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Disrupting cells and tissues by applying a force not inherent to the sample is considered a mechanical disruption method. Mechanical homogenization procedures generate lysates with characteristics different than those produced by chemical lysis. By avoiding detergents and chaotropes, many cytosolic proteins may remain intact following liberation from the cell. This is useful for protein isolation and enzyme assays. However, mechanical homogenization may simply be the tool used to rapidly disrupt cells and tissues with the use of denaturing reagents, especially during RNA isolation procedures. Regardless of the mechanical approach, whether it is to beat, grind, shear, or explode cells, they are tools that can be applied in many different ways to sample preparation.
For simplicity, the methods used for sample disruption have been divided into four groups: grinding, shearing, beating, and shocking. Many engineers may cringe by this delineation, but we are approaching the topic practically and as biologists. Foremost, it must be highlighted that many methods make use of more than one force, as with conical homogenizers which grind and shear. We attempt to note this where it happens. Additionally, there are many tools and methods which are not discussed simply due to a lack of time and resources to examine all options. This section focuses on the more widely used methods.
In our effort to better understand mechanical disruption, many of the homogenizers and methods have been compared by analyzing samples following homogenization. In certain instances, such as with yeast and muscle tissue, microscopic observation can be very useful for determining the extent of disruption. In other instances, a measurement of the DNA, RNA, or protein released into the supernatant has been used. In many processes, especially where more than one homogenizer was used to process a sample, relative efficiencies of homogenization was performed by measuring lactate dehydrogenase (LDH) released from the cells/tissues. Theses comparisons will be noted throughout the following sections.
Grinding relies on creating friction by sandwiching the sample between two hard surfaces that slide against each other. Forces on the sample are two-fold, namely downward pressure accompanied by a tangential shearing force. Grinding causes tearing and ripping of samples, much like shearing, but differs in that there is direct contact between sample and homogenizer. Mortar and pestle is the best known tool for grinding, but others are grain mills and certain types of glass homogenizers. With adequate patience, solids can be reduced to very fine particles by grinding, part of which is dependent upon the topology of the grinding surfaces.
In its various forms grinding can be used on wet, dry, and frozen samples, however it is most efficient on solid samples. One key characteristic of grinding is that friction generates heat and at times can be significant. Consequently, the heat tolerance of the analyte should be considered when selecting a grinding method. Frequently samples are frozen prior to grinding, commonly with liquid nitrogen, which can be used for chilling both sample and homogenizer. This cryogenic grinding makes the sample brittle and fracture easily, but it also preserves analytes that are heat labile or which may rapidly degrade upon liberation, such as RNA. Large traditional mortar and pestle are useful for cryogenic grinding as the mass of the homogenizer acts as a cold reservoir.
Certainly the oldest tool used for grinding is the mortar and pestle, making its debut long before the dawn of civilization. It is still a popular grinding tool in the lab, being used for some of the most advanced analytical processes. A miniaturized version of the mortar and pestle, the CryoGrinder™, is used for scaled down cryogenic grinding of small samples in the milligram range. Several of the glass tissue homogenizers, such as conical glass and Tenbroeck, use grinding forces to effectively disrupt cultured cells and tissues.
Mortar & Pestle: Mortar and pestle is still widely used for sample homogenization. In life science labs, their widest use is for grinding tissue frozen with liquid nitrogen (see below). However, they are also indispensable for grinding solids at room temperature. For the single occasional sample it is sufficient, but when throughput is necessary, alternative tools such as the mixer mills are more practical.
Strengths – Mortar and pestle is easy to use and relatively inexpensive to purchase. With dry grinding, it is possible to generate very small particles.
Limitations – Throughput with mortar and pestle is low. Contamination issues may also be a concern as the grinding will generate dust. Though sturdy, many sets are made of glass or porcelain, which can chip or crack if dropped.
Cryogenic Grinding with Mortar & Pestle: Grinding frozen samples with liquid nitrogen using a mortar and pestle is a widely used method. The mortar and pestle are cleaned and placed in a Styrofoam tub or cooler where liquid nitrogen is poured or dispensed onto the mortar and pestle. Care is needed to avoid splattering liquid nitrogen when the mortar and pestle first start chilling. After several minutes the set will be cooled and a fog will usually settle over the apparatus. The sample may already be frozen or it can be snap frozen by dropping it into a beaker of liquid nitrogen (use plastic beakers). If the sample is taken from a -80°C freezer, let it sit on the surface of the mortar to chill further. To grind, hold the pestle with a gloved hand (use a protective glove) and firmly press on the sample while twisting. The sample will typically shatter into small pieces, some of which may fly from the mortar so use added caution with biohazardous materials. The fragmented pieces of the sample will continue to get smaller as the sample is ground using a circular motion with downward pressure. Once the grinding is completed, the ground sample must be tapped or scraped from the pestle. The sample must then be transferred into a receiving vessel using a pre-chilled spatula. If the sample is to be subsequently stored frozen, pre-chill the tube or vial that will hold the ground sample.
Strengths – The mortar and pestle, whether it is used for grinding at room temperature or with liquid nitrogen, is a good standard method for reducing samples into small particles. The apparatus is relatively inexpensive and is available in ceramics to metals. The relatively large mass of the mortar and pestle serve as a cold reservoir which helps to prevent sample thawing.
