Chapter 1. Methods of Study

This chapter should help the student to:

• Know the mathematical relationships among the units of measure used for histologic specimens.
• Name the instruments and techniques used to prepare and study histologic specimens; know the attendant advantages and limitations of each.
• Know and compare the basic steps in preparing specimens for light and electron microscopy.
• Select appropriate methods to reveal specific microscopic features and substances in cells and tissues.
• Describe the basic principles of histochemistry.
• Know the substances of biologic interest that can be localized by histochemical techniques.
• Name the classes of histochemical techniques and describe the advantages and limitations of each.
• Name and describe methods appropriate for the isolation and study of specific cells and tissues from intact organisms.
• Name and describe methods appropriate for the isolation and identification of specific proteins and nucleic acid sequences from cells and tissues.

1. List, in order, the basic steps in preparing histologic sections for microscopy (Fig 1–1) and briefly describe the purpose of each step (Table 1–1).

2. Name the type of microscopy that best visualizes unstained living cells and tissues and serves as a basic tool for tissue culture (IV.D.3).

3. Compare light and electron microscopy in terms of:

1. Methods of fixation, embedding, sectioning, and staining (Tables 1–1, 1–2, and 1–3)

2. Thickness of sections (Table 1–1)

3. The support on which sections are mounted (Table 1–1)

4. The type, source, wavelength, and path of illuminating beams (IV.A and B; V.A and B)

5. Magnification (V.A)

4. Compare transmission and scanning electron microscopy in terms of:

1. Methods of preparing tissue for observation (Table 1–1; Fig. 1–1; V.D.2)

2. Electron path (V.A, B, and D.2)

3. Image obtained (V.D.1 and 2)

5. Explain cryofracture, and its advantages over conventional tissue preparation for electron microscopy (Table 1–4)

6. Describe autoradiography, and the type of information it provides (Table 1–4)

7. List the types of cell and tissue components that can be identified by histochemistry (VII.A-F).

8. The Periodic acid–Schiff (PAS) reaction allows the localization of which substances (VII.E)?

9. Describe a typical enzyme histochemical reaction (VIII) in terms of:

1. How the location of a particular enzyme is marked

2. Special considerations in preparing tissue for sectioning

10. Name three types of markers used to make antibodies visible with a microscope (IX.B).

11. List the advantages of studying isolated cells, tissues, or organ rudiments in culture rather than in the intact organism (XI).

12. What is the purpose of cell fractionation and how is it accomplished (XII)?

13. Name three types of column chromatography and describe the protein characteristics targeted by each (XIII; Table 1–5).

14. Explain how one-dimensional gel electrophoresis is used to estimate a protein's molecular weight (XIV.A).

15. What determines a protein's isoelectric point? How can differences in this characteristic be used to separate proteins (XIV.B)?

16. Under what circumstances would two-dimensional gel electrophoresis be preferable to one-dimensional gel electrophoresis (XIV.C)?

17. Name the type of molecule that is identified in Northern, Southern, and Western blotting (XV).

18. What characteristic of a DNA strand determines the site of restriction nuclease activity (XVI.A)?

19. Describe the structure of a plasmid and several characteristics that make it useful as a cloning vector (XVI.C.1).

20. After the 10th cycle, how many copies of a DNA sequence would be present in a PCR reaction mixture for each copy present after the first cycle (XVI.C.2)?

21. Explain the advantages of real-time (RT) PCR (XVI.C.3).

22. Explain how small-animal imaging exploits both histologic and molecular biologic methods to extend our understanding of tissue structure and function (II.B.7).

I. General Features of Histology & Its Methods

A. Goals of Histology

Histology, the study of tissues, is largely a visual science that relies on microscopy and other imaging modalities to reveal cell, tissue, and organ substructure. Its goals include:

1. Understanding tissue structure at levels not visible to the unaided eye, including three-dimensional relationships among the biochemical constituents.

2. Understanding the relationship between tissue structure and function.

3. Establishing a basis for learning histopathology, which involves the relationship between abnormal tissue structure and functional defects.

4. Providing a basis for treating diseased and injured tissues.

B. Histologic Methods

Histology is a subdiscipline of anatomy (from the Greek “cutting apart”). Its methods involve dividing tissues and organs in preparation for microscopic examination and chemical analyses.

1. Microscopy. Tissue analysis by light microscopy (IV) and electron microscopy (V) is the goal of these methods.

2. Tissue preparation for microscopy. The optics of each microscope type makes certain demands on tissue preparation (III). Common procedures include sectioning to produce thin, translucent slices and staining to reveal otherwise transparent substructures.

C. Advantages and Limitations of Histologic Methods

Understanding these advantages and limitations allows better interpretation of the information they provide. Advantages include making small and complex structures and processes observable. Limitations exist because the methods themselves, especially those that require dividing an organism into pieces, often halt the life processes being studied.

D. Tissue Structure and Function

These are so closely related that neither can be fully understood without appreciating the other. Structure-function relationships should be the primary focus of your study and your review.

E. Units of Measure

Measurements of cell and tissue components allow comparisons of relative sizes. The metric system is used. Common units of measure in histology are the millimeter (mm, 10−3 m), the micrometer (mm, 10−6 m), and the nanometer (nm, 10−9 m).

