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Isolation and Culture of Bioluminescent Bacteria

March 11, 2010 in Uncategorized | No comments

Update: Keep your bacteria in the dark. I had been doing this without meaning to, and was confused to hear a fellow microbiologist growing a derivative of my cultures in her lab and getting no glow. When she grew them in the dark, they glowed again! Brief exposures have no effect. It’s the prevailing light conditions that seem to decide whether to stay active. This may be due to the light breaking down the little peptide the bacteria use to detect whether there are enough of them to glow meaningfully, or it might be a deliberate evolutionary adaptation. Who knows? :) I will update the document soon to add this consideration. Added pictures to this post, meanwhile. Finally got a good camera and took some.

In the logical follow-up to my last guide, which illustrated the production of homebrew bacterial media, here is a protocol for isolating bioluminescent bacteria from fresh seafood. The project was inspired by Mac Cowell’s successes. It is heavily based on the Indiana Biolabs protocol, differing in that it offers more background and suggests streaking your cells right away (I floundered for a while with mixed, impure cultures before finally streaking them out correctly on a petri dish and getting a pure culture).

It was written in a hurry, so if there are errors, omissions, or if it doesn’t make enough sense or explain things sufficiently, please let me know.

The document is available as a PDF and as an editable Openoffice.org document. It is released under a Creative Commons Attribution, Sharealike 3.0 license.

Again, please let me know if you use this protocol, and tell me how it worked out for you!

In related news, now might be a good time to point everyone in the direction of Applied Micro Systems, who sell lab equipment, supplies and mushroom cultivars for people interested in homebrew microbiology and mycology.

Several repurposed jars full of bioluminescent bacteria

Some of my P.phosphoreum cells growing on tissue partially immersed in liquid broth. They glow brightest when grown like this or on a long agar slant.

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Cargo Cults and Electrophoresis

February 2, 2010 in Uncategorized | 3 comments

Toward the latter half of WWII, landing strips and airbases were being built all over the Pacific on small, otherwise totally isolated islands. Little contact was made with the natives to explain what was going on, and in some cases the locals were incredulous to see the new arrivals performing strange rituals on their tarmac runways and receiving airdropped “gifts” from the gods/ancestors.

When the bases were abandoned, in some cases the locals started replicating the structures and activities of the base personnel, performing marches and drills, talking into replica radios, and waving lit firebrands on the runways and waving them. Elements of Christianity were often woven into the practise in an attempt to attract the same blessings as the military staff had. These culturally contaminated pseudoreligions are called “cargo cults“, and these words have since been applied to any misapplication of a procedure to an end without understanding how it works.

It might be premature of me to describe modern Electrophoresis as it is performed in most labs as “Cargo-cultish” until I’ve properly demonstrated that I have a better alternative, but having read this paper over coffee this morning, I was really reminded how little me or my fellow lab staff really bother to understand Electrophoresis, a fundamental, essential part of our daily work with DNA and the part whose quality is usually what ends up being scrutinised on peer reviewed papers!

Ask someone which buffer to use and you’re always offered one of three options: 1x TAE (Tris-Acetate-EDTA), 1x TBE (Tris-Borate-EDTA) or 0.5 TBE for those edgy folks who’ve been doing this for a while. TBE offers better resolution when you go to look at your results, and can be run at higher temperatures, so people often favour TBE for routine use. Some people in the know will suggest TAE where you’ll need to do something to the DNA afterwards, because the Borate in TBE is high enough concentration that it can mess up DNA extractions from the Gel and enzymatic reactions later. I only learned this upon reading the aforementioned paper, and I’ve frequently lost my DNA upon gel extraction, which I now realise may be down to my selection of buffer.

As the paper intricately explains, the use of Tris buffers is a holdover from DNA electrophoresis being borrowed from protein electrophoresis. What followed after its adoption was a brief spree of experimentation, followed by a trend towards standardisation that lead to everyone adopting TAE or TBE so they could all understand one another’s protocols. The assumption was that any effort put into improving gel resolution was wasted effort, as only marginal gains would be made.

