The ODIN Needs Your Help - T4 DNA Ligase Testing - Free Ligase

[Josiah Zayner]

My company, The ODIN(http://www.the-odin.com) is starting to ship restriction enzymes and ligase! However, enzymes are pretty heat stable. T4 DNA ligase is another matter, it requires ATP, ATP hydrolyzes. I need a few people to test out some T4 DNA ligase and reaction buffer that has been shipped. Shipping of Taq and other Master Mixes has worked and they contain dNTPs so I assume this should work out just fine but I need testers to make sure.

If you goto the website(http://www.the-odin.com/t4-dna-ligase-5ul/) and use coupon code FREETHELIGASE I will ship you some ligase for free as long as you promise to provide feedback on how it works after shipping.

Thanks,
     Josiah Zayner
     The ODIN
     ca...@the-odin.com

[Koeng]

How about the raw ligase? I buy ATP directly from NEB to do goldengates with, and some ligase from a non-commercial source might be interesting. 

Also, how are you getting the ligase? Expression of it in house? Or resale? Either or, I'm sure it'll be a great offer, but I am curious if you had the vector in house for some in vivo ligations I'd like to test out. Anyway, useful site! It's awesome that you've gotten more enzymes up, but I think BsaI/BsmBI are also a real important couple. Once you get materials to do GoldenGate, it's a pretty wonderful system, just wish I could get around to making some basic parts for DIY vectors...

-Koeng

[Josiah Zayner]

Yeah it is just a resale. Most people don't have ATP I imagine so I want to make sure it works as is, ya' know?

I will look at BsaI BsmbI but they are less used enzymes so they are freaking expensive.

You can do like 10 times the cloning with NcoI/EcoRI/HindIII on price maybe even more.

Same thing with Biobricks enzymes. I am selling those for iGEM folks but they are expensive. Someone should go back and start a cloning system that uses EcoRI and HindIII, the two least expensive enzymes.

Also, Koeng, you forgot to use the coupon code so I refunded you shipping!

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[Koeng]

True true, EcoRI and HindIII are favorites of mine because of how much they supply. However, they are 6bp cutters... So I would actually argue that NotI and SbfI would be better for a new cloning system! I guess it depends on how much it would cost to remove the sites vs the actual enzyme cost.

I've noticed that at least at an academic lab people always use the same volume of enzyme... Like 1µl is 20 units of EcoRI but just 5 units of BtgZI... anyway it's a pretty wasteful approach, so if anyone is reading this is using restriction enzymes, go by units.

Then again, I haven't even cloned anything with classic restriction enzymes in a year or so because gibson is so good. However, gibson can cost an arm and a leg if you buy it directly from NEB (price for quality, their stuff gives me ~5-10xs better efficiency than our normal stuff). I know that SLiCE ( http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3333860/ http://www.ncbi.nlm.nih.gov/pubmed/24395368 ) would probably be very cheap to make in a DIY lab if created in mass, which might be useful for you. I'm curious what the efficiency is, but if it's good, I'd be willing to switch my home experiments over if it meant I wouldn't have to buy mixs anymore.

-Koeng

(btw: Thanks! I knew I forgot something)

[Bryan Jones]

Koeng, SLiCE sounds interesting. Do you know where to get your hands on the PPY E.coli strain that they use? Is it sold anywhere (a quick google search didn't turn up anything)? I guess I could try to contact the authors or those papers. I'd like to give it a try in the academic lab I'm working in. If it works as well as they claim it might be a good alternative to both traditional restriction enzyme/ligase cloning and Gibson Assembly, especially for the DIYbiologist on a tight budget.

[Koeng]

http://www.sciencedirect.com/science/article/pii/S2405580815000850

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4453199/pdf/12896_2015_Article_162.pdf

 It appears that JM109 can do SLiCE just fine. Not sure about comparisons of efficiency thou

-Koeng

[Koeng]

http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0137466 

May be interesting

[BraveScience]

Same here. Ligation dependente cloning using biobricks sucks, given the volumes and the amount of steps you have to necessarily go through, doesn't turn out to be that efficient. 

I got interested as well in Gibson. Indeed price\quality of commercial kit vs homebrewed one is a tough one.

Anyway I stumbled across the BASIC system. Yes, you need a restriction enzyme (just one, BsaI) and work through ligation. But you can shuffle your parts around as you wish.

http://pubs.acs.org/doi/abs/10.1021/sb500356d

It's a one-pot reaction.

