Cells Can Be Isolated from a Tissue Suspension and Separated into Different Types

The first step in isolating cells of a uniform type from a tissue that contains a mixture of cell types is to disrupt the extracellular matrix that holds the cells together. The best yields of viable dissociated cells are usually obtained from fetal or neonatal tissues. The tissue sample is typically treated with proteolytic enzymes (such as trypsin and collagenase) to digest proteins in the extracellular matrix and with agents (such as ethylenediaminetetraacetic acid, or EDTA) that bind, or chelate, the Ca²⁺ on which cell-cell adhesion depends. The tissue can then be teased apart into single living cells by gentle agitation.

Several approaches are used to separate the different cell types from a mixed cell suspension. One exploits differences in physical properties. Large cells can be separated from small cells and dense cells from light cells by centrifugation, for example. These techniques will be described below in connection with the separation of organelles and macromolecules, for which they were originally developed. Another approach is based on the tendency of some cell types to adhere strongly to glass or plastic, which allows them to be separated from cells that adhere less strongly.

An important refinement of this last technique depends on the specific binding properties of antibodies. Antibodies that bind specifically to the surface of only one cell type in a tissue can be coupled to various matrices—such as collagen, polysaccharide beads, or plastic—to form an affinity surface to which only cells recognized by the antibodies can adhere. The bound cells are then recovered by gentle shaking, by treatment with trypsin to digest the proteins that mediate the adhesion, or, in the case of a digestible matrix (such as collagen), by degrading the matrix itself with enzymes (such as collagenase).

One of the most sophisticated cell-separation technique uses an antibody coupled to a fluorescent dye to label specific cells. The labeled cells can then be separated from the unlabeled ones in an electronic fluorescence-activated cell sorter. In this remarkable machine, individual cells traveling single file in a fine stream pass through a laser beam and the fluorescence of each cell is rapidly measured. A vibrating nozzle generates tiny droplets, most containing either one cell or no cells. The droplets containing a single cell are automatically given a positive or a negative charge at the moment of formation, depending on whether the cell they contain is fluorescent; they are then deflected by a strong electric field into an appropriate container. Occasional clumps of cells, detected by their increased light scattering, are left uncharged and are discarded into a waste container. Such machines can accurately select 1 fluorescent cell from a pool of 1000 unlabeled cells and sort several thousand cells each second (Figure 8-2).

Figure 8-2

A fluorescence-activated cell sorter. . A cell passing through the laser beam is monitored for fluorescence. Droplets containing single cells are given a negative or positive charge, depending on whether the cell is fluorescent or not. The droplets are (more...)

Selected cells can also be obtained by carefully dissecting them from thin tissue slices that have been prepared for microscopic examination (discussed in Chapter 9). In one approach, a tissue section is coated with a thin plastic film and a region containing the cells of interest is irradiated with a focused pulse from an infrared laser. This light pulse melts a small circle of the film, binding the cells underneath. These captured cells are then removed for further analysis. The technique, called laser capture microdissection, can be used to separate and analyze cells from different areas of a tumor, allowing their properties to be compared. A related method uses a laser beam to directly cut out a group of cells and catapult them into an appropriate container for future analysis (Figure 8-3).

Figure 8-3

Microdissection techniques allow selected cells to be isolated from tissue slices. This method uses a laser beam to excise a region of interest and eject it into a container, and it permits the isolation of even a single cell from a tissue sample.

Once a uniform population of cells has been obtained—by microdissection or by any of the separation methods just described—it can be used directly for biochemical analysis. A homogeneous cell sample also provides a starting material for cell culture, thereby allowing the number of cells to be greatly increased and their complex behavior to be studied under the strictly defined conditions of a culture dish.

Cells Can Be Grown in a Culture Dish

Given appropriate surroundings, most plant and animal cells can live, multiply, and even express differentiated properties in a tissue-culture dish. The cells can be watched continuously under the microscope or analyzed biochemically, and the effects of adding or removing specific molecules, such as hormones or growth factors, can be explored. In addition, by mixing two cell types, the interactions between one cell type and another can be studied. Experiments performed on cultured cells are sometimes said to be carried out in vitro (literally, “in glass”) to contrast them with experiments using intact organisms, which are said to be carried out in vivo (literally, “in the living organism”). These terms can be confusing, however, because they are often used in a very different sense by biochemists. In the biochemistry lab, in vitro refers to reactions carried out in a test tube in the absence of living cells, whereas in vivo refers to any reaction taking place inside a living cell (even cells that are growing in culture).

