A hypothetical protocol for DIYBIO DNA synthesis

[CodeWarrior]

As some of you may know a means of synthesising DNA that is accessible to the ambitious and well informed amateur has been a long term intellectual pursuit of mine. A kind of theoretical moby dick. Here I'd like to present something like a paper (for a certain value of paper) containing my latest thoughts. If these ideas pan out it would permit synthesis of DNA fragments potentially 100s of nucleotides long with out any pre made oligos (or at least only non gene specific ones), with out invoking potentially toxic and hard to obtain phosphoramidite based chemistry or any thing similar. A method based almost entirely on enzymatic processes. Of course there are a lot of ifs. However I'm not in a position at the moment to take the theory any further into actual experimentation. So I'm opening the idea up to the community in the hope that you can collectively make something of it. It's not really a better alternative for those of you who can already easily order DNA, it's not nearly as efficient as the phosphoramidite method not even in theory. It's a significantly more complex process. However for those of you who can't get synthesised DNA or who want to make your own as a project this may be of interest.

[Felipe Tanaami]

Veeery interesting! I have a printer acumulating dust here, because it's head is clogged. Now I have a good reason to try to clean it!
It's funny, I was researching exactly the same thing, I read that Cooper's igem team paper last week and I was thinking on a way to make long sequences with it. The microarray was a great idea! What do you think about the photolabile protected nucleosides? It can simplify the work a lot, I imagine. Keep us updated!

[CodeWarrior]

in my opinion the bottleneck in DNA synth is chemistry not engineering. Photo liable nucleotides mean you can dispense with the piezoelectric print head and use a projector system instead. but unless you can find somewhere to buy them or you can synthesise them easily yourself I'd say the chemical difficulty involved wouldn't justify the effort.

[Nathan McCorkle]

My first thoughts were that the reversible terminator nucleotides
(whether heat or photo or acid labile/activated) are still too
expensive and single-source for this to be interesting for the effort,
at least in a DIYbio setting. I think the solution of the problem is
at the seam between chemistry and engineering... I am not sure if that
is simply materials science or chemical engineering at that point, but
it sounds pretty close.

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[Felipe Tanaami]

Well, what if we protect the 3'-O enzymatically? I didn't study in depth, but it seens promising. There are some papers on the field, like this one:

http://sci-hub.cc/10.1007/978-3-7091-6310-8_4

(I've just saw the pictures =D) Particularly on the page 12, scheme 17. Scheme 20 seens good too, because it has regioselectivity. Forgive me if I am missing something...

[CodeWarrior]

Scheme 20 is nice it adds an acyl group. Rutgers 2014 igem project used an acyl group to protect dNTPs. They used ammonia to remove it. I believe ammonia does not require protecting groups on nucleotide bases in fact it's sometimes used to remove them. However scheme 20 only works for bases a and t. I wonder if you could modify the enzyme to work on c and g. Scheme 17 only works on a u base.

Well I've thought about this at length actually. Two posable strategies for alternative blocking groups have occurred to me. The first it may, just may, be posable to use a disaccharide synthase to add a sugar based compound to the 3 prime position of a dNTP. You can see scheme 31 in that paper shows a way of using a disaccharide synthase to add at the 5 prime position but there are a lot of disaccharides in nature and lots of different enzymes to make them. You'd have to screen a lot of disaccharide synthases till you found one with week activity for 3 prime addition then do directed evolution to get something that could give you dNTPs blocked by joining another monosaccharide to the ribose. To the best of my knowledge all disaccharides break down into monosaccharides in sufficiently hot water. So you could make your own thermal liable blocked dNTP. It won't be cheeper to do this than buy cleanamp.

The other option might be to use dirty ends. When dna breaks it doesn't always leave clean 3 prime ends. Often you get a lot of 3 prime ends with PO4H2 groups on them. You can't polymerise off this. But there is an enzyme PNKP that nips off the PO4H2 group so polymerisation can continue. If you used dirty dNTPs and used PNKP instead of heat to deblock that might be an option. However the only simple way I can think of to get dirty dNTPs is to get a DNA sample, subject it to lots of oxygen, possibly ozone. Use exonucleases to break it down into clean and dirty dNMPs painstakingly purify out the dirty dNMPs turn them into dirty dNTPs using NDP / NMP kinases then separate them by base. This will also not be cheeper or easier than buying cleanamp.

