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
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)