II. General Features of Cell Biology & Its Methods

A. Goal of Cell Biology

Cell biology is the study of cell structure and function. Its goal is to reveal the cellular and molecular basis for normal and abnormal biologic functions. Most of its methods involve disassembling intact organisms to examine the functional roles of their individual components.

B. Methods of Cell Biology

Cytology, the study of cells, once referred primarily to analyses of cell structure. With the advent of many new and powerful methods for studying living organisms at the cellular and molecular level, the expression “cell biology” now is more commonly used. Cell biology has blurred the borders between traditional basic and clinical biomedical sciences. Its methods are invaluable to histologists seeking to understand tissue biology, just as histologic methods have become basic tools for cell biologists in all biomedical subdisciplines.

1. Cell, tissue, and organ cultures involve isolating components of organisms in controlled conditions and thereby allowing observation of the effects of treatments without interference from the homeostatic mechanisms in intact organisms (XI).

2. Cell fractionation involves mechanically breaking cells and subsequently separating their minute components by centrifugation for electron microscopic or biochemical analysis (XII).

3. Chromatography involves separating molecules based on their physical or chemical properties, enabling their purification for further study (XIII).

4. Electrophoresis involves separating charged molecules using an electrical field. Both molecule size and charge can contribute to the separation process (XIV).

5. Membrane transfer and blotting methods allow the identification of molecules separated by electrophoresis (XV).

6. Genetic technology (e.g., DNA cloning), exploits advances in our understanding of gene structure and function to explore the basis for health and disease in cells and tissues (XVI).

7. Small-animal imaging (SAI) allows analysis of tissue and organ structure and function in living organisms. Most animal experimentation still involves sacrificing animals for experimental analyses. Advantages of SAI include examining biologic processes from the molecular to the organ system level in living animals and repeated measurements on individual animals for time-course studies. SAI technology currently supports live-animal imaging using powerful modalities including magnetic resonance imaging (MRI), nuclear magnetic resonance (NMR), positron emission tomography (PET), single photon emission computer tomography (SPECT), bioluminescence, and others. Advances in this field are already improving our understanding of human biology and disease as well as creating new approaches to medical diagnosis and treatment.

III. Preparing Tissues for Microscopy

Light and electron microscopy share the basic preparative methods outlined in Figure 1–1 and described in Tables 1–1, 1–2, 1–3 and 1–4. Each method has limitations and associated artifacts to be considered in interpreting histologic images. Pay particular attention to the similarities and differences in tissue preparation for light and electron microscopy. Using thin sections may seduce one into thinking of three-dimensional structures in two-dimensional terms. To overcome this problem, tissues and organs are sectioned in several planes or prepared as serial sections to allow conceptual (often computer-assisted) three-dimensional reconstruction. Because even complex staining mixtures cannot reveal every tissue component, adjacent sections may be treated with different stains.

IV. Light Microscopy

A. Light Source

Light microscopes usually are illumined by bulbs that emit white light of varying intensity. White light comprises a limited range of wavelengths (average = 550 nm). Halogen bulbs with tungsten filaments emit intense white light and are commonly used in compound bright-field microscopes (IV.D.1).

B. Microscope Lenses

Light microscopes have glass lenses. The condenser lens lies between the light source and the specimen. It collects light from the source and projects it as a cone through the specimen. The objective lens consists of one or more lenses located between the specimen and the ocular lens. It enlarges and resolves the specimen's image and projects it toward the ocular lens. Several objectives, each providing a different magnification, typically are mounted on a rotating turret. The ocular lens is located beyond the objective. It further enlarges the image and projects it onto the observer's retina, a screen, or a photographic emulsion. Increasingly, research microscopes are equipped with imaging systems that convert analog images obtained through glass lenses into digital data to be recorded as computer files.

C. Optical Properties of Lenses

1. Magnification, a property of both objective and ocular lenses, increases the specimen's apparent size and makes it appear closer. The total magnification value is obtained by multiplying the power of the objective lens by that of the ocular lens.

2. Resolution determines microscopic image clarity and richness of detail. It measures how close two objects can be and still appear separate; the smaller the value, the greater the resolution. The resolution of the human eye is 200 μm; of a light microscope, 0.2 μm; and of an electron microscope, 0.002 μm. Increased magnification is useless without improved resolution. Resolution (R) is thus independent of magnification and is calculated from the numerical aperture (NA) of the objective and the wavelength (λ) of illumination:

   

3. Numerical aperture (NA) is related to the width of the lens opening (aperture). The greater the NA, the greater the resolving power.

4. Refractive index measures the comparative velocity of light in different media. Owing to the change in refractive index at air–glass interfaces, the air between the lens and the coverslip bends some of the light projected through the specimen. At high magnifications, the accompanying loss of resolution reduces image quality. Using immersion oil (same refractive index as glass) between the coverslip and an oil immersion objective lens maintains the refractive index, thereby improving resolution.

