Cell disruption

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Cell disruption is a method or process for releasing biological molecules from inside a cell.

Methods

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.

Major factors

Several factors must be considered.

Sample size of cells to be disrupted

If only a few milliliters or milligrams of sample are available, care must be taken to minimize loss and avoid cross-contamination. Disruption of microbial cells, when hundreds or even thousands of liters of material are being processed in a production environment, presents different challenges. Here, throughput, efficiency, and reproducibility are key factors.

Number of samples to be disrupted at one time

Frequently when sample sizes are small (10 mg to 10 g (wet weight)), methods and equipment are available to process many samples at the same time. Mechanical cell disrupters are available that can batch process 192 samples at a time. Other machines are capable of automated sequential processing of multiple samples. Some issues to consider when processing multiple samples are cross contamination, speed of processing, equipment availability and cost and ease of cleaning and decontaminating of equipment between cell samples.

Toughness of cells to be disrupted

Some cells are relatively easy to disrupt (e.g., E. coli, blood cells, brain tissue)). More difficult samples (e.g., yeast, fungi, animal connective tissue), often require increased mechanical power or more aggressive chemical treatments. The most difficult samples (e.g., spores) may require mechanical forces combined with chemical or enzymatic methods. Samples with a strong extracellular matrix, such as animal connective tissue, biopsy samples, venous tissue, cartilage, seeds, etc., are often disrupted by impact pulverization in liquid nitrogen (see external link below). This technique, also known as cell lysis in liquid nitrogen, is based on the fact that samples containing water become very brittle at extremely cold temperatures.

Efficiency of cell disruption method

Disruption conditions may impact the desired product. For example, if subcellular fractionation studies are undertaken, it is often more important to have an optimal yield of intact subcellular components, while sacrificing overall disruption efficiency. In another example, extreme extraction conditions such as high or low pH, heat formation, or the presence of detergents and other denaturing chemicals may increase the yield of disrupted cells but destroy the intracellular component being sought.

For production scale processes, the timing of disruption and the reproducibility of the method become more important factors.

Stability of the molecule(s) or component being isolated

In general, the cell disruption method is closely matched with the material that is desired from the cell studies. It is usually necessary to establish the minimum force of the disruption method that will yield the best product. Additionally, once the cells are disrupted, it is often essential to protect the desired product from biological degradation processes (e.g., proteases), from oxidation or other chemical events and from putrification.

Purification methods to be used following cell disruption

It is rare that a cell disruption process produces a directly usable material; in almost all cases, subsequent purification events are necessary. Thus, when the cells are disrupted, it is important to consider what components are present in the disruption media so that efficient purification is not impeded.

Is the sample or its cell contents biohazardous?

Preparation of cell-free extracts of pathogens or recombinant cells expressing potentially toxic material presents unique difficulties. Several mechanical disruption techniques are not suitable because of potential biohazard problems associated with contamination of equipment and the generation of aerosols during processing.

Mild Lysis

For easily disrupted cells such as blood cells and insect or animal cells grown in culture media, a mild osmosis-based method for cell disruption (lysis) is commonly used. Quite frequently, simply lowering the ionic strength of the media will cause the cells to swell and burst. In some cases it is also desirable to add a mild surfactant and some mild mechanical agitation to completely disassociate the cellular components. Because these mild lytic methods are performed under chemically mild conditions, they are often used for subcellular fractionation studies.

Most biological cells are more difficult to disrupt. This includes most bacteria, yeast, algae and many plant and animal tissues. In these cases, mild lysis methods such as osmotic shock are insufficient to open the cell. Further, cost and relative effort to grow and harvest these cells, combined with the often small quantity of cells available to process, have favored cell disruption methods utilizing laboratory-scale manual mechanical devices such as bead mills (beadbeaters), rotor-stator homogenizers, ultrasonicators or high pressure homogenizers. These, and other stronger cell lysis methods are discussed below.

Laboratory-scale methods

Enzymatic method

The use of enzymatic methods to remove cell walls is well-established for preparing cells for disruption, or for preparation of protoplasts (cells without cell walls) for other uses such as introducing cloned DNA or subcellular organelle isolation. The enzymes are generally commercially available and, in most cases, were originally isolated from biological sources (e.g. snail gut for yeast or lysozyme from hen egg white). The enzymes commonly used include lysozyme, lysostaphin, zymolase, cellulase, mutanolysin, glycanases, proteases, mannase etc.

