether Cassadine
has cut and led to mutagenesis
this is a really easy to perform I say
and it's highly sensitive any base
change at all will inactivate a
restriction site and change the digest
and pattern which you can then quantify
so this is the most sensitive assay I
think that's out there in terms of a
sort of a gel based assay however you do
need to design a specific RFLPs a for
each target that you're going to do so
if let's say you wanted to compare the
performance of four different guides
then you will need to test four
different restriction enzyme digestion
patterns so it can get quite complicated
and quite hard work pretty quickly also
the interpretation of these gels can can
be difficult for some it's something
that I found so the easiest assay out
there that everyone's pretty much
everyone's using is called the tied
assay and so again this is just a
genomic PCR of your parental cells and a
genomic PCR of your CRISPR treated cells
and you don't do any nucleus digestion
or any agarose gels you just simply send
them for Sanger sequencing so normally
you should get to put this nice clean
sequence if cells are treated with
CRISPR then obviously at some point
close to your Pam sequence the sequence
is going to fall apart and become
seemingly random and because you've got
all those different insertions and
deletions and you often have to tell
your secrets in company that you're
expecting the sequence to sort of
degrade at a particular position and
that this is an experiment and you want
that sequence because often the sequence
company will think they've done a bad
job and won't send you this chromatogram
and we'll just sequence your your
fragment again and again and again and
again so you need to tell them that
you're expecting it to look bad you then
take the two chromatograms here your
parental and your CRISPR treated one and
you upload them
at the tide website the nki in Amsterdam
so this is from bas van stencils lab
laboratory it's a tool is developed for
everybody and what this tool does is
compare these two Kamata grams and it's
able to be convolute the differences
between them and and call whether
insertions or deletions have occurred so
this is obviously a very easy Universal
assay for standard CRISPR and what I
mean is CRISPR that just uses a single
guide it does underreport mutagenesis
slightly I would say 20 to 30 percent
underreporting when compared with RFLP
essays and it certainly can't really
deal with complex CRISPR approaches
using julna cases and to guides which I
get to talk about later on so in summary
genome editing with CRISPR you could
consider this now the third generation
of genome editing it's incredibly simple
and easy to use because you do not need
to engineer any proteins and test them
out you just need to create these
different RNAs to program the cos 9
cassadines are very efficient nuclease
and it seems to work in all the species
tested so far it's very easy to make
multiple cuts and edits at the same time
just by expressing more than one guide
RNA the problem with CRISPR is that it
comes from an adaptive immune system and
by that the CRISPR must make mistakes so
for example if a bacteriophage is
normally been attacked by CRISPR if it
mutates its genome such that the proto
space that does not match anymore
that back to your phage strain could
then take over and win the battle
against the bacterium that is infecting
so
that bacterium needs to cut that new
phage sequence to some degree so it
needs to have a low degree of error and
when it cuts that phage genome by a
mistake that mismatched genome by
mistake it will then record it and now
that new bacterial strain is then
resistant to that new phage strain so
there's this constant battle between the
phage and the bacterium and both need to
be error-prone to keep duking it out
with each other so that's great for
CRISPR in prokaryotes but that's not
very good for us molecular biologists
where we want caste and specificity to
be perfect and it certainly is not
gypsy studies have found that cast nine
stabili interacts with hundreds to
thousands of Pam sequences across the
genome with any given guide RNA so not
just the arm target but lots and lots of
off targets cast ein tends to not cleave
most of these targets but a small number
of them will be cleaved and we'll end up
getting mutated and sometimes they will
get mutated just as efficiently as the
on target sometimes more these off
target sites typically have one two
three mismatches in the proto spacer
region so there can be tens to hundreds
of off target sites for any given guide
we've only got 20 of bases of
specificity to begin with so the fact
that it accepts it tolerates mismatches
of between one and three bases does mean
that there are lots of potential off
target sites out there large-scale
studies of mutagenesis I found that
while some guides are better than others
there's no really reliable rules for
predicting what will be a more specific