5. Lens-related artifacts. Objective lenses comprise a series of glass lenses. The first (frontal) lens is spherical or hemispherical and magnifies the image. Others correct for aberrations or artifacts of lens curvature. Spherical lenses bring light of shorter wavelength into focus closer to the retina than light of longer wavelength, resulting in multiple blurred images. This chromatic aberration can be avoided by using achromatic or apochromatic lenses. Optical properties at the center of a spherical lens differ from those at the periphery. Apochromatic lenses correct for this spherical aberration. With spherical lenses either the center or the periphery of the field is out of focus. Planar lenses correct this curvature of field and provide flat-field focus. The best objective lenses are planar apochromatic lenses with a high numerical aperture.

D. Light Microscope Types

1. Compound bright-field microscopes are the most common tool of histology and histopathology. The term compound (versus simple) refers to a series of lenses; bright-field means the entire field is illuminated by an ordinary condenser. Specimens must be translucent and stained to provide contrast.

2. Dark-field microscopes use a special condenser to provide contrast in unstained material. A disklike shield excludes the center of the light shaft from the condenser so that the specimen is illuminated from the sides. Objects that deflect light into the objective lens are visible and appear bright on a dark background.

3. Phase contrast microscopes use a special lens system to transform invisible differences in phase (light speed) retardation (caused by the different refractive indices of specimen components) into visible differences in light intensity. Fixation and staining are unnecessary. Because these microscopes allow living specimens to be visualized, they are basic tools for tissue culture. Specimens must be thin and translucent. High resolution is difficult to obtain.

4. Polarizing microscopes allow selective visualization of birefringent (anisotropic) structures— repetitive or crystalline structures such as collagen fibers or myofibrils. Staining is not required. Light traverses a polarizing filter, the condenser projects the polarized light onto the specimen, and birefringent structures in the specimen rotate the polarized light. The objective lens projects the image through a second polarizing filter, which is oriented so that only light waves oscillating in planes different from that leaving the first polarizing filter can enter the ocular lens and be seen. Birefringent structures appear as bright, often colored, objects on a dark background.

5. Fluorescence microscopes allow the localization of substances that are labeled with fluorescing compounds (fluorochromes, such as fluorescein or rhodamine). When stimulated by light of a proper wavelength, fluorochromes emit light of a longer wavelength. An ultraviolet light source is commonly used, and the emitted light is in the visible spectrum. An excitation filter between the light source and the specimen filters out all wavelengths except that needed to stimulate the fluorochrome. A barrier filter between the objective and ocular lenses protects the eyes from ultraviolet rays and projects only the emitted light.

6. Interference microscopes combine optical features of phase contrast and polarizing microscopes to provide contrast in unstained material. Relying on differences in refractive index (IV.C.4), they can measure the phase retardation induced by components of a specimen. Unlike standard phase contrast microscopes, they can compare the refracted light with an unimpeded reference beam and provide an electronic readout of the data. Because refractive index and phase retardation are proportionate to mass, these instruments can be used to calculate the mass of cellular components. Modified interference optics, pioneered by Nomarski, are used in differential interference contrast (DIC) microscopes.

7. Confocal scanning microscopes allow the appreciation of three-dimensional structure in a specimen without cutting sections. Laser optics and computerized imaging methods provide optical sections of thicker specimens. To penetrate the thicker specimen, a narrow laser beam is focused at a specific depth (focal plane) and scanned across the specimen. The imaging system stores the scanned image, point by point (as in a computerized axial tomography [CAT] scan), building up a sharp image of the focal plane while excluding out-of-focus images from other parts of the specimen. A series of optical sections at different depths can be stored digitally, ultimately permitting reconstruction of a sharp three-dimensional image.

8. Virtual microscopy involves computerized optical scanning of a prepared slide or specimen, typically using modified brightfield or fluorescence microscopes. A digital image of the entire section is created and stored as a digital file. That file can then be accessed and displayed on a computer screen by software using algorhythms similar to those used for satellite mapping. During viewing, the software accesses only a portion of the image and allows the user to zoom in or out to view the image at greater or lesser magnifications. Virtual microscopy is increasingly used by pathologists for web-based histopathology consultations. One disadvantage is that the focus during scanning is only in a single plane. Thus image generated is exclusively two-dimensional and does not permit the observer to focus in the Z-axis to bring out-of-focus areas into sharper focus or to transition between different planes of focus to obtain a three-dimensional appreciation of the scanned tissue's structure.

9. Laser capture microscopy and microdissection allows microscopic identification, computer-assisted selection, and laser-assisted isolation of a cell or small cell cluster from a tissue section, or a cell culture or smear. The details of the selection and isolation process depend on the instrument and software employed. The sample selected can then be analyzed for specific DNA, RNA, or protein content using cell biology methods. Currently, effective protein analysis requires selection from frozen sections rather than fixed, paraffin-embedded specimens.

V. Electron Microscopy (EM)

A. General Principles

The equation for resolution is the same as for light microscopy (IV.C.2). An electron beam (wavelength = 0.005 nm) is used instead of visible light (wavelength = 397–723 nm), giving electron microscopes much greater resolving power and allowing for magnification as great as 200 times that of light microscopes. Glass lenses are opaque to wavelengths shorter than 400 nm, but the negatively charged electron beam can be deflected and focused by electromagnets as it travels through a vacuum.