So the question in my mind is how hard would it be to make a dirty dNTP chemically with out an enzyme. Is there a reagent I could add to dNTPs and phosphoric acid that would attach PO4H2 to the 3 prime location with out the base or 5 prime end needing protecting groups. There are 4 OH groups hanging off the phosphors at the 5 prime end so can we expect some PO4H2 groups added here? What if I turned the phosphoric acid into a free radical before adding it using electrical discharge? All things I've considered. Basically I'm a sucky chemist and it takes a very talented chemist to do serious organic synthesis only using reagents you can purify out of things you can buy at your local supermarket or buy from amazon.

And again none of this is going to be cheeper or easier than cleanamp. But as I said a cheep solution is unrealistic this method is aimed at those who a) want a learning experience or b) can't get industry made DNA sequences.

[CodeWarrior]

And interesting question is what would happen if we mixed a short oligo say AAA and add TDT and P3O8H5. Could TDT add a phosphate with out an attached ribose. Could we modify it so it does? Could we get 5'-AAA-3'-PO4H2? If that works add an exonucliase and you've just converted about a 1/3 of A nucleotides to ones with 'dirty' ends.

[CodeWarrior]

My apologies I've said something stupid. That should be P3O10H5 not P3O8H5.

[Koeng]

I'm not a chemist by any standard, but I do occasionally purify my own enzymes. Is there any solution for mass production of the enzyme itself? Purifying the enzyme itself may be more work than the system itself (not thermophilic), and modified versions of the enzyme required would likely be patented, preventing a large scale distribution network (other than for the people who patented it, and they likely won't be too DIY friendly). I'm not qualified to ask about most of the chemistry, but this is my first impression of one of the difficulties.

-Koeng

[CodeWarrior]

I don't understand why the enzymes would need to be thermophilic except for the assembly PCR step which you can use off the shelf enzymes for. Personally the only enzyme purification I've done is with hist tags and nickel ion chromatographic columns. It's scalable in the sense that you can always get a bigger column or a larger broth tank for culturing bacteria.

[Koeng]

Just that thermophilic enzymes are easier to purify. 

Nickel columns are pretty easy as well, but the issue is is that the enzymes are likely patented, so you couldn't actively sell them. Of course, people could always purify it themselves, but that takes a lot of work. Even though I have his tagged most of the enzymes I commonly work with,  I don't purify them, because it's too much work. I fear people would think it's too much work to get their own synthesis enzymes as well.

-Koeng

[CodeWarrior]

I can't honestly say I've ever noticed that I've never tried to purify a thermophilic enzyme. I'm assuming you use heat to break down other enzymes and use some basic size based chromatography to seperate out from unfolded peptides etc?

So your consern is that if you needed a custom enzyme people wouldn't be bothered to do it? Perhaps. But you'd only need to do that if you couldn't buy cleanamp in which case you might not be able to buy lots of enzymes like tdt etc.

I wanted to put some thought into how people could make a dna synthesis technique work with minamal cooperation from the existing biotech industry. I don't know how the Americans feel but here in the uk the goverment and the established biotech industry doesn't seem to see much potential in the diybio movement but does see the potential for harm. The diybio movement has been compared to the coding movement that's gave birth to the raspberry pi. If so DNA would be the code and dna synthasis the compiler. Imagin if every raspberry pi came with no compiler. Instead you have to send your source code off to a company that might compile it, if they approve of it, for a charge. That situation would be unthinkable but it's more or less where we are with diybio.

So as I said I wanted to look into how you might make dna assuming virtually no cooperation from industry. But if you can order off the shelf enzymes I don't see why you wouldn't. Unless you were making enough for economies of scale to apply., I've been thinking about it and making it using bogstandard DNAse II and endonucliases might even work better.

[CodeWarrior]

As a means of making dirty dNTPs I mean

[CodeWarrior]

My apologies for perpetually spamming and bumping this thread. One more simplifying idea has occurred to me. Suppose the synthesised TdT enzyme had a long tail at one terminal which was then covalently bound to the glass slide along with the short oligo seeds pre DNA synthesis. No enzyme would then need to be deposed in each new cycle the existing TdT could be reused. Using this system and clearAmp less enzyme would be used and the cheeper four colour epson print heads could be used.

[Nathan McCorkle]

On Fri, Jul 15, 2016 at 10:20 AM, CodeWarrior <code.w...@gmail.com> wrote:
> My apologies for perpetually spamming and bumping this thread. One more
> simplifying idea has occurred to me. Suppose the synthesised TdT enzyme had
> a long tail at one terminal which was then covalently bound to the glass
> slide along with the short oligo seeds pre DNA synthesis. No enzyme would
> then need to be deposed in each new cycle the existing TdT could be reused.

