Fast, Accurate and Green PCR

Attending the 2009 American Society of Microbiology meeting this week in Philadelphia, I decided to take my own advice and visit the exhibits to see whether any of the exhibiting companies had any really innovative and unique products on display.

I focused mainly on products I might use in my own lab, such as PCR reagents and basic lab products. And there was one company that had something that really made me stop and say “wow!”. ----------- Suzanne

That company was Finnzymes, and the “wow” came from three products that are individually impressive but together make a superb PCR ensemble that is compact, versatile and offers excellent speed and accuracy.

A Compact Thermal Cycler…

First up was the Piko™ Thermal Cycler, which is the smallest thermal cycler. Amazingly, this tiny instrument was around the size of a vortex mixer; with a footprint of only 16cm wide by 17 cm deep and 23 cm high and a weight of only 4 kg. Check out the picture on the right for a comparison of footprint sizes for the Piko™ and a conventional thermal cycler.



Even better, the Piko™ uses a 1/4 of the power of a normal thermal cycler, so this is one way for our experiments to be a bit greener. And it offers signficantly reduced reaction times. More on that later.

The instrument ranges in price from $4500-$6000 depending on the blocks you purchase (there are several options), so it also costs significantly less than traditional thermal cyclers.

A thermal cycler that won’t hog your bench space ticks one box. But your PCR is only as good as your polymerase, and Finnzymes have thought of that too..


A Speedy, Accurate Polymerase…


David Unger, Managing Director of Finnzymes USA, explained that the power behind Phusion™ comes from the unique dsDNA binding domain fused to a Pyrococcus-like proofreading polymerase, which results in a very tight association of the enzyme with the template DNA.

“Taq polymerases work by moving on and off the template. This slows down the enzyme and leads to difficulty if inhibitors are present.

With Phusion™, the enzyme stays linked to the template and so has a faster processivity along with the ability to work in inhibiting conditions such as 20% whole blood in the PCR reaction.”

The enzyme is twice as fast as Taq (<15 second/kb extension time) and can amplify long templates, up to 20 kb with less enzyme. Just as importantly, the error rate is 50 fold lower than Taq and 6 fold lower than standard Pyrococcus furiosus DNA Polymerases.

Sounds great (and it is). Together, Piko™ and Phusion™ offer fast, accurate and economical PCR.

But throw in Finnzymes’ patent pending UTW™ (Ultra Thin-wall) tubes and plates and the whole thing gets a lot more powerful.

96 x 5ul Reactions in 10 minutes…

The UTW™ PCR plates are, as the name suggests, ultra thin, which dramatically cuts down on the time taken to heat and cool the samples.

Piko™ is available with with a 24-well block for UTW™ 0.2 ml tubes, but for maximum output the instrument can be equipped to take Finnzyme’s 96-well UTW™ Piko™ PCR Plates.

These plates are about the size of a microscope slide and Piko™ can take up to 4 of them to make the equivalent of a 384-well plate for easy up and downstream processing with other lab equipment.

Using Phusion™, your 384 x 5 ul reactions can be completed in just 10 minutes flat. Now that’s bench-scale high throughput.

At that speed and size, even the smallest lab can forget about PCR backlogs or downtime.

Courtesy: Suzanne, BitesizeBio

RPM Does Not Equal RCF

RPM and RCF are two units that can be used to describe the speed of a centrifuge. Although they may look similar, they are oh-so-different and confusing them has resulted a disastrous end to many an experiment.

So let’s set it out in black and white to make sure you don’t succumb to the same mistakes as those who have passed through the labs of the world before you.

How Centrifuges Work

Centrifuges work by putting your samples in rotation around a fixed axis, thereby applying an accelerative force perpendicular to the axis as shown in the diagram on the right.

And of course, this force causes particles in your sample to accelerate toward the outer edge of the rotor. Conveniently, centrifuge manufacturers design their centrifuges so that the bottom of the sample tubes are at the outer edge of the rotor so the centrifugal force results in the sedimentation of particles in your sample to the bottom of of the tube.