B. Major Components and Operation of Electron Microscopes

Together, the cathode and anode are analogous to a light microscope's lamp. The cathode, a metallic filament, emits a spray of electrons when intensely heated by an electric current in a vacuum. The anode is a positively charged metal plate with a small hole at its center. The potential difference between the cathode and anode (60–100 kV) accelerates electrons toward the anode; some pass through the hole to form the electron beam. The condenser electromagnet deflects and focuses a cone of the beam on the specimen. The specimen typically is an ultrathin tissue section stained with electron-absorbing or electron-scattering substances to provide contrast. It is mounted on a wire mesh (grid) rather than a glass slide. The objective electromagnet deflects the electron beam that has passed through the specimen and grid to form and magnify the image. The one or two projector electromagnets, analogous to the light microscope's ocular lenses, further enlarge the image, and project it onto a fluorescent screen, photographic emulsion, or digital sensor. Electrons deflected or absorbed by the specimen do not reach the screen, whereas those that pass through the specimen do. The result is a transmission image formed by shadows of the specimen's electron-dense components.

C. Limitations of Electron Microscopy

The electron beam must travel in a high vacuum, and thus living tissue cannot be used. Components of the specimen overlying the wires of the mesh cannot be visualized. Tissue sections must be very thin, or they absorb or deflect the entire beam. The electron beam may alter specimen structure. The image cannot be visualized directly but helps create a shadow image only in shades of gray.

D. Types of Electron Microscope

1. Transmission electron microscopes (TEMs) permit visualization of the internal ultrastructure of cells and tissues as well as minute structures in cells or in intercellular spaces (limit of resolution = 0.2 nm). They operate as described earlier (V.B). Specimens are prepared as described in Table 1–1 and Figure 1–1.

2. Scanning electron microscopes (SEMs) permit visualization of surface ultrastructure (limit of resolution = 2 nm). After coating the specimen with a thin layer of heavy metal (Table 1–1), a narrow electron beam is scanned across its surface in a point-by-point sequence, generating two major signals. Secondary electrons are released from the specimen surface, collected on detectors, and converted electronically into an image that is displayed on a computer screen. This image provides a three-dimensional representation of the specimen surface. X-rays are generated when the electron beam strikes atoms heavier than sodium. Analysis of the X-ray signal can supply information about the concentration and distribution of certain elements in the specimen.

VI. Basic Principles of Histochemistry

Histochemistry marries histology with chemistry. Its goal is to reveal the chemical composition of tissues and cells beyond the acid–base distribution shown by standard staining methods without disrupting normal distribution. Histochemical analysis is guided by the following principles:

A. Preserving Chemical Distribution

The substance must not diffuse from its original site.

B. Preserving Chemical Composition

Reactive chemical groups must not be blocked or denatured. Unreactive groups must not become reactive.

C. Reaction Specificity

The reaction must be specific to avoid false-positive results.

D. Detectability

The reaction product should be colored, light emitting, or electron-scattering so that it can be visualized.

E. Insolubility

The reaction product should be insoluble to remain close to the substance it marks.

VII. Some Important Biologic Substances & Classic Methods for Detecting Them

A. Ions

Ions are difficult to localize owing to their small size and diffusibility. Some ions, however (e.g., iron in red blood cells and phosphate in bone), are immobilized by association with tissue proteins. Such ions can be localized and quantified by incubating specimens in chemical solutions that attach colored compounds to the ion of interest. For example, potassium ferrocyanide and hydrochloric acid yield dark blue ferric ferrocyanide at sites containing iron (useful in diagnosing hemosiderosis).

B. Lipids

Lipids are dissolved by organic fixatives or clearing agents, leaving gaps in the tissue, but are preserved in frozen sections. For light microscopy, lipids are demonstrated by dyes that are more soluble in lipid than in the dye solvents (e.g., Sudan black and oil red O). EM specimens are treated with reagents that react with lipids to form insoluble precipitates (e.g., osmium tetroxide). Such methods can reveal disease-related lipid accumulation (e.g., fatty change in the liver).

C. Nucleic Acids

Total DNA and RNA can be localized by specific and nonspecific methods. Feulgen's reaction uses HCl and Schiff's reagent to form a magenta precipitate in amounts proportionate to the DNA present. Acridine orange forms complexes with nucleic acids that emit fluorescence (yellow–green for DNA and red–orange for RNA) proportionate to the amount of nucleic acid present. Neoplastic and other fast-growing cells contain more RNA than slower-growing cells. Basic dyes. DNA and RNA stain nonspecifically with basic dyes. RNA's strong affinity for such dyes allows its distribution to be studied by subtraction. In this procedure, one of two adjacent sections is treated with ribonuclease (RNase) to remove RNA; subsequently both sections are stained with basic dyes (e.g., toluidine blue). RNA-containing structures (e.g., ribosomes) are basophilic in the untreated section but absent in the RNase-treated section. Hoechst dyes and DAPI fluoresce when they integrate into double-stranded DNA making them useful as a nuclear counterstain for fluorescent immunocytochemistry (IX). Their stoichiometric association with A-T pairs makes them useful for quantitating DNA. Molecular biologic methods (e.g., in situ hybridization, see section X) now allow specific nucleic acid sequences to be localized and quantified in tissue sections.

