Guide to the Disruption of Biological Samples - 2012

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Part VII: Mechanical Disruption Methods/ Beating


Beating involves crushing a sample with a projectile.  Most beating methods rely on placing a sample and projectile in a tube and rapidly shaking them back and forth.  The most common method is bead beating which uses grinding balls or small beads, though cylinders and irregular shapes have also been used.  Bead beating has been in practice for years for the disruption of microorganisms, originally using small glass beads and dental amalgamators (i.e., the shakers that dentists use to mix up the components of metal fillings). 

Bead beating can be quite effective if done correctly, though traditionally it was a bottleneck due to the limited number of samples that could be processed.  Some labs took it upon themselves to increase the throughput by adapting paint shakers to process samples.  At times these were effective, but not totally satisfactory.  To remedy this limitation, several companies developed bead beaters (formerly called mixer mills) that could handle racks of tubes or even microwell plates. 

Though simple in theory, bead beating can be quite complex as several factors must work together to effectively homogenize samples.  The tube, or microwell plate, used in bead beating is not just a sample holder, but actively participates in the homogenization.  The projectiles, henceforth referred to as balls or beads, must also be carefully matched to the sample.  The stroke length of the amalgamator or mixer mill is also critical, as is the physical motion it imparts on the sample tube.  Finally, sample mass, volume, and extraction buffer all impact the effectiveness of the homogenization.

Tubes/Plates:  The objective of bead beating is to sandwich a sample between a grinding bead, or ball, and a hard surface.  That hard surface can be either another bead/ball, or in many cases, the tube or deep well plate being used for the processing.  With microorganisms and micron sized beads, the colliding surfaces are more likely two different beads.  When processing tissues or seeds, the collision is between the tube and ball.  Balls and beads, as will be discussed below, are typically hard and dense materials like glass, ceramics, or steel.  Consequently, to get a good homogenate the tube must also be hard to push back on the grinding ball.  A metal tube would be ideal, but in reality, metal tubes (though they have been used) are impractical because of their high cost.  The alternative to metal is plastic tubes.

Not any plastic is suitable for bead beating as some are very brittle and will not hold up to the repeated impact of the ball (which is why glass tubes are not used).  For example, polystyrene tubes, though widely used for labware, crack very readily.  Polypropylene is much more durable, being the material of choice for microfuge tubes and deep well plates.  It holds up well to bead beating especially when small beads are used for lysing bacteria.  However, when used with large grinding balls, polypropylene tends to soften as the sample warms.  This dampens the impact between the grinding ball and tube.  High density polyethylene works relatively well with grinding balls.  It also has the added feature of being compatible with many organic solvents used for the extraction of small molecules from tissues during pharmacokinetic analysis.

Polycarbonate is the best plastic for tubes and vials used for bead beating.  It is extremely durable, clear, and impact resistant even at cryogenic temperatures.  The major limitation with polycarbonate is that it is incompatible with several solvents commonly used for extraction including: phenol, chloroform, acetone, and acetonitrile.  Consequently, for dry grinding or extraction using aqueous buffers, polycarbonate vials yield the best homogenate.

The last point about tubes and plates regards the closures used to seal them.  If samples are processed with liquids, then the cap must have some type of gasket (an O-ring or seal) to prevent liquid from leaking out.  Special care needs to be observed if the extraction buffer is an organic solvent, such as methanol or acetonitrile, as the vapor pressure of the solvent can push the solvent (and the analytes) out of the tube if the seal is insufficient.  Polypropylene microfuge tubes are often used for organic extractions, and experience has shown that all brands of tubes do not seal equally.  This is the same for deep well plates, as push-on sealing mats vary greatly on their tightness of seal.  We have observed that tight fitting polypropylene sealing mats, whether round or square, tend to seal much better than mats made of PTFE.  Furthermore, self-adhesive sheets do not work well, and heat sealed sheets are only suitable for very small beads (e.g., <600 micron) when used with deep well plates.

Balls/Beads:  Not discussing the beads used for bead beating would be similar to teaching cooking without referring to pots and pans.  The beads (and grinding balls) used during the homogenization process are the most important component of the process, thus discussing what products are available and how to use them is important.  First, bead beating not only uses beads, but grinding balls, ceramic satellites, and grinding resins.  Our lab usually refers to any spherical media of 3 mm or less as a bead.  Above that size it miraculously becomes a grinding ball.  The larger media are normally made of stainless steel while the smaller beads are either silica (glass) or ceramic (zirconium silicate or zirconium oxide). 