Fede

[BraveScience]

Koeng, i've tried a similar method to the one mentioned in the paper and it didn't work for me.

Yet i still believe it should be possible.

[Cathal (Phone)]

I first read "litigation dependent cloning" and had visions of a horrible dystopia.

--
Sent from my Android device with K-9 Mail. Please excuse my brevity.

[Koeng]

This reminds me of a goldengate PaperClip assembly. Very interesting however, once I get an opportunity I will read the entire paper! 

You tried the DH5a method or the SLiCE method?

-Koeng

[Koeng]

Have you attempted this cloning in conjunction with biobricks? It doesn't appear that it requires their standard but rather restriction enzyme sites on both sides. 

-Koeng

On Monday, October 19, 2015 at 3:42:46 AM UTC-7, BraveScience wrote:

[Koeng]

I read the paper, and if anyone's interested here is essentially the idea:

You take a gene with BsaI sites flanking it. Still gotta clone that plasmid, which honestly in my opinion destroys the point of leaving goldengate, but hypothetically you could use any plasmid with restriction enzyme sites. You then make 2 ssDNAs that anneal to each other and can ligate to the 4bp overhang of your part. 

         -------|---PART-----|--------------

-------------|-----PART---|------

One of the primers is small, while the other is large. The large one creates the bolded overhangs. These 21bp overhangs can be annealed to each other in an orthogonal fashion, like a goldengate reaction with long overhangs (and no ligase). That nicked product is then transformed into E coli, where in vivo the nicks are filled. 

Steps:

1-Digestion/ligation (of the specific overhangs)

2-DNA purification (gel?)

3-Assembly by annealing

Honestly, I find this to be an interesting way to clone, but I don't know about pragmatic implementation. If digestion/ligation could be combined with phosphorylation of the primers, I can see this working out to be quicker than my gibsons right now. However, it is clearly just about as efficient as gibson. Gibson also doesn't require a part vector, which is a huge plus. Their methylation stuff is cool, and honestly really cool when I think about it (it eliminates the need for BsmBI in goldengate!!! Essentially, to add the BsaI site to the new vector, they use primers which they have modified to be resistant to BsaI cutting by methylating a cytosine). 

However, it is clearly not as efficient as GoldenGate, and requires more setup than gibson. But it does win both in modularity. 

-Koeng

On Monday, October 19, 2015 at 3:42:46 AM UTC-7, BraveScience wrote:

[Brian Degger]

Cool ideas.

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[BraveScience]

Hi,

Yes, I've tried a micro-homology based approach but I must say that was a very rough try, with a gene potentially toxic, using home made chemical competent cells.
Plasmid itself was quite big, around 7kb.

I like BASIC concept, even though I'm an advocate of PCR based approach, but recently both PCRs and biobrick based cloning completely sucked. Moreover as we are looking into implementing an automated cloning system in our lab it makes total sense to shift toward an automatable system.

My feeling is that biobrick system is very expensive and, the ligation step, isn't that reliable or standardized as you use different blunt/sticky ends.
I'm pushing to use commercial cells as well.
Basically time you waste and number of try increase your end price incredibly.
Even if you use home made stuff.

So far that's what I see:
- multiple assembly in one-pot reaction
- less restriction enzymes
- avoid completely gel purification (different marker at level 0)
- highly plastic to permutation and combinations
- change adaptor when you need
- fast and cheap
- automatable

I believe it will be a pain to set everything up but it's definitely appealing.

Potentially this system has an advantage also for the diybio community.

Fede

[Edoardo Gianni]

Hello there,

I recently tried the SLiCE assembly at the London Biohackspace for our iGEM project and had quite good results with it. We put together some of the results on openwetware http://openwetware.org/wiki/SPLiCE and tested a version of the reaction with PEG8000, which seems to speed up the reaction (just like in the quick ligase buffer). We transformed the cells with pKD46 (which contains the lambda recombineering genes) instead of using the PPY strain and it seemed to work just fine. 

I am now trying to find the time to get a better idea of the efficiency as well as clone a biobrick version of the pKD46 as we synthesised the gam-bet-exo genes as gblocks. Feel free to add to the page if you have some comments :)

Edo

[Josiah Zayner]

That's an interesting idea.

A few questions:

Did you do a DpnI digest to remove the RFP pSB1K3 plasmid after the PCR or a gel purification of your PCR product?

Did you do any negative controls?

Have you sequenced or verified that it is in fact the pSB1C3 with an RFP insert?