However, there is a big problem with Tris buffers that haunts anyone performing Electrophoresis and causes all manner of problems, even when it isn’t recognised to be the cause of these problems. Heat generation due to the current passing through the gel can cause the gel to soften or even melt, and worse still as the heat increases the conductivity of the gel increases also, leading to runaway heating and poor outcomes.

Problems caused directly or indirectly by heat include:

  • “Smiling” bands,
  • Smeared or blurred bands due to uneven softening of gel surface
  • Band diffusion, forcing one to perform oversized gel extractions
  • Total Gel Meltdowns
  • Having to run at low voltages and missing dinner.

The reason behind this runaway reaction is the inclusion of Tris and excess Sodium from the preparation of the EDTA used in the buffer.

EDTA, they argue, isn’t even needed nowadays. Apparently they ran a gel using creek water without EDTA and had few issues; the enzymes used today have few unwanted activities under electrophoresing conditions. Or so we are told.

Tris, meanwhile, is too conductive and permits a lot of current through the Gel at a given voltage. As the gel heats, it lets even more through.

The authors experimented instead with a simple electrophoresis buffer composed of Sodium Borate at 10mMol (among other experimental buffers) and found it to be ideal on almost all fronts; Sodium Borate is cheap, can be prepared as a buffer from only one crystalline ingredient, gives excellent resolution, heats very slowly and can be used at very high voltages without issue (meaning faster gels).

If this is true, it means that Gel Electrophoresis as it is commonly performed is not only flawed but overpriced; a quick check of the price of 1L 10x TBE buffer from Bioplus shows that it costs $12.75. For 17.49/kg from the Science Company, I can make 52L of 10x Sodium Borate (1x SB Buffer is 1.907g/L Disodium Borate Decahydrate, the usual crystal form).

So the costs per litre of the two are $1.28 for TBE, and $0.04 for SB. For your investment of 4c you get better gel resolution at higher voltages (meaning less time waiting), which can apparently be more reliably purified and used downstream for enzymatic reactions such as ligation or PCR.

This is great if it’s true in my labwork at the CCRC, but it’s even better news for DIYbio folks worldwide: You can often find Sodium Borate on ebay where you’ll never see TAE/TBE, and the cost difference is pretty staggering.

More on this when I next run a gel: I have a bottle of 5x and 1x SB buffer next to me, just waiting for me to add Agarose and give it a try.

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Dremelfuge: It works, and it’s Available

December 29, 2009 in Uncategorized | 6 comments

Long Overdue Update: I’m very proud to say that, some time back, I updated the Dremelfuge design with better tolerances and a better shape to handle tubes. When I tested it (only once so far) at full speed on a dremel with two tubes full of fruit smoothie, it didn’t eject or break the tubes at all.

So there you go, Dremelfuge can now be considered the world’s cheapest midi-ultra-centrifuge, capable of putting about 52,000g on up to six 1.5ml eppendorf tubes. Warning; Lots of risk, don’t use this thing unless you have taken some serious precautions. Try to stay outside the plane of rotation.

Back to the original post:

Since my last post, I’ve been a very busy person.

Dremelfuge is now available for purchase on Shapeways from my shop there. There are two versions, one with an axle for chuck-fitting machines, and another with a bore into which the cutting-disc-holder from a standard dremel can be fitted. Price varies by location, but even at the $65 price which includes shipping to Ireland plus VAT, you could buy a Dremel to match it and still come in under €100 for a functioning centrifuge. I gather the price falls to $55 for American buyers.

Here’s a video of me demonstrating Dremelfuge. I tested it with standard microcentrifuge tubes, and found that it stably spins them anywhere from 5000g to somewhere above 20000g. I say “somewhere above” because the tubes shatterd somewhere between the third speed setting and the fifth on the dremel.
The math shows that the average force on a microcentrifuge tube quickly exceeds that of the commercial centrifuges I use in the lab. They go as high as 14,000g. Dremelfuge plus a Dremel 300 can put over 50,000g on a sample. Except that’s too much for the tubes so they shatter.

One nice bonus is that it seems to be very stable on a Dremel 300; there’s little to no vibration or rattling, even with highly unbalanced loads.

