DIYbio/FAQ/Equipment

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DIYbio FAQ v1.5: "The biohacker's FAQ"

This FAQ for DIYbio is actively maintained by it's editors, and by you! Edit your contributions directly or email updates to the DIYbio email list, diybio@googlegroups.com.

Major contributors (in alphabetical order):

• Bryan Bishop <kanzure@gmail.com>
• Jonathan Cline <jcline@ieee.org>

The contents of this FAQ are copyright under the OpenWetWare Copyright policy (Creative Commons Attribution-ShareAlike 3.0 Unported). When quoting any content of this FAQ elsewhere, include a full hypertext link back to the main FAQ page.

This topic Equipment and Consumables is part of the DIYBio FAQ

Please update this FAQ mercilessly with Q&A !

What equipment do I need to perform DIYBio-related projects?

• Basic biology equipment (see here for PCR equipment, however much of the equipment isn't actually required)
  • or generic tools used in other projects, just use it for biology (see MacGyverisms below)
• Basic computer equipment
• Perhaps some open source software development tools

What equipment do I need to preserve my microbes?

• Feed the microbes. For e. coli, use E. coli Media or similar.

• Freeze the microbes at low temperature storage, as follows:

On Oct 3, 9:42 am, Tom Randall <tarand...@gmail.com> wrote:
> On Oct 2, 4:59 pm, Ben Gadoua <ben.gad...@gmail.com> wrote:
> 
> >   E. coli is best snap frozen in LN2 but it keeps at -20C or -80C for years
> > and years, best to keep it at -20C instead of 4C because at 4C e. coli is
> > still slowly growning.
> 
> > Ben
> 
> -20C is fine. In any ordinary freezer. I recently revived some E. coli
> strains from a previous lab that I had simply stored in my kitchen
> refrigerator (freezer compartment of course, next to frozen OJ et al.)
> for 6+ yrs frozen in 50% glycerol/LB. Those with plasmids still grew
> up with amp selection and plasmid preps produced the right plasmid.
> Have since restocked, again in 50% glycerol/LB in a dedicated -20C
> freezer this time, again off the shelf, used, approx 6 cu non frost
> free freezer for stocks and enzymes. One reason for using E. coli K12
> even if not your final organism of interest, is that it is easy to
> maintain without much specialized storage. Maintaining competent cells
> is likely a more difficult proposition and unless you are willing to
> go the -70C route you will likely have to live with a shorter shelf
> life and some loss of transformation efficiency over time.

-- Ben Gadoua and Tom Randall on the DIYbio google group.

On Oct 2, 3:58 am, Cathal Garvey <cathalgar...@gmail.com> wrote:
> A note on unfreezing stock cells; try not to!
> 
> If you have cells frozen in glycerol/media, try to scrape a tiny bit off the
> surface, and use that to start a new culture. Put the rest right back in the
> freezer. The reason being, cells don't survive repeated freeze/thaw cycles
> very well at all. Best to keep them frozen all the time, and only thaw the
> tiny bits you scrape off the top.

-- Cathal Garvey on the DIYbio google group.

On Oct 1, 7:38 pm, Ben Gadoua <ben.gad...@gmail.com> wrote:
>   Most labs that I've seen keep some frozen stock of the basic e. coli with
> no transformation, frozen either as a stab culture or with DMSA/glycol mixed
> into the media+e. coli. After that labs unfreeze part of the e. coli
> culture, or take some, on the the tip of a pipetter and dilute it into your
> plasmid, do your heatshock or electroporation, and then add rich culture
> media and let it mature in a shaker incubator for around an hour, we aren't
> looking for confluency here, we just want the bacteria to be more
> concentrated, an hour or so is about 3 doublings, so however much bacteria
> we started with has now been multiplied by 8. (In rich media at optimal
> temperature with good aeration, e. coli doubles every 20 minutes) Then you
> do as Bryan alluded to, a streak plate, you could also do two plates, one
> with 90% of the culture spread out over the surface and another with 10% of
> the culture spread out over the surface. Both methods are valid, but if you
> have a very high concentration of culture, IE. you forgot about your bullet
> prep and left it for 4 hours, a streak plate may be better because it lets
> you dilute your culture 8-32 fold depending on how many times you streak.
> The plates that you'll use for your streaking will have an antibiotic in
> them that kills all of the bacteria that you hadn't transformed, because the
> plasmid that you put into the bacteria will have had an antibiotic
> resistence. Another reason for this antibiotic resistance is that any of the
> other stray bacteria that may have gotten into your culture the times that
> is was open will also be killed.
> 
>  So there you have it, e. coli culture in a nutshell.
> 
> Ben

-- Ben Gadoua on the DIYbio google group.