Non-spherical media is very effective at disrupting tough fibrous samples, such as muscles, skin, and sclera.  On the small size, garnet, cut wire, silica gel, and similar sharp grinding media can effectively slice resilient samples, particularly when used in conjunction with a larger grinding ball.  Satellites (shaped like Saturn) and cylinders can also be effective breaking up tough samples due to the sharpness of their edges.  We routinely use a combination of garnet (a hard, sand-like mineral) and zirconium oxide satellites for tougher animal tissues.

It is important to mix and match grinding beads/balls and sample types in order to get effective disruption.  Table 4 summarizes some of the more common configurations for balls, beads, and tubes/vials.

Table 4.  Grinding media and their applications.





100 µm Beads


Microfuge Tubes

Deep Well Plates

Tubes or wells should 1/6 volume of beads and 1/3 volume of cell suspension.

200 µm Beads


Microfuge Tubes Deep Well Plates

Amounts as above.  Good for small yeasts (Pichia) and larger bacteria.

400 µm Beads


Microfuge Tubes, Deep Well Plates

Best size for Saccharomyces.  Beads and culture broth should not exceed half volume.

500 µm Garnet


Polycarbonate Vials (4 ml)

Abrasive for tearing.  For tough samples use with a 6 mm satellite.  Use amounts as above.

800 µm Beads


Microfuge Tubes, Deep Well Plates

Good for pollen, mycelia, and algae.  Volume should be the same as 100 µm beads.

1.0-1.7 mm Beads

Leaf Tissue/Soil

Microfuge Tubes


Process up to several leaf punches.  Do not overfill tubes with buffer.

2.8-3.0 mm Beads

Plant Materials

Microfuge Tubes

Mass of sample should be kept to under 50 mg and buffer less than 500 µl.

5/32” Grinding Balls

Plant/Animal Tissues/Insects

Deep Well Plates

Animal tissues should be <100 mg while plants should be <50 mg.  Use 1 ball per well.

6 mm ZO Satellites

Plant/Animal Tissues

Deep Well Plates,
Polycarbonate Vials (4 ml)

Use as above.  When stainless steel may interfere with assays, zirconium oxide can be used.

3/8” Grinding Balls

Seeds/Animal Tissues

Polycarbonate Vials (4 ml),
Polyethylene Vials (4 ml)

Use for rice grains, kernels, other seeds, on tissues (<150 mg).  Do not overfill vials.

7/16” Grinding Balls

Pooled Seeds/ Organs/Tissues

Polycarbonate Vials (15 ml),
Custom Container

Good for large samples and pooled seeds for field trials. Container must be relatively large.  Use 2 balls for 15 ml vials or more for custom containers (up to 125 ml).

Grinding media, whether beads, balls or satellites, can be purchased in bulk or pre-dispensed in grinding tubes.  Beads can be purchased in a raw form, but these need to be cleaned prior to use.  Raw beads contain numerous contaminants from dirt, to skin cells, to hair, to insect parts.  Fine micro-particles in silica beads can bind analytes and lower yields upon homogenization.

Processed beads can be purchased which are acid washed to remove contaminants.  Molecular biology grade beads are also available which have been washed, baked, and packaged aseptically.  These beads are also quality control tested.  For homogenizing samples with very few cells, as with environmental samples with low cell counts, low binding beads can be used.  These beads are chemically treated so that less analyte bind to the beads upon liberation during bead beating.  These have been shown effective for detecting low levels of bacteria by quantitative PCR, increasing both linearity and sensitivity of assays.  Consequently, low binding beads have found use in tools used for biological weapons detection.

Amalgamators: Dental mixers, or amalgamators, have been used or adapted to bead beat microorganisms for years.  This simple instrument allows a tube to be locked into a small shaking arm, which then oscillates rapidly.  When bacteria, yeast, or molds are added to the tube with grinding beads, the amalgamator effectively grinds the cells in as little as 30 seconds.

These bead beaters are effective and relatively low cost.  Several vortexers have been modified to provide similar actions (but yield poorer results), with the added value of being able to process multiple samples.

Strengths – These are effective homogenizers at a very reasonable price.  Vortexer units are available with a pulsing feature, which helps to reduce the effect of heat generated during homogenization.  However, it must be emphasized that vortexers used for bead beating are not as effective as true oscillating amalgamators.

