Histology_Ross

Histology_Ross. OVERVIEW OF METHODS USED IN HISTOLOGY / 1 TISSUE PREPARATION / 2 Hematoxylin and Eosin Staining With Formalin Fixation / 2 Other Fixatives / 2 Other Staining Procedures / 3 HISTOCHEMISTRY AND CYTOCHEMISTRY / 3 Chemical Composition of Histologic Samples / 3 Chemical Basis of Staining / 5 Enzyme Digestion / 7 Enzyme Histochemistry / 7 Immunocytochemistry / 7 Hybridization Techniques / 10 Autoradiography / 12 MICROSCOPY / 13 Light Microscopy / 13 Examination of a Histologic Slide Preparation in the Light Microscope / 14 Other Optical Systems / 15 Electron Microscopy / 18 Atomic Force Microscopy / 20 Folder 1.1 Clinical Correlation: Frozen Sections / 4 Folder 1.2 Functional Considerations: Feulgen Microspectrophotometry / 7 Folder 1.3 Clinical Correlation: Monoclonal Antibodies in Medicine / 9 Folder 1.4 Proper Use of the Light Microscope / 11


Histology_Ross

Histology_Ross. The objective of a histology course is to lead the student to understand the microanatomy of cells, tissues, and organs and to correlate structure with function.


Histology_Ross

Histology_Ross. The methods used by histologists are extremely diverse. Much of the histology course content can be framed in terms of light microscopy. Today, students in histology laboratories use either light microscopes or, with increasing frequency, virtual microscopy, which represents a method of viewing a digitized microscopic specimen on a computer screen. In the past, more detailed interpretation of microanatomy was with the electron microscope (EM)—both the transmission electron microscope (TEM) and the scanning electron microscope (SEM). Now the atomic force microscope (AFM) can also provide high-resolution images, which are comparable in resolution to those obtained from TEM. Both EM and AFM, because of their greater resolution and useful magnification, are often the last step in data acquisition from many auxiliary techniques of cell and molecular biology.


Histology_Ross

Histology_Ross. These auxiliary techniques include:  histochemistry and cytochemistry,  immunocytochemistry and hybridization techniques,  autoradiography,  organ and tissue culture,  cell and organelle separation by differential centrifugation, and  specialized microscopic techniques and microscopes. The student may feel removed from such techniques and experimental procedures because direct experience with them is usually not available in current curricula. Nevertheless, it is important to know something about specialized procedures and the data they yield. This chapter provides a survey of methods and offers an explanation of how the data provided by these methods can help the student acquire a better understanding of cells, tissues, and organ function.


Histology_Ross

Histology_Ross. One problem students in histology face is understanding the nature of the two-dimensional image of a histologic slide or an electron micrograph and how the image relates to the three-dimensional structure from which it came. To bridge this conceptual gap, we must first present a brief description of the methods by which slides and electron microscopic specimens are produced. The routinely prepared hematoxylin and eosin–stained section is the specimen most commonly studied.


Histology_Ross

Histology_Ross. The routinely prepared hematoxylin and eosin–stained section is the specimen most commonly studied. The slide set given each student to study with the light microscope consists mostly of formalin-fixed, paraffin-embedded, hematoxylin and eosin (H&E)–stained specimens. Nearly all of the light micrographs in the Atlas section of this book are of slides from actual student sets. Also, most photomicrographs used to illustrate tissues and organs in histology lectures and conferences are taken from such slides. Other staining techniques are sometimes used to demonstrate specific cell and tissue components; several of these methods are discussed below. The first step in preparation of a tissue or organ sample is fixation to preserve structure.


Histology_Ross

Histology_Ross. The first step in preparation of a tissue or organ sample is fixation to preserve structure. Fixation, usually by a chemical or mixture of chemicals, permanently preserves the tissue structure for subsequent treatments. Specimens should be immersed in fixative immediately after they are removed from the body. Fixation is used to:  terminate cell metabolism, prevent enzymatic degradation of cells and tissues by autolysis (self-digestion), kill pathogenic microorganisms such as bacteria, fungi, and viruses, and  harden the tissue as a result of either cross-linking or dena turing protein molecules.