Tissue culture began in 1907 with an experiment designed to settle a controversy in neurobiology. The hypothesis under examination was known as the neuronal doctrine, which states that each nerve fiber is the outgrowth of a single nerve cell and not the product of the fusion of many cells. To test this contention, small pieces of spinal cord were placed on clotted tissue fluid in a warm, moist chamber and observed at regular intervals under the microscope. After a day or so, individual nerve cells could be seen extending long, thin filaments into the clot. Thus the neuronal doctrine received strong support, and the foundations for the cell-culture revolution were laid.

The original experiments on nerve fibers used cultures of small tissue fragments called explants. Today, cultures are more commonly made from suspensions of cells dissociated from tissues using the methods described earlier. Unlike bacteria, most tissue cells are not adapted to living in suspension and require a solid surface on which to grow and divide. For cell cultures, this support is usually provided by the surface of a plastic tissue-culture dish. Cells vary in their requirements, however, and many do not grow or differentiate unless the culture dish is coated with specific extracellular matrix components, such as collagen or laminin.

Cultures prepared directly from the tissues of an organism, that is, without cell proliferation in vitro, are called primary cultures. These can be made with or without an initial fractionation step to separate different cell types. In most cases, cells in primary cultures can be removed from the culture dish and made to proliferate to form a large number of so-called secondary cultures; in this way, they may be repeatedly subcultured for weeks or months. Such cells often display many of the differentiated properties appropriate to their origin: fibroblasts continue to secrete collagen; cells derived from embryonic skeletal muscle fuse to form muscle fibers that contract spontaneously in the culture dish; nerve cells extend axons that are electrically excitable and make synapses with other nerve cells; and epithelial cells form extensive sheets with many of the properties of an intact epithelium (Figure 8-4). Because these phenomena occur in culture, they are accessible to study in ways that are often not possible in intact tissues.

Figure 8-4

Cells in culture. (A) Phase-contrast micrograph of fibroblasts in culture. (B) Micrograph of myoblasts in culture shows cells fusing to form multinucleate muscle cells. (C) Oligodendrocyte precursor cells in culture. (D) Tobacco cells, from a fast-growing (more...)

Serum-free, Chemically Defined Media Permit Identification of Specific Growth Factors

Until the early 1970s tissue culture seemed a blend of science and witchcraft. Although fluid clots were replaced by dishes of liquid media containing specified quantities of small molecules such as salts, glucose, amino acids, and vitamins, most media also included either a poorly defined mixture of macromolecules in the form of horse or fetal calf serum, or a crude extract made from chick embryos. Such media are still used today for most routine cell culture (Table 8-1), but they make it difficult for the investigator to know which specific macromolecules a particular type of cell requires to thrive and to function normally.

Table 8-1

Composition of a Typical Medium Suitable for the Cultivation of Mammalian Cells.

This difficulty led to the development of various serum-free, chemically defined media. In addition to the usual small molecules, such defined media contain one or more specific proteins that the cells require to survive and proliferate in culture. These added proteins include growth factors, which stimulate cell proliferation, and transferrin, which carries iron into cells. Many of the extracellular protein signaling molecules essential for the survival, development, and proliferation of specific cell types were discovered by studies seeking minimal conditions under which the cell type behaved properly in culture. Thus, the search for new signaling molecules has been made much easier by the availability of chemically defined media.

Eucaryotic Cell Lines Are a Widely Used Source of Homogeneous Cells

Most vertebrate cells stop dividing after a finite number of cell divisions in culture, a process called cell senescence (discussed in Chapter 17). Normal human fibroblasts, for example, typically divide only 25–40 times in culture before they stop. In these cells, the limited proliferation capacity reflects a progressive shortening of the cell's telomeres, the repetitive DNA sequences and associated proteins that cap the ends of each chomosome (discussed in Chapter 5). Human somatic cells have turned off the enzyme, called telomerase, that normally maintains the telomeres, which is why their telomeres shorten with each cell division. Human fibroblasts can be coaxed to proliferate indefinitely by providing them with the gene that encodes the catalytic subunit of telomerase; they can then be propagated as an “immortalized” cell line.