Limitations – The individual tube bead beaters are rather effective, though throughput is low.  Some models can hold up to three tubes.  For labs running a few samples, such units might be adequate.  Vortexer units are less effective, but hold greater numbers of samples.  Depending upon the application, lower lysis efficiency may not matter.

Multitube Homogenizers:  The second generation bead beaters moved from single tube formats to homogenizers that can handle up to 24 microfuge tubes.  The first instrument in the category was the FastPrep®, which in many ways looks like a microtube centrifuge (a.k.a., microfuge).  Microfuge tubes placed in the round rack are oscillated at high speed (up to 8000 rpm) in very short vibratory-like motion.  A similar model is the Precellys®, which operates in much the same manner.  Variations on both machines have been made that allow for oversized microtubes (e.g., 7 ml) and even 15 and 50 ml conical tubes.

Strengths – The ability to process multiple samples make these machines very useful to the point where some of been used for clinical work.  For microorganisms these machines are effective.  For tissues, using course media such as garnet allows for many tissues to be shredded.

Limitations – The stroke or oscillating pattern of these machines are very small and thus they tend to work less effectively at homogenizing very resilient tissues, such as skin and sclera.  Samples also get very warm by the heat generated by the 8000 rpm motion.  Some machines have shutdown features due to overheating which frustrates many researchers.  Additionally, these units tend to be very expensive for their capacity, costing around $10,000.

High Throughput Homogenizers/Mixer Mills: With the rise of high throughput screening strategies, sample homogenization became the bottleneck.  To circumvent the logjam, various approaches were taken to homogenize samples en masse.  The simplicity and effectiveness of bead beating was applied to a larger format, resulting in high throughput homogenizers (also known as mixer mills).  These devices in the simplest configuration shake racks of tubes or microwell plates at speeds up to 1800 rpm.

Mixer mills are not new, but their adaptation to the SBS formats (i.e., Society of Biomolecular Screening standardized plate dimensions) is relatively recent as compared to other homogenization technologies.  Essentially a microwell plate, set of vials, or rack of tubes is locked into a moving platform.  Normally each well/tube has a sample with one or more grinding balls.  For processing, the homogenizer violently shakes the samples causing the grinding ball(s) to impact the samples against the tube walls.  Processing for 1-2 minutes thoroughly homogenizes most samples.  Similarly, microorganisms can be disrupted using beads.

Several brands of high throughput homogenizers are available, such as the Geno/Grinder®, HT Homogenizer®, and Retsch® Mixer Mill.  Though these can all hold plates and racks of tubes, they differ significantly in the method of operation.  The path or motion which the plates follow during processing is different for these homogenizers, with the Geno/Grinder® and HT Homogenizer® having a linear motion while the Retsch® Mixer Mill follows a figure “8” motion.  With linear motion homogenizers, all wells or tubes follow the same path, thus processing between samples is the same.  With figure “8” motion machines, like the originally modified paint shakers, wells on the outside of the plate (or tubes on the outside of a rack) follow a different path than samples towards the middle.  This leads to differential processing of samples and increased variation in analyte yield.  For analytical work, this could skew results based on the positioning of the sample during homogenization.  Indeed one user manual for a figure “8” homogenizer recommends flipping the plate halfway through processing to generate more uniform results.

High throughput homogenizers are versatile as they can be used for a wide array of sample types and sizes.  The most important aspect of resulting in good disruption from a high throughput homogenizer is to properly match the sample size (mass and volume) with a suitable well size and grinding media.  Generally, samples and buffers should take up no more than a third of the volume of the vessel.  Thus a leaf punch of 10 mg with 200 µl of extraction buffer can easily be processed in a 96 deep well plate (1.2 ml round well) using a standard 5/32” grinding ball.  Note that a round well was specified as square deep well plates afford samples to hide in the corners, avoiding homogenization.  Deep well plates should be used for small samples of 50 mg or less.  As sample size increases, polycarbonate vials are the next best choice for grinding.

Mouse muscle homogenized using a high throughput homogenizer.

