Beginner wants to create biolumineszent plant

[Mega]

Hello guys,

I'd like to 'create' trees that glow for making a bonsai. I could get
the lux-codon from vibrio fischerii bacteria extracted from seafish.
It'll surely be possible to get some agrobacterium tumorfaciens.

But how can I combine that, remove the bad plasmide from that
bacterium and insert the glowing genes (but not the saline genes,
because the tree shouldn't need salty water to live like the
bacterium).

I read about using an ultra-sonic cleaning device for making the
membranes pourosly. Would that be an option?


Or are there already manupulated agrobacteria including the lux genes
available on the market??


Yours

[Pieter]

You might want to take a look at last years Cambridge iGEM team
project E. Glowli: http://2010.igem.org/Team:Cambridge. Biobricks for
luciferase and luciferin regenerating enzyme are readily available
from the Registry. You can just order a plasmid from the Registry or
synthesize it with some adjustments to your host organism yourself at
a DNA mail order company. You might need to change the codon
optimalization and remove some bonsai specific restriction sites, etc

[Brian Degger]

Mega,
I like your idea, but there are more than just technical issues to this project.

I do want a society where gmo's are not a special regulated class of organisms. In Europe, what you suggest would be governed by rules, regulations and laws. It would be illegal to have in yr home. In America you can buy glowing gmo zebrafish for your child's aquarium.
Have you done biology? You obviously know some biology and have an idea of the process,  but I suggest a glowing bonsai is an advanced first project. What species is your bonsai?
Cheers b

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[Cathal Garvey]

Hey Mega,
You've clearly done a lot of good reading, kudos! About the only error there was "saline genes", there's no need to worry about those. Tolerance for salinity is an additives trait for most species, a combination of many genetic and physiological factors.

As to the plan. It's an ambitious goal, that much you ought to know from the outset! It will be much harder to achieve than a regular bacterial transformation, and more expensive.

First thing to consider is how to get the DNA you need. Rather than extracting DNA from vibrio, I recommend

[Cathal Garvey]

(whoops! ).. I recommend synthesising the DNA. To explain why, let's discuss the "language" of dna. You've heard it's a "universal code" right? That's not entirely true. The amino acid code is universal, although each species has a slightly different set of preferences. However, the rest of the 'code', that which instructs processes such as transcription (and the very first steps of translation), is far less universal. Bacteria and plants have evolved very different codes by now.

Also, bacteria have developed a clever way to automate complex pathways: they combine many protein-coding sequences under one promoter-terminator pair. That mightn't work for plants, and so you might have to cut the sequence up into many sub-genes.
That means that if you use wild vibrio DNA, it may not work until you have altered all of the following at least:
- New promoter
- New ribosomal binding site ("kozak consensus" instead of "shine dalgarno")
- Altering some troublesome codons.

And you may also need to subdivide the genes and possibly add a new terminator.

A lot of work. Years in fact, because biology *that* way is hard and prone to error.

Instead, if you design the."perfect" DNA on a pc and order it from a synthesis company, you will require far less time and effort... But lots more money!

As to agrobacterium, get a lab strain. Trying to recreate the traditional strain from scratch will be almost as much work as the above.! Alternatively look up Cambria, a company / organisation that have developed less patent - laden strains of "I can't believe it's not Agrobacterium!" for sort-of open source use.

Depending on which species you use, getting the DNA into the bacterium may be easier to do chemically. It's really worth reading deep into the literature and seeing what methods are easiest to do. For example if you asked how to transform E. Coli you might be told to use calcium chloride.. But if you read, you learn that peg3350+ mgso4 works great. In other words, Epsom salt and laxative or lubricant! Far easier.

For now that's all I can offer, bit busy over here! There's other stuff to consider to do with the pathways, regulation, etc. Hopefully soon I can help more, could even suggest ways to design the DNA for synthesis.

[Mega]

Thanks for all your comments. Soon this year I'll begin to study
biology, what i yet know about it comes from books I've read.

It may take some time to get started in practice...





Would it possibly be easier to 'modify' lichens? I imagine all I have
to do is to extract luminous (non-pathogenic) bacteria, and put them
together with some lichens in an ultra-sonic device... Some of the
bacteria may be exchanged or maybe even some genetical material
creating luminous lichens!

[Nathan McCorkle]

the flower-dip method was originally used with agrobacterium and
/Arabidopsis thaliana/

paper with protocol here:
http://faculty.salisbury.edu/~flerickson/protocols/floral%20dip%20transformation.pdf

Not sure how far this has come for use with other plants, but
basically if you want to avoid doing plant tissue culture to isolate
transformed plant cells, you try to transform the single-celled
embryos in the flowers.

Lots of work as others point out. Mainly because you have to
transform, then grow hundreds or thousands of plants, and check each
to see if they were successfully transformed. Lots of single-celled
embryos will die because the gene insertion is lethal, others may
cause mutations that stunt the plant too badly, others will not be
transformed and just create lots of 'noise'.

Other than the flower-dip, you could construct you DNA then shoot it
into plant cells with a 'gene gun'. Then you would have to dissociate
the plant cells, then dilute the solution enough to dispense one cell
into each of a tissue culture medium. Then screen each cell line for
transformation, when you find a positive, attempt to regenerate the
plant.

Hope this helps!

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[Bastian Greshake]

Hi,

Am 04.09.2011 um 22:29 schrieb Mega:

> I'd like to 'create' trees that glow for making a bonsai. I could get
> the lux-codon from vibrio fischerii bacteria extracted from seafish.
> It'll surely be possible to get some agrobacterium tumorfaciens.

I dunno which kind of plant you want to use for the bonsai, but remember that Agrobacterium tumefaciens works best in dicot plants (while you can make it also work for monocots in a lab).

Cheers,
Bastian

[Mega]

I thought of an maple bonsai.

Best

[Mega]

I came across a new idea:

It's very low problem to get some vibrio fischeri.
I can also get some EDTA and Lysozyme, and thus i can extract DNA and
RNA (and some proteins with that which don't affect the process).

So here I have the genes including the full lux-codon.