The amount of force required to move a particle depends on the size of the particle. Large particles (e.g. cells) require less force than small particles (e.g. precipitated proteins). For more detailed info on this, take a look at these Wikipedia articles on sedimentation and the Lamm equation.

Describing The Force

Relative Centrifugal Force (RCF) is the term used to describe the amount of accelerative force applied to a sample in a centrifuge. RCF is measured in multiples of the standard acceleration due to gravity at the Earth’s surface (x g). This is why RCF and “x g” are used interchangeably in centrifugation protocols.

The two variables that describe RCF are the radius and the angular velocity of the rotor. i.e. how wide the rotor is and how fast it is moving.

If the rotational speed is given in revolutions per minute (RPM) and the radius is expressed in centimetres (cm), then:



RCF or RPM?

From this it is clear that the correct unit for the amount of centrifugation to use in a given protocol is RCF (a.k.a. x g).

But most centrifuges, especially microcentrifuges, only have settings for RPM. So unless you are lucky enough to have a centrifuge with an RCF setting, you will have to work out the corresponding RPM that will be required for YOUR centrifuge to achieve the RCF set out in a protocol.

This can easily be done by taking your ruler, measuring the radius of your centrifuge rotor and plugging the numbers into an online converter such as this one from DJB Labcare. Alternatively, the same site also has a very useful nonograph that you can print out and keep on your bench to use for rcf to rpm conversions.

The take home message is that centrifugation speeds in quoted in RPM will only be constant for centrifuges with the same rotor radii. If you use an RPM setting from a protocol where someone used a centrifuge with a different radius from yours, you will get a different RCF. Often the difference will not be significant enough to affect the sample or the centrifuge, but sometimes, e.g. if you use an RPM setting that was originally meant for a microfuge in an ultracentrifuge, it can cause problems.

Courtesy: Nick, BitesizeBio

Southern, northern, western (and eastern?)

It’s official - biologists DO have a sense of humor, well some of them at least.
This is the story of how one of the most famous and quirky naming conventions in biology came into being. It’s a story of discovery, comedy and the triumph of people power over the establishment.
Read on to find out the story of how the Southern, northern and western (etc) blots got their names.