So here I have it: A centrifuge attachment for drills or rotary tools which spins them with even more power than the official thing, and costs a tiny fraction of the price to make and operate. I call that a success by every metric!

Thanks to Makerbot for making this possible in the first place, and my fiancee and family for their patience.

As always, I don’t endorse use of Dremelfuge as anything but an ornament, for reasons of liability.

Update: I’ve tested Dremelfuge in my lab with E.coli cells and HL60 human suspension cells. It pellets both excellently! I’ve already shown it to spin down Miniprep columns, and the math shows it hugely exceeds the power of a standard lab centrifuge when used with a Dremel 300 (€89 in Argos and useful for just about everything else, too).

So that’s it as far as I’m concerned: Dremelfuge is a fully functioning centrifuge. Can’t wait to see it in use on some cool projects!

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DremelFuge, First Go

December 24, 2009 in Uncategorized | No comments

I’ve had on my mind an idea for some time that I’ve wanted to try. Having a Makerbot has enabled me to experiment with mad science on a level I’ve not been able to before, so here it is: DremelFuge, a printable drill/rotary tool attachment that spins microcentrifuge tubes!

I uploaded a quickly mashed-together first draft to Thingiverse, but didn’t have a chance to print it that day as planned because I lost my laptop in town while Christmas shopping. Thankfully, I found the laptop since. Just tonight, I got an email from a friend in Washington telling me DremelFuge had been featured in Makezine, which blows my mind completely. Well, not being content to let it remain unprinted for a moment longer, I set to making it.

It was my hope that I’d be able to put it to immediate use and have something great to add right away, but unfortunately it doesn’t work just yet. However, that’s simply a matter of solving two design mistakes: Firstly, spacing the cavities further apart to increase the strength of the printed object, and secondly providing some means of actually loading the microcentrifuge tubes! Unfortunately as made, the object made no allowance for actually putting the tubes onto the rotor, which of course makes it impractical to use. I aim to fix this as quickly as possible.

Without further ado, the good news: It survived a full-speed test on the best drill we own, which tells me it should survive the rigours of actual use as well!

DremelFuge Speed Test on Powerdrill

More on this as I develop it!

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First Test Setup of OpenThermo Project

September 14, 2009 in Uncategorized | 10 comments

Long Overdue Update: This post has turned out to be one of the all-time most popular on my site, which surprises me to no end. Who’d have thought my crummy heatsink-and-tin thermal cycler would be cooler than isolating glowing bacteria or printing a 52,000g centrifuge?  But, who am I to question human interest. It’s not like my interests are particularly normal anyway. However, I do think this post needs updating, since people keep returning to it and asking questions.

The reason I haven’t pursued this project is largely because it’s being done far better by others more equipped to create a fully open-source thermal cycler. Look to http://openpcr.org to see this exciting project as it develops. As I write this, there are still 6 days left in the OpenPCR Kickstarter fundraising project, which has been wildly successful already. The more money they get through this fundraising, the more likely it is that they can include “bonus features” in the final model. If you donate $512 to the project, you can even choose to have them send you one of the first machines once they are ready! I have done so, and without doubt I’ll be reviewing the machine and working with it as soon as I have it, and sharing the data on this blog.

If OpenPCR works out, it’ll be the cheapest programmable PCR machine in the world. However, it’ll be far more than just the cheapest; with an open-source PC-based frontend, it’ll be the most customisable and future-proof PCR machine I’ll have yet encountered. I thoroughly look forward to making it part of my daily lab routine if so.

Back to the original post:

So, my last post (ages ago) was all about my dreams of designing an open lab toolkit for anyone to use, professionally or amateur. I haven’t been idle in the meantime! That said, I also haven’t had enough time to burn on this project, so it’s only just getting started. I’ve decided to start calling it OpenThermo, to get across the notion that it’s going to be Open Source and it’s, er, thermo.

