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ELIZABETH NOLAN: So where we left off last time,
we were talking about using antibiotics
as tools to study the ribosome.
And recall that antibiotics have many different structures,
can bind to the ribosome at different places.
And we closed with talking about this antibiotic, puromycin,
that can bind to the A-site and cause chain termination,
and also molecules that are derivatives
of puromycin, such as that more elaborate one with a C75 there.
And so the example of a system where
puromycin has been employed, and this
is just one of many, many examples,
but also gives us a little new information about players
in translation, involves studies of elongation factor peak, so
EFP.
And if you recall, where I closed last time
was with the comment that this EFP over the years
was implicated in a variety of cellular processes.
But its precise function remained unclear.
And so Rodnina and co-workers conducted
a series of experiments to ask, what
is the effect of EFP on peptide bond formation
when different dipeptides are in the P-site?
OK?
And their experiments were motivated by the fact
that there was some preliminary work out there suggesting
that EFP accelerates peptide bond formation,
but really, the details were unclear.
So we're going to look at the experiment,
their initial experiment they did,
which led to some new understanding about how
EFP affects the translation process.
So what is it that they want to do in this experiment
effectively?
Imagine we have our ribosome, and we have our three sites,
OK?
And so what they do in this experiment is
they have a dipeptide loaded in the P-site,
OK, where x is some amino acid.
OK, and then what they want to do
is have puromycin in the A-site and then effectively monitor
for peptide bond formation with or without EFP
added such that the product is effectively
a tripeptide, where we have fMat, the amino acid
and puromycin, OK?
And keep in mind, if this is what's being monitored,
there needs to be a step to hydrolyze
this tripeptide off the tRNA that's in the P-site, OK?
And throughout this work, how they monitored
this is that they have a radio label on the formal methionine.
So you can imagine that you can somehow separate and see
the dipeptide as well as this tripeptide-like molecule
with the puromycin attached.
So how to set up an experiment to test this?
So they do a stop-flow experiment,
so you heard some more about that method in recitation
last week.
And so in thinking about this, we
need to think about what will be mixed.
So what are the components of each syringe?
How will this reaction be quenched?
And so beginning to think about that,
the question is, how do we even get the ribosome
we need to start with in order to see the reaction?
Right?
So imagine that the goal is to have
a pre-translocation ribosome, so effectively that dipeptide
is in the P-site, and the A-site's empty.
And then that assembled post-translocation ribosome
needs to be mixed with puromycin such
that puromycin can enter the A-site and peptide bond
formation can occur.
OK?
So there's quite a bit of work that
needs to happen to even get this experiment set up,
because somehow that post-translocational ribosome
needs to be made.
OK, so if we think about this from the standpoint
of the experiment and using the stop-flow to rapidly mix,
we have syringe 1, and we have syringe 2,
and we have our mixer.
OK, so what are we going to put in syringe 1?
OK, so here, we're going to have the post-translocation
ribosome.
The A-site is empty, and the P-site
holds the dipeptide attached to the tRNA.
And then in syringe 2, we're going to have puromycin here.
OK, so before we get to EFP, I'm thinking
about how we're going to look at that in this reaction
and what it does.
How are we going to get here?
So what needs to be done to get this post-translocational
ribosome?
Is it in the sigma catalog?
Bio rad?
No way!
And even if it were, you would be broke needing
to purchase enough to do this experiment, right?
You talked about needing high concentrations in recitation
last week for these types of experiments.
So where does this come from?
What you need to do before even getting this into your syringe
here, to do a rapid mixing experiment?
AUDIENCE: You have to isolate it from cells?
ELIZABETH NOLAN: OK, so what is the likelihood of isolating--
well, what's it, what do you need to isolate from cells?
AUDIENCE: Well, you're going to need to modify it afterwards
because there'll be all sorts of other things.
ELIZABETH NOLAN: Right, but what's it?
AUDIENCE: The ribosome.
ELIZABETH NOLAN: OK, so we need a ribosome.
Right?
What else do we need?
So we need the ribosome, and we need
to get this into the P-site.