I read that under certain conditions - such as ethyleneglycole in the
medium - plant cells can take bacterial DNA and include that into
their own.

So I take the plant ( at first some very low plant like Bryophyte or
liches <- ok, at all no plant but the cyanobacteria may take the DNA
better).



Do you think that could possibly work?? I imagine that one
disadvantage may be that the lux codon has to be on a certain spot in
the DNA to work correctly. So it would be just luck (or accidentally)
that it would get inserted on the right spot in the plants DNA.

Another concern is quorum sensing -> will the cell density e.g. in
bryophyte be high enough to make the cells create light??



Yours,

[Cathal Garvey]

Just to clarify and assist some points:

You'll need more than EDTA and Lysozyme, but once you've got lysozyme the rest is certainly achievable, I think. I'll go look for my (as yet untested!) DIY instructions that should use only store-bought chemicals after the lysozyme/EDTA for a miniprep. Extracting Genomic DNA will be different to a miniprep, but not vastly different. Actually, the EDTA isn't even necessary, but it's strongly advisable; it serves to scavenge (chelate) the metal ions that DNA-chewing enzymes use to cut DNA, protecting it during extraction and preserving it for storage.
You could probably make do with citric acid if you buffered it correctly, also; citrate acts as a chelating agent, although I've had tremendous difficulty finding details on its precise properties; i.e., what pH ranges lead to maximum chelating potential, etc.. I've considered this a juicy unanswered question for a long time now, but IANAC (I am not a chemist!).

As to the DNA bit. If you lyse the cells correctly, you'll have raw DNA; possibly all in one piece, probably somewhat fractured. If you just bang that DNA into a somewhat-compatible host (let's say another bacterium, such as a cyanobacterium), you still won't get what you want; you're throwing thousands of genes into a cell and you're hoping that one of them sticks by accident; more likely, if any of the DNA runs at all it'll disrupt and kill the cell.

So you need to isolate just the DNA you want. Then, you have a number of things to consider, all of which are issues in themselves:

1. Will the DNA run as-is in my target cell? <--- BIGGEST question by far.
   
2. How am I delivering the DNA to the target cell? (e.g. Chemical transformation?)
   
3. On what vector will the DNA be maintained? (Plasmid, chromosome?)
4. Does my host cell support the method of DNA delivery and maintainance that I want to use?
5. How will I verify that the DNA arrived, and how will I troubleshoot if it doesn't work?

Questions 2-4 are a species-specific issue. What works for E.coli won't work for B.subtilis and vice versa. If you plan to use a plasmid to carry the DNA in the target cell, you must find a plasmid that works with the target cell, if one is available. If not, you'll have to investigate other options; has anyone previously used methods that get the DNA into the cells, and which method did they use?

Question 5 is a matter of good experimental practise. It's not enough to say "they're not glowing, the problem must be in the DNA delivery stage": how do you know that? Isn't it just as likely that the DNA arrived, but didn't work? This is why antibiotic selection is such a popular route to DNA transformation in bacteria; if they survived, then they *probably* have the DNA. Verification is possible thereafter by PCR or miniprep, but the selection step weeds out the huge background noise of cells that simply didn't take the DNA. I'm in favour of more responsible/cheap/accessible methods than using antibiotics at home for DIYbio stuff, but they're still the standard.

Question 1 though, is the biggie. DNA that runs just fine in one species mightn't run at all in a close neighbour on the evolutionary cascade, but you can be practically certain it will NOT run in a species belonging to another kingdom. i.e., adding V.fischeri DNA to a plant cell suspension will simply not work.

Instead, the DNA needs to be refactored by a little or a lot to get it to work. If you're working with bacteria only, as in V.fischeri -> Cyanobacteria, then you may need to change the promoter(s) only, possibly also tweak the Shine-Dalgarno (start codon region), and the rest might work O.K. as is. However, for plants, you might have to go a lot further. The promoters will need to be completely replaced, as will the start codon region in all likelihood, and codon bias (the "dialect" of the universal amino acid code) is likely to be much more distant. Also, you may have to hack in some additional regulatory bits at the end if a simple terminator isn't enough for the plant you're targeting, and you may need to pay attention to CG content to avoid the DNA being instantly flagged as "foreign" as some plants seem to do.

This is what synthetic biology is all about, and it's why it's making all the waves it's making; rather than doing the seemingly simple approach of extracting DNA and firing it back into a target cell, you're synthesising the DNA from scratch after designing it for its intended purpose. It's more expensive up front, but you're wasting less time, you require less expertise, and you can make all the changes you want for probably the same price. In the end, it's more likely to work, it's expensive if it doesn't, but at least you tend to know quickly.

The above isn't intended to be a discouragement. It's a primer, and there's as much additional assistance and advice as you'd like along the way if you go through with the project; I'd love to help make it happen. Expect it to be expensive and difficult, though; there's a reason we don't already have glowing houseplants or lichens, but now is a good time to be audacious when it comes to biotech!

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[Nick Beck]

This is a project super interesting to me, as it has been a dream of
mine as well. That said, I think there are some major challenges for
doing this in addition to the preceding points. Both metabolic flux
and bioluminescence triggers are extremely challenging when
synthetically constructing a multicellular host.

I'm not an expert on metabolic flux or the metabolic economics. That
said, assuming expert synthetic assimilation of the lux complex
(challenging per previous statements), you still have to address the
fact that you are adding a major energetic sink to your organism.
Balancing this against the very streamlined and highly conserved
metabolism of photosynthetic plants would be very complex. I think
your best bet would be to excise and substitute for a pathway that
produces a characterized natural product in leaves. There is no need
to bioluminesce in heartwood. You would need to siphon out energy
without compromising the other aspects of the hosts growth,
development, and metabolism. For this reason, cyanobacteria would be
orders of magnitude easier. Dinoflagellates photosynthesize and have
light/dark cycling built in, and are naturally bioluminescent. Check
out the links below, particularly the "Organisms" section of the
Bioluminescence website.

