DNA design questions

[Mega]

Hi,

When designing a gene cassette for synthesis:


Can you use LoxP - Gene Of Interrest - LoxP  without the fear that the target organism will make homologous recombination without the Cre gene?

I mean, they are completely the same sequence, so they may cut out my marker gene even without the Cre-recombinase, right??

If I remebmer correctly, they share 31 bp, so there's quite a nice possibility to recombinate, isn't it.

[Cathal Garvey]

LoxP sites, at about 31bp, aren't really that worrisome for
recombination. If there were lots of them, maybe.. but they've been used
for this purpose for ages in producing strains of recombinant or
knock-out mice and I don't hear many complaints. Mind you, I'm not
listening to that area of research, so perhaps you're right! :)

I'd say don't worry about natural recombination. Just make sure your
loxP sites are in the right orientation to excise the DNA and not just
flip it backwards! The relative orientation of LoxP sites can lead to
excision, flipping, or crossover. You want excision, of course!

Very excited to see where you go with this. :)

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[Mega]

It's for my pGlowroplast design ;) When I have it synthesized that will add barely a few dollars to the costs.

I also contacted another lab, who may be willing to give away a chloroplast integr/expr. plasmid. Hope that finally this thing goes right!

[Mega]

Hi,

when you want higher expression levels and you already have used the strongest promoter, what can you do?

With 35S Promoter, there's the double 35S promoter which does the job.


But can you just duplicate any promoter to get better expression? Won't the second promoter be transcribed into mRNA when you've bad luck?

[Cathal Garvey]

The second promoter would get transcribed, yes, but this wouldn't be a
problem; RNA promoters don't do anything, unless they contain lots of
secondary structures (which would be a problem).

I don't think doubling up on promoters is likely to help, though; in a
highly active promoter, you're already saturating the transcript. You
could replace the promoter with a viral promoter and express a viral
polymerase, but they're usually strong enough to kill the cell; look at
the T7 promoter in E.coli, for example!

Sometimes, excess expression can actually ruin an expression system; if
you make too much protein, it can start forming inclusion bodies.
Inclusion bodies, like crystals, can "seed" the inclusion of other
proteins, if I recall correctly, meaning you actually get less net
effect. And while it may be possible to re-purify functional protein
from inclusions after cell disruption, that's no use for bioluminescence
in-cell.

I would suggest trying to get a working system initially without
worrying too much about excess expression level. Once it's working, you
have a platform to experiment further with by changing codon usage,
promoters, or by editing the amino acid sequence of the protein to find
mutants with higher activity, stability at higher expression levels, etc.

And at that point, if you have a somewhat-glowing-plant working and want
to improve it, it'd be easy to raise crowdfunding for the mutagenesis
step to make it even cooler, because you can tell your funders that
worst-case-scenario they only get a somewhat-glowing-plant! ;)

[Mega]

Does anyone by chance know how long an amino acid chain can be without getting problems with transcription and/or translation?

Say you want to make a 6kbp long fusion protein  (you clone the genes in frame and remove the stop codons) .  That would make 2000 amino acids(!), and those sequences I've seen so far have around 400 amino acids. 

Will there be problems with the RNA polymerase or during translation? 

(Google didn't hit any adequate results, unfortunately)

 

[Pat]

The number of amino acids coded for shouldn't have an effect on translation or transcription. For instance titin a protein found in our bodies is ~30,000 amino acids long.

Increasing the size of the protein however will have an effect on protein folding and subsequent activity if the protein is an enzyme.

[Bryan Bishop]

On Mon, Oct 29, 2012 at 6:40 AM, Pat <ele...@gmail.com> wrote:

The number of amino acids coded for shouldn't have an effect on translation or transcription. For instance titin a protein found in our bodies is  ~30,000 amino acids long

I can't tell if proteins with subunits exist because of assembly problems, or if because different proteins co-evolved into some mutual function. Probably both, but designing new and working subunits isn't as easy. So, stick with 30k amino acids, for now.

- Bryan
http://heybryan.org/
1 512 203 0507

[Cathal Garvey]

The longer the chain, the more chances there are for a weak link to
break it.

Issues of transcriptional stalling or translational abortion/stalling
could end up killing the whole system, and it's hard to debug. At least
if you have several smaller fusions, you can try to identify which part
of the system is failing and fix just that.

I'd suggest several smaller fusion proteins rather than one giant one.
Fuse the critical pairs or sets that form "bottlenecks" in the system:
LuxAB are cofactors, so fuse them. LuxCDE are steps in a synthetic
chain, so fuse them.

[Andreas Sturm]

Sounds interesting thanks!!

But if I do take Promoter -LuxAB - Trmnr - Promoter- LuxCDEG- Trn, the chance of homologous recombination wil be there.

And I must take the strongest promoter to get any visible light ammount, I'd have to take it twice...


If I include the viral 2a oligopeptide and *hope* it works, that will circumvent the problem with possible weak links?

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[Nathan McCorkle]

homo recomb from what though? just use diff promoters and terminators
if that's what you were concerned about.... or the same promoter with
different codon usage

--
-Nathan

[Andreas Sturm]

Other promoters won't be sufficient, because it have to be the strongest available. And there's only one strongest.

The same promoter with different codon bias? Sounds great, but are you sure that works? I have never heard of that before. But it may be true.