So how are we going to get that dipeptidyl tRNA
into the P-site?
AUDIENCE: You need an mRNA.
ELIZABETH NOLAN: We need an mRNA,
and we're going to design that mRNA based on what amino acids
we're interested in.
So we need to come up with an mRNA.
What else do we need?
So think back to the whole cycle.
AUDIENCE: You need EF-Tu, GTP.
You need everything necessary to form the fMat to x-peptide.
ELIZABETH NOLAN: Yeah.
So what does that mean first?
And when does that bond form?
That's the next thing, right?
So can we deliver this species to the P-site,
based on what we understand about translation
in the past four or five lectures?
No.
Right?
So first, the initiation complex needs
to be prepared in lab, which means
you need initiation factors, a ribosome, mRNA, the initiator
tRNA.
And then that initiation complex needs to be purified,
which is done by a type of sucrose gradient
centrifugation.
OK, and then what?
Once that initiation complex is formed,
there needs to be a round of elongation, where
the ternary complex of EF-Tu.
The amino acid and GTP comes in to deliver that x-tRNA x
to this A-site, and then have peptide bond formation occur.
OK?
And then, we also need the help of EFG
to move that to the P-site, right?
So that whole cycle we've talked about
from a fundamental perspective needs
to be done at the bench in order to get here.
So there's a lot of factors that need
to be purified and obtained, quite
a bit of effort to just even set this experiment up.
OK?
So always think about where these things come from.
So we have this.
We have puromycin, right?
And then we want to look at the effective EFP.
So the idea is, are there differences
in peptide bond formation?
Is it accelerated in the presence of EFP,
as how some of this preliminary data indicated?
And if so, is that for all amino acids?
Or is it specific for certain amino acids, right?
So we need to include EFP.
And in these experiments, it was either omitted or included
in each syringe.
And something just to think about when
thinking about these rapid mixing experiments
is what happens in the mixer, right?
If you're having the same volume, which
is the case coming from syringe 1 and 2,
you're going to have a dilution in here
of all of the components.
Right?
So these are going to be rapidly mixed in the absence
or presence of EFP.
There'll be some time to allow for reaction to occur.
And then, in this case, the reaction
is going to be quenched.
So it's the quench flow-type setup
that came up in the recitation notes from last week.
So in this case, we're going to have a syringe
3 with a quencher.
And in this particular work, they used base,
so sometimes it's acid.
Sometimes it's base.
And this was a solution of KOH.
OK, so then after some time, OK, we
can have the reaction quench.
OK, and then there'll be some sort of workup and product
analysis.
OK?
So in this case, they chose to hydrolyse the peptidyl tRNA's
and look at the peptide fragments.
So you can imagine you need a method that's
going to separate fMet x, whatever amino acid x
is, from that product there.
And then the radio label on the fMat
is used for quantification.
So what happens here?
And I'll just give a summary, and then we'll
look at it in more detail.
So what they did in these experiments--
and recall that JoAnne talked about in recitation
last week, when doing these kinetic experiments,
you have to tweak them quite a bit
to get the exact good conditions to observe
what you want to see.
So imagine that happened.
We have our k observed, and I'm going
to show these on a log scale.
So always keep in mind, paying attention to what type of scale
the axes are in.
And so what we're going to look at
is the k observed for formation of this tripeptide,
depending on amino acid.
And I'm going to generalize a bunch of the data here,
and then we'll look at all the individual cases.
OK?
So here, we have x does not equal proline OK,
and here, not colored in, is no EFP.
And shaded is k observed for the reactions conducted
in the presence of EFP.
OK?
So what was observed in these studies,
looking at having many different amino acids here?
With that, many of these amino acids
showed negligible difference, whether or not
EFP was included in the reaction.
OK?
And we can look at that data in more detail
from the paper on the slide.
What was very striking about these initial experiments
was what happened in this case, when x equals proline here.
So effectively, what they observed in this case
was about 90-fold rate acceleration.
Effectively, if we compare the k observed for peptide bond
formation in the absence of EFP, we
see it's significantly diminished
for proline if EFP isn't there.