D. Proteins and Amino Acids

Older methods of protein identification are specific for particular amino acids. For example: Million reaction for tyrosine, Sakaguchi reaction for arginine, and tetrazotized benzidine reaction for tryptophan. Classes of enzymes can be detected by enzyme histochemistry (VIII). Specific proteins are typically localized by immunohistochemistry (IX).

E. Carbohydrates

Complex carbohydrates (i.e., polysaccharides and oligosaccharides) can be localized by histochemical methods. The Periodic acid-Schiff (PAS) reaction is commonly used to demonstrate polysaccharides, particularly glycogen. This reaction stains many complex carbohydrates, thus glycogen localization requires enzymatic subtraction using amylase. This method helps distinguish among glycogen-storage diseases. Other carbohydrate stains include alcian blue (specific for sulfated glycosaminoglycans at pH 1), rutheniumred (nonspecific polyanion stain forming a colored and electron scattering precipitate with polysaccharides), lectins (specific sugar-binding proteins including mannose-binding Concanavalin A and N-acetyl-d-glucosamine-binding wheat-germ agglutinin). Some carbohydrates are immunogenic owing to their large size or their presence as covalently linked components of glycoconjugates (e.g., proteoglycans, glycoproteins, glycolipids); these can be analyzed by immunohistochemistry (IX).

F. Catecholamines

The catecholamines, including epinephrine and norepinephrine, fluoresce in the presence of dry formaldehyde vapor at 60°C. This reaction is used to study catecholamine distribution in nerve tissue.

VIII. Enzyme Histochemistry

Enzyme histochemistry relates structure and function. It is used to locate many enzymes, including acid phosphatase, dehydrogenases, and peroxidases. Because fixation and clearing inactivate enzymes, frozen sections are commonly used. Sections are incubated in solutions containing substrates for the enzymes of interest and reagents that yield insoluble colored or electron-dense precipitates at sites of enzyme activity. (see Table 1–5)

IX. Immunohistochemistry

Immunohistochemistry uses labeled antibodies to localize specific cell and tissue antigens and is the most sensitive, specific, and widely used histochemical method. Because many targeted antigens are proteins whose structure may be altered by fixation and clearing, frozen sections are often used. In some cases, water-soluble plastics and waxes can be used for embedding.

A. Raising Antibodies

Injecting antigens (proteins, glycoproteins, proteoglycans, and some polysaccharides) causes an injected animal's B lymphocytes to differentiate into plasma cells and produce antibodies (Chapter 11). Members of a lymphocyte clone (descendants of a single cell) produce a single type of antibody, which binds to a specific antigenic site, or epitope.

1. Polyclonal antibodies. Large complex antigens may have multiple epitopes and may elicit several antibody types. Mixtures of different antibodies to a single antigen (obtained through fractionation of an injected animal's serum) are called polyclonal antibodies (commonly raised in rabbits and goats).

2. Monoclonal antibodies. Antibodies specific for a single epitope and produced by a single clone are called monoclonal antibodies and are commonly raised in mice. Lymphocytes from the spleen of an antigen-injected mouse are mixed with myeloma cells (lymphocyte-derived tumor cells) under conditions that cause the lymphocytes and myeloma cells to fuse. Each resulting hybridoma cell has the myeloma's capacity for continuous cell division in culture and the lymphocyte's capacity for unique antibody secretion. An isolated hybridoma gives rise to a large clone that produces large quantities of pure antibody.

B. Labeling Antibodies

Antibodies are not visible with standard microscopy and must be labeled in ways that do not interfere with their binding specificity. Common labels include fluorochromes (e.g., fluorescein, rhodamine), enzymes demonstrated by enzyme histochemistry (e.g., peroxidase, alkaline phosphatase), and electron-scattering compounds for electron microscopy (e.g., ferritin, colloidal gold).

C. Antigen Localization

1. Direct immunohistochemistry. For this method, antigen-containing tissue is incubated in a solution of labeled antibody. The antibody binds directly to the antigen, and the label appears at the antigenic site.

2. Indirect immunohistochemistry. For this method, antigen-containing tissue is incubated in a solution containing unlabeled antibody. This primary antibody, which binds directly to the antigen in the tissue, is named according to its antibody class and the animal that produced it (e.g., mouse IgG). The tissue is subsequently incubated with a labeled secondary antibody, which is an antibody to the primary antibody (e.g., rabbit antimouse IgG). Multiple labeled secondary antibodies bind to each primary antibody, and thus more label accumulates at the antigenic site than with the direct method. The indirect method is therefore more sensitive and avoids the risk of altering the primary antibody's binding specificity by attaching a label.

