Säkert användande av centrifuger

Centrifugering är en teknik som hjälper till att separera blandningar genom att tillämpa centrifugalkraft. En centrifug är en anordning, vanligtvis drivs av en elektrisk motor, som sätter en object, t.ex. en rotor, i en rotationsrörelse runt en fast axel.

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Basics in Centrifugation

Centrifugation is a technique that helps to separate mixtures by applying centrifugal force. A centrifuge is a device, generally driven by an electric motor, that puts an object, e.g., a rotor, in a rotational movement around a fixed axis.

A centrifuge works by using the principle of sedimentation: Under the influence of gravitational force (g-force), substances separate according to their density. Different types of separation are known, including isopycnic, ultrafiltration, density gradient, phase separation, and pelleting.

Pelleting is the most common application for centrifuges. Here, particles are concentrated as a pellet at the bottom of the centrifuge tube and separated from the remaining solution, called supernatant. During phase separation, chemicals are converted from a matrix or an aqueous medium to a solvent (for additional chemical or molecular biological analysis). In ultrafiltration, macromolecules are purified, separated, and concentrated by using a membrane. Isopycnic centrifugation is carried out using a ”self-generating” density gradient established through equilibrium sedimentation. This method concentrates the analysis matches with those of the surrounding solution. Protocols for centrifugation typically specify the relative centrifugal force (rcf) and the degree of acceleration in multiples of g (g-force). Working with the rotational speed, such as revolutions per minute (rpm), is rather imprecise.

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Important definitions

In general, applications for centrifugation specify the degree of acceleration to be applied to the sample rather than specifying a specific rotational speed such as revolutions per minute. The acceleration is typically given in gravity [× g] (or multiples of x g or g-force), the standard acceleration value due to gravity at the Earth’s surface (9.81 m/s²). The distinction between rpm and rcf is important, as two rotors with different diameters running at the same rotational speed (rpm) will result in different accelerations (rcf).

Why?

As the motion of the rotor is circular, the acceleration force is calculated as the product of the radius and the square of the angular velocity. Historically known as “relative centrifugal force” (rcf), this is the measurement of the acceleration applied to a sample within a circular movement. This process is measured in units of gravity
(× g).

Example*

	Rotor A	Rotor B
Speed	14,000 rpm	14,000 rpm
Radius	5.98 cm	9.50 cm
Gravity	13,100 × g	20,817 × g

*using the formula above

As mentioned, when using
rotors with different radii for centrifugation, the same rcf
(g-force) should be used.

Labteamet_csm_Figure-2_Relative-centrifugal-force-f_

Both centrifuges can spin a rotor with 1.5/ 2 mL tubes at the same speed (14,000 rpm) but the acceleration applied to the samples is very different: 13,100 × g versus 20,817 × g, resulting in different results. To make life easier and to better reproduce the data, some centrifuges have buttons directly on the operating panel for automatic conversion between rpm and rcf. If your centrifuge does not have an rpm-rcf converter, you may use the formula, the rpm-rcf converter found on the homepages of centrifuge suppliers, or a nomogram for conversion. The k-factor is a parameter for the sedimentation distance in a test tube. This factor is also called clearing factor and represents the relative pelleting efficiency of a centrifugation system at maximum rotational speed. In general, the k-factor value is used to estimate the time, t (in hours), required for complete sedimentation of a sample fraction with a known sedimentation coefficient measured in s (svedberg).

A small k-factor represents a more rapid separation. The value of the k-factor is primarily determined by the rotor diameter. Compared to rpm/rcf, the usage of the k-factor has become less important for general centrifugation processes. Especially for ultracentrifugation, the k-factor is still relevant.