Histology_Ross

Histology_Ross. Formalin, a 37% aqueous solution of formaldehyde, at various dilutions and in combination with other chemicals and buffers, is the most commonly used fixative. Formaldehyde preserves the general structure of the cell and extracellular components by reacting with the amino groups of proteins (most often cross-linked lysine residues). Because formaldehyde does not significantly alter their three-dimensional structure, proteins maintain their ability to react with specific antibodies. This property is important in immunocytochemical staining methods (see page 7). The standard commercial solution of formaldehyde buffered with phosphates (pH 7) acts relatively slowly but penetrates the tissue well. However, because it does not react with lipids, it is a poor fixative of cell membranes. In the second step, the specimen is prepared for embedding in paraffin to permit sectioning.


Histology_Ross

Histology_Ross. In the second step, the specimen is prepared for embedding in paraffin to permit sectioning. Preparing a specimen for examination requires its infiltration with an embedding medium that allows it to be thinly sliced, typically in the range of 5 to 15 m (1 micrometer [ m] equals 1/1,000 of a millimeter [mm]; see Table 1.1). The specimen is washed after fixation and dehydrated in a series of alcohol solutions of ascending concentration as high as 100% alcohol to remove water. In the next step, clearing, organic solvents such as xylol or toluol, which are miscible in both alcohol and paraffin, are used to remove the alcohol before infiltration of the specimen with melted paraffin. TABLE Commonly Used Linear Equivalents1.1 1 picometer (pm) 0.01 angstrom (Å) 1 angstrom 0.1 nanometer (nm) 10 angstroms 1.0 nanometer 1 nanometer 1,000 picometers 1,000 nanometers 1.0 micrometer ( m) 1,000 micrometers 1.0 millimeter (mm)


Histology_Ross

Histology_Ross. When the melted paraffin is cool and hardened, it is trimmed into an appropriately sized block. The block is then mounted in a specially designed slicing machine—a microtome—and cut with a steel knife. The resulting sections are then mounted on glass slides using mounting medium (pinene or acrylic resins) as an adhesive. In the third step, the specimen is stained to permit examination.


Histology_Ross

Histology_Ross. In the third step, the specimen is stained to permit examination. Because paraffin sections are colorless, the specimen is not yet suitable for light microscopic examination. To color or stain the tissue sections, the paraffin must be dissolved out, again with xylol or toluol, and the slide must then be rehydrated through a series of solutions of descending alcohol concentration. The tissue on the slides is then stained with hematoxylin in water. Because the counterstain, eosin, is more soluble in alcohol than in water, the specimen is again dehydrated through a series of alcohol solutions of ascending concentration and stained with eosin in alcohol. Figure 1.1 shows the results of staining with hematoxylin alone, eosin alone, and hematoxylin with counterstain eosin. After staining, the specimen is then passed through xylol or toluol to a nonaqueous mounting medium and covered with a coverslip to obtain a permanent preparation.


Histology_Ross

Histology_Ross. Formalin does not preserve all cell and tissue components. Although H&E–stained sections of formalin-fixed specimens are convenient to use because they adequately display general structural features, they cannot elucidate the specific chemical composition of cell components. Also, many components are lost in the preparation of the specimen. To retain these components and structures, other fixation methods must be used. These methods are generally based on a clear understanding of the chemistry involved. For instance, the use of alcohols and organic solvents in routine preparations removes neutral lipids. To retain neutral lipids, such as those in adipose cells, frozen sections of formalin-fixed tissue and dyes that dissolve in fats must be used; to retain membrane structures, special fixatives


Histology_Ross

Histology_Ross. FIGURE 1.1 • Hematoxylin and eosin (H&E) staining. This series of specimens from the pancreas are serial (adjacent) sections that demonstrate the effect of hematoxylin and eosin used alone and hematoxylin and eosin used in combination. a. This photomicrograph reveals the staining with hematoxylin only. Although there is a general overall staining of the specimen, those components and structures that have a high affinity for the dye are most heavily stained−for example, the nuclear DNA and areas of the cell containing cytoplasmic RNA. b. In this photomicrograph, eosin, the counterstain, likewise has an overall staining effect when used alone. Note, however, that the nuclei are less conspicuous than in the specimen stained with hematoxylin alone. After the specimen is stained with hematoxylin and then prepared for staining with eosin in alcohol solution, the hematoxylin that is not tightly bound is lost, and the eosin then stains those components to which it has a high affinity. c. This


Histology_Ross

Histology_Ross. and then prepared for staining with eosin in alcohol solution, the hematoxylin that is not tightly bound is lost, and the eosin then stains those components to which it has a high affinity. c. This photomicrograph reveals the combined staining effect of H&E. 480.


