The centrifuge is an instrument used in nearly every biomedical research lab across the globe.
Centrifugation is a process by which a centrifuge is used to separate components of a complex mixture.
By spinning laboratory samples at very high speeds, the components of a given mixture are subjected to centrifugal force, which causes more dense particles to migrate away from the axis of rotation and lighter ones to move toward it. These particles can sediment at the bottom of the tube into what’s known as a pellet, and this isolated specimen, or the remaining solution, the supernatant, can be further processed or analyzed.
The principle component of a centrifuge is the rotor, which is the moving part that spins at high speeds.
Rotors can be fixed in position, or a centrifuge can use multiple rotors fixed atop part called the spindle.
Usually, the centrifuge rotor will have a lid that is screwed down tightly to prevent samples from flying out.
Many centrifuges will have a refrigeration unit that allows the internal temperature to be controlled during the spin.
They also have knobs or buttons for inputting the parameters for each run, which can include the duration of the spin, the temperature, and the magnitude of the spin in terms of speed or relative force.
Spin speed is measured as RPM, or revolutions per minute. RPM is tricky value, because it’s not the speed of the centrifuge that causes particles to separate from a mixture, but the force acting on these particles.
The force acting on a particle is related to the radius of the centrifuge rotor, and since different centrifuges have different rotor sizes, different forces can be applied at the same speed, or RPM.
Centrifugation strength can also be quantified as Relative centrifugal force, or RCF. RCF is generally presented as a multiple of earth’s gravitational acceleration.
RCF is expressed as the product of the radius of the rotor and the square of the angular velocity divided by earth’s gravitational acceleration.
RCF can be related to RPM by the following equation, where r stands for the radius of the centrifuge measured in centimeters.
Using this equation can mean the difference between a failed or successful experiment, but you don’t have to apply this calculation for every new procedure. Centrifuges often come with nomograms that can help convert RCF to RPM quite easily. Use a ruler to connect the radius of the centrifuge and a given RPM value, in order to obtain RCF.
To begin spinning your samples consider temperature. If you are using a refrigerated centrifuge, then you will want the machines internal temperature to reach the desired value before starting the spin, or you can find other ways to cool the rotor.
Immediately before a spin ensure all of the caps on your tubes are tightened and secure.
When you loading your tubes, ensure that each sample is counterbalanced with another sample directly across from it.
If you’ve only got one tube, then make another tube that can act as a counterweight.
If you’ve got three tubes, you can arrange them in a triangle so they are equidistant from each other.
Balancing weights in a centrifuge is critical. Centrifuge rotors reach high speeds and have a lot of kinetic energy. If improperly balanced, the entire centrifuge unit can be propelled from its resting place and do serious damage.
Once you confirm that the rotor and lid are secure, start the centrifuge and hang around until it’s reached the desired speed. If you notice a problem call and experienced member of the lab to come help you.
When your centrifugation is complete, you should be able to see your biological specimen at the bottom of the tube in a pellet, which has separated from the rest of the solution, or supernatant.
The supernatant can be removed by either decanting it – a fancy name for pouring it off, or it can be aspirated – a fancy term for using suction to remove it.
The purified specimen can then be returned to a solution via a process called, resuspending. Repetitions of centrifuging, or spinning cells, followed by aspirating cells, and resuspending in buffer, is often referred to as, washing cells.
Now that you’ve seen some basics of centrifugation, its time to have a look at some of the types of centrifuges out there and the procedures you can carry out with them.
Fixed angle rotor centrifuges are probably the most common type of this instrument you’ll encounter in the lab. Many table top centrifuges fit in this category.
These centrifuges, in which tubes sit in a fixed and angled position, are used in differential centrifugation protocols. In these protocols, a series of centrifugations at different speeds can be used to purify biological specimens like animal cells. Typically, these protocols involve several cell washing steps.
In contrast to fixed angle rotors, swing rotors have flexible tube holders allow samples to rotate outward. These rotors are beneficial in applications like density gradient centrifugation, where biological samples migrate to distinct layers of gradient media. This type of centrifugation is useful for quickly isolating one cell type from another, or for isolating individual organelles.
Lastly, the ultracentrifuge is the big brother of all the centrifuges you’ll find in the lab. It can spin in excess of 70,000 rpm, which makes it well suited for the isolation of small particles, like DNA or viruses.
Because of the high speeds of this centrifuge extra care should be taken to ensure that the loads are properly balanced and that both the rotor and lid are secure.
You’ve just watched JoVE’s introduction to Centrifugation. In this video we reviewed: what a centrifuge is and how it works, how to operate and run a centrifuge, some safety precautions, and different applications of your centrifugation. Thanks for watching and remember to balance your tubes.