I've considered using nanopores, or a matrix of them, like a
strainer... they're cheap and easily orderable too for about $32 each
(they even sent me a free sample), from places like:
http://www.temwindows.com/product_p/sn100-p20q05.htm

[CodeWarrior]

I'm curious how you would exploit the nanopore to achieve synthesis?

[Nathan McCorkle]

On Fri, Jul 15, 2016 at 3:47 PM, CodeWarrior <code.w...@gmail.com> wrote:
> I'm curious how you would exploit the nanopore to achieve synthesis?

oh, just a way to retain something like tDt or other enzyme for
repetitious reactions, instead of tethering it. A chunk of rock (the
silicon nitride with holes in it) seems like less variant/dynamic than
a coupling system in use (i.e. we don't have to do any covalent
chemistry)... plus it doesn't go bad sitting on the shelf like
biotin/streptavidin coupling kits might.

[Bryan Bishop]

(Don't mind me, I'm just looping in the https://groups.google.com/group/enzymaticsynthesis people.)

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[CodeWarrior]

lodging it in a pore wouldn’t work, it needs to move about to interact with the fairly short seed oligos you’d start with.

In an effort to improve the method by finding a way not dependent on special reagents like cleanamp dNTPs I’ve been looking at synthesis by sequencing type methods. Consequently I’d like to propose yet another protocol.

1. a oligo sequence we will call A is introduced to TdT and ordinary dNTPs

2. the resulting extended oligos are separated by length

3. the oligos are introduced to micro beads suitable for manipulation with optical tweezers covered with covalently bound (at the 5’ end) oligos of the A sequence and a second sequence we shall call B. By shear chance a small number of the oligos will have the B’ sequence (we use ‘ to indicate the complimentary sequence) and will anneal to a small fraction the beads.

4. beads will be separated into wells and subjected to PCR.

5. both A and B sequence oligos bound to fluorophores at the the 5’ end are introduced and used to mark those beads which are moved into a separate pool and washed clean on the tagging oligos.

6. a super thin layer of metal is evenly deposited on an optically flat side of a prism and A and B sequence oligos are covalently bound to the surface at the 5’ end in spall regular patches.

7. the separated micro beads are deposited on the surface one on each patch and PCR takes place after which the beads are discarded.

8. the surface is washed and placed in a surface resonance imaging frame and a reference reading of the surface is taken.

9. primer A (or B) is added and allowed to anneal, a second reference image is taken.

10. a PCR mix containing only one nucleotide (dA, dT, dC or dG) is added and extension takes place (but no heating occurs)

11. the surface is washed and dried and another reference reading is taken, the difference in the surface plasmon resonance effect between rounds should indicate how many nucleotides have been added.

12. 10 and 11 are repeated with a different nucleotide (dA, dT, dC or dG) in cycles until no more extension takes place.

13. for extra proof reading 10 to 12 can be repeated with the alternate primer (B or A) to read the complimentary sequence.

We now have a metal surface with many patches with bound oligo nucleotides of the sequences metal-5’-ANB’-3’ and metal-5’-BN’A’-3’ and we know what the sequence N is for each patch. if we have enough patches we likely have every short sequence posable. This is a reusable resource. this metal surface can be used to synthesis DNA sequences by transferring the sequence back to micro beads (coated in A and B oligos) placed on it by using PCR. These beads can be moved and brought together in sequence, treated with type type iis restriction endonuclease specific to the A or B sequences to release the bound sequence where necessary or leave only the sense sequence intact for annealing to other sequences for ligation or PCR. By these methods longer sequences could be assembled.

Of course this depends on the surface plasmon resonance being sensitive enough to detect a single added nucleotide  Since all SPR really cares about is the mass on the surface having lots of A and B primers initially bound on the metal should give a suitably strong SPR effect. ... I hope. I'm no SPR expert.

[Nathan McCorkle]

On Sat, Jul 16, 2016 at 10:21 AM, CodeWarrior <code.w...@gmail.com> wrote:
> lodging it in a pore wouldn’t work, it needs to move about to interact with
> the fairly short seed oligos you’d start with.

Right, I said nothing about lodging it.