Some human cells, however, are not immortalized by this trick. Although their telomeres remain long, they still stop dividing after a limited number of divisions because the culture conditions activate cell-cycle checkpoint mechanisms (discussed in Chapter 17) that arrest the cell cycle. In order to immortalize these cells, one has to do more than introduce telomerase. One must also inactivate the checkpoint mechanisms, which can be done by introducing certain cancer-promoting oncogenes derived from tumor viruses (discussed in Chapter 23). Unlike human cells, most rodent cells do not turn off telomerase and therefore their telomeres do not shorten with each cell division. In addition, rodent cells can undergo genetic changes in culture that inactivate their checkpoint mechanisms, thereby spontaneously producing immortalized cell lines.

Cell lines can often be most easily generated from cancer cells, but these cells differ from those prepared from normal cells in several ways. Cancer cell lines often grow without attaching to a surface, for example, and they can proliferate to a very much higher density in a culture dish. Similar properties can be induced experimentally in normal cells by transforming them with a tumor-inducing virus or chemical. The resulting transformed cell lines, in reciprocal fashion, can often cause tumors if injected into a susceptible animal. Both transformed and immortal cell lines are extremely useful in cell research as sources of very large numbers of cells of a uniform type, especially since they can be stored in liquid nitrogen at -196°C for an indefinite period and retain their viability when thawed. It is important to keep in mind, however, that the cells in both types of cell lines nearly always differ in important ways from their normal progenitors in the tissues from which they were derived. Some widely used cell lines are listed in Table 8-2.

Table 8-2

Some Commonly Used Cell Lines.

Among the most promising cell cultures to be developed—from a medical point of view—are the human embryonic stem (ES) cell lines. These cells, harvested from the inner cell mass of the early embryo, can proliferate indefinitely while retaining the ability to give rise to any part of the body (discussed in Chapter 21). ES cells could potentially revolutionize medicine by providing a source of cells capable of replacing or repairing tissues that have been damaged by injury or disease.

Although all the cells in a cell line are very similar, they are often not identical. The genetic uniformity of a cell line can be improved by cell cloning, in which a single cell is isolated and allowed to proliferate to form a large colony. In such a colony, or clone, all the cells are descendants of a single ancestor cell. One of the most important uses of cell cloning has been the isolation of mutant cell lines with defects in specific genes. Studying cells that are defective in a specific protein often reveals valuable information about the function of that protein in normal cells.

Some important steps in the development of cell culture are listed in Table 8-3.

Table 8-3

Some Landmarks in the Development of Tissue and Cell Culture.

Cells Can Be Fused Together to Form Hybrid Cells

It is possible to fuse one cell with another to form a heterocaryon, a combined cell with two separate nuclei. Typically, a suspension of cells is treated with certain inactivated viruses or with polyethylene glycol, each of which alters the plasma membranes of cells in a way that induces them to fuse. Heterocaryons provide a way of mixing the components of two separate cells in order to study their interactions. The inert nucleus of a chicken red blood cell, for example, is reactivated to make RNA and eventually to replicate its DNA when it is exposed to the cytoplasm of a growing tissue-culture cell by fusion. The first direct evidence that membrane proteins are able to move in the plane of the plasma membrane (discussed in Chapter 10) came from an experiment in which mouse cells and human cells were fused: although the mouse and human cell-surface proteins were initially confined to their own halves of the heterocaryon plasma membrane, they quickly diffused and mixed over the entire surface of the cell.

Eventually, a heterocaryon proceeds to mitosis and produces a hybrid cell in which the two separate nuclear envelopes have been disassembled, allowing all the chromosomes to be brought together in a single large nucleus (Figure 8-5). Although such hybrid cells can be cloned to produce hybrid cell lines, the cells tend to lose chromosomes and are therefore genetically unstable. For unknown reasons, mouse-human hybrid cells predominantly lose human chromosomes. These chromosomes are lost at random, giving rise to a variety of mouse-human hybrid cell lines, each of which contains only one or a few human chromosomes. This phenomenon has been put to good use in mapping the locations of genes in the human genome: only hybrid cells containing human chromosome 11, for example, synthesize human insulin, indicating that the gene encoding insulin is located on chromosome 11. The same hybrid cells are also used as a source of human DNA for preparing chromosome-specific human DNA libraries.