guide than another
unfortunately so there's there's no real
computational way of saying this is
going to be a better guide than another
other than to say one guides got less
potential off targets for then another
what has been found is that the first
three bases of the protis beds proto
spacer surprisingly are dispensable for
on target cleavage so there's only 17
basis of specificity required and the
first three of these can be mismatched
and in general it's been found that
cache line tolerates more mismatches at
the five prime end of the protis base
and these twelve bases here with some
some groups have called the seat region
are more specific and are more important
and certainly are going to be involved
in those early stages of of unwinding
the genomic duplex and informing the
CRISPR Complex
so there are lots of tools out there for
measuring off-target mutagenesis so
groups have done chip seek to see where
cast nine binds in cells they've been
done in vitro site selection experiments
or they prepare genomic DNA and cleave
it with cast iron in vitro to find our
targets and that does indeed stack up
with what actually happens in vivo or
you can do very clever methods which
capture DNA breaks and cells and you can
pair prepare DNA libraries of those
breaks and then do high spirit
sequencing and therefore sequence what's
actually happening in cells and all of
these methods have been really powerful
and helping to describe off target
events in general but i would argue it's
not practical to perform off target
assays like this in most of your
experiments you really need to avoid off
top off target activity in the first
place if you can so there are design
tools out there for the defense talons
and CRISPR web tools out there that like
generate lists of potential off target
sites and so some reach researchers
usually because reviewers of in certain
journals have asked them to do it will
perform target specific assays on a
small number of those off target sites
typically those that reside in protein
coding exons to confirm whether the the
CRISPR mutated cells don't have
mutations at some other off target
alternatively you could do lots of
genomic PCR assays on all this different
oft are
sighs peel them and send them for deep
sequencing and then profile that
mutagenesis on quite a large scale so as
I said I don't think this is practical
for most people I think if you're
developing medical strategies so you do
in some of gene therapy style approach
and you're doing gene correction in in a
patient stem cells then certainly
there's a lot of pressure on you to to
look at off target activity and confirm
that it's not occurred in your
experiments but I think for most people
this is it's too time-consuming so we
need to avoid off target activity as
much as possible
so how can we make CRISPR more specific
there's a lot of different ways so the
first and most obvious one is do not
express caste 9 for too long or for to
higher-level the off target sites
generally are inefficient sites the
caste 9 complex might not be very stable
at these off targets or the cleavage
activity may not be very high at these
targets so obviously the more caste 9
you have and the longer you express it
you increase the chance of cleaving and
mutating these weaker sites so if you
can use kattiline mrna our protein in in
your most important experiments another
quite clever approach is you could use
so in the sort of DNA vector based
approaches you could use an additional
guide RNA in addition to the ones that
you need for your experiment you can use
one against the caste 9 expression
vector itself and so obviously caste 9
will be expressed and will function for
a few hours but when it is functional
that vector itself gets cleaved and may
well get lost but certainly you won't be
making any more caste 9 messenger RNA so
after that initial burst of caste 9
expression and function casting will be
lost
and so what you end up with is quite low
levels of and low persistence of of
CRISPR when you use a self targeting
guide that's quite clever another
approach is to change the guide RNAs so
as I mentioned the first three bases of
the proto spacer are not essential for
on target activity so you can use what's
called a truncated guide RNA where the
guides just have 17 18 or 19 nucleotide
proto spaces the vast majority of these
work at your on target but those off
targets are substantially weakened now
and so you tend to find off target
mutagenesis falls quite considerably at
almost all of your off target sites so
that's quite a clever approach another
approach is to use the dual nikkei
strategy which I'll should go through
and describe or you could use an
enhanced specificity mutant of caste 9
so this is an engineered version of
caste 9 where they've engineered out is
it's loose specificity and again I shall
describe that later on the dual nikkei's
approach for CRISPR takes advantage of