[Nathan McCorkle]

it would work better than having your sequences homo recomb'd out....
also remember what Cathal said about too strong being bad in some
cases. certainly 1000 polymerase per second vs 900 is not much
different... i don't know what average polymerase per second flux is
for low, medium, strong, strongest promoters... does anyone actually
use polymerase per second??? (other than the igem folks or whoever
created the term)

is poloymerase per second (PoPS) actually a good way to judge a
promoter? It seems decent, but it doesn't address overexpression
leading to inclusion bodies, etc....

[Cathal Garvey]

I don't *think* that sequences around the length of a normal promoter in
such close proximity will be a big recombination hazard.. but it's
possible. One way to try to address this would be to read up as much as
possible on the promoter, and run BLAST on the promoter (accepting only
close relatives which are still active for comparison), and try to
identify which parts of the promoter are most important for function.
Then, make some changes to the apparently "unimportant" parts of the
promoter.

This is much harder in eukaryotes than bacteria, because promoter
structure is usually less "logical": in E.coli for example, the
consensus promoter for Sigma Factor 70 (the "constitutive")
transcription factor is:
TTGACANNNNNNNNNNNNNNNNNNNTATAATNNNN[CDS]

Whereas in Eukaryotes and Archaea you get common motifs such as the
TATAAA and the B-recognition sequence SSRCGCC* close to the CDS but with
less stringent spacing, and then you can have enhancers and such up to
several kilobases away. Regardless, this means in Eukaryotes, you have a
similar immediate promoter structure consisting of highly important and
unimportant DNA sequences, but unless you understand which is which you
can't as easily edit things to reduce similarity between otherwise
functionally identical promoters. At least, not without potentially
sacrificing promoter strength.

Something to consider: if I recall correctly, the limiting factor in
luminescence is luciferin availability. That is, having the same
promoter strength for LuxAB and LuxCDE should be a lower priority than
having LuxCDE under strong expression, assuming you can provide LuxCDE
with what it/they need to produce lots of Luciferin. Which is:
tetradecanal. Can't recall if LuxCDE is enough to convert Tetradecanol
to Tetradecanal, or would you need another enzyme to catalyse that..I
think not.

* that's IUPAC notation: S means "strong" = G or C, R means puRine = A or G

[Andreas Sturm]

That's interesting. LuxG (or fre) was the limiting factor (in eucaryotes).
Next it could be Decanal (CDE).

Don't know if there is abundant luxAB. I read that this is not really an enzyme, because it reacts and is then junk (1 proper enzyme catalyses 10'000 molecules substrate, as you know) .

So that would make Promoter1-LuxAB Kanamycin-Term- Prom2- LuxCDEG. So basically, there's not much diference in lenght.

[Andreas Sturm]

That's interesting. LuxG (or fre) was the limiting factor.

Next it could be Decanal (CDE).

Don't know if there is abundant luxAB. I read that this is not really an enzyme, because it reacts and is then junk (1 proper enzyme catalyses 10'000 molecules substrate, as you kno .

[Cathal Garvey]

Got a reference for the LuxAB only working once? That's really weird
sounding! Canonically, luciferases work as catalytic enzymes, have never
heard of them being exhausted so rapidly..

[Andreas Sturm]

I'm going to look that up. I was surprised too, because luciferase is always refered to as an enzyme.

It may also be wrong, I just read it once.

[Mega]

Hi, 

for bbacterial / plastid expression: 

Is a terminator ( 3' UTR)  really neccesary? 


Because, bacteria can do polycistronic translation, so the next gene will also be on the same mRNA and bring its own terminator...

[Mega [Andreas Sturm]]

Hi all,

I am designing primers to get a kanamycin resistance gene.

My question is, the kanamycin gene (nptII)  is out of pGreenII for plants. I want it to work in E.Coli. Will the this eukaryotic RBS also work in E. Coli?

TGACGTTCCATAAATTCCCCTCGGTATCCAATTAGAGTCTCATATTCACTCTCAACTCGATCGAGGCATGATTGAACAA

As I need a BamHI site anyway, I could also mutagenize it into this...

TGACGTTCCATAAATTCCCCTCGGTATCCAATTAGAGTCTCATATTCACTCTCAACagGATCcAGGCATGATTGAACAA

The agga-motive rather looks like prokaryotic beings will accept it?


Or just leave it as it was?

[Koeng]

I don't think Eukaryotes use RBSs...

http://en.wikipedia.org/wiki/Five_prime_cap

[Andreas Sturm]

Wikipedia (yeah, not that scientific source) After finding the ribosome binding site in eukaryotes, the ribosome recognizes the Kozak consensus sequence and begins translation at the +1 AUG codon. ( http://en.wikipedia.org/wiki/Ribosomal_binding_site )

Btw,
http://www.invitrogen.com/site/us/en/home/References/Ambion-Tech-Support/translation-systems/general-articles/ribosomal-binding-site-sequence-requirements.html

 Our data demonstrate that in contrast to the E. coli ribosome, which preferentially recognizes the Shine-Dalgarno sequence, eukaryotic ribosomes (such as those found in retic lysate) can efficiently use either the Shine-Dalgarno or the Kozak ribosomal binding sites.

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[Koeng]

Nice! I will keep that in mind. However eukaryotes still need that 5 prime cap :(