And along those lines, it was known
before that proline attached to its tRNA
is a poorly reactive tRNA.
So different aminoacyl tRNA's react differently
in the ribosome.
So there's that layer of complexity
we haven't really talked about in this class yet here.
So if we take a look at all these different examples,
this one is the outlier.
OK?
So what these data indicated is that EFP has some special role
in accelerating peptide bond formation for peptide bonds
that contain a C-terminal proline residue here for that.
And so these experiments were just
a starting point for many additional experiments
that ended up showing EFP is really critical for helping
the ribosome translate sequences that have
consecutive prolines in a row.
So either three prolines or maybe a PPG sequence here.
And in the absence of EFP, what can happen
is that the ribosome stalls.
So these aminoacyl tRNA's are not very reactive,
and the ribosome just gets kind of stuck.
And you can imagine that's not good for the cell.
And then if we bring these observations back around
to some of these early works that were suggesting
EFP has a role in a diversity of different cellular processes,
what might we ask?
We might ask, well, where do the sequences
of multiple prolines come up?
So what types of proteins have three prolines
in a row some place in their sequence?
Or something like PPG.
And so they took a look at that.
And if we think about E. coli, there's
about 4,000 different proteins, and there's
a subset of around 270 that have these types of sequences
in them.
So not hugely common, but they exist.
And so then ask, what do these proteins do?
Right?
Provided a function is known.
And so what we see is within that subset of about
270 proteins, there's examples of proteins
that are involved in regulation, in metabolism, you know,
important cellular processes.
So you can begin to understand why
it might be that this protein got
implicated in all these different types of phenomena,
right?
But in terms of the details, it's
really back here in terms of how this translation factor is
helping the ribosome make a certain subset of peptide bonds
there.
So if you're curious about this, the paper's really wonderful.
There's a number of additional interesting experiments
that are done and additional methods to these kinetics
there.
I'm happy to point you in that direction.
So yes?
AUDIENCE: Does this rate of the reaction affect upon
ribosome folding?
ELIZABETH NOLAN: It could.
I mean, basically, you're talking
about what happens as the polypeptide extrudes
from the ribosome, right?
And if you're stalled and have some piece
of this nascent polypeptide on the outside.
Ribosomes stalling, yeah, what does
that do in terms of how trigger factor, for instance,
interacts.
That's something we'll talk about in the next module,
and we'll be getting there on Wednesday,
I hope, if not Friday.
So with that, we're going to close discussions of module 1
in the ribosome with looking at some biotechnology
and thinking about how we can use
this fundamental understanding of the ribosome
to do some new things.
And so we're going to talk about re-engineering translation
and ways to use this machinery to incorporate
unnatural amino acids.
And so to begin thinking about this,
we can just consider some questions.
And so many of us in this room are chemists or chemistry
majors.
We can think about organic chemistry, so 5.12,
5.13, and all of the different organic transformations
that are presented.
So if we think about all these organic transformations
and how they're available to synthetic chemists,
we see a lot of versatility.
And we can simply ask ourselves, can such versatility
be achieved for protein modification?
What is the toolkit?
How can that toolkit be expanded?
And then thinking about this further,
can we use the translation machinery?
So is it possible to modify the translation machinery
to allow us to make peptides or proteins that
have unnatural amino acids?
So amino acids are moieties that are not the canonical ones.
And can we do this in cells?
Can we do this in a test tube?
And if we can, what does that provide us
with in terms of possibilities?
So the answer is yes, and we're going
to focus on the how and strengths
and limitations in terms of our discussions of this machinery
here.
I also note-- I believe, JoAnne, this will come up.
Will you be talking about this in the nucleotide parts, too?
JOANNE STUBBE: If we get that far.
ELIZABETH NOLAN: If we get that far.
So in addition to here, this may come up again
towards the end of the course, as a tool.
So hopefully we'll get that far, because that's exciting.
So let's think about re-engineering translation.
And we can think about two things.
We can think about the genetic code here,
and we can think about the ribosome.
And so I'll just present you with the questions.
If we consider the genetic code, what
can be done to this genetic code to change
an amino acid in a protein?