Efficiency seems awfully high which makes me wonder if what you are seeing is just the pSB1K3 or something else.

Also, the need for electrocompetent cells, an electroporator, multiple sets of primers and a commercial lysis buffer kind of removes any sort of money savings. In fact, it makes this method much more expensive then a standard cloning reaction.

JZ

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[Edoardo Gianni]

Hi Josiah, good points. I had similar worries after the first good results.

I did DpnI treatment but no gel purification, that would be ideal especially if you have the same selection barker.

I did transform the original RFP in pSB1K3 and plated on chloramphenicol, got no colonies.

No sequence verification but we sent another part assembled this way to iGEM, which should sequence it soon. It would be interesting to see if you get some errors at the recombination site.

Efficiency is high as I am using quite a lot of DNA (100ng of vector in the ligation, 30ng used for transforming), plating all the cells from the electroporation and using highly competent electrocompetent cells. I would expect far fewer colonies in an assembly with less DNA and with chemically competent cells, I would assume a similar efficiency to a digestion/ligation - definitely not the efficiency of a Gibson superoptimised kit.

The method doesn't need electrocompetent cells, it's just that we got an electroporator donated and it works well for us. In addition we were working on an automated competent cells maker, which would simplify the process of making the cell. Regarding the primers, for a two part assembly the normal biobrick prefix and suffix are long enough to allow homologous recombination with the prefix and suffix in pSB1C3. For multiple part assembly the primers would have similar costs to a gibson assembly, although you could think of some standard ends to simplify the process. 

I didn't think about the lysis buffer, I'd like to try DIY alternatives though (i.e. soap? french press? glass beads?). On the other hand you only need to make the lysate once and it lasts for hundreds/thousands of reactions. The other part I thought would be expensive is the reaction buffer, but it comes at about 20$ for 6ml from NEB (it's the same one you are trying to test for the T4 ligase).

Btw regarding the T4 ligase the enzyme is what would worry me, more than the ATP. My understanding is it's not as stable as taq/restriction enzymes, but the london biohackers are happy to betatest if you ship overseas :)

Edo

[Josiah Zayner]

Most commercial ligase documentation suggests that the biggest issue is ATP and DTT degradation in the buffer from temp.

I know T4 ligase activity is temperature sensitive but do you know of any work on its temperature stability? I couldn't find much. Most commercial documentation suggests heating at greater than 65C for 10+ minutes to inactivate the enzyme, though this could just be a generic number I assumed it was empirical(if you look at restriction enzymes they often have different temperatures for heat inactivation.

Does iGEM sequence parts? I thought they only sequence things that they send out in their parts distribution?

What kind of cells did you use for the transformations of the recombination reactions? for the pKD46? Did you grow the pKD46 overnight at < 30C?

You might want to go a little more in depth on your webpage as most of this information is not on there. Also, showing pictures of the negative control plates, i.e. transformation without extract, transformation without insert, transformation without vector would be reasonable info to provide.

All of these things would make your protocol alot more repeatable and trustworthy.

It is almost $30 to ship overseas so I don't think I can pay that.

Thanks,

     Josiah

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[Josiah Zayner]

Oh yeah also, concentration of Cm in the plates and anything else I forgot that would be helpful.

[BraveScience]

I've never tried SliCE but I heard reports of increased mutation rate. Is that possible?

As in the article shared by koeng, cloning should be simple, DH5 alpha should have enough recombinogenic activity/ligase to splice your fragments, if used for the production of the extract.

Second question: can you make this extracts and sell/ship them?

Fede

[Edoardo Gianni]

It's just legacy information from my previous lab "keep ligase the least possible outside of the freezer as it looses activity more than the others." 

iGEM started sequencing everything you send them after 2012 I believe. 

DH10b for all the work, and yes O/N @30C. Thanks for all the other tips, I'll try to implement them as soon as I can. All the work happened short to the iGEM deadline and is not complete. I'd like to test the method further and I'll update the openwetware page as I do.

Don't worry about the ligase and keep us updated on the stability :)

@fede in the SLiCE paper they suggest it's stable at -20 for months but no idea about shipping it. I would guess that with all the other crap in the lysate that can interact it may loose efficiency, but it's worth trying - I am happy to send some if you want to test it.

Edo

[Koeng]

http://www.sciencedirect.com/science/article/pii/S2405580815000850

Use a triton x100 and Tris-HCl buffer. It's cheap and extremely easy to make (Made some yesterday, going to make my own SLiCE friday)

Interesting is that the difference is that the cells grow 3x more concentrated in the JM109 method. May have to do with DNA repair genes getting turned on during late growth.

