We have had a rush on time and money saving techniques on Bitesize Bio in the last few weeks. Ways to re-cycle electroporation cuvettes, reduce gel buffer costs, do fast restriction digests and re-cycle midiprep columns have all been suggested.

In this article I’ll add the possibility of using a microwave to sterilize or decontaminate growth media. From the outset I’d like to say that I am not too sure about this, but I’ll make the case and you can tell me what you think.

Normally, growth medium is sterilized or decontaminated using an autoclave. Autoclaves are generally expensive, energy-hungry beasts that (in my experience) break down a lot so I would be very happy to use them less if I could.

Decontamination using microwaves.

The case for using microwave ovens for decontamination of cultures or materials was made back in 1977 by Latimer and Matsen. They showed that 1-5 minutes in a conventional microwave was sufficient to decontaminate 5mL cultures or petri dishes of common clinical pathogens including E. coli, S. aureus and K. pneumoniae. B. subtilis spores proved a bit more subborn, requiring more than 10 minutes of microwaves to wipe them out.

Border and Rice-Spearman backed this up with a 1999 study that showed materials contaminated with various bacteria and yeast strains were completely decontaminated by one minute in the microwave (I guess their microwave was better). And in 2006, Silva et al, investigating the decontamination of dentures, showed that 6 minutes in the microwave sterilised S. aureus and C. albicans but only partially disinfected P. aeruginosa and B. subtilis.

Sterilisation using microwaves

A 2001 Biotechniques paper by Weiss and Galande showed that LB plates made from microwave-sterilised LB-agar were apparently sterile (control plates were no detectably contaminated by microorganisms), had a similar shelf-life to autoclaved plates and supported bacterial growth as normal. The plates were prepared from dry powders dissolved in distilled water and aliquoted into 50mL tubes.

This is a very fast way to make plates and has the added advantage that the antibiotics can be added in from the start as they are not destroyed by microwaves. Invitrogen have a product that takes advantage of this. ImMedia is LB medium provided as sachets of dried, weighed power containing all of the required media components (including antibiotics). It is designed so that you can just add the sachet contents to water, microwave and your media is ready. But Weiss and Galande’s method is just as good and much cheaper.

My view

My take-home from this is that microwaves are are reasonably good at decontamination but more stubborn microorganisms (e.g. the spore-forming B subtilis) are not effectively disinfected. So the method does not sound too reliable to me. Also, filling the lab with smelly fumes from contaminated stocks does not seem to be a good idea. For easy liquid culture decontamination I think I will stick with Virkon, and for solid media, autoclaving seems to be the only good option.

The microwave media prep method is certainly interesting. Weiss and Galande’s results seem to be pretty robust and I would consider this method for an emergency media prep - if I need to start an E. coli culture last thing at night and there’s no sterile media available. Although the decontamination results show that microwaves don’t kill everything, E. coli grows so quickly that for routine purposes, a low level of contamination by slower growing organisms can be tolerated.

But I would not use this method for anything other than routine cultures and certainly not for slow growing organisms. Maybe that’s just me being a typical scientist, reluctant to take on new methods as Liam suggested. The microwave method is also limited by the fact that only small volumes (50ml) can be sterilised so it is never going to replace the autoclave for batch media production.

That’s my view - what’s yours?

If you have ever worked out the price of an electroporation cuvette you will realise that, at several dollars each, they are worth recycling.

Accounts on how amenable electroporation cuvettes are to recycling vary, but I find that as long as you treat them well it is possible to use single cuvette many times.

It’s the metal parts of the cuvette you need to worry about the most - you need to get them clean of DNA and cells and dry again quickly to prevent corrosion.

So the key is to wash and dry as soon as possible after transformation. Read more »

I often wonder why it is that molecular biology researchers stubbornly refuse to change 40 year old methods that, while they work, are not as good as newer, faster and cheaper methods out there.

I suppose rational scientists often have irrational superstitions.

One example of an old method that could be improved is the growth media used for plasmid preparation.

The majority of us, throughout our university careers, have used either SOC, LB or TB, for recombinant plasmid propagation, typically in E. coli. LB or Luria-Bertani broth has been in use for almost 60 years or thereabouts, while SOC has certainly been in use for 2 decades.