Histology_Ross

Histology_Ross. containing heavy metals that bind to the phospholipids, such as permanganate and osmium, are used (Folder 1.1). The routine use of osmium tetroxide as a fixative for electron microscopy is the primary reason for the excellent preservation of membranes in electron micrographs. Hematoxylin and eosin are used in histology primarily to display structural features.


Histology_Ross

Histology_Ross. Hematoxylin and eosin are used in histology primarily to display structural features. Despite the merits of H&E staining, the procedure does not adequately reveal certain structural components of histologic sections such as elastic material, reticular fibers, basement membranes, and lipids. When it is desirable to display these components, other staining procedures, most of them selective, can be used. These procedures include the use of orcein and resorcin-fuchsin for elastic material and silver impregnation for reticular fibers and basement membrane material. Although the chemical bases of many staining methods are not always understood, they work. Knowing the components that a procedure reveals is more important than knowing precisely how the procedure works. Specific chemical procedures can provide information about the function of cells and the extracellular components of tissues.


Histology_Ross

Histology_Ross. Specific chemical procedures can provide information about the function of cells and the extracellular components of tissues. Histochemical and cytochemical procedures may be based on specific binding of a dye, use of a ﬂuorescent dye–labeled antibody with a particular cell component, or the inherent enzymatic activity of a cell component. In addition, many large molecules found in cells can be localized by the process of autoradiography, in which radioactively tagged precursors of the molecule are incorporated by cells and tissues before fixation. Many of these procedures can be used with both light microscopic and electron microscopic preparations. Before discussing the chemistry of routine staining and histochemical and cytochemical methods, it is useful to examine briefly the nature of a routinely fixed and embedded section of a specimen. Chemical Composition of Histologic Samples The chemical composition of a tissue ready for routine staining differs from living tissue.


Histology_Ross

Histology_Ross. Chemical Composition of Histologic Samples The chemical composition of a tissue ready for routine staining differs from living tissue. The components that remain after fixation consist mostly of large molecules that do not readily dissolve, especially after treatment with the fixative. These large molecules, particularly those that react with other large molecules to form macromolecular complexes, are usually preserved in a tissue section. Examples of such large macromolecular complexes include:  nucleoproteins formed from nucleic acids bound to protein,  intracellular cytoskeletal proteins complexed with as sociated proteins,  extracellular proteins in large insoluble aggregates, bound to similar molecules by cross-linking of neighbor ing molecules, as in collagen fiber formation, and  FOLDER 1.1 Clinical Correlation: Frozen Sections


Histology_Ross

Histology_Ross. Sometimes, the pathologist may be asked to immediately evaluate tissue obtained during surgery, especially when in-stant pathologic diagnosis may determine how the surgery will proceed. There are several indications to perform such an evaluation, routinely known as a frozen section. Most commonly, a surgeon in the operating room requests a frozen section when no preoperative diagnosis was available or when unexpected intraoperative findings must be identi-fied. In addition, the surgeon may want to know whether all of a pathologic mass within the healthy tissue limit has been re-moved and whether the margin of the surgical resection is free of diseased tissue. Frozen sections are also done in combination with other procedures such as endoscopy or thin-needle biopsy to confirm whether the obtained biopsy material will be usable in further pathologic examinations. Three main steps are involved in frozen section prepa-ration:  Freezing the tissue sample. Small tissue samples are frozen


Histology_Ross

Histology_Ross. obtained biopsy material will be usable in further pathologic examinations. Three main steps are involved in frozen section prepa-ration:  Freezing the tissue sample. Small tissue samples are frozen either by using compressed carbon dioxide or by immersion in a cold fluid (isopentane) at a temperature of 50 C. Freezing can be achieved in a special high-efficiency refrigerator. Freezing makes the tissue solid and allows sectioning with a microtome.  Sectioning the frozen tissue. Sectioning is usually per-formed inside a cryostat, a refrigerated compartment containing a microtome. Because the tissue is frozen solid, it can be cut into extremely thin (5 to 10 m) sec-tions. The sections are then mounted on glass slides.  Staining the cut sections. Staining is done to differen-tiate cell nuclei from the rest of the tissue. The most common stains used for frozen sections are H&E, methylene blue (Fig. F1.1.1), and PAS stains. The entire process of preparation and evaluation of frozen