Second, is triggering. Many bioluminscent organisms respond to tactile
stimulation, endogenous (neuro/chemical) signalling, or exogenous
chemical queues. This is particularly true in marine organisms. Some
bioluminescent fungi do appear to be passively 'on' once the gene
begins expression. They are less studied, but may actually be a
superior starting point for you. Fungal genomes have their own
complications owing to the fungal life-cycle, though. You would need
to find a way to trigger bioluminescence. If it is always "on," you
would need to account for that perpetual energy cost. In a plant,
always "on" would be a pointless drain, since plants typically need to
photosynthesize, and the photon emissions of bioluminescence would be,
at best, extremely dim in photosynthetically significant illumination.
So, you would need a light/dark switch or some sort of exogenous
queue, which would require a sensing pathway. Sensing and real-time
pathway regulation have been achieved numerous times, but typically in
"lower" organisms (E. coli) with far fewer morphological and metabolic
complexities than higher plants.

For more on bioluminescence, and fluorescence, see these two excellent
websites. Personally, I would strongly recommend fluorescence as a
starting point. Once you get that going you could apply much of the
success to bioluminescence in v2.0/3.0. Transgenic fluorescence is
well characterized across kingdoms, so it would be more tractable.
Also, your exogenous stimuli would just be an appropriately tuned
lamp. LEDs are cheap and can be tuned to the absorption spectra of
your pigment.

Bioluminscence website: http://www.lifesci.ucsb.edu/~biolum/
Latz lab @ Scripps: http://siobiolum.ucsd.edu/biolum_intro.html

Cheers,
Nick

On Oct 1, 2:43 pm, Cathal Garvey <cathalgar...@gmail.com> wrote:
> Just to clarify and assist some points:
>
> You'll need more than EDTA and Lysozyme, but once you've got lysozyme the
> rest is certainly achievable, I think. I'll go look for my (as yet
> untested!) DIY instructions that should use only store-bought chemicals
> after the lysozyme/EDTA for a miniprep. Extracting Genomic DNA will be
> different to a miniprep, but not vastly different. Actually, the EDTA isn't
> even necessary, but it's strongly advisable; it serves to scavenge (chelate)
> the metal ions that DNA-chewing enzymes use to cut DNA, protecting it during
> extraction and preserving it for storage.
> You could probably make do with citric acid if you buffered it correctly,
> also; citrate acts as a chelating agent, although I've had tremendous
> difficulty finding details on its precise properties; i.e., what pH ranges
> lead to maximum chelating potential, etc.. I've considered this a juicy
> unanswered question for a long time now, but IANAC (I am not a chemist!).
>
> As to the DNA bit. If you lyse the cells correctly, you'll have raw DNA;
> possibly all in one piece, probably somewhat fractured. If you just bang
> that DNA into a somewhat-compatible host (let's say another bacterium, such
> as a cyanobacterium), you still won't get what you want; you're throwing
> thousands of genes into a cell and you're hoping that one of them sticks by
> accident; more likely, if any of the DNA runs at all it'll disrupt and kill
> the cell.
>
> So you need to isolate just the DNA you want. Then, you have a number of
> things to consider, all of which are issues in themselves:
>

>    1. Will the DNA run as-is in my target cell? <--- BIGGEST question by
>    far.
>    2. How am I delivering the DNA to the target cell? (e.g. Chemical
>    transformation?)
>    3. On what vector will the DNA be maintained? (Plasmid, chromosome?)
>    4. Does my host cell support the method of DNA delivery and maintainance

>    that I want to use?

>    5. How will I verify that the DNA arrived, and how will I troubleshoot if

> glowing houseplants or lichens, but now is a *good* time to be audacious

> when it comes to biotech!
>

[Mega]

"If you just bang
that DNA into a somewhat-compatible host (let's say another bacterium,
such
as a cyanobacterium), you still won't get what you want; you're
throwing
thousands of genes into a cell and you're hoping that one of them
sticks by
accident; more likely, if any of the DNA runs at all it'll disrupt and
kill
the cell. "

This brute-force approach was exactly my plan ;)

Doing that with one cell won't work - I know for sure. But using some
thousands of them at least a few may be correct.


"[It's a waste of energy for the] plant" - Well I don't care.
evolution shall take care of that, because it always does.


Well, then I'm going to have a look at fluorescence first.

[Nathan McCorkle]

On Mon, Oct 3, 2011 at 2:25 PM, Mega <masters...@gmail.com> wrote:
> "If you just bang
> that DNA into a somewhat-compatible host (let's say another bacterium,
> such
> as a cyanobacterium), you still won't get what you want; you're
> throwing
> thousands of genes into a cell and you're hoping that one of them
> sticks by
> accident; more likely, if any of the DNA runs at all it'll disrupt and
> kill
> the cell. "
>
> This brute-force approach was exactly my plan ;)
>
> Doing that with one cell won't work - I know for sure. But using some
> thousands of them at least a few may be correct.

you should start learning how to design a microfluidic device that
would do what you want. You basically need to use a CAD program to
design the fluid traces, and potentially you would have a few layers
that interconnected, not unlike electronic circuit design.

I am working on making microfluidic prototyping easier, but there
aren't designs for the actual fludic systems open/popular online yet.

Shearing bacterial DNA and transforming it into plant cells would be
pretty easy, but you'd need to do it hundreds of thousands of times...
and you'd need to watch the cells grow too to detect if something was
working. That said, sheared DNA will generally only integrate into the
genome if there are some matching sequences on the termini, to get
homologous recombination going. Adding those to your sheared DNA is
also possible... but then the bioluminescent system is probably
relatively new compared to the rest of the organism's metabolism, so
the metabolic pathways to create light might not be found in a sheared
genomic fragment... this would be a problem if intermediary molecules
weren't found in the plant. Getting the DNA to express in a plant is
another problem, which I think Cathal went through pretty well....
Sure plants could mutate the DNA and it could eventually start
working, but that would take meiosis, not mitosis, and meiosis in
plants involves pollen and eggs, which are found in flowers, whole
plants are not in the realm of parallelized microfluidics.