the fact that caste 9 has these two
separate nucleus domains that cleave
either the top or the bottom strand of
your genomic targets and mutation of
either one of these nucleus domains
would lead to caste 9 being in the case
so for example the d-10 a mutant which
mutates the catalytic domain of the
rough sea nucleus domain means that
caste 9 can only Nick the target strand
and a wok won't cut the Pam strand
alternatively the H egg 40 a mutant
takes out the catalytic domain of the
H&H nucleus and so now this enzyme can
only Nick the pam strand so now we have
a Nick Nick's are repaired quite
faithfully in mammalian cells
so we're nick in itself doesn't do very
much so if you Nick your genomic target
or enough target it will get repaired
faithfully you shouldn't get any
mutagenesis however if you use this
strategy where you use to guide RNAs and
I've just called them a and B and they
need to be arranged in this orientation
so the pram sites are facing out and
there are not as it stands these two
guide RNAs will recruit the nikkei's at
the same time hopefully so in this case
we're using the d-10 a mutant and it
will Nick the target stand in each case
so you've got Nick's on opposite strands
if those two casts tiny cases bind a
Nick at the same time then what you end
up with is a double strand break but
instead of it being a blunt break it's
now gots a long overhang between those
two Nick points so in this case we'll
have a long five-prime overhang between
this point here all the way to there and
on the opposite strand to fire prime
overhang here and so if used the d-10 a
mutant you create five prime overhangs
if use the H a forty a mutant you create
three prime overhangs using the same
guide RNAs so you have this flexibility
now so I'm just going to show this again
but in a more realistic scenario where
the strands are unwound so we have
coincident binding of two casts nines
that are by next to each other and they
Nick at the same time because these
strands are already unwound by caste 9
the complex will fall apart creating a
double strand break with these overhangs
so it's been found that the tail to tail
orientation as it were of these
complexes is or Pam out orientation of
to guide RNAs is the most reliable way
of creating double strand breaks using
the dual knickers strategy and this
offsets quite important if the two
guides are too close to each other the
cast nines cannot bind at the same time
because they're getting in each other's
way
if the offsets too large then the chance
of the the two bubbles the the unwound
bases in one come in let's say you're a
complex and you be complex merging into
one large bubble will reduce and so you
again you won't get a double strand
break you'll just get two Nick's so
there's an optimal window in which this
works but there are tools out there that
have allow you to go and design these
guides these dual Nikkei strategies
which are quite straightforward so when
you use the dual Nikkei strategy you're
on target cleavage is often quite
similar to using cast nine wild type
cast name with a single guide it tends
to be far more mutagenic than or
conventional crisper because those
overhangs often get lost so you often
get larger deletions so in the case of
making knockouts it's a little bit
easier with the dual Nikkei strategy and
the key thing is you're off target
activity is near zero when people have
looked because the chance of these two
guides having off targets that are in
this arrangement somewhere else is is
miniscule so here I'm going to show you
a typical approach to making knock out
cell lines using a dual neck ace and
that's this is something that an
undergraduate student did in our
laboratory so it's very easy to do so
firstly design your mutation strategy
then this very much depends on your gene
of interest in this case this was a
transcription factor gene so we targeted
the
first Exxon which had the DNA binding
domain encoded within it so this was
Exxon - if it's an enzyme you might
target the catalytic domain the only
thing to really consider is whether your
your gene of interest has multiple
promoters or multiple splice forms
because if you target the ATG or star
site of one splice form you may not
knock out another spice form which might
take over so you might not get a knock
out that you expected so all you need to
design up initially is where to place
your guide so you have an a nav guide
and you've used a design tool to help
pick those out for you you design you
genomic PCR primers that flank that and
you should always do your genomic PCR
assay first and just make sure that
you're able to amplify that region
cleanly and perform your your assays and
in this case for dual Nikko's we prefer
to use an RFLP ese to look at the
efficiency of mutagenesis you then need
to so this is a plasmid based approach
you then need to clone other girls that
encode the pro 2 spaces for your guide
RNAs and you typically clone them into