And if we think about the ribosome, what
can be done to the ribosome to change
an amino acid in a protein?
And effectively, can we expand the genetic code
to encode something other than what it's supposed to encode?
So can this code allow us to encode an unnatural amino acid?
And from the standpoint of the ribosome,
is it possible to design new ribosomes?
So can we make a new ribosome that
can incorporate unnatural amino acids into proteins?
So these are separate but related,
and we're going to first discuss basically reassigning--
is it possible to reassign a codon?
So why would we want to do this?
And let's think about that for a minute.
And what do I mean by expanding the genetic code?
So if we think about the genetic code,
we all know that it encodes these 20 amino acids building
blocks, there's the start codons and the stop codon.
And effectively, the codons are all used up, right?
There aren't extra codons floating around
that we could poach and assign to something else here.
So can we overcome this?
And why would we want to do that?
Just broadly, if we think about being able to put something
other than a natural amino acid in a protein at a specific
location-- so exactly where we want it--
that opens up many possibilities for experiments.
And we can think about those experiments
both happening within a cell or outside of a cell.
And these are experiments that just wouldn't be
so easy or feasible otherwise.
So maybe we'd like to study protein structure.
What could we do?
So fluorine is used in NMR quite a bit.
Imagine if you could site-specifically
label an unnatural amino acid that
has a CF3 group, for example, and use
that in your NMR studies.
So that's something you'll get to think about in the context
of problem set two.
Ways to study protein function, protein localization.
So for instance, instead of attaching
GFP, which is big, to a protein of interest,
maybe it's possible to incorporate
a fluorescent amino acid that lets you
see that protein in the cell.
Protein-protein interactions.
And maybe we'd like to make a new protein that has
some desired characteristic.
So there's a lot of possibilities
to such technology.
Just to keep in mind, what do many of us do?
Many of us are familiar with site-directed mutagenesis,
where we can change an amino acid in a protein.
And we learn many, many things from this,
but it is limited to naturally occurring amino acids.
Right?
So we'd like something more versatile.
If we think about strategies also just a little
bit, backing up here.
OK, the first thing I'll just point out
is that how I'm going to divide this,
just in case this wasn't clear, is considering
the native ribosome and then considering
engineered ribosomes.
And this is where we're going to focus today.
And if we consider strategies, other strategies
to incorporate unnatural amino acids,
and I guess I'll call these standard,
we can imagine chemical and biosynthetic.
And I'm not going to go over a plethora of examples
for either route.
There'll be some slides included in the posted
lecture notes that gives examples and pros and cons.
But one example I will give here is just
thinking from the standpoint of a chemical modification, what's
an example and why we might want to do better.
OK, so this is independent of something
like site-directed mutagenesis, where you're
having an organism do the work.
So if we just consider an example of a chemical
modification, there's certain amino acid side chains that
are amenable to modification.
So imagine you purify a protein, and you
want to somehow tag that or label it, right?
One option is to modify cysteine residues.
And so iodoacetamide and related reagents are commonly employed,
so imagine that you have some cysteine.
You can react this with iodoacetamide
that has some R group, right?
What happens?
Here, OK.
You can get a covalent modification,
and maybe this is a fluorophore or something else, right?
So this is terrific, but what are some potential problems?
AUDIENCE: Sorry, would this be a way
to modify the amino acid before it's
incorporated into the protein?
Or would this be something you would
do to modify the cysteine in an assembled protein?
ELIZABETH NOLAN: Yeah, this would be after the fact.
So imagine you have some protein.
You've isolated your protein, and you have some cysteine.
Right, and you'd like--
for some reason, you'd like to modify this protein.
So maybe a fluorophore to see it.
Maybe you know, a CF3 group for NMR here,
which then gets to the point, what are
possible problems with this?
AUDIENCE: Do you have to use a mild base to be deprotonated,
or is it maybe deprotonated based
on where it is in the protein?
ELIZABETH NOLAN: Yeah, so that gets to an initial issue, which
is what's required to have this chemistry to happen?
Right?
The cysteine needs to be deprotonated.