>
> In an effort to improve the method by finding a way not dependent on special
> reagents like cleanamp dNTPs I’ve been looking at synthesis by sequencing
> type methods. Consequently I’d like to propose yet another protocol.
>
> 1. a oligo sequence we will call A is introduced to TdT and ordinary dNTPs
> 2. the resulting extended oligos are separated by length

My idea is essentially looping over those two steps, except you do it
recursively (feed output back into input), and stop at some N-number
of length and reserve it for a pool to do sterically-constrained
gibson synthesis (in a reaction chamber less than the polymer
persistence length, so to prevent secondary and tertiary structure
from screwing with ligation). If you command 10-additions with
sufficiently dilute nucleotide solution (dilute enough to
statistically say there is 1 nucleotide per volume introduced to the
reaction chamber), then just discard any molecules that don't meet
your target length (indicating an addition failed or there were more
nucleotides introduced than expected, during a reaction period). You
don't need sequencing either, since you control the input nucleotide.

[Nathan McCorkle]

On Jul 16, 2016 12:39 PM, "CodeWarrior" <code.w...@gmail.com> wrote:
>
> Ah that's workable for supper short >sequences but the yield will be low and >the needed input masive. Imagine you >tune your TdT reaction perfectly and get >a 30% yield of single nucliotide aditions

30% sounds far from perfect... wouldn't perfect be 100%? Also why do you assume 30% when phosphoramidite chemistry is close to 99% already, and we know enzymes work and don't have similar error issues, less stringent  chemistry (enzymes are all water based, phosphoramidite synthesis hates water).

> (which I expect is optimistic). Then let's >argue your seperation procedure is >100% lossless (highly unlikely). After 8 >rounds you'll have less than 0.007% of >the oligos you started with. You've also >had to do 8 seperation a probably 8 >consecutive PAGE procedures.

Oh, i am talking about nanofluidic channels, where you don't need gel or anything more than maybe counter-ion buffers.

>It's just not efficient to add 8 nucliotides >to a sequence. After that you need to >jam a primer on the end and do some >PCR or you'll be working with a sample >diluted out of existence.

Meh, just shove the at-least single molecule into an e.coli with electroporation... 'low' yield 'problem' solved. Seems a boon to me to be able to require at minimum a single molecule of output. Also, I am talking lab-on-a-chip tech, not garage-scale reaction apparatus.

[CodeWarrior]

sorry just to be clear your going to use a lab on a chip to perform TdT reactions, purify out single nucleotide addition oligos, transfect them into e coli, extract them from e coli and repeat the whole process several times? Ok I won't say this is impossible but to construct such a lab on a chip would be a good deal harder than even doing something as ambitious as building your own surface plasmon resonance imager. Also wouldn't it be far easier to use basic PCR than e coli to amplify the sequence.

actually I did a mathematical analysis based on modelling TdT reactions as a system of ODEs and the theoretical maximum efficiency is 100*e^-1% for one nucleotide additions which is approximately 36.7879%.

[Nathan McCorkle]

On Sun, Jul 17, 2016 at 5:15 AM, CodeWarrior <code.w...@gmail.com> wrote:
> sorry just to be clear your going to use a lab on a chip to perform TdT
> reactions, purify out single nucleotide addition oligos,

yep, on-chip, using one of the few single-molecule 'is there' analytic
techniques (impedance/stripping voltage, dye like YOYO-1 or gelred,
SERS raman photospec)

> transfect them into

eventually, either doing gibson/SLICE first in-vitro, or possibly
transforming fragments into something like b.subtilis that is known to
chew and re-assemble fragmented DNA during transformation. something
like that (getting to on-chip oligos would be an interesting enough
start for me!!)

> e coli, extract them from e coli and repeat the whole process several times?

I didn't really think of extraction, at the point post-transformation,
just let the cell divide for a few hours then flush into the macro
world to a user in a micro-fuge tube (or maybe there are a few
chambers on the chip that get progressively larger).

> Ok I won't say this is impossible but to construct such a lab on a chip
> would be a good deal harder than even doing something as ambitious as
> building your own surface plasmon resonance imager.

I have to definitely disagree with that.

> Also wouldn't it be far
> easier to use basic PCR than e coli to amplify the sequence.

Well e.coli basically does this kind of thing for you, and I think has
better overall fidelity than a single polymerase (or the more advanced
versions). At least they seem cheaper than in-vitro kits.

>
> actually I did a mathematical analysis based on modelling TdT reactions as a
> system of ODEs and the theoretical maximum efficiency is 100*e^-1% for one
> nucleotide additions which is approximately 36.7879%.