What equipment is in a basic biology lab

• A note to readers.. please add more answer to this question

• Using ebay to set up a molecular bio lab for less than $1k USD (see also [1] and [2])

Where can I find auctions for biotech?

See DIYBio FAQ Main Page, Appendix 2, List of Equipment Suppliers

What Equipment can I build myself ?

See DIYBio FAQ: Projects for a growing list of equipment to build right now.

• Cheap technology for biology
• CO-2 rich environment in a ziplock baggie
• gel electrophoresis in a straw (keiki gels)
  • Keiki gels: Gel electrophoreis in a straw; described by one of the original 'idea posts'. very basic protocol which was featured on boingboing.
• inkjet, laser printer, wax, ice-lithography, sharpie, or other forms of diy microfluidics
• 5 minute DNA extraction in a shot glass

How can I make a spectrophotometer?

See DIYBio FAQ: Projects

How can I make an atomic force microscope (AFM)?

• How to use plastic printed parts to do a DIY AFM
• AFM != STM

How can I make a sterile environment?

Using Ultraviolet (UV) for sterilization

Here is a table of how much UV exposure it takes to kill various organisms and bacteria. Note units are in microwatt-seconds per cm2. Specs for UV lamps usually give emission as microwatts per cm2 measured at a distance of 1 m from the lamp.

Do not leave UV lamps on for long periods of time because the bulbs "wear out". Specifically, the amount of UV produced will decrease over time (see manufacturer spec sheets). With extended use, it may seem that the bulb is creating a sterile environment when actually it is not producing enough UV.

Warning: UV spectrum of sterilization bulbs are bad for you. Do not expose yourself to the UV. If you build a UV "hood", add a switch which always turns the light off when the door is opened.

What Equipment have other DIYBio members made? Can I buy one from them? Where are the plans?

See DIYBio FAQ: Projects

Chemicals and Reagents are expensive. How can I make my own? What can I substitute?

Substitutes for Expensive Agarose

It is possible to use substitutes for Agarose depending on the experiment and the need for precision in the result.

"Agarose seems expensive (500g for $300) however this makes almost 25 litres of 2% solution, enough for approx. 1000 gels. 30 cents a gel ain't that bad compared to precast costs."

-- Ben Gadoua on DIYbio google group

Using Agar instead of Agarose

"We have recently attempted to find an inexpensive alternative to agarose for analytical purposes. We observed that agar is an adequate support medium for gel electrophoresis (Fig 1). [...] These included Difco Bacto-Agar (Detroit, U.S.A.), Merck Agar-Agar (Darmstadt, Germany), Oxoid agar (London, UK.), Biolab Agar (Halfway House, R.S.A.), Bitek Agar (Detroit, U.S.A.) and bulk agar obtained from New Zealand. All these agar formulations gave good separation of our DNA ... Conclusion: ... agar can be used as a gel matrix in place of agarose in many instances. We do not recommend using agar if the DNA is to be purified from a gel. We have attempted experiments involving the southern blotting technique using agar as our gel matrix. Our results were, however, unsatisfactory. We therefore recommend that agar gels can be used as a cheap altelnative to agarose to check the purity, size and amount of DNA in a sample."

From: AGAR, AN ALTERNATIVE TO AGAROSE IN ANALYTICAL GEL ELECTROPHORESIS, BIOTECHNOLOGY TECH, CD. Viljoen, B.D. Wingfield* and M.J. Wingfield Volume 7 No.10 (October 1993) pp723-726. DOI:10.1007/BF00152620

"The other option is to wash with two changes of EDTA (think it was 25mM) to remove divalent metals and sulfonated (non-gelling) agar. There is also a process based on alcohol washes - but can't find my reference to it."

-- Abizar on DIYBio google group

"Most agar has a lot of positively charged groups on it that interfere with electorphoresis if DNA. That's why we use agarose, which is either derived from a source very low in these groups or chemically modified. I worked for a DNA fingerprinting company and had to lead a troubleshooting team once to find out why our DNA bands were all smeary on gels- turned out our agarose supplier had switched where they sourced their agarose from, and it was too full of sulfhydryl groups to run the large slab gels well (although apparently it was OK for smaller gels with shorter DNA fragments). Agar is normally much worse, since they don't expect anyone to use it for gels. Acrylamide is toxic until it polymerizes. You can buy precast gels to minimize your risk, but they are expensive."

