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The production of biologically interesting molecules using cloning and culturing methods allows the study and manufacture of relevant molecules. Except for excreted molecules, cells producing molecules of interest must be disrupted. This page discusses various methods.
Another common laboratory-scale mechanical method for cell disruption uses tiny glass, ceramic or steel beads mixed with a sample suspended in aqueous media. First developed by Tim Hopkins in the late 1970s, the sample and bead mix is subjected to high level agitation by stirring or shaking. Beads collide with the cellular sample, cracking open the cell to release intercellular components. Unlike some other methods, mechanical shear is moderate during homogenization resulting in excellent membrane or subcellular preparations. The method, often called "beadbeating", works well for all types of cellular material - from spores to animal and plant tissues. It is the most widely used method of yeast lysis, and can yield breakage of over 50%. It has the advantage over other mechanical cell disruption methods of being able to disrupt very small sample sizes, process many samples at a time with no cross-contamination concerns, and does not release potentially harmful aerosols in the process.
In the simplest example of the method, an equal volume of beads are added to a cell or tissue suspension in a test tube and the sample is vigorously mixed on a common laboratory vortex mixer. While processing times are slow, taking 3-10 times longer than that in specialty shaking machines, it works well for easily disrupted cells and is inexpensive.
In most laboratories, beadbeating is done in sealed, plastic vials, centrifuge tubes, or deep well microtiter plates. The sample and tiny beads are agitated at about 2000 oscillations per minute in specially designed vial shakers driven by high power electric motors. Cell disruption is complete in 1–3 minutes of shaking. Machines are available that can process hundreds of samples simultaneously inside deep well microplates.
Successful beadbeating is dependent not only design features of the shaking machine (which take into consideration shaking oscillations per minute, shaking throw or distance, shaking orientation and vial orientation), but also the selection of correct bead size (0.1–6 mm diameter), bead composition (glass, ceramic, steel) and bead load in the vial.
All high energy beadbeating machines warm the sample about 10 degrees/minute. This is due to frictional collisions of the beads during homogenization. Cooling of the sample during or after beadbeating may be necessary to prevent damage to heat sensitive proteins such as enzymes. Sample warming can be controlled by beadbeating for short time intervals with cooling on ice between each interval, by processing vials in pre-chilled aluminum vial holders or by circulating gaseous coolant through the machine during beadbeating.
A different beadbeater configuration, suitable for larger sample volumes, uses a fluorocarbon rotor inside a 15, 50 or 200 ml chamber to agitate the beads. In this configuration, the chamber can be surrounded by a static cooling jacket. Using the same rotor/chamber configuration, large commercial machines are available to process many liters of cell suspension. Currently, these machines are limited to processing monocellular organisms such as yeast, algae and bacteria.
Samples with a tough extracellular matrix, such as animal connective tissue, some tumor biopsy samples, venous tissue, cartilage, seeds, etc., are reduced to a fine powder by impact pulverization at liquid nitrogen temperatures. This technique, known as cryopulverization, is based on the fact that biological samples containing a significant fraction of water become brittle at extremely cold temperatures. This technique was first described by Smucker and Pfister in 1975, who referred to the technique as cryo-impacting. The authors demonstrated cells are effectively broken by this method, confirming by phase and electron microscopy that breakage planes cross cell walls and cytoplasmic membranes.
The technique can done using a mortar and pestle cooled to liquid nitrogen temperatures, but use of this classic apparatus is laborious and sample loss is often a concern. Specialised stainless steel pulverizers generically known as Tissue Pulverizers are also available for this purpose. They require less manual effort, give good sample recovery and are easy to clean between samples. Advantages of this technique are higher yields of proteins and nucleic acids from small, hard tissue samples - especially when used as a preliminary step to mechanical or chemical/solvent cell disruption methods mentioned above.
For nitrogen decompression, large quantities of nitrogen are first dissolved in the cell under high pressure within a suitable pressure vessel. Then, when the gas pressure is suddenly released, the nitrogen comes out of the solution as expanding bubbles that stretch the membranes of each cell until they rupture and release the contents of the cell.
Nitrogen decompression is more protective of enzymes and organelles than ultrasonic and mechanical homogenizing methods and compares favorably to the controlled disruptive action obtained in a PTFE and glass mortar and pestle homogenizer. While other disruptive methods depend upon friction or a mechanical shearing action that generate heat, the nitrogen decompression procedure is accompanied by an adiabatic expansion that cools the sample instead of heating it.
The blanket of inert nitrogen gas that saturates the cell suspension and the homogenate offers protection against oxidation of cell components. Although other gases: carbon dioxide, nitrous oxide, carbon monoxide and compressed air have been used in this technique, nitrogen is preferred because of its non-reactive nature and because it does not alter the pH of the suspending medium. In addition, nitrogen is preferred because it is generally available at low cost and at pressures suitable for this procedure.
Once released, subcellular substances are not exposed to continued attrition that might denature the sample or produce unwanted damage. There is no need to watch for a peak between enzyme activity and percent disruption. Since nitrogen bubbles are generated within each cell, the same disruptive force is applied uniformly throughout the sample, thus ensuring unusual uniformity in the product. Cell-free homogenates can be produced.
The technique is used to homogenize cells and tissues, release intact organelles, prepare cell membranes, release labile biochemicals, and produce uniform and repeatable homogenates without subjecting the sample to extreme chemical or physical stress.
The method is particularly well suited for treating mammalian and other membrane bound cells. It has also been used successfully for treating plant cells, for releasing virus from fertilized eggs and for treating fragile bacteria. It is not recommended for untreated bacterial cells. Yeast, fungus, spores and other materials with tough cell walls do not respond well to this method.
- Liquid Nitrogen Cryo-Impacting: a New Concept for Cell Disruption. Richard A. Smucker, Robert M. Pfister. Appl Microbiol. 1975 September; 30(3): 445–449.