Disadvantages include:

  • Not always reproducible.

In addition to potential problems with the enzyme stability, the susceptibility of the cells to the enzyme can be dependent on the state of the cells. For example, yeast cells grown to maximum density (stationary phase) possess cell walls that are notoriously difficult to remove whereas midlog growth phase cells are much more susceptible to enzymatic removal of the cell wall.

  • Not usually applicable to large scale.

Large scale applications of enzymatic methods tend to be costly and irreproducible.

Bead method

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%.[1] 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.

A number of manufacturers produce machines that can be used for bead beating. These products include the BeadBeater and the FastPrep-24.

Sonication

Main article: Sonication

Another common laboratory-scale method for cell disruption applies ultrasound (typically 20–50 kHz) to the sample (sonication). In principle, the high-frequency is generated electronically and the mechanical energy is transmitted to the sample via a metal probe that oscillates with high frequency. The probe is placed into the cell-containing sample and the high-frequency oscillation causes a localized low pressure region resulting in cavitation and impaction, ultimately breaking open the cells. Although the basic technology was developed over 50 years ago, newer systems permit cell disruption in smaller samples (including multiple samples under 200 µL in microplate wells) and with an increased ability to control ultrasonication parameters.

Disadvantages include:

  • Heat generated by the ultrasound process must be dissipated.
  • High noise levels (most systems require hearing protection and sonic enclosures)
  • Yield variability
  • Free radicals are generated that can react with other molecules.

Detergent methods

Detergent-based cell lysis is an alternative to physical disruption of cell membranes, although it is sometimes used in conjunction with homogenization and mechanical grinding. Detergents disrupt the lipid barrier surrounding cells by disrupting lipid:lipid, lipid:protein and protein:protein interactions. The ideal detergent for cell lysis depends on cell type and source and on the downstream applications following cell lysis. Animal cells, bacteria and yeast all have differing requirements for optimal lysis due to the presence or absence of a cell wall. Because of the dense and complex nature of animal tissues, they require both detergent and mechanical lysis to effectively lyse cells.

In general, nonionic and zwitterionic detergents are milder, resulting in less protein denaturation upon cell lysis, than ionic detergents and are used to disrupt cells when it is critical to maintain protein function or interactions. CHAPS, a zwitterionic detergent, and the Triton X series of nonionic detergents are commonly used for these purposes. In contrast, ionic detergents are strong solubilizing agents and tend to denature proteins, thereby destroying protein activity and function. SDS, an ionic detergent that binds to and denatures proteins, is used extensively for studies assessing protein levels by gel electrophoresis and western blotting.

In addition to the choice of detergent, other important considerations for optimal cell lysis include the buffer, pH, ionic strength and temperature.

Solvent Use

A method was developed for the extraction of proteins from both pathogenic and nonpathogenic bacteria. The method involves the treatment of cells with sodium dodecyl sulfate followed by extraction of cellular proteins with acetone. This method is simple, rapid and particularly well suited when the material is biohazardous.[2]

Simple and rapid method for disruption of bacteria for protein studies.

Disadvantages include:

  • Proteins are denatured

'cell bomb'

Another laboratory-scale system for cell disruption is rapid decompression or the "cell bomb" method. In this process, cells in question are placed under high pressure (usually nitrogen or other inert gas up to about 25,000 psi) and the pressure is rapidly released. The rapid pressure drop causes the dissolved gas to be released as bubbles that ultimately lyse the cell.

Disadvantages include:

  • Only easily disrupted cells can be effectively disrupted (stationary phase E. coli, yeast, fungi, and spores do not disrupt well by this method).
  • Large scale processing is not practical.
  • High gas pressures have a high risk of personal hazard if not handled carefully.

Cryopulverization

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.[3] 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.

High-shear mechanical methods.

High-shear mechanical methods for cell disruption fall into four major classes: rotor-stator disruptors, valve-type processors, fixed-geometry processors and fixed orifice and constant pressure processors. (These fluid processing systems also are used extensively for homogenization and deaggregation of a wide range of materials – uses that will not be discussed here.) These processors all work by placing the bulk aqueous media under shear forces that literally pull the cells apart. These systems are especially useful for larger scale laboratory experiments (over 20 mL) and offer the option for large-scale production.