So probably the pH of your buffer
is going to be elevated some.
Does your protein or enzyme like that or not?
Maybe, maybe not.
Yeah?
AUDIENCE: You can also run into selectivity issues--
I mean, having free cysteine residues isn't common,
but it could be a potential problem.
ELIZABETH NOLAN: Yeah, so you need--
well, it will depend on the protein, right?
Is the cysteine free or a disulfide?
Is it a native cysteine, or have you
done site-directed mutagenesis first to put this cysteine
in the position you want?
Right?
And then what happens if your protein has multiple cysteines
building on what Rebecca said, and you
want to have this label at a site-specific location?
Right?
What are you going to do about that?
Are you going to have non-specific labeling?
Are you going to mutate out the other cysteines?
If you do that, what could that mean for your protein
fold or function?
There's a number of caveats that need to be considered.
Nonetheless, it's a possibility to do.
In terms of time, this is a pretty extreme example,
but I'll just show one example here
in thinking about this whole process and what you do,
which also builds upon Rebecca's question.
So imagine a protein with two subunits.
And subunit 1 has a cysteine, and subunit 2 doesn't.
So for some reason, you want to do this labeling.
This is actually a protein from my group.
And we wanted to stick a fluorophore on it.
So we have a cysteine on one of the two subunits.
You can run this reaction and get this fluorophore modified
form here.
And then you can see that's the case, looking at SDS-PAGE.
So here we're looking at Coomassie stain that
shows us total protein, and we see
there's two subunits, 1 and 2.
So the molecular weights are a little different,
and we can separate them on this gel.
And then if we look in the fluorescence channel,
what do we see?
We only see fluorescence associated with subunit 1
and not subunit 2, which tells us our labeling strategy
has worked well.
Like, what we're showing in this equation.
But what's everything that needs to be done?
Well, we need to overexpress the protein in some organism.
In this case, E. coli.
We need to purify the protein.
And once we have this purified protein in hand,
we need to do the chemical reaction for the labeling.
And then we need to purify that product somehow,
and that's going to depend on the system you're working at.
And then it needs to be analyzed, right?
You always want to know what you're working with, right?
So was this reaction to 100%?
Did we end up with a mixture?
If it's a mixture, what to do about that?
So what does this mean in terms of time?
And this is not for all cases, OK?
This is for this exact case involving this protein shown
as a cartoon here.
So it takes about six days from start to finish to overexpress
and purify it.
Steps 2 to 4, based on the purification,
we do another four days, right?
So that's 10 days from start to finish, just
to get this protein you'd like to use in your experiment.
Right?
And you can imagine if somehow a label
could be put on in vivo, during this initial step
here, that that would save some time at the end of the day.
So before moving on to what's done
for unnatural amino acid incorporation
by what we'll call the Schultz method out of Professor Peter
Schultz's group, just to think about biosynthetic methods
for a minute.
So some common ones are done for structural studies.
So for instance, you can imagine feeding an organism something
like selenocysteine or selenomethionine.
Another example is labeling nitrogens or carbons
for NMR, where the organism is fed,
say, a labeled amino acid, maybe with N15 or C13 there, right?
So that's just a biosynthetic method,
where you're changing the growth conditions, rather
than doing something to manipulate the genetic code
or the ribosome.
So what's the conclusion here?
What we want is we want a method of site-specific incorporation
of unnatural amino acids in vivo.
So in a cell and in a desired organism, depending on what
you want to do with high efficiency and also fidelity,
so getting back to that idea and before.
OK, so why do we want to do this in vivo?
It allows for studies within cells,
and you also can purify protein from cells,
so you can do in vitro experiments as well.
OK, and you can imagine, if you could
have all of the pieces of this machinery in a cell,
maybe there's some technical advantage to that.
So this is what we're going to consider here.
So to this question, can the ribosome
incorporate unnatural amino acids into proteins?
Effectively, what do we need to think about?
One, we need to think about relaxing
the substrate specificity of the aminoacyl tRNA
synthetase to accommodate some unnatural amino acid, right?
Somehow that unnatural amino acid
needs to get to the ribosome.