two separate expression vectors which
have the polymerase three promoters in
them you can then choose to if you want
to so then lift over these guide RNA
expression units that you've created so
for your a and your B guide and sub
clone them into your cass nine
expression vector to make an all-in-one
vector the reason you would do that is
that the efficiency of transfection
of one plasmid is far higher than to
three or four plasmids so with every
additional plasmid you you add to a
transfection your transaction efficiency
will fall quite considerably so there
are quick cloning tools to clone indoors
protospace the sequences and to move
over polymerase 3 promoter units into
another vector so this takes
a couple of weeks to do all together and
I'm obviously it's only a small portion
of your time in those couple of weeks
you then transfect those into cells
we're using electroporation in this case
and our cast iron expression vector Co
expresses GFP and so we can look for the
fluorescence of GFP and so this tells us
what our transaction efficiency is and
in this case it's about 90% delivery
into the cells so we know that at most
90% of the alleles of our targets could
get mutated we can't get a hundred
percent of most it's 90 percent so if
you're only getting say 60% transfection
you need to then adjust your mutagenesis
that you'll see later on to that sixty
percent number you need to normalize the
transaction efficiency to understand
your CRISPR efficiency and then we do
these RFLP essays and as I warned before
they can be sometimes confusing to
understand but in this particular case
we have this PCR product and we're
cutting it with one of two different
enzymes so individual digests this one
cuts three times some crates for
fragments this one costs once and grace
to fragments and we can see in the case
of here that guide a with wild-type
casts nine works well because we've lost
this restriction site that cut these two
fragments up so these two bands
disappear and they lost here and you
sometimes see the join of of these two
fragments been joined together as in
here but often this is mutated as well
so it will be a smear it won't be a
clean band so the way to quantify this
is to quantify the loss of these bands
here and then alternatively guide B
works because the production of these
two fragments here is reduced and so
we've got lots of uncut product and then
you can compare those digestion patterns
when you're using guides a and B
together with the cast nine nikkei's
and so from all of this you can then
quantify what percentage of mutagenesis
you've got so 90% of the guide a target
site was mutated
when using guide a so in other words
this guide was a hundred percent
efficient guide B we got 75% mutagenesis
with guide B as compared with 90%
transfection so this was a pretty good
guide but not perfect and then when we
used guides a and B together with the
nikkei's we saw 70% mutagenesis of both
alleles so basically the dual nikkei's
is as good as your worst guide and then
to find your knockout cells so if you
can you try and clone your polyfill on a
mix of cells so you do the crisp you do
the CRISPR transfection three days later
you then dilute out the cells to make a
single cell per well or if
colony-forming cells you would then
plate out in a dilute way such that
individual colonies will grow up from
individual cells you then pick those
colonies or lines and then you screen
them and it's easiest to screen them for
what you're interested in first so in
this case we're interested in the
knockout of the protein that were
looking at which was there's F 1 in this
case and we can see that of these 9
lines tested in this first run here 2 of
them have lost their f1 protein
expression and then we've probably got
two that are quite normal and then we've
got another 5 where we seem to have
either heterozygous mutation or sort of
frame or mutations which haven't created
a frameshift knockout but certainly have
really screwed up expression of those f1
so it's best to identify cells which
have lost your expression first and then
you do the genotyping on those so you do
your RF
P and you do your Sanger sequencing on
those so indeed these were knockouts on
both alleles and these these were our
sir also mutants on both alleles just
not knockouts on both it's a knockout
and mutant heterozygous so another
approach and very sensible approach now
how to do in CRISPR is to used an
enhance specificity mutant so as I said
before class 9 has evolved to be
error-prone and that enables adaptive
immunity to new challenges and the way
it does this is that Caston makes many
backbone DNA contacts to stabilize
complexes prior to cleavage so cassadine
uses many positively charged residues
like lysines and arginines
to bind to the negatively charged
phosphate backbone of your genomic DNA
target stand and of the non target
strand to hold them in place and
researchers used the crystal structure