In 1975 when Ed Southern invented his method of using a radiolabeled DNA probe to detect a specific DNA sequence within a DNA sample (e.g. a fractionated genome) and named it after himself - the Southern blot - I’m sure that he had no idea about what he had started.
Two years later, J.C. Alwine, a biologist with a sense of humor, developed a technique analogous to the Southern blot, this time for the identification of a specific RNA within a complex RNA sample using a radio-labelled DNA probe. Alwine couldn’t resist the temptation to call his technique the northern blot in an allusion to Southern’s technique, raising a chuckles in labs everywhere.
Then W. Neal Burnette, a post-doc working in the Nowinski group at the Hutchinson Cancer Center in Seattle, started the real fun.
Burnette was searching for a way to combine the powers of radio immunoassay and SDS-PAGE electrophoresis so that he could pinpoint specific antigens in a complex protein mixture, such as a cell extract.
After some “laughably naive” attempts to visualise the interaction between antibodies and the separated proteins in the gels, he was inspired by Alwine’s nothern blot method (so indirectly by the Southern blot) to make a solid phase replica of the gel. So he developed the method of using electrophoresis to blot the protein onto nitrocellulose paper and after some further work, perfected the technique of blocking non-specific binding sites and visualising the specific radioimmunolabelled antigens using an X-Ray film.
In a historic, but mostly forgotten conversation with Nowinski, Burnette coined the name “western blot” for his technique. What fun. Like nothern blotting, “western blot” was also an allusion to the Southern and nothern techniques, but Burnette had upped the ante by throwing in a geographical reference to location of the Nowinski lab. So if the Nowinski lab had been in New York, we would all be doing “eastern” blots.
A quick aside for the pedants among us. Note that among these techniques, only the Southern blot should be capitalised since it refers to Southern’s name, the others - nothern, western etc - are not proper nouns, so should not be capitalised. Try pulling your boss up on that one next time he is in mid-flow talking about a “Northern blot” in a departmental presentation.
Anyway, back to our story. Unfortunately for Burnette no sooner had he perfected his technique than a paper describing a very similar method, also inspired by nothern blotting, was published by Towbin et al working at the Friedrich Miescher Institute in Switzerland.
Burnette was dejected, but nonetheless, convinced that his methodology was sufficiently different to Towbin’s, he decided to submit a manuscript on his western blot method to the Analytical Biochemistry journal.
The reviewers hated it, they hated the name even more - obviously humor was not high on their agenda - and the manuscript was rejected.
But despite this, the popularisation of Burnette’s technique, and particularly the name “western blot” still happened even without the assistance of the literary establishment. It happened through the sense of humor of the researchers who were doing the work, through people power (assisted by Xerox power).
It happened because researchers, besides being interested in the technique itself, were tickled enough by its quirky name to make copies and send it to their friends. In Burnette’s words…
“…the few preprints I had sent to colleagues seemed to have undergone logarithmic Xerox multiplication. I began receiving phone calls from researchers unable to read the umpteenth photocopied generation of the pre-print, a sort of technical samizdat that I had to endlessly interpret”
A few years later, Burnette eventually coaxed Analytical Biochemistry into accepting his paper and it was published in 1981, but by then, word of mouth had already beaten them to it. Ironically, considering the people power that was doubtless (at least partly) responsible for it’s eventual publication, Burnette’s paper is available only to Analytical Biochemistry subscribers. *end of open access rant*
Bowen and colleagues continued the naming convention in 1981 with their publication of the southwestern blot, a technique for identifying DNA-binding proteins in nuclear protein extracts using specific oligonucleotide probes. The “south” in the name refers to the use of DNA probes, while the “west” refers to the protein blot.
Interestingly, Bowen’s paper alludes to Burnette’s western blot even though it was published before Burnette’s paper, which shows just how strongly word-of-mouth actually publicised the western blot.
And in 1998, Ishikawa and Taki published their far-eastern blotting method, no doubt a reference to their geographical location, for the analysis of lipids by TLC separation followed by blotting onto a PDVF membrane.
Finally, there is one blot that deserves mention. Legend has it that Ethan Signer coined the phrase “eastern blot” for the tantric practice of willing a failed gel into show bands. Apparently, you take your blank gel, meditate, repeat the mantra, and the bands appear…
…if only!
If you’re a biologist with a sense of humor, join in by telling us about your favorite quirky naming conventions in the comments section.

Courtesy: Nick, BitesizeBio

Low cost DNA gel documentation

Equipment for photographing DNA gels stained with ethidium bromide (or other fluorescent dyes), doesn’t have to cost thousands of dollars. These days, great pictures can be obtained with a standard digital camera and an orange filter. Here’s how.

You will need:
• A digital camera
• A Cokin orange filter 002A or similar (This does not have to fit onto the camera, a square filter will do.)
• A polystyrene ice bucket with a thick (3-5cm bottom)
• A UV transilluminator

To build it:
1. Cut a hole in the bottom of the box that is big enough for the camera lens to fit into.
2. Tape the filter over the hole, inside the box.
3. Set the camera flash to off and the mode to black & white.
4. Push the camera’s lens tube into the hole (or place the camera lens over the hole if there is no lens tube). Depending on the shape of your camera, you may have to modify the box to ensure that the camera is pointing straight into the hole.
5. Place your DNA gel onto the transilluminator and put the box over the gel.
6. Turn the transilluminator and the camera on. You should now be able to see the stained DNA on the gel in the viewfinder of the camera.
7. To get a quality picture and good detection limits you will have to play around with the ISO, shutter speed and aperture. This will be specific to your camera, but the setting ISO 200, Shutter speed 1/3, Aperture 8.0 worked for me so may be a useful starting point for you.

Update: Here’s a schematic diagram that shows how the whole thing should fit together. Let me know if you have any comments or questions.




Courtesy: Nick, BitesizeBio