Rotor-stator Processors and blenders

The rotor/stator homogenizer is commonly used for small volumes of tissue suspended in 3 to 10 times its volume of homogenizing media (1-100 ml total). Larger volumes of tissue in homogenization media (100-2000 ml total) are processed either by larger rotor/stator machines or by blade blenders (often called a drink blenders). Both of these homogenizers rely on rotory cutting and/or chopping action using compact blades or paddles turning at speeds of 10,000 to 30,000 rpm.

Disadvantages include:

  • Does not work with microorganisms like bacteria, yeast and fungi and most monocellular tissue cultures.
  • Often variable in product yield.

Valve-type processors

Homogenizing valve, a method to homogenize at high pressure.

Valve-type processors disrupt cells by forcing the media with the cells through a narrow valve under high pressure (20,000–30,000 psi or 140–210 MPa). As the fluid flows past the valve, high shear forces in the fluid pull the cells apart. By controlling the pressure and valve tension, the shear force can be regulated to optimize cell disruption. Due to the high energies involved, sample cooling is generally required, especially for samples requiring multiple passes through the system. Two major implementations of the technology exist: batch processors French pressure cell press and pumped-fluid processors.

The French pressure cell press uses an external hydraulic pump to drive a piston within a larger cylinder that contains the sample. The pressurized solution is then squeezed past a needle valve. Once past the valve, the pressure drops to atmospheric pressure and generates shear forces that disrupt the cells. Disadvantages include:

  • Not well suited to larger volume processing.
  • Awkward to manipulate and clean due to the weight of the assembly (about 30 lb or 14 kg).

Mechanically pumped-fluid processors function by forcing the sample at a constant volume flow past a spring-loaded valve.

Disadvantages include:

  • Requires 10 mL or more of media.
  • General sample heating. Very high transient heating at valve interface.
  • Prone to valve-clogging events.
  • Due to variations in the valve setting and seating, less reproducible than fixed-geometry fluid processors.

Fixed-geometry fluid processors

Fixed-geometry fluid processors are marketed under the name of Microfluidizer processors, which is equipped with Y-Type Interaction Chamber. In these chamber, the flow stream is split into two channels that are redirected over the same plane at right angles and propelled into a single flow stream. High pressure promotes a high speed at the crossover of the two flows, which results in high shear, turbulence, and cavitation over the single outbound flow stream.The Y-type interaction chamber is more powerful than valve and orifice type processors in spite of block tendency in the high viscosity condition. The processors disrupt cells by forcing the media with the cells at high pressure (typically 20,000–30,000 psi or 140–210 MPa) through an interaction chamber containing a narrow channel. The ultra-high shear rates allow for:

  • Processing of more difficult samples
  • Fewer repeat passes to ensure optimum sample processing

The systems permit controlled cell breakage without the need to add detergent or to alter the ionic strength of the media. The fixed geometry of the interaction chamber ensures reproducibility. Especially when samples are processed multiple times, the processors require sample cooling.The Microfluidics Corp(USA) and Genizer LLC(USA) are two providers for Y-type interaction chamber. The Y-type interaction chamber with cooling function are provided by Genizer LLC(USA).

Fixed Orifice and Constant Pressure

Constant Cell Disruption Systems by Constant Systems part of Score Group plc - these systems are fully contained and operate using a finely controlled hydraulic system powered by electricity only. The sample is taken in and instantly pressurised up to a maximum of 40,000 PSI before being passed through a very small and fixed orifice and then returned to atmospheric pressure. As the sample is being processed this type of cell disruptor ensures that the pressure is maintained throughout the process, ensuring repeatability throughout the sample run.

Both fluid and non fluid samples can be processed through this type of cell disruptor, plant leaves and skin samples being a good example of non fluid samples. Having a maximum process pressure achievable of 40,000 PSI enables this type of unit to process more difficult sample types with fewer repeat passes. A built-in cooling jacket ensures temperature control of the sample (Water Bath or Chiller Unit is required)

See also

External links

References

  1. ↑ http://www.embl.de/pepcore/pepcore_services/protein_purification/extraction_clarification/cell_lysates_yeast/
  2. ↑ S Bhaduri and P H Demchick (1983). "Simple and rapid method for disruption of bacteria for protein studies.". Appl Environ Microbiol 46 (4): 941–3. PMC 239492. PMID 6639038. 
  3. ↑ Liquid Nitrogen Cryo-Impacting: a New Concept for Cell Disruption. Richard A. Smucker, Robert M. Pfister. Appl Microbiol. 1975 September; 30(3): 445–449.
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