-- EJ on DIYbio google group

" As a suggestion, if you're using Agar instead of Agarose, you can prerun it for maybe half an hour to try and get rid of many of the positive contaminants. You could also wash the agar in alcohol and then in ketone to try and get some of the impurities out. You'll have to try it and see, there's no other way to tell, if you want to use agar you might want to use smaller dna fragments, they run better on less pure gels. "

-- Ben Gadoua on DIYbio google group

Using Other Substitutes for Agarose

Table lists starches & flours:

Table: Alternate to Agar

"Silica gel is another possible solidifying agent. Likely totally inert, and stable at high temperatures for thermophiles. "

-- Tom Knight, diybio google group

"None of those (in the 'starches & flours' table) work (well). Agar is clear and indigestable by bacteria, none of those replacements have those properties. Guar gum is the only substitue worth trying, but from what I understand it's a LOT harder to work with. For the price of food grade agar it just isn't worth trying to save money. Food grade works great for everything I've tried and you can get it pretty dang cheap online."

-- Jake, diybio google group

How do you purify agar?

"here is a good washed agar protocol, cleans up contaminants. It involves acetone, easy to find at home depot, but dont be smoking. Also the washed agar will gel at a lower concentration, 1% instead of 1.5% or 2%. As store bought stuff is less pure than Difco, it would certainly benefit more from cleaning. I have only done the second protocol." http://www.fgsc.net/neurosporaprotocols/How%20to%20wash%20agar.pdf -- Tom on the diybio google group

"Soak agar shreds or granules in "several changes" of distilled water (DI). Make a 4% gel, slice, dialyze or electrodialyze it, then use that to make a more dilute gel (no specified dilution) through reheating. Alternatively- dry and dissolve flakes later as needed. Dissolve agar "in the solvent to be employed," and hot-filter through several layers of "lintless gauze, coarse filter paper, shredded paper or diatomaceous earth, or centrifuge at high speed (eg, 5000 g) for 10 mins in a rotor pre-heated to 80 C" (small volume technique: pull into 10 mL pipette with loose cotton plug. Remove plug and deliver to plate or slide). Make agar gel from this, then chil, freeze, and thaw it to disrupt gel and "express the water and dissolved impurities." The reader is referred to Crowle 1961 (first edition of Immunodiffusion) for more detailed explanations."

From Crowe's "Immunodiffusion," 2nd ed., 1973

-- AJ, diybio google group

Preparation of agarose with cetylpyridinium chloride "or other tertiary ammonium compounds."

From Clausen in "Laboratory Techniques in Biochemistry and Molecular Biology" (volume 1, part 3, edited by Burdon and Knippenberg)

-- AJ, diybio google group

"There was another text- it mostly consisted of making slabs of gel that are then allowed to sit in distilled water, which is changed every day for a week or more."

Maybe from "Handbook of Immunoprecipitation-in-Gel Techniques," edited by Axelson (1983).

-- AJ, diybio google group

• Much of the above is distilled from the thread: agar purification recipes

Staining DNA when performing Electrophoresis using Agar/Agarose Gels

For DIY environments, staining DNA can be done with "GR Safe" or "SYBR Safe", both work well. For other substitutions keep reading.

• Note: EtBr is not used in a DIY environment due to safety issues.

"The shelf life (of GR Safe / SYBR Safe) is also longer than 6 months, provided you keep it in its container (out of the light!). I've used tubes that were several years past their prime, with no problem."

--Kay Aull, diybio google group

Methylene Blue stain

Inexpensive, "harmless" dye. Sold in pet stores, usually in the aquarium section. Can also be found on ebay.

"The disadvantage being, of course, that the resolution of Methylene Blue is pretty bad; you need a lot of DNA for it to show up. To a lesser extent, this is a problem for all alternative dyes"

-- Cathal Garvey, diybio google group

How to Make Taq

"Taq is super easy to purify, you don't need a column. If you want Taq to use in PCR, you can just grow the plasmid with the DeltaTaq insert (available from ATCC), and heat the crude lysate. If you want to clean it up a bit more, for example for an enzymatic study, try this protocol: Rapid purification of high-activity Taq DNA polymerase Pluthero Nucl. Acids Res..1993; 21: 4850-4851"

--Stacy, diybio google group

Stains

The following are simple stains which can be used for microbes.

"Gentian violet is easily had at drugstores as an antifungal."

--mlp, diybio google group

"Eosin is OTC at any pharmacy."

--mlp, diybio google group

Retrosynthesis: how do I synthesize this chemical compound?