Figure 8-5

The production of hybrid cells. Human cells and mouse cells are fused to produce heterocaryons (each with two or more nuclei), which eventually form hybrid cells (each with one fused nucleus). These particular hybrid cells are useful for mapping human (more...)

Hybridoma Cell Lines Provide a Permanent Source of Monoclonal Antibodies

In 1975 the development of a special type of hybrid cell line revolutionized the production of antibodies for use as tools in cell biology. The technique involves propagating a clone of cells from a single antibody-secreting B lymphocyte so that a homogeneous preparation of antibodies can be obtained in large quantities. The practical problem, however, is that B lymphocytes normally have a limited life-span in culture. To overcome this limitation, individual antibody-producing B lymphocytes from an immunized mouse or rat are fused with cells derived from an “immortal” B lymphocyte tumor. From the resulting heterogeneous mixture of hybrid cells, those hybrids that have both the ability to make a particular antibody and the ability to multiply indefinitely in culture are selected. These hybridomas are propagated as individual clones, each of which provides a permanent and stable source of a single type of monoclonal antibody (Figure 8-6). This antibody recognizes a single type of antigenic site—for example, a particular cluster of five or six amino acid side chains on the surface of a protein. Their uniform specificity makes monoclonal antibodies much more useful for most purposes than conventional antisera, which generally contain a mixture of antibodies that recognize a variety of different antigenic sites on a macromolecule.

Figure 8-6

Preparation of hybridomas that secrete monoclonal antibodies against a particular antigen. Here the antigen of interest is designated as “antigen X.” The selective growth medium used after the cell fusion step contains an inhibitor (aminopterin) (more...)

The most important advantage of the hybridoma technique is that monoclonal antibodies can be made against molecules that constitute only a minor component of a complex mixture. In an ordinary antiserum made against such a mixture, the proportion of antibody molecules that recognize the minor component would be too small to be useful. But if the B lymphocytes that produce the various components of this antiserum are made into hybridomas, it becomes possible to screen individual hybridoma clones from the large mixture to select one that produces the desired type of monoclonal antibody and to propagate the selected hybridoma indefinitely so as to produce that antibody in unlimited quantities. In principle, therefore, a monoclonal antibody can be made against any protein in a biological sample.

Once an antibody has been made, it can be used as a specific probe—both to track down and localize its protein antigen and to purify that protein in order to study its structure and function. Because only a small fraction of the estimated 10,000 –20,000 proteins in a typical mammalian cell have thus far been isolated, many monoclonal antibodies made against impure protein mixtures in fractionated cell extracts identify new proteins. With the use of monoclonal antibodies and the rapid protein identification methods we shall describe shortly, it is no longer difficult to identify and characterize novel proteins and genes. The major problem is instead to determine their function, using a set of powerful tools that we discuss in the last sections of this chapter.

Summary

Tissues can be dissociated into their component cells, from which individual cell types can be purified and used for biochemical analysis or for the establishment of cell cultures. Many animal and plant cells survive and proliferate in a culture dish if they are provided with a suitable medium containing nutrients and specific protein growth factors. Although many animal cells stop dividing after a finite number of cell divisions, cells that have been immortalized through spontaneous mutations or genetic manipulation can be maintained indefinitely in cell lines. Clones can be derived from a single ancestor cell, making it possible to isolate uniform populations of mutant cells with defects in a single protein. Two cells can be fused to produce heterocaryons with two nuclei, enabling interactions between the components of the original two cells to be examined. Heterocaryons eventually form hybrid cells with a single fused nucleus. Because such cells lose chromosomes, they can provide a convenient method for assigning genes to specific chromosomes. One type of hybrid cell, called a hybridoma, is widely employed to produce unlimited quantities of uniform monoclonal antibodies, which are widely used to detect and purify cellular proteins.