How to select the right centrifuge for your application

If you follow a given protocol, make sure to use the same type of rotor and apply the given relative centrifugal force (rcf) as well as the same temperature and running time. In general, the following major parameters have to be determined for a successful centrifugation run:

A: Type of sample

B: Vessel selection

C: Type of centrifuge

D: Type of rotor

E: Determination of desired relative centrifugal force

F: Defined temperature during centrifugation

Fixed-angle or swing-bucket rotors

The most common rotors in laboratory centrifugation are either fixed-angle or swing-bucket rotors. Only a few applications require special rotors such as continuous-flow rotors, drum rotors, and the like. Flow-through rotors enable continuous flow collection of precipitates. These systems are used, e.g., in harvesting fermenters or for juice production in the food industry. Special customized versions, optimized for the specific application, are necessary.

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Fixed-angle rotor

The obvious advantage is the lack of moving parts in the rotor. This results in lower metal stress (longer lifetime), a higher maximum g-force is possible and for many applications, faster centrifugation times can be realized. The limited capacity (less flexibility) of the fixed-angle rotor is the only drawback. The position of the pellet strongly depends on the angle of the tube, it is located from the side to the bottom of the tube when spinning. Most rotors have a 45° angle for the tubes. The larger the angle for the tubes, the tighter the pellet. Smaller rotor angles result in more spread out pellet areas.

Swing-bucket rotor

This kind of rotor is highly flexible for using different tube formats, including SBS-format plates, based on a broad range of adapter systems and a high sample capacity. The moving swing-bucket parts result in increased metal stress for the rotor and the buckets as the bucket weight places a load on the two pivots and grooves. Compared with a fixed-angle rotor, therefore, a swing-bucket rotor is limited to a lower maximum g-force, which leads to longer centrifugation times. Based on the swing-bucket principle, the pellet is located in the bottom of the tube (horizontal position of tube during the run). The recovery by the user is facilitated compared to pellets located at the side of the tube.

The centrifuge

Floor-standing centrifuges

Floor-standing centrifuges free up bench space but do need at least one square meter of lab floor space. They are a good choice for high-speed or high-capacity protocols. Among floor-standing centrifuges, choices include ultracentrifuges, super-speed centrifuges, and low-speed centrifuges. An ultracentrifuge is a device for exceptionally high speed. These refrigerated centrifuges have an evacuated chamber to enable a rotational speed of up to 150,000 rpm. The g-force is about 300,000 to 1,000,000 × g. Special vessels that are placed within the rotor or attached to a special rotor are necessary. When g-forces of 40,000 to 60,000 × g are needed, super-speed floor-standing centrifuges are to be used. Low-speed floor-standing devices are generally used for applications like cell culture or blood with less than 10,000 rcf as the maximum g-force.

Bench-top centrifuges

Bench-top centrifuges are available in different sizes:

Microcentrifuges

Microcentrifuges are optimized for low-volume tubes, have a small footprint, and provide 14,000 to 30,000 × g for up to 48 microtubes. Some devices can even be used for a few 15 mL or 50 mL conical tubes or 2 SBS-format plates. Many suppliers offer non-refrigerated and refrigerated versions and different sizes of devices based on their tube capacity.

Multipurpose centrifuges

Offering a bigger rotor chamber, multipurpose centrifuges allow a broad range of rotors to be used (highly versatile). In addition to a flexible rotor system, specific adapter systems enable use of a wide variety of different kinds of tubes and bottles (from 0.2 mL to 1,000 mL) as well as plates. The maximum speed heavily depends on the vessel’s characteristics.

Centrifuge Safety

Safe operation

Imbalances

What causes an imbalance?

Rotors spin very rapidly and generate extreme forces. It’s therefore crucial to properly balance rotors during runs, especially when rotors are only partially loaded with tubes or plates. Balancing is always important (in order to not decrease the lifetime of the rotor), but especially so when centrifuging at higher speeds. Despite your efforts, however, imbalance errors caused by unbalanced sample loads can occur.

What risks do I face when exposed to an imbalanced load?

Incorrect loading can reduce the lifetime of the rotor, and uncontrolled, heavy vibration can lead to permanently damaging the centrifuge. More importantly, however, an imbalanced load can injure you or someone else. In the worst case, an imbalance can lead to a rotor crash.

