The Gel Integrated System is done.  I have packed an electrophoresis power supply, illumination, and casting system into one tiny box.  All the parts, other than the enclosure, are off the shelf parts.  It can be shipped flat and assembled with only a screwdriver.  This all comes in at a sale price that is an order of magnitude lower than commercial systems, with a bench footprint an order of magnitude lower than commercial systems.

It has been tested, and it is completely functional.  The gel is bright enough that you can take pictures of it with your cell phone.  If you want to get one, I will be selling the first batch of ten here for $200 a piece.  Use that link to contact me if you need a large batch, or find me on the diybio mailing list.  I will hopefully have an assembly video (and feature video) soon, but until now you can see my instructable on assembly here.

Almost ready to test the latest prototype!  Everything electrical is go, but I managed to loose my LED panel in my moves from showing my prototype in CA, visiting the bay area, and them moving from an apartment in Somerville back to school.  Oops.

One of the major improvements in this prototype is simplified wiring.  I did away with the ammeter (since it was on the fritz anyways), and now it is very easy to assemble with only a screwdriver.  The improvement is that lots of wires don’t need to go into screw terminals now- there is a max of only two wires per terminal, and if you wrap stranded wires around solid wires, you have a pretty solid connection.

I am really looking forward into getting these into peoples hands!

This is the newly designed tray for the gel system.  It is a major improvement over the last tray, and it comes with some other major revisions to the box.  The big upgrades to the tray are:

• about 2x cheaper
• built-in diffuser
• tighter slots for the dams (orange things in that picture) by .005″

This makes the tray system pretty solid, and the low profile saves gel and buffer, and makes it even harder to get your digits into anything that is electrified.  It also makes the box 100% cuttable, with no need to drill any holes, which was an issue since the laser cutters I use can’t focus on an object that is 3″ high.

The other major revision that is coming is an adjustable LED light source.  I love the EL panel, but it is tough to get a good photo without a decent camera, and it is just not as bright.  Even though the EL panel is great for me, I expect that some people will just want to use a cell phone, which is not feasible with the EL panel as it is. Heck, my phone is my lab notebook, and carrying around a big camera is inconvenient, so most of the time it is all I have (I am advocate of the lab having a camera).

With the dam sorted and the illuminator design tested, I am ready to call the next revision the minimal viable product and ship it out.

Some folks were having trouble with microcentrifuge a.k.a. eppendorf tubes escaping the rotor of their dremelfuge.  I compiled a handy list of dimensions for the tubes we have at BOSSLAB, and stuck it a PDF.  I might have to build a dremelfuge now.

The brands and sizes measured are:

Carolina biological colored tube- 1.5ml

Eppendorf- 1.5ml

ABI Geneamp tubes- .5ml

Biologix flat top PCR tube- .2ml

tube_sizes.pdf

The second prototype of the Gel Integrated System is done.  I ran a gel on it (wet tested it) today, and I am happy to say that I am ready to sell, build, and ship the first few units.  There are a few minor revisions that will go out in the first few units, like a clear “lid”, and slightly different dimensions for the wire cutouts, but mostly they will be the same as what I tested today.

Here is a brief rundown of how the test went.

Pouring the gel started out ok, but I need to get the seals a little wider and the slot that they go into a little thinner, so they seal.  The leakage I experienced should be a non issue for future users.

It is literally impossible to run this box backwards by accident, since you can only put the tray in in one direction.  If you wanted to run it “backwards” you could pull the electrodes out and move them to the other side, but otherwise your DNA will always be loaded on the right side.  The wells need to be wider and thinner, but setup for this part is slick and simple., and changing the width of the wells is as simple as cutting the comb out of thinner plastic.

Here is the post-run gel, in regular light  You can barely see the loading dye on the bottom, since I didn’t quite get the sample in the well, but the lanes ran very evenly.

This picture shows two things: one, I might need a pre-filter if I want to do photography.  Two, you can barely see that the the gel did run, but that the sample wells need to be broader so you get the nice crisp bands that everyone loves.  In real life it is quite a bit more visible.  We will see what people want!

Overall, I  would call it a success.  A few tweaks need to be made on the units to be shipped out, but it’s time to try to get this thing out there, and have people use it!

The second prototype has seen large improvements in fabrication.  Other than the obvious problems, like the box being too small, there have been a few big improvements.

On the left is the cut sheet from the previous prototype, and on the right is the cut sheet of the latest prototype.  The latest prototype is actually larger by square inches, but it takes almost half the time to cut!  And there is less waste of material between the parts.  The improvement here was manually aligning pieces to share edges, and then deleting (again, manually) one of the shared edges.  If you do this, deleting the extra edge is very important.  If you don’t the laser will hit it again, and sometimes create a very small “peel” of plastic between the two laser lines.  This piece of plastic is very thin, and will be insulated by air on post sides, which will cause it to catch on fire!  So delete the extra line.

This is the “mouse door” cutout that lets you route cables with big connectors through small slots in mortise joints.  This one is still slightly too small, but it still works pretty well!