So if this can be done, and we can
make a tRNA that has an unnatural amino acid attached
to it, can this aminoacyl tRNA get to the A-site
and do the work?
So this is the method we're going to talk about
in some detail for the rest of today
and into Wednesday, this Schultz method.
So the idea is that there's a tRNA that's dedicated
for this unnatural amino acid.
We see this unnatural amino acid shown here, where
the UAA is indicated by probe.
We need an aminoacyl tRNA synthetase
that will take this unnatural amino acid
and attach it to the three-prime end of the tRNA to give us
this aminoacylated tRNA.
And then what?
Imagine this tRNA can make its way to the ribosome.
What happens?
We need a codon for this aminoacyl tRNA.
It needs to carry the anticodon, and we're
going to talk about this in some more detail in a minute.
So we can have a plasmid that has
the DNA with the gene of interest in it, right?
This plasmid DNA can be transformed into, say, E. coli
that has this machinery here.
We can have transcription to give the mRNA that
is from this plasmid DNA.
And then imagine translations such
that this unnatural amino acid is incorporated.
So effectively, where we're going
is that we need a general method.
We want this method to be broadly useful,
where we can genetically encode this unnatural amino acid
and have it incorporated in response to a unique triplet
codon, here.
So in thinking about that, what are the pieces that we need?
And we'll think about E. coli for the moment,
but this could be other.
So yeast, mammalian cells, right?
Let your imagination run wild with this here.
When the incorporation of the UAA in response
to a unique triplet codon.
So if we're going to do this, what do we need?
OK, effectively, we need some new components of the trans--
like protein biosynthetic translation machinery, right?
So we need to rewind and think about the whole translation
process.
OK, so the first order of business
is that we need a unique codon.
Right?
So this only designates or uniquely designates the UAA.
And so we need to ask, where does this come from?
Because we just went over the fact
that the codons are used up for amino acid start and stop.
We need a new tRNA.
OK, so this tRNA needs to be specific for the unique codon.
OK, and we need the corresponding aminoacyl tRNA
synthetase, right?
And we need this to load the unnatural amino acid
onto the unique tRNA here.
So what is a key feature of all of this?
A key feature is that if we want to do this in some organism,
we need this machinery to be orthogonal to the machinery
in that organism.
We cannot have cross-reactivity, because then there's not going
to be any selectivity of this incorporation.
So no cross-reaction.
So what do we need to consider, in terms of these?
We need to think about all of the machinery, right?
And I just list some considerations here.
So this new tRNA can only allow for translation
of the codon for the UAA.
It can't be a substrate for any of the endogenous aaRS,
because then it will become loaded, potentially,
with the wrong amino acid.
So think back to lectures 2 and 3.
This new aminoacyl tRNA synthetase
can only recognize the new tRNA and not endogenous tRNA.
So cross-reactivity again.
This unnatural amino acid also can't be a substrate
for endogenous enzymes.
And also keep in mind, there needs
to be some way to get this unnatural amino acid
into a cell if we want to do this in a cellular context.
So there's just transport issue that needs to be kept in mind.
Will this unnatural amino acid get into the cell?
OK, so what we're going to do is consider these requirements
and what was done to build up this methodology
during initial work.
So the first issue is this unique codon,
and what is its identity here?
And so if we consider the 64 codons,
they're used up with the 20 common amino acids.
We have the three stop codons and the one start codon.
And so in thinking about this, can ask,
do we really need three stop codons?
We certainly need our start, and we need
codons for the amino acids.
But is there some wiggle room here?
And so in terms of these stop codons,
we have TAA, TAG, and TGA.
And these all have names.
Ochre, amber, and opal.
OK, and so the idea we're going to see is just the question,
can we reassign a stop codon?
And can we reassign a stop codon such
that it's the codon for the unnatural amino acid?
And so basically, if we want to reassign a stop codon,
how do we choose?
Right?
So two things to consider.
One, how frequently is each stop codon used?
So what do we know about that?
And then does this stop codon terminate essential genes?