>
>
> "[It's a waste of energy for the] plant" - Well I don't care.
> evolution shall take care of that, because it always does.
>
>
> Well, then I'm going to have a look at fluorescence first.
>

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[Brian Reichholf]

The lux operon is a tricky one due to it requiring quorum sensing to work sufficiently. I do not think that simply transforming cells with it would suffice to allow easily detectable bioluminescence.

From what I've read bioluminescence is often facilitated using the luciferase gene from fireflies. From what I've read eukaryotic expression of V. fischeri derived luciferase requires a suitable promotor and a fusion protein of luxA and luxB (the latter could probably be quite simply achieved using PCR, as they are neighbouring genes).

GFP on the other hand would be a good place to start, as it can be handled fairly easily and has been established to work in higher plants such as Arabidopsis thaliana too.

[Mega]

http://www.terrificscience.org/lessonpdfs/GlowingEcoli.pdf

Hey guys, in this pdf the speak about a strange 'plasmide lux' that
can simply be inserted in e.coli... But Google shows me no reasomable
results for that term....

Are such plasmides with the lux-operon available anywhere???


Yours sincerely







On 3 Okt., 19:39, Nathan McCorkle <nmz...@gmail.com> wrote:

> the metabolic pathways tocreatelight might not be found in a sheared

> genomic fragment... this would be a problem if intermediary molecules
> weren't found in the plant. Getting the DNA to express in a plant is
> another problem, which I think Cathal went through pretty well....
> Sure plants could mutate the DNA and it could eventually start
> working, but that would take meiosis, not mitosis, and meiosis in
> plants involves pollen and eggs, which are found in flowers, whole
> plants are not in the realm of parallelized microfluidics.
>
>
>
> > "[It's a waste of energy for the] plant" - Well I don't care.
> > evolution shall take care of that, because it always does.
>
> > Well, then I'm going to have a look at fluorescence first.
>
> > --
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[Cory Tobin]

> Hey guys, in this pdf the speak about a strange 'plasmide lux' that
> can simply be inserted in e.coli... But Google shows me no reasomable
> results for that term....
>
> Are such plasmides with the lux-operon available anywhere???

http://www.addgene.org/14080/

-cory

[Mega]

Thank you for that.

Does it include quorum sensing or has this been cut out (I hope so)?

Let's assume I just put this plasmide in an e.coli. Will it just
become luminiscent?
Or do you have to grow it on a substrate containing luciferin?

[Cathal Garvey]

Generally, the research plasmids are either re-jigged to remove quorum sensing (i.e., by replacing the natural promoter/operator with a constitutive promoter and no operator), or have the quorum producing protein under constitutive control.

Usually the former. i.e., for a popular research plasmid such as the one on addgene, you should just get glowing colonies. However, the degree of glowing may not be up to your expectations, so caveat-emptor. Many labs use plasmid systems that produce detectable but not visible levels of bioluminescence; they use instruments instead to measure bioluminescence as a reporter of XYZ. Might be an idea to google the plasmid name and see if anyone has uploaded pictures or information on its actual activity.

Alternatively, compare the sequence of the operon on that plasmid to the iGEM plasmids used by the (Cambridge?) team last year; they created a few variant plasmids and had some nice videos showing the bioluminescence in action, it was clearly visible to the naked eye. If the research plasmid isn't too different to the iGEM plasmid in terms of operon structure, codon optimisation index of the CDS's, promoter strength... it should be fine.

Some plasmids have a rijiggered operon which has a big impact on the outcome; by having the luciferin-synthase genes on a separate expression casette from the luciferase subunits, you can get far brighter outputs and much better control over bioluminescence through exogenous systems. So, if you see a two-operon system with LuxAB controlled separately, that might be better.

All the above should be taken as hearsay and speculation; I've only ever played with the wild-type Photobacterium phosphoreum, and while it's lovely and bright I don't know how it measures up against E.coli.

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[Cory Tobin]

> Does it include quorum sensing or has this been cut out (I hope so)?

No quorum sensing. It's just the luciferase expressed from pBBR1,
which is a vector compatible with most Gram negative bacteria. Medium
copy number.

> Let's assume I just put this plasmide in an e.coli. Will it just
> become luminiscent?
> Or do you have to grow it on a substrate containing luciferin?

You will need to add luciferin.

-cory

[Cathal Garvey]

I'm not sure there's cause for pessimism here. The same bioluminescence
system functions in Vibrio/Photobacterium phosphoreum, a psychrophile,
at 4C without issue; the lack of warmth in the average plant shouldn't
be a barrier to bioluminescence.

Further, LuxCDE handle reduction of tetradecanoic acid (AKA Myristic
acid) to tetradecanal; the fatty acid precursor is pretty common and
occurs abundantly in species such as nutmeg:
https://en.wikipedia.org/wiki/Myristic_acid

The challenge is to ensure that LuxCDE colocate in a subcellular
fraction where they will encounter lots of tetradecanoic acid, and that
your LuxAB and FMNH2 likewise colocate with the "prepared" tetradecanal.

Perhaps the best route is to get your operon into the bacteria already
present in the plant cell; the chloroplasts. You can either tag each
protein with a chloroplast localisation peptide, which instructs the
plant cell to direct the finished proteins to the chloroplast upon
ribosomal synthesis, or you can transfect the chloroplast itself with
the transgenes.

Plastid transfection can be accomplished at some useful efficiency using
PEG3350 (Miralax):
http://onlinelibrary.wiley.com/doi/10.1111/j.1365-313X.1993.00729.x/abstract
(Locked behind paywall)

Here's a sample protocol using PEG 4000 (Miralax should work), mannitol
and Calcium Chloride:
"All constructs were used to transfect Arabidopsis protoplasts, isolated
from 3 - 5 weeks old Arabidopsis leaves grown under long day conditions
(23°C, 16/8 hr light/dark), essentially as described [40]. Cell density
was adjusted to 2 × 106/ml. 100 μl protoplasts were transfected with 20
μg plasmid DNA in 40% polyethylene glycol 4000, 0.8 M mannitol, 1 mM CaCl2."
From: http://www.biomedcentral.com/1471-2148/10/379

Would it be easy? Hell. No. Tobacco is really easy to engineer compared
to other plants, so you won't know until you try whether the PEG/CaCl
method will work at all in your target species.. and by then, you might
have wasted money on ordering DNA that will work in the target species.