and sort of mass mutagenesis assays to
find out which ones of those contacts
could be mutated so you substitute that
lysine with an alanine for example and
see where the cast iron still functions
or not and then of all those were
mutants which still functionally then
combine the mutations together to see
whether they could get 2 with 3 & 4
residue mutations and still have cast
iron activity and indeed this is what
they have so there are 4 different and
hand specificity mutants of cast iron
out there now that have recreated in
different ways either through sort of
rational design or through a sort of
screening based approach and this slide
just shows two of them so for example
high fidelity casts 9 disrupts 4
contacts between casts 9 and the non
target strand sorry and the target stand
this one here
and enhance specificity cast 9es cast
nine has three mutations that disrupt
contacts with the with the non targets
tandem we have used this one so a key
feature of using these enhance
specificity mutants is now they do not
tolerate any mismatches whatsoever and
they need a full 20 base proto spacer so
all 20 bit basis of that protospace are
key to on target activity so truncated
guides do not work extended guides if
you've made a longer guide for some
reason 21 22 bases they don't work and
mismatch guides obviously don't work so
our target activity is almost completely
reduced there are some off targets in
the sort of worst designed guides out
there which might still have some
activity but in general off target
activity is lost when you use these
mutants so if you can get hold of
expression vectors for these enhance
specificity mutants or use recombinant
proteins with these mutants then
certainly try them there are a small
number of target sites where Catalan
activity will be reduced when using
these enhanced specificity mutants but
by and large this is still a very potent
system so in summary CRISPR has rapidly
overtaken Zetas fans and talent
technologies as new targets can be
programmed without making any new
proteins at all so that that barrier to
entry of doing genome editing is now
gone and really anybody in any field
even people with quite limited molecular
biology expertise can now perform genome
editing CRISPR is becoming widely
adopted in most molecular and cell
biology fields crispers very efficient
but specificity is a major concern
especially if you use wild-type Castner
so high specificity casts 9 meters are
now available and they're also some very
clever strategies to do in crisper as
well which mean that your specificity is
greatly improved it's possible to modify
multiple targets in parallel and of
course you can perform functional
genomic screens so this is an extremely
powerful approach for CRISPR there are a
number of CRISPR applications that
require precise edits so we're not just
talking about knocking out and and
destroying and disrupting according
region of a gene or the important parts
of a gene regulatory element so there
are situations where you want to
integrate trans genes or short sequences
like epitope tags so these are very
specific sequences and you will want to
put them in specific places also if
you're trying to correct human genetic
disorders you wanted to make very
specific single double or triple based
changes and you don't want random
mutagenesis to occur and a key thing to
point out is if you look at the NCBI
clinvar database and download the 75,000
or so human disease variants that are
out there the vast majority of them are
very small in length so a lot of them
are so thirty percent of them are
transition point mutations so single DNA
base changes a transition mutation just
just for reference is where you've
swapped to pyrimidines for a pyrimidines
so for example in a for a tea a tea for
an A or a G for a C or C for a G a
transversion point mutation occurs in
about 20 percent of human genetic
disease
and that involves an exchange of a
pyrimidine for a purine base or vice
versa so for example an a for a T or a G
for an a 20.per 626 percent of human
genetic diseases involve short deletions
and then the reduplication zone
obviously angular larger changes such as
copy number changes and large insertions
and deletions when you look at the
deletions and duplications and
insertions the vast majority of them are
less than 25 bases in length so most
human disease associated mutations are
very short in length and so if you've
got a method to make small precise edits
potentially you could use that to
correct the majority of human genetic
disorders so how do we make precise
edits with CRISPR so the first option
and one that most groups are using to
date is to use homology directed repair
of the cell so this involves creating a
double strand break and obviously you
can do very large insertions when you
use someone's you directed repair but
you can make small edits as well the
second option is to use a new method
called base editing we're using an