I'm interested in more details... I don't really understand what you
mean in this sense. In my arrangement, if tDt didn't add, I'd just
tell it to try again (or wait longer between cycles, or reduce chamber
volume to increase relative concentration), and it it added somehow
too many, either you could waste the molecule... or have an
alternative reaction loop that is some sort of really slow exonuclease
that only works on the growing end. Am I missing something about
efficiency? Regardless of assembly, etc, downstream?

[CodeWarrior]

sadly I’m a bit busy but I’ll try to work through the maths quickly.

so we have 4 populations start oligos f0, oligos with one nucleotide additions f1 and oligos larger than that fm we also have a finite supply of nucleotides n. our system of ODEs is

dn/dt=-a n f0-a n f1-a n fm

df0/dt=-a n f0

df1/dt=a n f0-a n f1

dfm/dt=a n f1

the terms in turn are, a n f0 reactions where a nucleotide is consumed changing a start oligo to an oligo with one nucleotide, a n f1 reactions where the desired one nucleotide addition oligos are turned into longer many nucleotide added oligos. a n fm reactions where many nucleotide oligos grow still larger. Notice all reactions are equally likely as TdT doesn’t care about oligo length once you get past oligos a few bases long. The non linear system can be simplified by noticing that df0/dt+df1/dt+dfm/dt=0 implies f0+f1+fm=I the initial number of start oligos. this means we can solve for n and substitute getting a linear system. if you solve this for initial condition then find the time t-max where f1 reaches a maximum and then find f1(t-max) you get I/e. all the other variables cancel out.

adding an exonuclease would be an interesting alteration. in vivo TdT works against exonuclease activity to produce short oligo additions maybe you can get past the 36% yield limit that way.

[Koeng]

I'd say that at the size of about 400-500bp you need to transfer into E coli. Otherwise, it is better to in vitro. The reason is is that ~400-500bp is enough to reliably get in a single sequencing run while being small enough to reliably PCR. I've noticed over 5kb PCR begins to get unreliable. E coli has extremely high fidelity for non-toxic sequences, to the level where we often don't even sequence restriction enzyme cloning. For non-repeat sequences, check out LCR cloning (I like the simpler method more, there is the 'rapid and reliable' paper and 'rapid and simple' paper. The former has higher efficiency). You can assemble ~20 parts in E coli. If you can say, get 40bp of good synthesis, assembling genes in E coli could work pretty well. 

I'm pretty sure companies are already doing that, though.

-Koeng

[Nathan McCorkle]

On Sat, Sep 17, 2016 at 11:08 AM, Kent Kemmish <kemmi...@gmail.com> wrote:
> Hey Nathan,
>
> I've already built such a prototype (what you call "a strainer" I call "a demonpore array") if you want to talk about it? PM me.
>
> It took a year and half and involves a lot of stuff that's not obvious starting out. For boundary pushers I'd like to share it.
>
> -Kent

Hi Kent,
I've already got one of the SiMpore strainers, so I think I have
things under control on that front for now. I had some questions a
while ago about your tech, but it seemed like the device was still too
large to integrate onto another wafer-scale device.


Recently I met with a guy who has been in biotech-related
semiconductor gadgetry for more than 15 years, working on a lot of DNA
sequencing related tech and biosensors. He was more cautionary of
trying to work with single/low-count molecules than any specific
technique I talked with him about. This guy also mentioned some stuff
about efficiency of an enzyme polymerization, but I didn't have enough
time to really dive into exactly what his reasoning was. I think he
might have been too distracted to think about my scheme because I was
talking about low-count molecules. Overall that conversation was just
kind of confusing... on the one hand there is a lot of (hard) academic
work on single molecule manipulation and handling, but then there is
this other faction that seems to just shy away from non-bulk
reactions.

I am still trying to ingest what math CodeWarrior was talking about...
it still seems to be indicative of a bulk-reaction... while I feel
like the scheme I proposed would chop up any sort of efficiency
equations into a piecewise function, if you had to describe it
mathematically.




Latest work I've done on this project has been related to CAD...
almost at the point of being able to produce g-code or a FIB raster
file from 3D models... I've got simplistic model for a microfluidic
peristaltic pump... so now I really just need to pull the pieces
together and get to manufacture something (aiming to make a simple
'Hello DIYbio' using pumped colored water).