OTOH, in the USA you guys can always get an airgun and modify it into a
crappy biolistics gun to shoot DNA-coated particles at your plants.
There are protocols for coating particles with DNA out there.

It'll be expensive and difficult and it's *not* for beginners, but it's
certainly possible IMHO.

On 05/11/11 03:31, Somebody wrote off-list:
>> This is not true. the LuxCDABE cassette encodes a pathway that produces light independent
> of luciferin. LuxCDE reduce fatty acids to aldehydes, which are oxidized by LuxAB making
> light. LuxAB genes are active alone in making light with the addition of long chain
> aldehydes, such as decanal. See Genbank AF170104
>
> The similar cassette from Photorhabdus luminescens works better than the more common
> cassette from Aliivibrio fischeri, in that it is active at 37C, while the fischeri cassette
> will not function above about 30C.
>
> Neither of these cassettes will likely function at all in plants without a LOT of work.

[Bacter]

Oh man that modified airgun would be super-handy though...

[Bacter]

Also, I'm not sure that brute-force will be sufficient here. If like Cathal said, you can isolate specifically the DNA you want, then sure give it a couple million events and one of them would probably stick somewhere that doesn't break something, but expecting just the right bit of DNA to be broken off, never MIND the inclusions of any promoter/enhancer regions, AND in that same cell stick to somewhere profitable, well... I think that would probably take a lot of tries.

A lot like high two-digit superscript after a 10 kind of lot. Why not find the easiest plant to transfect and start there? I guess the growth times for tobacco would be pretty irking?

[Patrik]

On Nov 6, 3:13 am, Cathal Garvey <cathalgar...@gmail.com> wrote:
>
> OTOH, in the USA you guys can always get an airgun and modify it into a
> crappy biolistics gun to shoot DNA-coated particles at your plants.
> There are protocols for coating particles with DNA out there.
>
> It'll be expensive and difficult and it's *not* for beginners, but it's
> certainly possible IMHO.

Yeah, I definitely think we should put some thought into a DIY gene
gun. That seems entirely feasible, and could actually circumvent a lot
of genetic engineering hassles.

Cathal, when you say that we could do this "in the USA" - what would
be the main impediment for you guys? Gun regulation issues? Or gene
gun / genetic engineering issues? Just wondering.

[Mega]

Ok, what I learned so far...

Firstl, it's no big deal for a pro to insert a lux operon containing
plasmide in an e.coli cell to make it glow. the antibioticum
ressistance and lacZ' will make it selectable.
Usuall, there's no more quorum sensing because the have cut it out.

- ''Just'' insert the plasmide into a bacterium and it glows.

To engineer a plant on the other hand is a much more difficult issue.


But what about a path in the middle (don't know if this term is used
in english ;) )?

Lichens consist of a fungus and canobacterium.



So, i take a plasmide that contains lux and ampecillin resistance.

Now i transform very small pieces of the lichen. Some of the
Cyanobacteria will be transformed and ressistant to ampecilline.

The fungus should either be ressistant to ampecilline on its own
(true??) or the cyano spread the ampecilline-degenerating substance
within the whole lichen.

Polyethleneglycol will help the endoctose to take place...

Then I'll grow them on a Lamp plate to select the lichens that include
ressistance. Of the bacteria inside the lichen, the ones containing
the most ampecilline-degenerating substances, these will grow fastest
and so they will mitose much more often.


I think that could be a way to go?

[Cathal Garvey]

Legal impediments; you need a license to own a gun in Ireland, and AFAIK
that does include low-caliber air-guns.

Which is fine in my opinion; we Irish had a storied history of shooting
one another, and the lack of guns in the public domain mean our police
force is likewise unarmed. It's a nice compromise.

However, this does lead to my focusing on chemical or biological methods
of gene transfection, which is probably good really: given the choice
between a chemical and instrumental route, the former is usually easier
for a toe-dipping beginner to invest in and try out. The latter is often
more reproducible provided everyone's using the same equipment, but
that's not true of a "Make your own Gene Gun!" situation. You'd need the
"OpenPCR of Gene Guns". And patents would probably get in the way of
something awesome along those lines.

[Nathan McCorkle]

Why not use electroporation for transfection... I've got this book:
http://pages.towson.edu/jsaunder/Saunders%20Publications/48.Guide%20to%20Electroporation%20and%20Electrofusion.pdf

and its like the holy-grail for electroporation. Techniques and
reviews of systems, and it even talks about building your own (the
advantages and disadvantages mainly).

"The exponentially decaying wave generator gave high rates of both
uptake and expression; however,
the pulse field strength working range was very narrow. Regardless of
which wave generator is used, it is clear that the
experimental protocol must be optimized for each cell type
that is being examined. The optimization often involves the
use of different electroporation chambers. The cuvette-style
electroporation chamber (13) increases the ease and simplicity
in the handling of cells during electroporation and has evolved
as an industry standard"
from:
http://www.plantphysiol.org/content/99/2/365.full.pdf

Exponential decay is what capacitors do, and I know we've got some
Electrical Engineers active on this list... I'm no electrical expert,
but I think I remember reading that the capacitance changed the time
of decay, so really configurable electroporators basically just have
lots of different sized capacitors.

With some thought and dedication, I think we could build an Open Electroporator.

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[Cathal Garvey]

On Tue, Nov 08, 2011 at 06:02:02AM -0500, Nathan McCorkle wrote:
> Why not use electroporation for transfection... I've got this book:
> http://pages.towson.edu/jsaunder/Saunders%20Publications/48.Guide%20to%20Electroporation%20and%20Electrofusion.pdf

I'm not sure that electroporation works well for plants, but I'd totally get behind an open electroporator anyway. For bacterial protocols it'd be amazing.