X. In Situ Hybridization

In situ hybridization is a method of analyzing the tissue distribution of particular nucleotide sequences in DNA (e.g., specific genes) and RNA (e.g., specific mRNAs). Hybridization refers to the binding of complementary nucleotide sequences to one another with high specificity. Recombinant DNA technology permits copies of selected single-strand nucleotide sequences to be synthesized in large numbers. Synthetic sequences complementary to the RNA or DNA sequence, an investigator wishes to localize, are termed probes and can be labeled with radioisotopes (e.g., 32P), biotin, or digoxigenin. Radiolabeled probes are demonstrated by autoradiography. Biotin-labeled probes are demonstrated with enzymes (e.g., peroxidase) or fluorochromes covalently linked to avidin, a molecule with high affinity for biotin. Digoxigenin-labeled probes are demonstrated by indirect immunohistochemistry using antidigoxigenin primary antibodies. Labeled probes were first used to analyze nucleic acids isolated from cell or tissue homogenates. The term in situ refers to the application of this technique to tissue sections, smears of cells, cultures, or even whole embryos; when such specimens are incubated with labeled probes, the probes bind to and reveal the distribution of their complementary sequences.

XI. Cell, Tissue, & Organ Cultures

These methods are used to study living cells and tissues without the interference of the organism's homeostatic mechanisms. They facilitate the controlled manipulation of the environment. Cells and tissue isolated and grown in culture are referred to as in vitro (in glass) and those in the intact organism as in vivo (in the living). Cells may react differently to in vitro and in vivo treatments.

A. Culture Medium

The medium in which cells and tissues are grown substitutes for the plasma that normally bathes them in vivo. It consists of a buffered isotonic saline solution to which are added nutrients (amino acids, vitamins, hormones, carbohydrates) of rigidly controlled composition. The development of serum-free, defined culture media, has decreased the need for serum and tissue extracts of less well-defined composition in media. Antibacterial and antifungal agents are often added.

B. Culture Types

1. Cell culture. In suspension culture, cells are suspended in medium either free or on floating beads. In some cases cells are suspended in semisolid blocks of extracellular matrix or agarose. In plate culture, cells attach to plastic or glass tissue culture dishes. The dishes may be coated with substances that improve attachment and cell function, including gelatin, collagen, polylysine, albumin, or extracellular matrix extracts. Plate-cultured cells behave differently at different densities. Cells may be cultured in confluent monolayers (i.e., the entire culture surface is covered with cells in contact with one another) or at clonal densities (i.e., seeded at low densities to avoid cell–cell contact). The latter method allows the growth of individual cell colonies, or clones.

2. Tissue and organ culture. Tissues or organ fragments are isolated and grown as explants, usually at the air–medium interface. This method is used to study embryonic differentiation and morphogenesis away from the complex environment of the embryo.

C. Isolation and Study of Pure Cell Strains

Individual cell types may be isolated and studied in vitro to explore their separate contributions to tissue and organ function. Cell suspensions are commonly obtained from tissues by enzymatic digestion (e.g., with trypsin or collagenase) of the components that hold the cells together. Cell types may then be separated on the basis of size and mass through specialized forms of centrifugation (e.g., elutriation, density gradient centrifugation). Other methods of isolating a cell type from a heterogeneous population in suspension exploit the differential binding of cells by antibodies attached to a culture surface; others use a fluorescence-activated cell sorter, which separates cells labeled with fluorescent antibodies from cells lacking the label.

XII. Cell Fractionation

This procedure is used to isolate and collect cellular components in quantity to study their contributions to cell function. It begins with the mechanical homogenization of cells and tissues to break plasma membranes and release the cell contents (e.g., individual organelles). These are then separated by size and density, using either of two centrifugation methods. In differential centrifugation, components are separated by their characteristic sedimentation rates, using different amounts of centrifugal force for various periods. In density-gradient centrifugation, the homogenate is layered onto a gradient of solute and centrifuged until its components come to rest in portions of the gradient with densities similar to their own.

XIII. Column Chromatography

Because they are the principal mediators of cell function, proteins have been targeted by many methods to isolate and analyze the content of cell homogenates (XII). Chromatography involves packing a hollow column with a semipermeable material (often tiny resin or agarose beads) and applying a cell or tissue homogenate, or a centrifugal fraction of such a sample, to the top of the column. After the sample percolates into the column, solvent is added and allowed to flow through. Timed fractions of the flow-through material (eluate), containing different molecules from the homogenate, are collected. Release (elution) is retarded by interactions with the packing material and results in separation. Three standard strategies have evolved owing to their proven usefulness (Table 1–6). Almost unlimited variations in packing materials and solvents are available and may be used individually or in combination to isolate molecules of interest.

XIV. Electrophoresis

A. One-Dimensional Gel Electrophoresis

An electric field applied to a solution of proteins, or other large molecules, causes them to move in a direction and at rates reflecting their net charge. When the movement occurs in a semipermeable gel (e.g., polyacrylamide, agarose), migration in the gel is limited by pore size as well. In native gel electrophoresis, proteins migrate in physiologic buffers and retain their native structure and folding; thus the shape of the molecule (e.g., globular or filamentous) also affects its migratory rate. More commonly, protein solutions are heated in the presence of a detergent (sodium dodecylsulfate, or SDS) and a reducing agent (mercaptoethanol) before they are subjected to polyacrylamide gel electrophoresis (SDS-PAGE). The reducing agent breaks –S–S– linkages and, together with SDS, promotes unfolding. SDS further coats the protein with negative charges such that each protein's net charge is negative, and all have essentially the same charge density. When applied to a polyacrylamide gel and electrophoresed in buffers containing SDS and mercaptoethanol, proteins migrate toward the positive pole at rates that depend largely on their size. Smaller molecules are retarded less by the gel and move faster. When proteins of known molecular weight are used as standards, this method allows the molecular weights of individual proteins to be estimated. Typically, SDS-PAGE is carried out in flat or slab gels, in which multiple samples are run in parallel lanes for comparison. After electrophoresis, the gel is removed from the apparatus and stained (e.g., with Coomassie blue) to reveal the position of the proteins and thus their molecular weight. Nucleic acids may be separated in either polyacrylamide or agarose gels. DNA and RNA gels are typically stained with ethidium bromide, which complexes with the nucleic acids and causes the bands to fluoresce on exposure to ultraviolet light.

