Choosing The Best Cell Disruption Method for Research and Pharmaceutical Production

The production of biologically interesting molecules using cloning and culturing methods allows the study and manufacture of relevant molecules for research and pharmaceutical production. In spite of the ubiquity of these methods, a challenge remains in maximizing production efficiency and workflow. Except for excreted molecules, cells producing molecules of interest must be disrupted. This page provides guidance for the selection of your optimum cell disruption or cell rupture method.

These methods include:

• Lysis
• Enzymatic Method
• Bead Method
• Sonication
• Cell Bomb
• Rotor-Stator
• French Press
• Mechanically Pumped-Fluid Processors
• Microfluidizer® Processors

Tiny molecules. Big factors.

Several factors must be considered in selecting your optimum method for cell disruption.

1. What is the volume or sample size of cells to be disrupted?

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

2. How many different samples need to be disrupted at one time?

   This needs to be considered in conjunction with sample size because frequently when sample sizes are small, there are many samples. As sample sizes increase, fewer samples are usually processed. In any event, issues of sample cross contamination, speed of processing, and equipment cleaning become critical.
   

3. How easily are the cells disrupted?

   As the difficulty of disruption increases (e.g. E. coli), more force or stress is required to efficiently disrupt the cells, and the processors that impart the force increase in sophistication. For even more difficult samples (e.g. yeast), higher levels of force are required with a parallel increase in the processor power and cost. The most difficult samples (e.g. spores) require a combination of extreme mechanical forces combined with chemical or enzymatic efforts, often with limited disruption efficiency.
   

4. What efficiency of disruption is required?

   While high disruption efficiencies are generally desirable, over-disruption may impact the desired product. For example, if subcellular fractionation studies are undertaken, it is often more important to have intact subcellular components while sacrificing disruption efficiency. For production scale processes, the time to rupture the cells and the reproducibility of the method become more important factors.
   

5. How stable is the molecule(s) or component that needs to be 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 normal biological processes (e.g. proteases) and from oxidation or other chemical events.
   

6. What purification methods will be used following cell disruption?

   It is rare that a cell rupture 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.
   

Disruption influencers

Disruption methods vary considerably in the severity of the disruption process, the equipment and/or reagents needed, and in ease of use. These methods are usually matched to the difficulty in disrupting the cells under study and the amount of material being processed.

Lysis

For easily disrupted cells such as insect and mammalian cells grown in culture media, a mild 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 or sonication to completely disassociate the cellular components.  Due to the cost and relative effort to grow these cells, there is often only a small quantity of cells to be processed, and preferred methods for cell disruption tend to be a manual mechanical homogenizer, nitrogen burst methods, or ultrasound with a small probe.  Because these methods are performed under very mild conditions, they are often used for subcellular fractionation studies.

For cells that are more difficult to disrupt, such as bacteria, yeast, and algae, hypotonic shock alone generally is insufficient to open the cell and stronger methods must be used. These organisms have cell walls that must be broken to allow access to intracellular components.  These stronger 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). 

Disadvantages include:

1. 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.
   
2. Not usually applicable to large scale.
   Large scale applications of enzymatic methods tend to be costly and irreproducible.
   
3. The enzyme must be removed (or inactivated) to allow cell growth or permit isolation of the desired material.

Bead Method

A second common laboratory-scale mechanical method for cell disruption uses glass or ceramic beads and a high level of agitation.  At the lowest levels of the technology, the beads are added to the cell suspension in a tube and the sample is mixed on a common laboratory vortex mixer.  This process works for easily disrupted cells, is inexpensive, and multiple samples can be conveniently processed.  The more sophisticated level, bead-based methods use a closed container holding the sample and the beads with an electric motor to provide vigorous agitation.  When larger samples are processed, some form of cooling is provided (typically liquid CO₂) as the sample heats significantly due to the extreme agitation.  Another configuration uses an inert, rapidly rotating rotor in a small container containing the cells and beads. 

Disadvantages include:

1. Limited ability to scale to larger samples.
   
2. Variability in product yield and purity
   
3. Occasional problems with foaming and sample heating, especially for larger samples
   
   

Sonication

A third 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 high 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:

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

The "Cell Bomb"

A fourth 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:

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

High-shear mechanical methods.

High-shear mechanical methods for cell disruption fall into three major classes: rotor-stator disruptors, valve-type processors, and fixed-geometry 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

Most commonly used as tissue disruptors.

Disadvantages include:

• Do not work well with difficult-to-lyse cells like yeast and fungi
• Often variable in product yield.
• Poorly suited for culture use.
  

Valve-Type Processors

Valve-type processors disrupt cells by forcing the media with the cells through a narrow valve under high pressure (20,000-30,000 psi).  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: the French press and pumped-fluid processors.

1. French press technology 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:

   1. Not well suited for larger volume processing.
      
   2. Awkward to manipulate and clean due to the weight of the assembly (about 30 lbs/14 Kg).
   
   
2. Mechanically pumped-fluid processors function by forcing the sample at a constant volume flow past a spring-loaded valve. 
   
   Disadvantages include:
   
   1. Requires 10 mL or more of media.
      
   2. Prone to valve-clogging events.
      
   3. Due to variations in the valve setting and seating,  less reproducible than fixed-geometry fluid processors.
      

Fixed-Geometry Fluid Processors

The patented technology of Microfluidics' fixed-geometry fluid processors are marketed under the name of Microfluidizer® processors.  The processors disrupt cells by forcing the media with the cells at high pressure (typically 20,000-30,000 psi) through a proprietary interaction chamber containing a narrow channel that generates the highest shear rates commercially available.  The ultra-high shear rates allow for:

1. Processing of more difficult samples
   
2. Fewer repeat passes to ensure optimum sample processing

It should be noted that for the lysis of larger volumes of cells (over 20 ml), the Microfluidizer® systems provide a highly reproducible, convenient, and efficient method for cell lysis.  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:

1. Day-to-day reproducibility
   
2. Machine-to-machine reproducibility
   
3. Direct scalability from laboratory scale (20 ml to several liters) to production scale (10s of liters per minute) 

Disadvantages included:

1. In many circumstances, especially when samples are processed multiple times, the Microfluidizer® processors do require sample cooling, and sample cooling is available.

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