enzyme to swap basis over the advantage
of that is that there's no DNA break
created and then the third option is to
use a brand new method which came out at
the end of 2019 so there's only a couple
of months old
it's called prime editing and there
there's no double strand DNA breaks
created and it is unlike all the other
approaches is capable of making all
kinds
vedat edits of less than 80 bases in
length but a very high efficiency and a
very high specificity so I should take
you through each of these three so as I
mentioned in the overview at the
beginning homology directed repair
requires the donor DNA template and so
for the in the case of a large
alteration like inserting a trans gene
that's often a plasmid or a viral vector
with your transgene flanked by quite
long homology arms of 750 bases or more
if you wanted to make point mutations or
very small edits then your HDR
experiment would involve a single
stranded DNA donor template so just an
old agony that I'd were the homology
arms are just 40 bases in length so both
of these systems work and you can get
precise editing of a target the problem
is is that error-prone non-homologous
end joining repair come finds these HDR
strategies because it is the most
dominant repair pathway for the double
strand breaks that you create so the
majority of cells that you that you
create after doing an HDR experiment
will be random mutants and in the
background is a minority of accurate HDR
Corrections and a lot of your correct HD
ourselves will be correct on one allele
and randomly mutated on another so this
can be extremely frustrating and
requires a lot of work to filter through
each of these cell clones that you will
make to find your yourselves with the
edits that you're interested in so it's
important again to look right back at
the beginning where I that
non-homologous end joining occurs
throughout the cell cycle but is also
very high in g1 and s phase whereas HDR
occurs primarily in in g2 so one trick
that can be used with great effect is to
make a fusion of casts 9 with part of a
protein called jamming in jamie-lynn is
approach
that is cell-cycle regulated its
transcription an expression is not cell
cycle regulated so it's expressed
throughout the cell cycle but it is
targeted for degradation by the
proteasome but in g2 jamming in becomes
phosphorylated and this domain that gets
targeted for degradation is now
resistant and so jamming in can then
accumulate and function in g2 and M
phase so this is a post translational
regulation of cell cycle regulation of
Germany so you can take that domain of
jamming in that gets targeted for
degradation but is protected by
phosphorylation and stick that on to
cast nine and this now means that the
cache line jamming in fusion again is
expressed throughout the cell cycle but
gets degraded can't function apart from
in g2 when it becomes phosphorylated and
protected so when you use this you get a
big shift in the bias of hgr experiments
towards HDR events non-homologous end
joining still happens but no perhaps is
a minority event so now maybe two-thirds
HDR on one third non-homologous end
joining so this is a very powerful
modification to HDR strategies so this
is just showing an example from one
recent paper of researchers who are very
good at HDR and these are the kinds of
outcomes that you might get so here that
targeting a variety of genes with single
base changes or yeah I think yeah or
here's a triple base insertion so
different HDR edits and in blue bars you
can see this is a percentage of cells
that have got their HDR edits and then
an in gray are the percentage of last
the percentage of alleles that have got
random insertions and deletions on them
because of non-homologous end joining
so this shows the extent of the problem
so base editing is an approach where you
can make very small changes often just
individual bases using an enzyme that
will exchange that will modify the DNA
base and when it's replicated it would
lead to that that that sequence being
changed so in this case they're using a
fusion of Castine and often leaves and
the nikkei's mutant d-10 a which I'll
describe in a minute as to why they do
that but the they use cast nine fused to
a deaminase and there are different
denominators out there so for example
there's aid and there's Apple Beck and
what these D rnases or ABC 7 and what
these do lasers do is deaminate for
example an adenosine base and convert it
to an inner scene base when in a scene
is replicated it gets replicated as
guanosine base so what you end up with
here then is an A to G change
alternatively you could deaminate a
cytosine base to a D you and that gets
replicated as a T so then you get a c to
t change these the d-10 any case and
they're very clever about the design of
which strand they put their guide RNA on
and this is all to make sure that only
the base edits on the strand that
they're interested in getting cooperated