Retrosynthetic analysis is a method of beginning with a target compound that you wish to synthesize, and working backwards from some otherwise ridiculously hideous and expensive compound, to more simple elements and compounds that you may be more likely to have available. In 1990, E. J. Corey won the Nobel Prize in Chemistry for his work in retrosynthetic analysis. While it is possible to manually generate a retrosynthesis tree, computational tools can assist in this laborious task. At the moment, however, there are no free software tools for retrosynthesis. The pydaylight library is a wrapper to the Daylight toolkit and might serve as a good start. Please contact Bryan Bishop if you want to collaborate on this software.

Are there any plans for a DIYbio-friendly, open source database system for biology protocols, how-tos and hardware/equipment construction?

This is work in progress. You should consider contacting Bryan Bishop about this.

• Updates from Austin, TX related to diybio packaging standards (2009-08-13).
• An index of threads related to open manufacturing and standardized packaging of open source hardware projects
• Recipe representation
• Recipe representation as it concerns DIYbio
• More on recipe representation (especially re: an open thermocycler project)
• More on protocols and hardware packaging formats (2009-04-15) (see also Meredith's notes)
• please expand this list

Can the software tell me what equipment I need to run a particular experiment?

This is work in progress. This is known as dependency checking. The handy "checktools" program hopes to do this. ((Note: the idea here is that once the pcr.xml file makes a few friends with other protocols, software can then be written to extract a list of tools from the standardized protocol format. But this doesn't exist yet, since we only have "pcr.xml".))

Keiki gels (gels-in-a-straw) MiniFAQ

• one of the original 'idea posts'
• Keiki gels: Gel electrophoreis in a straw (it works!)
  • boingboing'd
• very basic protocol

Do all of the straws run at the same rate?

"I think the key there will be making sure that all the straws are exactly the same length -- each straw behaves like a resistor, so just like any other resistive material, a greater amount of material will mean a higher resistance (and thus lower current at constant voltage)." -- Meredith

How do you stain the DNA in a straw?

"Easiest way to (stain the DNA) would be to use a stain that you add to the warm agarose before pouring, such as SYBR Safe or GR Safe (or ethidium bromide, but the cool kids don't use that anymore). I suppose you could slit the straw open with a razor blade if you wanted to use methylene blue, but that sounds like a huge pain." -- Meredith

DIY Genetic engineering

• Basic questions on genetic engineering
• Notes on gene vectors and miRNAs (gene silencing)
• Gene naming, homologous sequence similarity
• Genetic engineering (in general)

Discussion of Organisms for DIY Genetic Engineering

See the group discussions and the diybio model organisms list.

Candidates are:

• Psychomitrella patens (a moss that is naturally competent)
• Halobacterium NRC1 (grows in very salty media)
• Acinetobacter baylyi ADP1

ADP1

ADP1 has been considered a good candidate for DIY Genetic Engineering because it is naturally competent.

"I developed ADP1 as a model organism for simple genetic engineering while at Scripps. The paper appears under my name in Nucleic Acids Research (5780¿5790 Nucleic Acids Research, 2004, Vol. 32, No. 19 doi:10.1093/nar/gkh881). When I did the work, ADP1 was considered A. calcoaceticus, and was given a clean bill of health (biosafetly level 1). Later, to my dismay, it was collapsed into A. baylyi, grouping it with nasty pathogens and making it thereby less accessible. My interpretation: they probably are all the same species, technically, but Acinetobacter's predisposition for collecting genes from outside sources (which is exactly what makes it so useful - read the paper), led some strains to collect a bunch of virulence factors and become superbugs, like the ones that plague hospitals. If you can get some ADP1, I would consider it as safe as Ec K-12, but be very careful with less well-characterized strains of this species, because it can and will pick up genes that offer an adaptive advantage in it's environment, so you never know what a wild-type Acinetobacter might be capable of."

-- dmetzgar, diybio google group

DNA synthesis MiniFAQ

• DIY DNA synthesis and bibliography
• DLP projector based gene synthesizer proposal
• Some ranting notes on schemes for microfluidic DNA synthesizers
• Sequencing and synthesis predictions for 2010 from 2003
• Synthesis of unnatural nucleic acids (like GNA)
• Short bibliography of DIY-friendly DNA synthesis techniques
• Notes towards DIY DNA synthesis and the synthesis of photolabile protecting groups from kitchen household products
• Origins of oligonucleotide impurities and errors

Can I order DNA over the internet?

mrgene, e-oligos, geneoracle, etc.

What are oligonucleotides?