Histology_Ross

Histology_Ross. nuclei from the rest of the tissue. The most common stains used for frozen sections are H&E, methylene blue (Fig. F1.1.1), and PAS stains. The entire process of preparation and evaluation of frozen sections may take as little as 10 minutes to complete. The total time to obtain results largely depends on the transport time of the tissue from the operating room to the pathology laboratory, on the pathologic technique used, and the expe-rience of the pathologist. The findings are then directly com-municated to the surgeon waiting in the operating room. aba b FIGURE F1.1.1 • Evaluation of a specimen obtained during surgery by frozen-section technique. a. This photomicrograph shows a specimen obtained from the large intestine that was prepared by frozen-section technique and stained with methylene blue. 160 b. Part of the specimen was fixed in formalin and processed as a routine H&E preparation. Examination of the frozen section revealed it to be normal. This diagnosis was later confirmed


Histology_Ross

Histology_Ross. blue. 160 b. Part of the specimen was fixed in formalin and processed as a routine H&E preparation. Examination of the frozen section revealed it to be normal. This diagnosis was later confirmed by examining the routinely prepared H&E specimen. 180. (Courtesy of Dr. Daniel W. Visscher.)  membrane phospholipid–protein (or carbohydrate) complexes.


Histology_Ross

Histology_Ross. These molecules constitute the structure of cells and tissues— that is, they make up the formed elements of the tissue. They are the basis for the organization that is seen in tissue with the microscope. In many cases, a structural element is also a functional unit. For example, in the case of proteins that make up the contractile filaments of muscle cells, the filaments are the visible structural components and the actual participants in the contractile process. The RNA of the cytoplasm is visualized as part of a structural component (e.g., ergastoplasm of secretory cells, Nissl bodies of nerve cells) and is also the actual participant in the synthesis of protein. Many tissue components are lost during the routine preparation of H&E–stained sections.


Histology_Ross

Histology_Ross. Many tissue components are lost during the routine preparation of H&E–stained sections. Despite the fact that nucleic acids, proteins, and phospholipids are mostly retained in tissue sections, many are also lost. Small proteins and small nucleic acids, such as transfer RNA, are generally lost during the preparation of the tissue. As previously described, neutral lipids are usually dissolved by the organic solvents used in tissue preparation. Other large molecules also may be lost, for example, by being hydrolyzed because of the unfavorable pH of the fixative solutions. Examples of large molecules lost during routine fixation in aqueous fixatives are:  glycogen (an intracellular storage carbohydrate common in liver and muscle cells), and  proteoglycans and glycosaminoglycans (extracellular complex carbohydrates found in connective tissue).


Histology_Ross

Histology_Ross. These molecules can be preserved, however, by using a non-aqueous fixative for glycogen or by adding specific binding agents to the fixative solution that preserve extracellular carbohydrate-containing molecules. Soluble components, ions, and small molecules are also lost during the preparation of paraffin sections. chapter 1 TABLE Some Basic and Acidic Dyes1.2 Dye Color Basic dyes Methyl green Green Methylene blue Blue Pyronin G Red Toluidine blue Blue Acidic dyes Acid fuchsin Red Aniline blue Blue Eosin Red Orange G Orange


Histology_Ross

Histology_Ross. Intermediary metabolites, glucose, sodium, chloride, and similar substances are lost during preparation of routine H&E paraffin sections. Many of these substances can be studied in special preparations, sometimes with considerable loss of structural integrity. These small soluble ions and molecules do not make up the formed elements of a tissue; they participate in synthetic processes or cellular reactions. When they can be preserved and demonstrated by specific  At a slightly acidic to neutral pH (5 to 7), sulfate and phos phate groups are ionized and available for reaction with the basic dye by electrostatic linkages.  At low pH (below 4), only sulfate groups remain ionized methods, they provide invaluable information about cell metabolism, active transport, and other vital cellular pro cesses. Water, a highly versatile molecule, participates in these reactions and processes and contributes to the stabilization of macromolecular structure through hydrogen bonding.