[BraveScience]

Hi all,

This is an awesome project. Kind of puzzled why I don't see much publications on the topic.
It's possible it has been already shifted to patent applications? (Found one using acid decapping meanwhile)

I saw pretty much a lot of ideas thrown out there, is there anyone that started testing? Ideas are running wild here, but about the basic experiments to validate the procedure?

Microfluidics seems the most straightforward approach, especially if you look for solid state synthesis. I'm thinking to give a try to the theory with the digital microfluidic device we've built at digi.bio. Using magnetic beads fuctionalized to oligonucleotides would allow programmable exposure of elongating chain to different reagents, concentrations and incubation times.

Something like elongate>wash>exonuclease>wash>decap>wash>elongate

If I make enough devices I can test probably 100x different reaction protocols and map the search space for optimal conditions.

@codewarrior I was discussing your proposal at the diybio meetup in amsterdam, I was pretty surprised to see so much involvement. We thought starting a community project there.

Best,
Fede

[Felipe Tanaami]

There is already a patent describing the enzymatic synth of oligos, but not using microfluidics or inkjet printer. I can't find it right now (my internet is very unstable), but it describes the best parameters, including using the pyrophosphate kind instead of another nucleosides. But this is not the reason I didn't try your protocol, I am just without time and money, but as soon I solve this problems, I will try. Your idea is great, and I am seeking this goal for very log time too, I think this will be a holy grail and will change everything, at least for the amateur scientists. The first thing I will try when solving my personal problems is your protocol, he wasn't forgotten :) (sorry for my english)

[CodeWarrior]

My apologies for repeatedly flogging a dead horse of a thread but a new idea occurred to me this morning. If it could be made to work it would eliminate the need for special blocked dNTPs. Supose TdT could be modified to cause it to bind very tightly to DNA ends. So tightly that after nucliotide addition instead of advancing along the DNA strand it stays stubbornly in place blocking further addition. In the case the the dNTP no longer needs to be reversibly blocked as the TdT that has added a nucliotide now remains bound blocking further additions.

The TdT thus bound is still only non covalently bound to the DNA though. Even if an extreem PH solvent capable of removing it can't be found a sufficiently hot solvent will denature the protien alowing it to unbind but leaving the DNA intact.

One could invision a sequence in which one fist adds a modified TdT with a dNTP mix causing the majority of DNA strands to elongate. The solid suport would then be washed removing free TdT and dNTP. Then exposed to a solvent to denature / unbind TdT blocking strand elangation in preperation for further additions.

Does that sound viable? The question in my mind is how dificult might it be to protien engenear TdT incapable of progressing along or unbinding from a DNA strand end? I've no facilities to produce or test TdT mutants in a lab at the moment but given time I may be able to investigate the matter in silico.

[Bryan Bishop]

On Tue, Jan 10, 2017 at 8:00 AM, CodeWarrior <code.w...@gmail.com> wrote:

The TdT thus bound is still only non covalently bound to the DNA though. Even if an extreem PH solvent capable of removing it can't be found a sufficiently hot solvent will denature the protien alowing it to unbind but leaving the DNA intact.

Similar methods were described earlier in the thread (such as from Nathan):
https://groups.google.com/d/msg/diybio/FXaT-MjJQYI/tdHZwXqeCgAJ

DNA synthesis chemistry using TdT:
http://2014.igem.org/Team:Cooper_Union/TdT_project
http://2015.igem.org/Team:Cooper_Union/DeNovoSynthesis

There was some discussion about that team's use of TdT over here:
https://groups.google.com/d/msg/enzymaticsynthesis/DApMjXx8gS4/CmKEK9-vJlwJ

There was also an igem team that did some characterization work,
http://2014.igem.org/Team:Rutgers

even earlier mumblings about TdT from Nathan can be found here:
http://gnusha.org/logs/2013-07-13.log
http://gnusha.org/logs/2014-12-26.log

e.g. from a search he asked me to do a few years ago, http://diyhpl.us/~bryan/irc/nmz787-nucleotide.log.txt

I believe Cathal also had some ideas for TdT:
http://diyhpl.us/wiki/dna/synthesis/tdt/

TdT is certainly a useful critter to work with. A bunch of TdT chimeras and mutants would be really useful to investigate. By the way, I think that another area worth considering is DNA assembly; we can somewhat reliably get 100 bp oligos, and constructing longer stretches is a pain in the butt--- if we could somehow make that process simpler (such as using yeast homologous recombination) then we wouldn't need long DNA synthesis methods in the first place.