Wikipedia sez: "An oligonucleotide is a short nucleic acid polymer, typically with twenty or fewer bases. Although they can be formed by bond cleavage of longer segments, they are now more commonly synthesized by polymerizing individual nucleotide precursors. Automated synthesizers allow the synthesis of oligonucleotides up to 160 to 200 bases. The length of the oligonucleotide is usually denoted by "mer" (from Greek meros, "part"). For example, a fragment of 25 bases would be called a 25-mer. Because oligonucleotides readily bind to their respective complementary nucleotide, they are often used as probes for detecting DNA or RNA. Examples of procedures that use oligonucleotides include DNA microarrays, Southern blots, ASO analysis, fluorescent in situ hybridization (FISH), and the synthesis of artificial genes. Oligonucleotides composed of DNA (deoxyoligonucleotides) are often used in the polymerase chain reaction, a procedure that can greatly amplify almost any small piece of DNA. There, the oligonucleotide is referred to as a primer, allowing DNA polymerase to extend the oligonucleotide and replicate the complementary strand."

How are oligonucleotides synthesized?

Oligonucleotide synthesis is done via a cycle of four chemical reactions that are repeated until all desired bases have been added:

• Step 1 - De-blocking (detritylation): The DMT is removed with an acid, such as TCA (get it at Sigma-Aldrich), and washed out, resulting in a free 5' hydroxyl group on the first base.
• Step 2 - Base condensation (coupling): A phosphoramidite nucleotide (or a mix) (struct, synthesis of phosphoramidite building blocks [pdf]) is activated by tetrazole (get) which removes the iPr2N group on the phosphate group. After addition, the deprotected 5' OH of the first base and the phosphate of the second base react to join the two bases together in a phosphite linkage. These reactions are not done in water but in tetrahydrofuran (get) or in DMSO (get). Unbound base and by-products are washed out.
• Step 3 - Capping: About 1% of the 5' OH groups do not react with the new base and need to be blocked from further reaction to prevent the synthesis of oligonucleotides with an internal base deletion. This is done by adding a protective group in the form of acetic anhydride (get) and 1-methylimidazole (get)which react with the free 5' OH groups via acetylation. Excess reagents are washed out.
• Step 4 - Oxidation: The phosphite linkage between the first and second base needs to be stabilized by making the phosphate group pentavalent. This is achieved by adding iodine (go to local store) and water which leads to the oxidation of the phosphite into phosphate. This step can be substituted with a sulphorylation step for thiophosphate nucleotides.

(Note: this might be a good document to see how phosphoramidites can be ordered from suppliers.) Here are some oligo synth protocols in molecbio. Quantifying oligos from phosphoramadite synth. Note that you may not have to actually purchase phosphoramadites to start off with, but instead begin with a purified solution of nucleic acid??

What are the origins of oligonucleotide impurities and errors?

See here.

Microfluidics MiniFAQ

What are microfluidics?

Wikipedia sez: Microfluidics deals with the behavior, precise control and manipulation of fluids that are geometrically constrained to a small, typically sub-millimeter, scale. Typically, micro means one of the following features: small volumes(nl, pl, fl); small size; low energy consumption; effects of the micro domain (i.e., laminar flows, surface tension, diffusion, Marangoli forces, capillary forces, ...).

Even more on 'what are microfluidics'

See also:

• Microfluidics
• the original sharpie microfluidics post to diybio
• the rest of the thread.
• DIY microfluidics for continuous liquid flow using toner transfer (How To)

An example of microfluidics

The following is a run of the example microfluidics T-junction simulation in elmer, an open source CFD/FEM/FEA package. What you see here is the progression of an analyte due to electro-osmotic flow. There are two electric fields, three boundary conditions and a lot of wasted hours playing around with ElmerGUI and ElmerFront.

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What is a lab on a chip (LOC)?

A lab-on-a-chip (LOC) is a device that integrates one or several laboratory functions on a single chip of only millimeters to a few square centimeters in size. LOCs deal with the handling of extremely small fluid volumes down to less than pico liters. Lab-on-a-chip devices are a subset of MEMS devices and often indicated by "Micro Total Analysis Systems" (µTAS) as well. Microfluidics is a broader term that describes also mechanical flow control devices like pumps and valves or sensors like flowmeters and viscometers. However, strictly regarded "Lab-on-a-Chip" indicates generally the scaling of single or multiple lab processes down to chip-format, whereas "µTAS" is dedicated to the integration of the total sequence of lab processes to perform chemical analysis. The term "Lab-on-a-Chip" was introduced later on when it turned out that µTAS technologies were more widely applicable than only for analysis purposes.

Ultimately the idea is to have all of the typical components, procedures and processes of a laboratory available on a "chip", on a single perhaps disposable device, rather than having to build or purchase bulky equipment that sometimes tends to be hard to acquire or learn about.