But by adding in a few more ingredients or being more economical on others (especially yeast extract and tryptone) that you could get a higher plasmid yield, quicker and with less money. Read more »

Here’s a tip that you may find useful if you are expressing proteins in E.coli using a lac promoter-based expression system, e.g. pET, in LB medium (L-broth).

Lac expression systems are typically induced in the lab using IPTG (isopropyl-beta-D-thiogalacto- pyranoside), which is a non- hydrolysable analogue of lactose, the natural inducer of the lac operon.

Tight control of expression from the lac promoter, which is required if the protein being expressed is toxic to the E.coli host or for a variety of other reasons, is not possible when using LB because it contains lactose.

But how does lactose get into LB? Read more »

You know those ridiculously priced and throw-away DNA mini, midi and maxi-prep columns? Well the good news is that you can actually re-use them if you are reasonably careful at regenerating them, with this simple and cheap method described in detail by Nagadenahalli B. Siddappa in Biotechniques in 2007.

Apparently these columns can be reused up to 20 times… perhaps more a guesstimate than a real number, but hey, who’s complaining?

Read more »

Today I was browsing through the “new technologies” section on the Biocompare website. Apart from the amazing but super-expensive automation equipment that most of us unfortunately have little chance of getting our hands on (at least at the moment), five products caught my eye as being useful for improving techniques widely used by researchers. I hope you find some, or all, of them of interest to you… Read more »

All over the world, molecular biologists are tragically wasting hours of their life running DNA gels using tris-based conduction buffers like TBE or TAE.

These buffers are known to overheat at high voltages, causing problems with gel integrity, sample denaturation and more. Because of this, molecular biologists are forced to keep the voltage of their gels to a maximum of 5-10 volts/cm (e.g 100 volts for a 10 cm gel) and extend the running time, sometimes to hours.

Although long gel runs, like long restriction digests, are often used as a convenient coffee break opportunity they can also eat into the molecular biologist’s precious time, leading to longer and less efficient working days.

But, in 2004, a team of scientists from Johns Hopkins came up with solutions (pardon the pun) to this problem. They have developed and verified three conductive buffers that stay cool during electrophoresis, allowing the voltage to be racked up to a massive 35 volts/cm without any problem, reducing the time taken to run gels by up to 7 times. Read more »

Problems with DNA gel extraction can be a real show-stopper since this is such a routinely used procedure. But, even if you are having no particular problems, it’s always nice to try and pick up some information that might improve your technique just that little bit.

Probably for these very reasons, Suzanne’s article 10 Tips for better DNA Gel extraction proved very popular. It seems like many of us are keen to get all the tips we can on this procedure. Well, if it’s tips you are looking for, we are always happy to oblige.

By scouring the recesses of my brain, colleagues and the internet I have squeezed out 5 more tips on DNA gel extraction. Maybe one of them will make the difference for you. Read more »

For identifying positive clones from a plasmid cloning procedure, the routine of performing a mini-prep and then checking the putative clones by restriction digestion is most commonly used.

Of course, if you need to screen a large number of clones, another option is a colony PCR to identify positives, followed by restriction digests to confirm.

However, there is a quick & dirty screening method that is even faster than colony PCR.

Called colony cracking, it involves picking a colony straight into lysis buffer, running the extract straight onto an agarose gel, and then identifying positive clones based the electrophoretic mobility difference between the super coiled DNA with and without insert - plasmids with inserts will travel slower than plasmids without them (no-insert control).

Obviously, the larger the insert, the easier it will be to distinguish between positive and negative clones, but researchers claim to be able to detect inserts of only 200bp (see the alternate protocol links at the bottom of the page).

If you’d like to try this method, read further. Read more »

For routine procedures involving cell lysis, it’s good for the lysis to be… routine. Of course there are many good and freely available lysis buffer recipes but for convenience and reproducibility you can’t beat pre-made lysis buffers.

Focusing on lysis for protein extraction, here are some of the reagents available for fast and efficient lysis of some of the most common cell types you might be using. Read more »