B. Isoelectric Focusing

Owing to its amino acid composition, each protein has a characteristic pH (isoelectric point) at which it has no net charge and thus will not migrate in an electric field. Using special buffers to imbue a tube of polyacrylamide (tube gel) with a stable pH gradient along its length in an electric field, sample proteins can be induced to migrate to, and come to rest at, their own isoelectric points.

C. Two-Dimensional Gel Electrophoresis

Because many proteins may have the same molecular weight, complete separation on a one-dimensional gel may not be possible. In such cases, a sample may be subjected first to isoelectric focusing, after which the resulting tube gel may be laid atop a slab gel and subjected to SDS-PAGE. In this way, proteins that first separated based on their isoelectric points migrate into the slab gel and are further separated based on their molecular weight. The resulting separation yields spots of smaller amounts of individual proteins. Silver staining of two-dimensional gels is common owing to its greater sensitivity.

XV. Blotting & Electrotransfer

Exploiting the affinity of antibodies for specific proteins, and that of complementary nucleic acid sequences for one another, additional steps have been developed to analyze molecules first separated by electrophoresis. Gels used for electrophoresis, restrict the access of large native molecules, such as antibodies and large nucleic acids, to their target molecules. Thus methods have been developed to remove the separated molecules from these gels and to immobilize them on membranes in the same pattern conferred on them by the initial separation. Transfer from the gels to the membrane may be accomplished by blotting or by electric charge.

In blotting techniques, molecules in the gel are carried by the flow of a buffer across the gel and through the membrane. Flow is induced by capillary attraction through absorbent paper layers on the far side of the membrane. As the buffer flows through, the membrane traps the target molecules. In electrotransfer, target molecules migrate from the gel to the membrane in an electric field applied to the assembly. Immobilization is achieved electrostatically for proteins and by heating or exposure to ultraviolet light for RNA and DNA. Subsequently the membranes carrying the more accessible target molecules are incubated with labeled antibodies or complementary nucleic acid sequences (termed probes) to reveal the positions and relative amounts of the molecules of interest. Western blotting refers to the use of labeled antibodies to reveal the presence and amount of a specific protein on the membrane. Northern blotting refers to the procedure in which labeled probes reveal complementary RNA sequences. Southern blotting refers to similar procedures to localize specific DNA sequences. Methods used to label the antibodies and probes determine the detection methods employed. Antibody labeling and detection methods similar to those described for immunohistochemistry (IX) are often used for Western blots. In northern and southern blotting, probes are often labeled by incorporating nucleotides labeled with 32P or light-emitting molecules into the probe during its synthesis, followed by exposing X-ray film or photographic paper respectively to the labeled membrane at low temperatures and then developing them.

XVI. Genetic Technology

Advances in our understanding of DNA structure and function have yielded methods of unprecedented power for unraveling the intricacies of cell and tissue biology. Previously, viewed as mysterious agents of heredity, genes have become tools of potentially unlimited diagnostic and therapeutic power.

A. Isolating Genes and DNA Fragments

Studies of how bacteria inactivate foreign DNA led to the discovery of restriction nucleases (e.g., EcoR1, HindIII), a class of bacterial enzymes that are capable of cutting DNA into fragments by acting at very specific nucleotide sequence target sites. Different bacteria have restriction enzymes that are specific for different sequences, providing molecular biologists with an array of tools to isolate very specific DNA fragments, especially when the nucleotide sequence is known. When large DNA molecules are fragmented by a single restriction enzyme, the fragments generated are of different sizes, reflecting the positions of the target sites in the larger molecule. Because the overall nucleotide sequence is unique to each individual, the banding pattern revealed when the fragments are separated by size, using gel electrophoresis, is equally unique, providing a genetic fingerprint of an individual. Comparing banding patterns produced by several enzymes can determine with high confidence whether DNA in multiple samples came from the same individual. Knowing the nucleotide sequence of a gene, and those of its 3′ and 5′ flanking regions, allows individual genes and various upstream and downstream components to be obtained and isolated by electrophoresis. Cutting a band containing a fragment of interest from a gel, and its release from the gel material, allows the fragment to be purified and used for subsequent studies.

B. DNA Sequencing

Unknown nucleotide sequences of genes and other DNA fragments can be determined by several strategies. Commonly, isolated fragments are incubated with a DNA polymerase in conditions that halt assembly of the copy when a particular nucleotide is reached. The result is a mixture of subfragments terminating at each nucleotide in the sequence. Subfragment size depends on which point in the sequence it terminates. Electrophoresis of the mixture arranges the subfragments by size and therefore reveals the order in which each nucleotide occurs in the fragment.