To cut the long story short, I spent some time fiddling with thermistors to get a temperature sensing system working, only to discover that A) Thermistors are $#1+, and B) There’s an IC called an LM35 that accurately senses temperature and gives a clean, linear output based on the celcius value. That sped the development up a lot! I only had to get me one of these fine LM35s.. thanks to ebay, it arrived last friday.

The other area that I wasn’t looking forward to sorting out was the Peltier circuit; since the peltier heater/cooler units operate at a fairly high wattage, the Arduino wouldn’t be able to drive them on its own at all. Therefore, I had to sort out a way of getting high power through the peltier, under the control of the arduino, without burning out the latter by accident. Probably elementary to an electronics guru, but I’m not one of those. I discovered that, again, there are some wonderful ICs to do the job for me; in this case, the L293D motor controller chip apparently handles an isolated power source and high-power load while accepting low-voltage logic input. In other words, perfect.

I’ve therefore just started work on making the actual thermal cycler, finally. Shown below is my first setup for testing the L293D:

First setup of my homebrew thermocycler. Shown is a 12v battery bank, an L293D motor driver chip, my arduino, and a hacked-together peltier thermocycler rig using thermal tape, a heatsink and a little enzyme tin from the lab.

First setup of my homebrew thermocycler. Shown is a 12v battery bank, an L293D motor driver chip, my arduino, and a hacked-together peltier thermocycler rig using thermal tape, a heatsink and a little enzyme tin from the lab.

Quite the jury-rig. However, it seems to work. I would be testing it more now, rather than writing about it, except that I noticed a funny smell about 20s into the testing (a simple on/off/reverse/on/off script I’ll share later) and started looking for the source. I actually burned my finger slightly off the L293D chip! So.. Doesn’t like loads THAT high, it seems. Thankfully I come pre-armed with a solution: According to LadyAda on her Arduino Motor Shield instructions, you can just stack the chips to add twice the current capacity. I can’t do this just now, in part because I have more pressing business in the lab. That, and I’ll probably add a crude heatsink while I’m at it; definitely a DIY job for at-home time and not the lab.

Suffice to say I think it was working before I stopped it, but at the wattage I had it going it may have been about to burn out the chip.

Next task is to test out the LM35 chip, which should be elementary. I think I shall love the LM35, and want to make all sorts of funny things with it. At least the odds of my setting one on fire are far less than with the motor chip!

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On Arduinos and Homebrew Genetics

June 17, 2009 in Uncategorized | 4 comments

For a while preceding buying my own Arduino, I spent my time looking at all the cool projects people have used them for on the Make blog, Fashioning Technology and the Arduino wiki itself. Although I love all the projects that showcase the artistic use of electronics, and I’m impressed with the more utilitarian uses also, I see great untapped potential in the Arduino as a replacement technology for certain niches where equipment is prohibitively expensive.

One such niche I am intimately aware of is Biotechnology, particularly molecular biotechnology. For example, I work daily with a machine called a thermal cycler. These machines perform a seemingly simple duty; they accurately vary the temperature of a metal casing in which you place little plastic tubes containing your reaction mixes for molecular biology. You set them up to cycle through a variety of temperatures so as to help enzymes work at optimal efficiency, break the double-helix of DNA with heat, permit only certain molecules to adhere to one another during a reaction, or to inactivate enzymes through heat-treating. They are most often used for the Polymerase Chain Reaction, a cornerstone of modern genetics, and so they are often called PCR machines.

The odd thing about these machines is the cost. A thermal cycler can cost over €2,000 for a used robotic version, or as low as €200 for an ancient, damaged one. These machines almost universally have a poor interface, too; few can connect easily to a computer, and none are open-source.

What’s going on here? Am I missing some deep secret of thermal cyclers that should make them so expensive and limited? So far as I know, a modern thermal cycler technically only uses one basic technology; a Peltier thermoelectric device. These little semiconductor/ceramic plates get hot on one side and cool on the other when supplied with electricity, with the orientation depending on the polarity of the voltage. In short, the work like an electric heat pump, heating or cooling one side on demand. And, they cost less than €10 on ebay with free postage.