Does my centrifuge realize the load is imbalanced?

Many centrifuges have auto-imbalance detection and will decelerate or automatically shut off if they sense an excessively unbalanced load (sensors are specifically built into the centrifuge for this purpose). Mostly bigger models (large benchtop and floor-standing centrifuges as well as ultracentrifuges) have this option. Smaller benchtop models, on the other hand, do not create strong enough forces to cause harmful imbalances; with these models, you would just notice a slight vibration and/or a higher noise level. Be aware that auto-imbalance detection does not automatically compensate for an unbalanced load.

What do I have to do if an imbalance error occurs?

If the centrifuge begins to shake or wobble, it is off-balance and you should stop it immediately. A little vibration is normal, but excessive amounts can mean danger. Once you have stopped the centrifuge, first double check to see if you have correctly balanced the tubes or plates in the rotor or buckets. If they are correctly balanced and the wobbling still occurs, contact the manufacturer or dealer to get the unit serviced. Do not continue to run a centrifuge that visibly wobbles when the rotor is spinning.

Labteamet_csm_Illustration_SH_Beladung_01

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How do I avoid centrifuge imbalances in the first place?

Ensure that your work surface is level and firm. Do not use the centrifuge on an uneven or slanted work surface.

At high speeds, a centrifuge can easily become unbalanced if equal masses are not located opposite each other in the rotor:

For fixed-angle rotors, balance your tubes according to their weight. Load the rotor symmetrically and ensure the opposing tube is not only the same type of tube, but that it is also filled with the same mass. If the number of tubes with samples is uneven, counterbalance using water in an additional tube. Remember to balance the mass (weight) of the tubes, not the volume (size). Weigh the tube with your sample and record the mass. If you are spinning more than two tubes, only the tubes directly opposite each other have to be equal in mass.

For swing-out rotors, always load all rotor positions with buckets (incomplete loading of the rotor may reduce the lifetime of the rotor). The weight of the maximum load or maximum weight of the completely loaded bucket is specified (weight class) on the buckets. Do not exceed this weight. When loading the buckets, make sure the tubes or plates are placed symmetrically.

Always check to see that the buckets swing out smoothly. If they do not, clean the pivots and grooves and apply grease.

Loading

Make sure the tubes you’re using have been specified for use with your centrifuge rotor. If necessary, support the tubes with adapters and check to see that the tube length allows the buckets to swing out to a horizontal position. When using aerosol-tight buckets, check to see that the tube length fits with the lid. Some centrifuge suppliers offer special rotors dedicated for use in the centrifugation of spin columns without risk of tearing off the tube lids. Using a different rotor than recommended by the manufacturer can easily lead to spills within the centrifuge, which may result in the formation of aerosol. This can be dangerous to your health and the environment.

Inspect centrifuge bottles and tubes or plates for cracks before use. Because a centrifuge can spin at such high speeds, a liquid sample can easily become an aerosol if it is not properly contained. Cracked tubes can fracture at high speeds or, at the very least, leak the sample into the rotor. As a last step, be sure to securely fasten the rotor lid provided with the centrifuge or the aerosol-tight cap of the buckets.

Labteamet_csm_Illustration_SH_Beladung_02_RZ

Load swing-bucket rotor evenly

When working with a swing-bucket rotor that has not been completely loaded, position the tubes within the buckets in a pattern that results in the rotor pivots being evenly stressed.

Clearly label all your tubes for identification. You may know how you placed your tubes in the centrifuge before they started spinning, but at the end of the spin you will not be able to tell them apart. If you have a tube you’re using for balance, be sure you label it as such. It is best to label the tube directly instead of using a sticker. A sticker can fall off during the spin, making identification difficult.
Wipe the outside of the tube with disinfectant before placing it in the centrifuge. Wiping the tube is particularly important if you are working with a biohazardous material. You want to limit any possible spills or aerosol formation with your sample. The best prevention is to wipe down the tube with a proper disinfectant before the spin begins.
Ensure you have not exceeded the maximum filling quantity for the sample containers specified by the manufacturer.
Fill the sample containers outside the centrifuge. No liquid should be allowed to enter the centrifuging chamber during filling.