I'd see this working as a computer plug-in device like OpenPCR, such that the computer can display practical protocol instructions while the user pulses the cells. So, you'd get instructions like an automatic defibrillator; "Cool cells to 4C. Press Forward to continue." -> "Charging. Prepare recovery medium to add post-pulse. Press forward to pulse." -> "Cells pulsed. Add 200uL recovery medium." -> "Incubate cells"... etc. etc.

Because the equipment would be pretty standardised you could easily share protocols. That's not quite so possible with the sorts of crappy equipment people actually use in academic labs in my experience; appart from a few secret-inner-workings devices, the devices I've seen lack standardisation, so it can be hard to mimic someone else's protocol, and ultimately you can't know whether your device generates similar waveforms without splurging money on a waveform analyser. Also, the interfaces tend to be universally awful; the one we used to use in my prior workplace was all dials and click-counters, like a babbage difference engine with several kilovolts of power behind it.

Still, I'll carry on at chemical techniques; the more species you can transform easily with household or pharmacy-bought chemicals, the lower the barrier to entry for n00bs. I was delighted to find PEG3350-as-Laxative for that reason; PEG is essential to transforming Yeast, Plants and E.coli easily.

[Nathan McCorkle]

What would a good plant for DIY genetic manipulation be? If any of
these techniques are species dependent, we'd better choose a plant
platform that everyone likes. Would this be Arabidopsis or something
else for us? Do other plants have a more amenable stage in their life
cycle for transformation, or maybe a plant that recovers quickly from
a protoplast (or single cell equivalent, not sure what the plant
equivalent of trypsin is)

http://www.wiziq.com/tutorial/41707-Micropropagation-through-Plant-tissue-culture-Detailed-methodolog
http://csm.jmu.edu/biology/renfromh/pop/pctc/tcstart.htm
http://www.wiziq.com/tutorials/plant-tissue-culture
http://aggie-horticulture.tamu.edu/syllabi/689plantbreeding/Assigned%20Articles/Tissue%20Culture%20Applications/Crop%20improvement%20through%20tissue%20culture.pdf

this one is a really good FAQ from Harvard:
http://molbio.mgh.harvard.edu/sheenweb/faq.html

and others from tamu.edu:
http://tinyurl.com/83mqzc7

[Cathal Garvey]

Well, the two standards are Arabidopsis (grows quickly, convenient size,
studied intensively) and Tobacco (really easy to engineer).

Both are pretty useless to me: neither are useful and neither is
perennial. Also, neither is a tree. :)

The two standard methods, protoplast chemical
transfection/electroporation and biolistics, seem to cover most species,
but neither seems to cover the lot.

The equivalent of trypsin, which I interpret to mean "the enzyme that
makes protoplasts by digesting the cell wall", would be a blend of
cellulase and pectinase I think. That tends to give you nice
protoplasts, but you have to carefully buffer them with sugars or salts
to prevent lysis (cell explosion).

Once you have protoplasts, it's pretty easy to engineer compatible
species with PEG and a salt, but you can't leave them too long in the
PEG or they tend to lyse. So, not as easy as E.coli transfection, but
somewhere on the same scale certainly.

Actually, one of the big barriers to genetic manipulation isn't gene
delivery, but stable gene integration and expression. Plants don't
support episomal DNA so chromosomal integration is a must, and plants
have defences against viral DNA that tend to shut down "new" DNA at
unpredictable intervals after gene delivery. So you could select for
gene X, and get a lovely glowing plant or whatever, only to find that
Gene X is rendered non-operational within a generation and stays that way.

One way to avoid this is to avoid bacterial CpG methylation; special
strains of E.coli can be used to prepare plasmid DNA that don't use Dam
or Dcm, but those strains are unstable and grow awfully slowly because
of the Dam- phenotype. Another way is to use PCR, which will give
beautifully unmethylated DNA.. although that in itself might be
suspicious to a plant, for all I know.

Of course, avoiding Dam Methylation sites when synthesising your DNA
would be a good thing to try also, in addition to optimising the DNA to
"resemble" plant DNA as closely as you can.

--

[Nathan McCorkle]

On Tue, Nov 8, 2011 at 8:24 AM, Cathal Garvey <cathal...@gmail.com> wrote:
> Well, the two standards are Arabidopsis (grows quickly, convenient size,
> studied intensively) and Tobacco (really easy to engineer).

Sure, a feature some folks have talked about is having something that
could grow into a house or something large... poplar is fast growing
and a target of genetic investigation... maybe pumpkins would also be
interesting (a la Cinderella's giant pumpkin carriage)

--

[Cathal Garvey]

Not sure you could engineer pumpkins to get any larger than they already
do with careful cultivation, because as annuals they only have so long
to make root structures. Perhaps you could perform some cunning
engineering that leads each generation to make larger, better nourished
seeds; that way, your first generation would be a normal pumpkin, but
the next generation would grow far faster and have established roots and
leaves earlier - giving it more time to make a nice fat pumpkin. The
year after that, perhaps even moreso.

A gourd-house consisting of vastly enlarged pumpkins; the biotech
poster-boy that never was.

On 08/11/11 13:45, Nathan McCorkle wrote:
> On Tue, Nov 8, 2011 at 8:24 AM, Cathal Garvey <cathal...@gmail.com> wrote:
>> Well, the two standards are Arabidopsis (grows quickly, convenient size,
>> studied intensively) and Tobacco (really easy to engineer).
>
> Sure, a feature some folks have talked about is having something that
> could grow into a house or something large... poplar is fast growing
> and a target of genetic investigation... maybe pumpkins would also be
> interesting (a la Cinderella's giant pumpkin carriage)

--

[jlund256]

While the gene gun can be a 'gun', it is typically a potato gun--a
pulse of high pressure gas propels the particles.
Here is an article that describes the tech:

http://genegunbarrels.com/pdf/review_on_gene_gun_technology.pdf

[Idan Efroni]

I'm not sure the root structure is limiting for pumpkin size. And even
if it was, you can still, theoretically, make the pumpkin larger in
size while keeping the same biomass. However the genetic of fruit size
is quite complicated, as it is a systemic trait with lots of feedback
loops which we know (almost) nothing about. In such cases, traditional
"blind" breeding is much more efficient than engineering.