because NIC repair occurs on the non
edited strand so NIC repair doesn't
create obviously any mutations by itself
but it's been used to bias how the cell
either uses the edited or the or the non
edited strand so the advantage of this
approach is here using an enzyme
so the percentage of base editing is
very high indeed I guess a key
disadvantage is that depending on the
sequence that you're targeting it may
not be possible just to edit one base
only it might so for example we're
making a specific a2g change there may
be other a bases locally that will get
converted to a G as well and so we'll
get all the edits that we did not intend
so so it does very much depend on the
target is that how useful this base
editing is but in some situations it's
very very powerful indeed there's also a
risk that this base editor will modify
bases at off target sites and that those
will be quite hard to to discover so I'm
gonna end up with talking about prime
editing prime editing is an incredibly
powerful approach that uses a modified
CRISPR system but there are no double
strand breaks were created so we're not
reliant on hosts double strand break
repair it uses a cast 9 knee case to
locate this prime editing enzyme to your
target site and to prime the editing the
intended edits is carried on a guide RNA
template and reverse transcriptase is
used to copy that guide RNA template
onto your target and creates this edited
strand and then that edited strand is
then incorporated into your target so I
shall take you through this in detail so
the prime editor again involves a cast 9
a guide RNA and your target as before
but there are three key changes from
conventional CRISPR
so firstly they're using the h 840 a new
case of caste 9 so if this Knicks are
tough targets it will create to Nick but
doors won't lead to mutations and the
Nick at the on target by itself again
won't lead to any change either that in
itself is just an entry point as a
reminder the h 848 nikkei's nicks the
non target strand or the palmar strand
and so it will Nick here at this very
specific location 3 bases away from the
Pam the H 840 a new case is fused to a
mutant version of the FML V reverse
transcriptase domain this reverse
transcriptase is very well used in
molecular biology it's the one used in
reverse transcription reactions when
making C DNA's and molecular biology
labs it's something that's been studied
intensely there are lots of mutations of
it have been made to look at its
performance and so the mutant that
they're using of the reverse
transcriptase here they've selected for
one that functions very well in this
particular architecture here thirdly the
guide RNA which is the same as before so
normal guide RNA except that now it's
much longer so surprise
editing guide RNA or PEG RNA and this 3
prime extension carries the template for
the change that you want to make to your
genomic target so how does this work
so as a reminder you're all crisper
target sites consists of a Pam sequence
in the genomic DNA next to 20 bases that
match the pro 2 spacer in the guide RNA
and cast line cooks three bases away
from the Pam in this case it's just a
nick and the nicks occurring on the Pam
stone and that first base after there we
call +1 and what the prime editor can do
is change any sequence after this point
so all the sequence here including the
Pam is susceptible to the prime editing
so the reaction occurs as follows the
first step is the nicking of or binding
to the genomic target opening up of the
stands and the nicking of the Pam strand
so what you end up with is a 3 prime
flap here of your genomic target and it
is pairs up with your peg RNA so the
fact the very end of the peg RNA that 3
prime end pairs up with your proof to
space of targets and generally this
primer binding site is 12 bases long so
again you know what the sequence is it's
your proto spacer from the cut site on
which those 12 bases are placed at the
end of the Peck irony so all reverse
transcription reactions require a primer
of some kind and in this case this 3
prime overhang here is effectively a
primer for 5 prime to 3 Prime reverse
transcription so now in the next step a
verse transcriptase adds on the edit and
the edit just might be a single base
insertion or deletion doesn't matter
what it is here is this template so in
this case I've shown a 3 base change
in red so the original sequence in blue
and black the three based change in red
so it fills in that three base change
and then a short sequence that matches
the rest of your genomic targets which
you want to be to stay the same so the
the other side of your edit so this is
the nod edited sequence and then a key
thing is the reverse transcriptase
cannot go any further into the peg RNA
because it's bound up by caste 9 so the
reverse transcriptase doesn't copy a
guide RNA sequence into your genomic
target it only copies in what you've
intended in your edited region here so
it fills that in and then that's it