Labteamet_DW_14-07-24Wurzbach2046_Filter_rgb_klein

Label your tubes for identification

Load rotor symmetrically

For swing-out rotors, all buckets should be of equal weight when loaded. For fixed angle rotors, balance your tubes according to their weight. The rotor must be loaded symmetrically: This means opposing tubes should be the same type of tube and filled with the same mass. If you are spinning more than two tubes, only the tubes directly opposite each other have to be equal in mass. At high speeds, a centrifuge can easily become unbalanced if equal masses aren’t located opposite each other in the rotor.

Labteamet_csm_Illustration_SH_Beladung_01_RZ

Automatic rotor recognition

Many centrifuges (especially multipurpose benchtop centrifuges) have automatic rotor recognition. This feature detects a newly inserted rotor and automatically limits g-force (rcf)/speed (rpm) to the rotor’s maximum permissible value. In other words, automatic rotor recognition prevents a rotor from being used at a higher speed than it was designed for. However, smaller centrifuges with a limited selection of rotors, in particular, and similar types of rotors often do not have rotor recognition. In these cases, all rotors have been designed to work at the maximum speed of the centrifuge. The Eppendorf Centrifuge 5424 R is one example of this type of centrifuge: All of its rotors are designed to spin up to 15,000 rpm, thus no automatic rotor recognition is needed for this centrifuge.

Handling of Hazardous Material

Working in clinical diagnostic laboratories usually means working with potentially infectious samples like blood or other bodily fluids. But handling infectious microorganisms or harmful chemicals is quite common in research laboratories as well. To ensure the safety of laboratory personnel and to prevent laboratory acquired infections (LAIs) or other health hazards, reasonable precautions must be taken throughout the whole workflow.

Instead of examining the entire flow of work now, we will concentrate on the centrifugation step of infectious and hazardous samples as the dangers here are often quite underestimated. Statistics show that about 80% of all LAIs are unsuspected (aerosols). For further information, please see the additional literature listed at the end of this text.

Routes of laboratory acquired infections

Individuals who work with potentially infectious samples face many risks in both clinical diagnostic laboratories and in research laboratories. Because exposure to harmful or infectious substances occurs more often than expected, raising awareness about the risks and sources of LAIs as well as about the safety precautions lab workers need to take is very important. Certainly not every exposure leads to an infection, but the less the exposure, the lower the risk for acquiring an LAI.

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The most significant routes of laboratory acquired infections are:

Spills and splashes on skin and mucous membranes as well as contact with spills and droplets on surfaces (dermal contact)
Ingestion through mouth pipetting or touching the mouth (or eyes) with fingers or contaminated work equipment
Inhalation of infectious aerosols

Centrifuges as one source of aerosols

Due to their mode of operation, centrifuges can present a high risk if not operated correctly. To maintain a desired temperature, air-cooled and refrigerated centrifuges use an air ventilation system. This system discharges warm air from inside the centrifuge into the environment. Infectious agents present in the exhaust air will be dispersed quickly and broadly throughout the laboratory. Tube breakage poses an even greater risk, since this can produce a large amount of aerosol. The use of aerosol-tight lids or caps will reduce this risk significantly.

Taking precautions

To ensure lab workers are optimally protected, a couple of safety guidelines should be followed.

First, and very important, is factoring in human error:

Young, inexperienced workers are more likely to make errors; therefore, regular training and supervision are necessary.
A lack of mental alertness, high stress levels, and heavy workloads are factors that can lead to less careful behavior.
Suitable equipment must be provided and properly maintained; this includes both technical equipment and protective gear.