Limitations – A significant problem in cryogenic grinding with mortar and pestle is that small samples (e.g., 10-20 mg) can be essentially lost when ground into the surface of the mortar. This makes sample recovery difficult and leads to poor yields. Another major disadvantage of mortars and pestles is that the number of samples that can be processed is low. As the mortar and pestle may be in the -150°C range following grinding, they must be warmed to room temperature (slowly) between uses and cleaned. Consequently, if many samples are processed daily, many sets are needed. Labs that process significant numbers of samples cryogenically must dedicate significant shelf space to the mortar and pestle sets.
CryoGrinder™ for Small Sample Grinding: As small samples are difficult to recover from standard mortar and pestles, the CryoGrinder™ serves as an alternative tool for cryogenic grinding. Small samples, which are pulverized cryogenically using a mortar and pestle, are spread as a fine powder over the mortar surface. This is difficult to collect. The CryoGrinder™, which is essentially a miniature mortar and pestle, possesses a small well and associated pestle designed for samples less than 100 mg (P). Following grinding, collecting pulverized sample is more efficient. The CryoGrinder™ is used similarly to a standard mortar and pestle, in that the CryoGrinder™ is chilled and then samples are added to the well. The CryoGrinder™ is also powered by a handheld cordless wrench.
Cryogenic grinding is useful as a first step in preparing samples for chemical lysis or subsequent mechanical processing. Its true value is that samples can be reduced from large solid items to small particles without tremendous input of heat. With smaller particle size, the sample can be rapidly dissolved, as is done for RNA isolation. As compared to the mortar and pestle, the CryoGrinder™ does generate smaller particles as was determined by comparative enzyme liberation studies (see below).
| Strengths – The CryoGrinder™ is effective at grinding small samples while frozen. It is more effective than the mortar and pestle as measured by the release of LDH from muscle tissue homogenized by both methods. As the CryoGrinder™ generates smaller particles than the mortar and pestle the small particles will more readily dissolve into extraction buffers. Another advantage is that the CryoGrinder™ is motorized which allows for a greater number of samples to be processed without added fatigue. |
Figure 2. CryoGrinder™ is useful for the cryogenic grinding of very small samples, less than 100 mg. |
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Limitations –
Sample size for the CryoGrinder™ must
be small (100 mg or less) for the pestle to be pressed effectively
against the mortar. The mortars are also small and must be kept
within a liquid nitrogen reservoir (e.g., CryoCooler™) so that they remain cold.
Tissues Disruption with Glass Homogenizers: Original methods for homogenizing tissues made use of glass homogenizers. The tools available for this include ground glass homogenizers, such as the Potter-Elvehjem, conical, and Tenbroeck. These tissue grinders are closely related to the Dounce and Potter-Elvehjem (when used with a PTFE pestle), but the latter rely on shearing forces and will be discussed below. Glass tissue grinders have tight fitting mortars and pestles with ground glass surfaces. The surfaces are course like a very fine emery paper so that the pestles can dig into tissues being gripped by the mortar and shear the sample as it is turned. Tissues processed in glass tissue grinders are often chilled on ice. The Tenbroeck pestle, which is hollow, can be filled with cold liquid to cool from the inside. |
Figure 3. Comparison of LDH liberated by cryogenic Methods. Mouse muscle was cryogenic ground using mortar and pestle and CryoGrinder™. Greater LDH activity was harvested from CryoGrinder™ homogenized muscle. |
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The actual process of grinding is relatively simple and involves adding an extraction buffer and the tissue to the homogenizer tube, then slowly pressing the pestle on to the sample with a twisting motion. The piston is raised and lowered while twisting to help turn the sample to expose all sides to grinding. This action is repeated. Since they are glass, the homogenizers can be washed and sterilized before use. If residual detergents on the glass is a concern, then cleaning can be done with a 1% solution of sodium carbonate (which serves as a very nice wetting agent) followed by rinses with 3% acetic acid. The homogenizers can also be baked at 280°C to further decontaminate the glass if the application is for RNA or DNA isolation. |
Figure 4. Mouse muscle homogenized with CryoGrinder™.Though particles are slightly larger than other methods, homogenization was relatively complete. |
| Strengths – Glass tissue grinders are
inexpensive and easy to use. They work relatively well and generate
very fine homogenate. In single-step disruption experiments, conical
glass homogenizers liberate about half as much enzyme as compared to
larger more expensive high throughput homogenizers. Glass tissue
grinders are approximately 30-40% as efficient as the best methods (see
Fig. 20), but the relative cost is fractional ($70 vs. $15,000). They
are very easy to clean and decontaminate.
Limitations – Homogenizing with glass tissues grinders inevitably will leave fibrous and membranous components relatively intact. Certain tissues, even with prolonged grinding, are difficult to disaggregate. Throughput with these homogenizers is also low unless multiple units are available. Glass homogenizers are also prone to breakage. |
Figure 5. Mouse muscle homogenized with a conical glass homogenizer. Though the homogenate appears fine, significant connective tissue did not homogenize. In comparison to other methods, the conical glass homogenizer was 42% as efficient (see Fig 20). 
Figure 6. Tenbroeck (left) and conical (right) glass homogenizers. |