C. Copying Genes and DNA Fragments

Obtaining multiple identical copies of a gene or fragment of interest greatly facilitates further molecular analysis.

1. DNA cloning. One method of obtaining such copies involves inserting (cloning) the fragment into bacterial plasmids and allowing the DNA replicating machinery in growing bacteria to copy the entire recombinant DNA sequence of the plasmid as they copy their own DNA. Plasmids used as carriers (vectors) in such procedures are typically small, circular, double-stranded DNA molecules capable of entering living bacteria and maintaining their integrity while being copied separately from the bacteria's own genome. Cloning vectors typically include genes for antibiotic resistance; thus when bacteria are grown in media containing an antibiotic (e.g., ampicillin), only plasmid-containing bacteria survive and continue to grow. Vectors also have cloning sites, with target nucleotide sequences for one or more restriction enzymes that serve as the insertion site for the fragment of interest. Cutting the plasmid with a restriction enzyme linearizes the circular DNA and provides free ends for attaching the fragment. Digesting the fragment and the plasmid with the same enzyme yields matching ends, allowing the fragment to be covalently attached by the enzyme DNA ligase, which repairs and recircularizes plasmid. The result is a fragment-containing recombinant plasmid that is larger than the original and that can be separated electrophoretically. Often the cloning sites occur or are engineered into sequences for a marking enzyme (e.g., beta galactosidase). Inserting the fragment disrupts enzyme expression.

   Bacteria containing recombinant plasmids, when grown on media containing, a substrate for the enzyme that yields a colored precipitate in the presence of the enzyme, notably lack color and can be easily selected for further growth. Thus bacteria that are capable of growing in media containing the antibiotic, and that remain uncolored on media containing the enzyme substrate, most likely contain the desired recombinant plasmids. Colonies of bacteria-containing recombinant plasmids are inoculated into growth media and quickly multiply, making abundant copies of the fragment along with their own DNA and vector DNA. The bacteria can then be lysed, their DNA isolated, the recombinant plasmids separated from the larger bacterial genomic DNA by electrophoresis, and the fragments isolated from the plasmids by restriction enzyme digestion.

2. Polymerase chain reaction (PCR). A more rapid method for copying selected DNA fragments is available and more commonly used. PCR exploits the fact that cells use existing DNA sequences as templates for making new copies. Exponential doubling of copy numbers is thus possible if the copies made in each succeeding cycle of synthesis are used as templates in the next. Because PCR is carried out entirely in vitro, the transition from one replication cycle to the next can be greatly accelerated compared with rates possible in intact organisms, which (1) must copy their entire genome during each replication cycle and (2) cannot withstand the high temperatures used in PCR to separate the copy from the template DNA after synthesis. DNA sequences of interest can be selected using appropriate priming oligonucleotides (primers), short nucleotide sequences that hybridize to the beginning and ending portions of the target DNA sequence. The use of DNA polymerases that specialize in filling in the intervening nucleotides, but that stop copying when the other primer is reached, allows only the target sequence to be copied. Transient heating of the reaction causes the original and copy strand to separate (melt). As the reaction cools, additional primers hybridize to the original and copy strands, using both as templates for the next replication cycle and resulting in another doubling of the copy number. The number of cycles thus determines the number of copies generated.

   A key to developing this method was the discovery that bacteria living at high temperatures (Thermus aquaticus) have Taq polymerases that can withstand heat without denaturing. Another has been the development of thermal cyclers instruments that allow the programming of rapid and repeated temperature shifts so that the time between cycles can be shortened and can occur automatically without handling the samples. The method can also be used to amplify RNA sequences if viral reverse transcriptase is first used to transcribe the RNA into complementary DNA (cDNA) sequences, which are subsequently subjected to PCR (RT-PCR).

3. Real-Time PCR. More recently, methods have been developed to more quantitatively assess the amount of a particular mRNA species present in a sample, allowing the measurement of the effects of specific treatments on gene expression. The method exploits the fact that in the presence of an excess of the appropriate primers, and Taq polymerase, the more copies of cDNAs generated by reverse transcriptase from a particular mRNA species in a sample, the more rapidly its PCR products will accumulate. In one commonly used approach (TaqMan®, Applied Biosystems), a single-stranded oligonucleotide complementary to a unique sequence in the mRNA of interest is dual-labeled with a fluorescent reporter and a fluorescence quencher to create a probe. When the probe anneals to its target sequence, the reporter's proximity to the quencher prevents its detection. However, as the Taq polymerase fills in the sequence from the primer, it degrades the probe, liberating the reporter from the quencher and allowing it to be detected. Carried out in a thermal cycler equipped with a laser to stimulate reporter fluorescence and a fluorescence detector, cycle by cycle measurements of fluorescence reveal the rate of exponential amplification of the target mRNA in proportion to its abundance in the sample. The fact that these measurements are recorded during the amplification process rather than using its end products gives the process the name, Real-Time PCR. Because PCR amplification can eventually plateau at similar levels regardless of the initial concentration of the target mRNA, the ability to measure the process in real time during the exponential phase of the amplification provides a more sensitive and reliable measure of the initial concentration than an end-product assay.