In principal all that’s needed to make one, then, is a Peltier unit, a temperature sensor (60c in Maplin), a little breadboarding ( < €20) and an Arduino (€20). Connect it to a PC, optimise your code, and you have a highly flexible Thermal cycler for less than €50.

This ties into something of a broader interest for me, that of someday having my own lab wherein to do my research. I think it’s just part of my personality that I feel compelled to do stuff for myself, and tend to lose interest in projects not my own. I look forward to working on nutty genetics projects that I fancy might, just might, have the potential to help improve the world..or at least look nice growing in a pot by the window.

Setting up a Genetics lab is the big obstacle here; because you need such specialised equipment and reagents, it’s not a simple matter of building a shed and ordering from a catalog. (Update: I started a shop on Shapeways where I am already selling some cheap lab equipment: LabsFromFabs) Although a growing community of people are taking an interest in homebrew biology, their ability to do so will be constrained by the price of setting up and maintaining a working lab.

Thankfully, this is slowly changing. Much as electronic technology used to be a very exclusive field until prices plummeted and homebrew electronics became an affordable hobby, certain powers that be are working to bring Biology labs into the home for enthusiasts and clubs. Update: See the LavaAmp for one such project that is already near the market!

One such effort is OpenWetWare, a wiki project that aims to build a directory of Protocols, Tutorials and Essays to teach the knowledge and practices of Biology online, for free. Most homebrew clubs I’ve encountered maintain a page on OpenWetWare to showcase their work or share their experiences.

Another, potentially even more revolutionary example is the Biobricks Foundation, which aims to standardise genetic material to enable swift, easy construction of complex genetic tools and systems. Although they suffer from a terrible website design and a general lack of beginner-friendly information or support, the system they are using is seemingly reliable and robust. It should require no more specialised equipment than four enzymes and a special breed of e.coli to start making things that glow, smell like bananas or release special enzymes in response to something. Technically.

Best of all, existing genetic “parts” (read; the bits of whole genes such as the coding sequence, promoter, terminator, etc) can be modified so that they are themselves new “Biobrick” parts, and can be added to the Biobricks Registry of Standard Parts. If someone wants your part, they can ask you to mail some over to them!

Finally, and perhaps most importantly, those interested in genetics at home will be excited to hear that I firmly believe a service will be available within this decade where you can simply order a gene from scratch and have it delivered already transformed into e.coli (at least!). Mail order GMOs.

I say this because you can go right now over to Mr. Gene and order up to 3kb worth of DNA to be synthesised from scratch at a competitive price (€0.32 per nucleotide), which is enough to contain a whole functioning bacterial gene. Not only that, but their web-tools will help you easily add in all the special sequences you want, while excluding those you don’t, while maintaining the coding sequence and optimising it by species. That’s pretty much all the hard work.

The price is still expensive; although €960 should cover 3kb at 35c/n, it can cost a bit more for a useful gene to be made due to the relative difficulties in synthesising odd or difficult sequences. However, I have learned (through my own travails working in a genetics lab) that producing and stably assembling a gene by yourself is sometimes so time consuming (months!) and expensive that you’d be better off just paying the extra money and getting a guaranteed sequence in 15 days.

It is a small step from being able to order a complete gene and the e.coli separately, and transforming them on your benchtop, to having a company do all of this for you in advance and delivering the finished, genetically modified product by courier.

It would also be a small thing to order a modified strain of another bacterium, known as Agrobacterium tumefaciens. The significant difference here is, Agrobacterium is known for its ability to transfer DNA into plants, and is routinely used as the most simplistic means of creating GMO plants. So, mail order transformed Agrobacterium = Mail order modified plants.

I eagerly look forward to this day. For the moment though, and coming back to the beginning of this post, I aim to create an open, free and publicly available design for at least one piece of specialised equipment. With this and other similar projects, I’m hoping lab equipment stops being so damned exclusive and comes quickly to the Maplin Catalog. Update: See LabsFromFabs, my Shapeways shop where I’m already selling some lab equipment!

And if someone else reading this has the tech savvy to help (I’m an electronics n00b), please do!

Edit: The price of gene synthesis wasn’t €0.35, it was $0.35. Much cheaper than I quoted.

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