Labteamet_csm_Illustration_Safety_Equipment_RZ

For aerosol-tight centrifugation, both aerosol-tight caps or lids and suitable vessels must be used. In this regard, the following points need to be considered:

Material. Use nonbreakable materials such as PP, PE, and the like; avoid glass tubes.
Leak tightness. Make sure the vessel can be closed and remain leak-tight; use, for example, Eppendorf tubes.
Filling volume. Every vessel has a maximum filling volume, usually two-thirds or 80% of the total volume (called the “working volume”) that shouldn’t be exceeded; this ensures the liquid in the vessel does not touch the vessel lid.
Droplets. These can occur in the thread of the vessel lid or even on the outside of the vessel. These droplets can become airborne during centrifugation and thereby form aerosols.

Labteamet_csm_Illustration_SH_Tube_ueberfuellt_RZ

As mentioned earlier, aerosol-tight caps (for swing-out buckets) or aerosol-tight lids (for fixed-angle rotors) are essential for centrifuging hazardous samples. The following points outline proper handling and maintenance of the equipment:

Loading and unloading. Aerosol-tight caps and lids do not prevent formation of aerosols during centrifugation; rather, they ensure aerosols cannot leak from the closed system. Therefore, you must wait at least 10 minutes before opening the bucket or rotor. This gives the aerosols time to settle down. Also, you should load and unload the buckets or rotor in a biosafety cabinet (especially in virology and mycobacteriology) to minimalize the risk of escaping aerosols.
Disinfection and autoclaving. Buckets, caps, rotors, and lids are the main components that come into contact with aerosols. Correctly handling them does prevent aerosols from being dispersed within the rest of the device, but the parts themselves do need to be disinfected after each use. This is done using suitable disinfectants and following to the manufacturer’s recommendations (usually 70% ethanol works fine for rotors and buckets). You can also regularly autoclave rotors, rotor lids, buckets, and caps (please refer to the operating manual for parameters; autoclaving is usually carried out for 20 minutes at 121°C, 2 bar).
Seals. All aerosol-tight caps and lids are delivered with rubber seals. Together, the caps and seals form an aerosol-tight unit, which must be tested and certified by an independent test institute (e.g., Public Health England, Porton Down, UK). You need to regularly check the seals to ensure they are intact, nonporous, and correctly seated in the grooves. If any of these factors are not present, aerosol-tightness cannot be guaranteed. If necessary, grease or even replace the rubber seal.

Using centrifuges in a safe way

The centrifuge itself is another factor to be considered in the safety workflow:

Speed limits. Exceeding the speed limits of rotors can lead to tube breakage or even a rotor crash, therefore the rotor speed limit must be considered when the centrifuge is not equipped with automatic rotor recognition (a feature that ensures the maximum speed is not exceeded).
Tube breakage. Should a tube break or leak, do not open the centrifuge for at least 30 minutes after the run. Since this cannot always be detected before you open the buckets or rotor (a sudden imbalance can be a first sign of tube breakage), we recommend waiting at least 10 minutes at all times before you open the containers.
Maintenance. Good maintenance and regular checks of the centrifuge, rotors, and equipment is necessary for ensuring safety and preventing system failure (e.g., rotor crash, which leads to a large amount of aerosol escaping with the centrifuge exhaust). Please refer to the manufacturer’s instructions for maintenance information or watch our centrifuge maintenance video.

Personal protective gear is another crucial factor in avoiding contact with infectious or harmful substances (mostly through ingestion and dermal contact) and the following should be heeded:

Protective gear. Always wear a laboratory coat, safety gloves, and safety goggles when working with infectious or hazardous substances. This will minimize the risk of dermal or mucous contact with the material (splatter, droplets, etc.)
Clean hands. After working with infectious material, remove the used gloves and disinfect your hands before washing them thoroughly.
Clean gear. Be sure personnel protective gear, including laboratory coats, safety goggles, and the like, are regularly cleaned; replace if damaged.

Including these precautions in the workflow will provide a higher level of security during centrifugation as well as minimize the risk of LAIs.

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