On Nov 8, 8:56 am, Cathal Garvey <cathalgar...@gmail.com> wrote:
> Not sure you could engineer pumpkins to get any larger than they already
> do with careful cultivation, because as annuals they only have so long
> to make root structures. Perhaps you could perform some cunning
> engineering that leads each generation to make larger, better nourished
> seeds; that way, your first generation would be a normal pumpkin, but
> the next generation would grow far faster and have established roots and
> leaves earlier - giving it more time to make a nice fat pumpkin. The
> year after that, perhaps even moreso.
>
> A gourd-house consisting of vastly enlarged pumpkins; the biotech
> poster-boy that never was.
>
> On 08/11/11 13:45, Nathan McCorkle wrote:
>

[Idan Efroni]

The first thing you need to know that it has been done, at least in
tobbaco.
See here: http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0015461
(open access)

The basic idea was somewhat as outlined here - get the lux operon, add
flanking regions of chloroplast DNA, shoot the DNA in, and select for
positives. While one can think of different ways to do it, this is
certainly the simplest. It does require a) a gun, b) the sequence of
the plant chloroplast DNA.

Given this, I don't see a reason why this shouldn't work for any other
plant. As you may have already understood, transforming plants is
somewhat of an art, and if you want to venture away from the commonly
used ones, there is going to be a lot of trial and error. The
particular hormones and conditions of regeneration can vary to a large
extent, and some just won't regenerate (God knows I tried).

Lichens is an interesting but not so well investigated system.
Assuming it is a cyano-fungi (and not algea-fungi) lichen, it might
work. You will probably need a plasmid which is compatible with cyano,
as the e.coli ones are unlikely to work. There are plasmids out there,
but this means sub-cloning. Selection is bound be pretty hard on such
a slow growing organism, though.

How about green algea? There is bound to be some chlamydomonas
swimming happily in a pond in a park near you. They should be PEG
transformable. My guess is that the e.coli plasmid by itself wouldn't
replicate well in the chlamy chloroplast, so your best bet is to aim
for chromosome integration as in the plos one paper.

Idan

[Mega]

Do you think they will send me some seeds of that plant ;) ?
No, honestly, will they?

On 13 Nov., 03:55, Idan Efroni <ida...@gmail.com> wrote:
> The first thing you need to know that it has been done, at least in
> tobbaco.

> See here:http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.00...

[Cathal Garvey]

Not a hope in hell, I'm afraid!

That said, it's awesome to see that it worked. And that's with (AFAIK
after a quick skim of the paper) with a basically unoptimised operon;
you could seriously improve that bioluminescence with a little work. As
I mentioned previously, there's lots of work out there in improving
light production by de-coupling transcription of LuxAB from the rest,
and you could probably gain another factorial leap in brightness if you
somehow upregulated fatty acid synthesis in the target cells/organelles.

[Idan Efroni]

They can't, even if they want to. GM plants are regulated and you
can't just distribute them (that said, GM plants are being
distributed, but "under the radar" so to speak). Before you become too
disappointed, mind you that the intensity was very weak - 5 minute
exposure at 3200 ISO is quite dim. your eyes are extremely sensitive
so you would be able to barely see it in total darkness, but that's
it.
However, if you happen to live near NY, you can to write them an email
and try and visit the lab. Many times (but not always), scientists
will be happy to give you a tour and answer some questions.

Cathal has some interesting ideas for increasing the intensity, but
that would require multiple transformations - requiring either
multiple selection markers, or sequential transformation with excision
of the selection marker. Not a beginner project, for sure.

But you should think about algea. Even sea algea like Ulva can be
transformed.

[Nathan McCorkle]

The university of the first author is about an hour away from me... I'll try to get in contact

Sent from my mobile Android device, please excuse any typographical errors.

[Cathal Garvey]

> But you should think about algea. Even sea algea like Ulva can be
> transformed.

Any protocols out there for regenerating Volvox from single-cells?
Because apparently volvox can be seen with the naked eye, just about. So
if you transformed Volvox and regenerated to spheroid colonies, you'd
have a little vial of glowing "motes", which would be pretty attractive
to look at. :)

[Nathan McCorkle]

On Tue, Nov 15, 2011 at 5:52 AM, Cathal Garvey <cathal...@gmail.com> wrote:
>
>> But you should think about algea. Even sea algea like Ulva can be
>> transformed.
>
> Any protocols out there for regenerating Volvox from single-cells?
> Because apparently volvox can be seen with the naked eye, just about. So
> if you transformed Volvox and regenerated to spheroid colonies, you'd
> have a little vial of glowing "motes", which would be pretty attractive
> to look at. :)

hmm I like this idea

Nuclear transformation of Volvox carted
http://www.pnas.org/content/91/11/5080.full.pdf

Seems they're photosensitive, could cause problems...:
http://phgoods.info/content/11/8/1473.short

I've attached the full article (INFLUENCE OF THE CELL WALL ON
INTRACELLULAR DELIVERY TO ALGAL CELLS BY ELECTROPORATION AND
SONICATION)
Using C reinhardtii:
"
The primary objective of this study was to test the hypothesis that
electroporation primarily transports molecules across cell membranes,
because its mechanism is known to be specific to lipid bilayer
disruption ( [Jaroszeski et al 2000] and [Weaver and Chizmadzhev 1996]
), whereas sonication transports molecules across both cell membranes
and cell walls, because it nonspecifically disrupts cell-surface
barriers. The data from this study generally support this hypothesis.
The presence of a cell wall significantly reduced intracellular
delivery of BSA during electroporation, but generally did not reduce
BSA uptake during sonication.
"

>
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[Idan Efroni]

> Any protocols out there for regenerating Volvox from single-cells?

Not that I'm aware of, but could be. The paper Nathan has they
transform embryos.

[Mega]

Hey, what about that:

Scientists found out Gold nanoparticles make plants glow using a
mechanism I yet have to find on articles ion the web.

Copper has the same chemical property as gold becaus it's in the same
'row' as gold. May that be a viable row? To use copper nanoparticles?