that's all this enzyme does so it binds
opens nix primes and fills in and that's
all it's doing so now we're hoping for
the cell to incorporate this edited
strand over the non edited strand so we
have a 3 prime flap containing the edit
in red and 3 Prime single strand Enzo a
relatively stable in the genome there
are no enzymes that will choose 3 prime
to 5 prime so this free 3 prime flap is
there and the NIC is there now there is
going to be some breathing this Nick
creates instability in this duplex so
these bases will unzip at some to some
level and so there will be some
equilibrium between the 3 prime flap and
a 5 prime flap now this will be the most
dominant form this won't happen very
often but when it does this 5 prime flap
is now available for digestion
there are many enzymes that choose
single standard DNA 5 prime to 3 prime
there are many enzymes that recognize 5
prime flaps and just cut the flap off so
the idea is that host enzymes will
excise this fire prime fly
and your three prime flap your editor
flap will they come on in and hybridize
through the basis that you've kept the
same but now you'll have a mismatch a
mismatch between your edit and the
original non edited sequence so you've
got a 50/50 chance of the cell when
repairing that mismatch of incorporating
your edit now the clever thing in the
prime meditating is that the have used a
second guide RNA but now it's a
conventional guide RNA doesn't have this
three prime extension on so it's the
second guide RNA to target your same
prime editor enzyme to Nick the non
edited stand okay so this guide RNA
hopefully it's going to Nick after the
primary things happened if it nicks
before the primary things happened
nothing really is going to happen then
it gets filled in it's fine so some of
the time this Nick is going to happen
after the prime in it when it does the
the its previously been find that by
adding a Nick into a mismatch sequence
so this was a why it's used in the the
base editing system as I've just
mentioned before is it biases the the
repair of this mismatch to remove this
strand and retain the edited strand so
if you Nick the non edited strand you
will lose the non edited sequence and
retain your edit okay so in the best
form of primary thing or to guide RNAs
one of them is delivering your edit via
reverse transcription and the second
guide RNA is biasing the repair of the
edit towards your edited sequence and so
you don't end up going back to your
original sequence
so this is data from as Prime editing
paper so as I've shown you before these
are the kinds of ratios of specific
edits you get from homology directed
repair using single strand illegals
so quite low levels of precise edits and
high levels of background non-homologous
end joining when used the prime editor
you often get around 50% of yourselves
contain your precise edit and it is
precise
there's no note in Dells associated with
it when the primary that goes in and
there's a very low background of indels
associated with mismatch repair that
that there hasn't been accurate but
occur at your target and depending on
how you design that second guide RNA to
cut your target strand if you can design
that towards your original target then
and you can refer to the paper for this
then this background goes down even
lower so primary things a is a huge
breakthrough in the CRISPR field it's
got broad application because it's more
efficient than conventional CRISPR for
creating precision edits and there's got
a very low level of error as it
currently stands and and obviously it's
people are going to work on this and
probably optimize it even more a key
thing is it can mediate all four
transition point mutations and all eight
transversion point mutations and base
editing the enzymatic base editing I
talked about before cannot do that it
can only do transition mutations it's
been possible so far to insert up to 45
bases using prime editing so adding
epitope
tags onto the ends of genes and also
they've been able to delete up to 80
basis with prime editing and actually I
think they've sort of discovered what
the limit of it is yet so
on paper this can correct around 90% of
human pathogenic genetic variants in
principle so it's very very exciting for
everybody so in summary I've told you
about Zelda fans and talents and showed
you that nucleases can stimulate editing
of genomes and living cells via host a
double strand DNA break repair Zelda
fans and talons are most useful when
small proteins are required
so when delivery to primary tissues is
difficult but for most applications
CRISPR is far easier faster and widely
applicable and conventional CRISPR
remains the choice for gene knockouts
and performing screens but now CRISPR
can be performed with high specificity
and prime editing is probably the best
choice for the precision editing of
genomes well thanks for watching I hope
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