Figure 13. High throughput homogenizer with deep well plate and 5/32” grinding ball produced very fragmented muscle and a relative disruption efficiency of 52% (see Fig. 20). (P)

Any impact resistant tube or container that can be locked into a high throughput homogenizer can be used for disrupting samples.  Indeed short 60 ml polycarbonate jars have been used to homogenize complete animal organ systems.  Most samples however are less than 1 gram.  When sample size is larger than 100 mg, a 24 well format is an effective format to use.  Polycarbonate vials (4 ml) are available in the SBS 24 well format, which allows for liquid handling of homogenate.  In this format, larger grinding balls (3/8”) are used.  Polycarbonate is the choice material to use for homogenization vials as it is hard and pushes back on grinding balls.  Though polypropylene vials will work, as with the deep well plates, it tends to soften as the tube heats with processing.  With harder samples, like seeds, the ball can wedge in the tube with the sample.  For samples that require organic solvents that melt polycarbonate, such as phenol and chloroform (including Trizol®), polyethylene vials are available that are solvent compatible.

High throughput homogenizers can be used for wet or dry grinding.  Wet grinding as primarily described above, is just that, homogenizing with solvent.  Dry grinding is popular in the analysis of seeds and plant materials.  In this approach, the seed is processed with the ball without solvent.  The key factor in dry grinding, especially seeds, is that the samples tend to be very hard and as such require a disproportionately large grinding ball.  For instance, a grain of rice is only about 20 mg, and easily fits in a well of a deep well plate, but the 5/32” grinding balls used in that format have insufficient mass to crack the rice.  To pulverize the rice requires a 4 ml polycarbonate vial with a 3/8” grinding ball.  A comparison of soybean and rice processed in deep well plates and 4 ml vials can be seen in Table 5.

Table 5. Comparison of Rice and Soy Processed in Deep Well Plates and Polycarbonate Vials.

Rice and soybean ground using a high throughput homogenizer.

Samples in plates were ground with 5/32” stainless steel grinding balls while larger 3/8” balls were used with 4 ml vials.

Many agricultural biotechnology labs analyze seeds from field trials (P) by pooling seeds prior to homogenizing.  To accomplish this, larger polycarbonate vials (15 ml) are available that can hold up to 15 corn kernels and numerous smaller seeds (several grams).  Using a larger vial allows for larger grinding balls, and in the case of corn, two 7/16” balls are used (P).  This process generates corn meal that can be used for a range of tests from starch composition to genetic analysis.  The larger vials reduce throughput using platform homogenizers, but it still is preferable to using coffee grinders which required cleaning after each sample.  For instance, in a laboratory using coffee grinders to process corn, throughput was increased from five samples per hour to 100 samples using high throughput methods.

These larger polycarbonate vials have found new application in biopharmaceutical labs and are now being used more extensively for high throughput analysis of animal tissues for pharmacokinetic analysis.  Residual drug levels are often assessed from dosed animals, a labor intensive process that requires homogenization of tissues and organs, usually with rotor-stator homogenizers.  Organs and tissues can be homogenized in 15 ml vials very effectively in a one-step process that produces very fine homogenate.  The larger 7/16” balls used with 15 ml vials provide additional force that differentiates the degree of homogenization, even when compared to proportionally smaller samples in deep well plates (Figure 14).

Muscle homogenized using 15 ml vials on a high throughput homogenizer.

Figure 14. High throughput homogenizer with 15 ml vials and two large 7/16” grinding balls produced a fine muscle homogeneous lysate with a relative efficiency of 81% (see Fig. 20).

Strengths - High throughput homogenizers are designed for processing hundreds of samples daily, but their overall effectiveness makes them useful for lower throughput operations as well.  The fact that the grinding balls and vessels (i.e., plates, vials, and tubes) are separate from the mechanical action used to power the disruption minimizes cleaning and cross-contamination issues.  Most researchers treat the plates, vials, and grinding balls as disposable items (though balls and some vials can be reused) which also helps to minimize contamination/clean-up issues.

Limitations – High throughput homogenizers require an initial investment of $7000 to more than $15,000.  The use of consumables may increase sample processing costs, but that must be measured against reduced labor and overhead charges.

Not all high throughput homogenizers shake plates the same way, as some use a figure “8” paint shaker motion while two designs have a linear motion.  The paint shaker type homogenizers tend to yield different lysis efficiencies between wells as not all samples follow the same shaking path.  Linear motion homogenizers yield comparable processing for each sample.

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[1] A general comment for all samples is that extraction buffers should not contain detergents as foaming will prevent beads and balls from moving freely during processing.