So we can imagine that if we were to reassign a stop
codon that's used frequently by E. coli or another host,
or if we were to reassign a stop codon that's
important for terminating the synthesis of essential genes,
in either case, the outcome could be pretty bad, right?
So what was found in thinking about those issues is
this amber stop codon, TAG, one, it's the least frequently used.
And just for an example, about 9% in E. coli and about 23%
in yeast for terminating genes.
And additionally, it rarely terminates essential genes.
OK, so based on this, it was decided
to reassign TAG as the codon for the unnatural amino acid.
OK, so we've gotten through to here.
So the question is now, what about requirements 2 and 3?
So yeah?
AUDIENCE: I have a question.
So it seems interesting to choose a stop codon
to change because if the stop codon messes up,
it seems more catastrophic to us all than one
of the other redundant amino acids.
Like, why use a stop codon?
I think it's interesting.
ELIZABETH NOLAN: Yeah, so there is a risk.
There's certainly a risk, right?
But these considerations were made
to try to diminish that risk.
Right?
So you could make the argument that maybe all of these stop
codons aren't essential, right?
And what is more deleterious?
Will it be to try to use a stop codon that's infrequently used,
or to reassign a codon that's for an amino acid that comes up
in many, many different proteins in the cellular pool?
Right?
So there's a judgment call there.
But if we consider in E. coli, this TAG stop is for about 9%
of proteins.
How does that compare to, say, reassigning one of the codons
to incorporate a lysine or a valine.
I don't know, but that was just want
to think about, how frequently is that codon used?
Because certainly there's different codon usage
in different organisms.
Do you have something to say?
JOANNE STUBBE: So it depends on what
you want to put the unnatural amino acid in for.
So if you want it in endogenous levels, it could be a problem.
But if you're overproducing your protein--
ELIZABETH NOLAN: Yeah, it may not be a problem.
JOANNE STUBBE: Then it's not a problem, because you induce,
and then you flood it with that and you
get high levels of cooperation.
So it depends on what your purpose is.
ELIZABETH NOLAN: Yeah, so is JoAnne's point
clear to everyone?
So you could imagine expressing at an endogenous level, right?
Or you could imagine causing the cell
to overexpress the proteins, like off a plasmid,
like what many of you have done in lab class
or maybe in research there for that.
Are there many examples of reassigning a different one?
JOANNE STUBBE: I think it's really tough.
I mean, inside the cell, you really
do have problems if you don't re-engineer,
because you get truncations also.
ELIZABETH NOLAN: Yeah, we're going to talk about that.
So there is a big problem about the stop
that we're going to talk about, once we
get to how we have this done, which is premature termination.
AUDIENCE: I'm kind of confused more or less at that point,
because stop codon, we're just using because it's not
currently-- it doesn't go for anything,
it just ends, like, endogenous sequences that
are not the UAA so if we replace it, we might not get those.
But we'll get the one that we're trying to synthesize.
ELIZABETH NOLAN: Right, so we need a codon
for the unnatural amino acid.
And right now, we're limiting our space
to triplet codons, which is what was initially
done when this type of methodology was developed.
So the question is, we have four options in terms of bases.
AUDIENCE: That are not coding, right?
ELIZABETH NOLAN: No, no, in terms of our codons, right?
So three, right?
Four, three, so there's 64 codons,
and they're all used up.
AUDIENCE: Yeah.
ELIZABETH NOLAN: Right, so there's not some extra.
AUDIENCE: Yeah.
ELIZABETH NOLAN: So then what can be reassigned?
AUDIENCE: These stop-- so the stop.
ELIZABETH NOLAN: Well, yeah, well, we
can't reassign the start.
Then there's no proteins, right?
There's only one start codon.
So the thinking was, is a stop codon dispensable?
Right?
And then a decision was made, based
on basically the frequency of use of the stop codon
and whether or not the stop codon terminates
essential genes.
Is this something foolproof?
No, there's major problems in terms
of yields that come up as a result.
And we'll see that on Wednesday, right?
But you need a starting point to get a method underway.
So where will we begin tomorrow is
talking about where this tRNA and the aaRS
come from to do this.