I also changed the box configuration.  On the left is the new box, which is thinner and more affordable to fabricate.  It eliminates the on-edge holes, which I normally have to drill.  Originally I wanted to turn the tray on-edge and just laser cut the holes, but the cutter was not tall enough for that.

Thats all for this note!

One more cut of the gel box.  It is looking really good, and there are only a couple of tight spots where the wires enter/exit the box.  It was assembled with only a screwdriver!  Tomorrow I will do a “wet test” of the system, and see how it fares.

The year is 2013.  Why is it when you Google “How do I align my forward and reverse sequences” the two methods that come up are using ClustalW (Clustal Omega), which requires a reference sequence, or some kind of awful witchcraft involving Microsoft word and the search function?  I guess there are also some programs out there that you have to pay a large sum to use (squencher etc.) but I certainly don’t have that kind of money.

So I decided to write a little python script to help me do alignments.  The output, which you see above, is not very user friendly yet, but it at least gives you a place to start before you devolve to microsoft word (of all things!  I will probably use libreoffice though).

Let me explain whats going on up there.  The script, which is called DNACrusher, takes the forward and reverse reads from your plasmid, and takes the reverse complement of the reverse read.  Then it overlaps the ends that should overlap (the end of the forward read, and the beginning of the reverse complement) by an increasing number of basepairs and counts two things: the longest matching run, and the number of matching base pairs, and stores them in a tuple that remembers how big the overlap is.

The blue line represents the raw number of agreeing basepairs.  As you would imagine, as you mash more bp together, the number of agreeing pairs generally go up, as does the noise on that read.  I haven’t computed any statistics, but that looks about right.  The only anomaly is right before 200 bp overlap, where you have a spike.  That turns out to be at 172 bp.

The green line represents the longest run of agreeing bp.  This line, as you can see, is relatively flat except for a corresponding peak at 172bp.  This makes me fairly certain that there is an overlap of about 172 bp, which seems reasonable.  When run against clustalW with the same sequences and a reference sequence, the overlap is about 160 bp, including some gaps in one strand or the other.

You can get the python at ye olde github account.  Included the two reads I used to create the graph above.  There are only a few methods in the little script, here is a breakdown of how to use them:

Import the DNACrusher module and pyplot

>>>import DNACusher

>>>import matplotlib.pyplot as plt

easy peasyImport your data from a Fasta File

Make a file object from your fasta file

>>>sequence=open(‘Fasta_File.txt’,’r’)

ta-da!

Convert Your File Object to A String

>>>sequence=DNACrusher.fasta_to_string(sequence)

now sequence is a string, which is way easier to deal with

Reverse Complement

>>>sequence=DNACrusher.reverse(sequence)

>>>sequence=DNACrusher.complement(sequence)

‘atgcaata’->’tattgcat’

Overlap N Base Pairs

>>>DNACrusher.crush(forward,reverse,N_BasePairs)

This will return the count of matched bps and the longest run of pbs in a tuple (count, longest)

Overlap Up to N Base Pairs

>>>align=DNACrusher.allign(forward,reverse,N_BasePairs)

This will return a list of tuples [ (pairs_overlapped,(count,longest) , …]

Now to Plot It

>>>plt.plot([ x[1][0] for x in align])  #plot the number of agreement

>>>plt.plot(x[1][1] for x in align]) #plot the record # of runs

>>>plt.show() #show the graph!

Now if you do this with the sequences I provided, you should see the above graph!

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Man, this is a cliche thing to print, but it is really useful.  Thats a picture of it on my Amscope 10-30x scope.  Now that I have the dimensions down, I am going to re-print it with a rest for my phone.  I would provide the model, but it was actually 5% “too small” although it could be that the printer just printed it a little small-I need to check the calibration and see how close it is.

You can see here how I trimmed the length a bit too, since I didn’t know how long it should be.  I just used some flush cutters to start the removal, and then the layers peeled off from the Z axis.  Its kind of an adjustable length print, since the walls are just two layers thick.

Anyway, the results are great!  This was taken at 10x, and you can probably already see why it is easier to solder through a ‘scope than with just your eyes.  And sometimes you need to get tiny, so that is what 30x mode is for.

Here is a shot at 30x.  You can see the grain of the plastic!  It is also fantastic for soldering, and checking for shorts in across SMD components.

The adapter makes it easy to take pictures by making sure the camera sensor is where (or near where) your eye normally is when you look at the scope.  If you have ever had trouble looking into a scope (tiny picture, weird black crecents), it’s probably because you were too close or too far away.  The eyepieces help a lot with this, but they don’t fit my phone, which makes it really hard to photograph stuff- hence the adapter.

Now that I have a scope and an adapter, I definitely want to take some 3d stereo-microscopy pictures, and videos of what soldering looks like at 30X.  I think it would be really neat for teaching people to solder SMD components to be able to show exactly what it looks like when solder wicks onto a pad instead of just talking about it.