-Koeng

[Bryan Jones]

I tried SLiCE last week with pretty good results. My goal was to minimize cost overall. To this end, I used regular DH5a without labmda red, lysed with homemade Triton X100 buffer as Koeng suggested, and transformed into chemically competent cells.

I cloned a PCR product (30ng) into a linear vector (120ng). Molar ratio of ~2:1. The pieces had ~40 base overlap. 
As a control I ran a Gibson Assembly (well, NEBuilder, which is basically a tweaked version).

I transformed with 45ng of DNA, or 30% of the reaction (22.5/15% for the NEBuilder control).

The efficiency of the SLiCE was significantly less than NEBuilder. I got 57 colonies from NEBuilder and 14 from SLiCE.

I'll sequence a few of the colonies to make sure I got what I wanted.

All things considered, I think the reaction efficiency was pretty good (>10% of the efficiency of Gibson), and could probably quite a bit better with the addition of lambda red and a bit of optimization.

[BraveScience]

@Edo thanks, but I think I can make by own. I'll give it a try one of these days in the university lab and then try to replicate the results at the openwetlab @WaagSociety

[Edoardo Gianni]

Hi Bryan,

Awesome job. I'll try the Triton X100 the next time I need to make some lysate. If you wanna add more data to the SLiCE page on openwetware feel free to edit it, I guess the more the protocol is robust the more other people can use it. 

Btw I thought this paper might be of interest, it definitely minimises costs the most, literally "aquacloning" http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0137652

Edo 

[Koeng]

I tried out my homemade SLiCE mix. I put 100ng of both insert and vector in for a 2 piece assembly, for 30 minutes. I compared it to some homemade Gibson mix.

On my plates, I got 13 colonies on the SLiCE plate and 42 colonies on the Gibson. I am extremely impressed with 1/3 considering just the simple strain and simple lysis procedure.

-Koeng

[Josiah Zayner]

This is pretty cool. If people try and repeat these experiments can they post detailed details(heh) of what they did so others can follow suite.

Also, it doesn't seem anyone is running the control of everything but the SLiCE mixture. If you are running an experiment can you do that and let us know how that goes?

Thanks,

     Josiah

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[Koeng]

Yea, my apologies for the non-controlled experiment, I didn't set it up to be very controlled because I didn't think I'd get that good of efficiency. I will be running a few more in the next few weeks so I will post the results. 

Unfortunately enough, it appears that SLiCE is patented ( http://www.google.com/patents/US20130045508 ), and the company that owns the patent charges an arm and a leg ( http://slicecloning.com/SLiCE.html , more than HiFi mix per reaction). I guess the ODIN can't sell the mix, but possibly sell a kit to make it yourself.

-Koeng

[Bryan Jones]

I'm not an expert on reading patents, but it looks like the patent covers the method of the cloning procedure, not the production of SLiCE extract, nor the composition matter of SLiCE extract. Selling the SLiCE mix might not actually violate the patent (that'd be up to the user to do or not do). It also looks like the the patent only covers assembly of fragments that have "20 bp to 52 bp that are homologous". So using 19bp or 53bp homologous regions might be a way around the patent. Maybe a bit less efficient, but it looks like that would still work.

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[Josiah Zayner]

Yeah, don't know if I am interested in selling a SLiCE mix because I still don't know how efficient it is in a DIY setting. That is why I am interested in the results you all are having. And what you all are using, i.e. homemade competent cells vs. high efficiency competent cells, &c. Is just some blunt end ligation occurring or is it an actually legit Gibson like assembly? I will test it out in a few weeks.

Patents don't really hold me back from biology stuff, maybe they should? They say the patent applies to every RecA- bacterial strain but have only tested 5-10? The patent seems flimsy with so many ways around it. Just don't list the bacteria I use? or Use a bacterial strain isolated from my skin that 16s is not an identical match for any known bacteria. They seem like a small company that probably doesn't have enough money to sue and would probably gain very little in damages. But who knows.

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[Koeng]

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4453199/ It makes a lot of sense that these people recommend 19 base pairs. 

It's also definitely not just blunt end ligation, you can check out the data just in that paper, where they did control for that. I agree that I would need more controls to get decent results, I simply didn't think that it'd work this well. I'll update once I get those results (in the next few weeks I am definitely going to do more controlled experiments).

-Koeng