[Simon Quellen Field]

I think you mean the same column as gold.

So is silver.

But both will oxidize quickly as nanoparticles, unlike gold.

-----

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[Cathal Garvey]

Also, it only makes the plants fluorescent, not bioluminescent. i.e. you
still need to install a UV lamp to "power" them, which defeats the point
entirely. Looks cool, but really just an expensive gimmick.

Besides, some plants are already fluorescent; loose chlorophyll
fluoresces orange under UV, although with an intact photosynthetic
cascade the energy is handed off to other molecules before it can
fluoresce. Some plants decouple this handover mechanism at times, and
they'd probably fluoresce when this occurs.

Alternatively, mashing up the leaves a bit would achieve the same
result; try shining a UV light on some good green pesto to see this in
effect. The olive oil and basil leaves render good pesto a bit
fluorescent orange.

[Mega]

Hey, my newest project is to make e. coli glow with a plasmide called
pVIB.

This plasmid has the complete lux operon and also a ampicilline
ressistance. But now I got some concerns about the amp-r

Couldn't it be that some pathogen bacteria may take the plasmid from
e.coli and get ressistant?

So if one gets ill he maybe cannot be cured with ampicilline and
related substances. Is that true? Maybe kanamycin would be better, but
I don't find a plasmid with it.

[Cathal Garvey]

Changing the subject because this had nothing to do with the old
bioluminescence discussion.

Mega: Totally with you on this one. While antibiotic resistance isn't
likely to be an issue with genetically modified plants or animals
(incompatibility with microbes in that encoding format), with bacteria
you can rest assured that they will share DNA when it is convenient.

For that reason, I generally advocate sterilising your cultures *after*
use, so that their contents (DNA etc.) are already starting to degrade
even before disposal. Taking this further, I'm trying to work on
platforms that don't require antibiotics, or at least platforms that use
medically insignificant, legally available antibiotics.

I'm sure it will be said by others so I'll say it now (although the
actions of others are no excuse for sloppiness on our part): The
greatest source of antibiotic resistance is stupid use of antibiotics by
livestock farmers and factory farms. The second greatest source of
resistance is patients abusing antibiotics.

On 19/12/11 17:33, Mega wrote:
> Hey, my newest project is to make e. coli glow with a plasmide called
> pVIB.
>
> This plasmid has the complete lux operon and also a ampicilline
> ressistance. But now I got some concerns about the amp-r
>
> Couldn't it be that some pathogen bacteria may take the plasmid from
> e.coli and get ressistant?
>
> So if one gets ill he maybe cannot be cured with ampicilline and
> related substances. Is that true? Maybe kanamycin would be better, but
> I don't find a plasmid with it.

--

[Mega]

""I'm sure it will be said by others so I'll say it now (although the
actions of others are no excuse for sloppiness on our part): The
greatest source of antibiotic resistance is stupid use of antibiotics
by
livestock farmers and factory farms. The second greatest source of
resistance is patients abusing antibiotics. ""

I agree 100% on that... Well my fear was that using ampecillin was
totally ethically incorrect from my point of view.... But as you said,
farmers are abusing them in huge amounts - so if I sterilize
everything, that shouldn't affect mother nature so much...

Do you know whether farmers use ampicillin?

Kanamycin shall be the best antibiotic for transformation: ressistance
is common in bacteria. So when they conjugate and share ressistance
plasmids - that could happen also in nature quite likely. Ampicillin
on the other hand is used as a broad-range (german word translation,
correct i hope) antibioticum.

As my first project i want to make something very impressive, yet non-
pathogen(!) and (in comparison) easy. pVIB seems to be easy because
that is a readily prepared plasmid.
I 'just' have to use a transformation methode and see the results. Of
course, if in pVIB there was kan-r instead of amp-ressistance, it
would be a lot better (in case they escape into the environment, there
would be no danger to anyone)

[Mega]

Ok,

I got another question:

After transformation of the bactieria I have to plate them on lb +
agar + ampicilline.

How much ampicilline do I have to get for one petri dish??

Thx

[Nathan McCorkle]

the table here lists the working concentration of ampicillin to
commonly be 100ug/mL, kanamycin @ 50ug/mL:
http://www.qiagen.com/plasmid/bacterialcultures.aspx#tab3

so you'd mix up a stock solution of ampicillin (they recommend
50mg/mL)... so in 50mL of distilled, sterile water, add 2.5 g
ampicillin

if you make 250mL of LB agar solution, after its been sterilized and
allowed to cool to less than 55 degrees C, you'd add (250mL*100ug/ml)
25mg ampicillin. Since your stock solution was 50mg/mL, you'd add
0.5mL of this stock solution to the media

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[Mega]

http://www.livescience.com/10307-transplanted-genes-plants-light.html

The company wanted to sell autoluminescent plants... But google has no
good results for 'Bioglow inc"
Else it maybe would have been possible to make contact and by a
prototype

On 14 Jan., 21:13, Nathan McCorkle <nmz...@gmail.com> wrote:
> the table here lists the working concentration of ampicillin to
> commonly be 100ug/mL, kanamycin @ 50ug/mL:http://www.qiagen.com/plasmid/bacterialcultures.aspx#tab3
>
> so you'd mix up a stock solution of ampicillin (they recommend
> 50mg/mL)... so in 50mL of distilled, sterile water, add 2.5 g
> ampicillin
>
> if you make 250mL of LB agar solution, after its been sterilized and
> allowed to cool to less than 55 degrees C, you'd add (250mL*100ug/ml)
> 25mg ampicillin. Since your stock solution was 50mg/mL, you'd add
> 0.5mL of this stock solution to the media
>
>
>
>
>
>
>
>
>
> On Sat, Jan 14, 2012 at 12:07 PM, Mega <masterstorm...@gmail.com> wrote:
>
> > Ok,
>
> > I got another question:
>
> > After transformation of the bactieria I have to plate them on lb +
> > agar + ampicilline.
>
> > How much ampicilline do I have to get for one petri dish??
>
> > Thx
>
> > --
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