Histology_Ross

Histology_Ross. Chemical Basis of Staining Hematoxylin and eosin are the most commonly used dyes in histology. An acidic dye, such as eosin, carries a net negative charge on its colored portion and is described by the general formula [Na dye ]. A basic dye carries a net positive charge on its colored portion and is described by the general formula [dye Cl ]. Hematoxylin does not meet the definition of a strict basic dye but has properties that closely resemble those of a basic dye. The color of a dye is not related to whether it is basic or acidic, as can be noted by the examples of basic and acidic dyes listed in Table 1.2. Basic dyes react with anionic components of cells and tissue (components that carry a net negative charge).


Histology_Ross

Histology_Ross. Basic dyes react with anionic components of cells and tissue (components that carry a net negative charge). Anionic components include the phosphate groups of nucleic acids, the sulfate groups of glycosaminoglycans, and the carboxyl groups of proteins. The ability of such anionic groups to react with a basic dye is called basophilia [Gr., base-loving]. Tissue components that stain with hematoxylin also exhibit basophilia. The reaction of the anionic groups varies with pH. Thus:  At a high pH (about 10), all three groups are ionized and avail able for reaction by electrostatic linkages with the basic dye. and react with basic dyes. Therefore, staining with basic dyes at a specific pH can be used to focus on specific anionic groups; because the specific anionic groups are found predominantly on certain macromolecules, the staining serves as an indicator of these macromolecules.


Histology_Ross

Histology_Ross. As mentioned, hematoxylin is not, strictly speaking, a basic dye. It is used with a mordant (i.e., an intermediate link between the tissue component and the dye). The mordant causes the stain to resemble a basic dye. The linkage in the tissue–mordant–hematoxylin complex is not a simple electrostatic linkage; when sections are placed in water, hematoxylin does not dissociate from the tissue. Hematoxylin lends itself to those staining sequences in which it is followed by aqueous solutions of acidic dyes. True basic dyes, as distinguished from hematoxylin, are not generally used in sequences in which the basic dye is followed by an acidic dye. The basic dye then tends to dissociate from the tissue during the aqueous solution washes between the two dye solutions. Acidic dyes react with cationic groups in cells and tissues, particularly with the ionized amino groups of proteins.


Histology_Ross

Histology_Ross. Acidic dyes react with cationic groups in cells and tissues, particularly with the ionized amino groups of proteins. The reaction of cationic groups with an acidic dye is called acidophilia [Gr., acid-loving]. Reactions of cell and tissue components with acidic dyes are neither as specific nor as precise as reactions with basic dyes. Although electrostatic linkage is the major factor in the primary binding of an acidic dye to the tissue, it is not the only one; because of this, acidic dyes are sometimes used in combinations to color different tissue constituents selectively. For example, three acidic dyes are used in the Mallory staining technique: aniline blue, acid fuchsin, and orange G. These dyes selectively stain collagen, ordinary cytoplasm, and red blood cells, respectively. Acid fuchsin also stains nuclei.


Histology_Ross

Histology_Ross. In other multiple acidic dye techniques, hematoxylin is used to stain nuclei first, and then acidic dyes are used to stain cytoplasm and extracellular fibers selectively. The selective staining of tissue components by acidic dyes is attributable to relative factors such as the size and degree of aggregation of the dye molecules and the permeability and “compactness” of the tissue. Basic dyes can also be used in combination or sequentially (e.g., methyl green and pyronin to study protein synthesis and secretion), but these combinations are not as widely used as acidic dye combinations. A limited number of substances within cells and the extracellular matrix display basophilia.


Histology_Ross

Histology_Ross. A limited number of substances within cells and the extracellular matrix display basophilia. These substances include:  heterochromatin and nucleoli of the nucleus (chiefly because of ionized phosphate groups in nucleic acids of both), cytoplasmic components such as the ergastoplasm (also because of ionized phosphate groups in ribosomal RNA), and  extracellular materials such as the complex carbohydrates of the matrix of cartilage (because of ionized sulfate groups). Staining with acidic dyes is less specific, but more substances within cells and the extracellular matrix exhibit acidophilia. These substances include:  most cytoplasmic filaments, especially those of muscle cells, most intracellular membranous components and much of the otherwise unspecialized cytoplasm, and most extracellular fibers (primarily because of ionized amino groups).


Histology_Ross

Histology_Ross. Certain basic dyes react with tissue components that shift their normal color from blue to red or purple; this absorbance change is called metachromasia. The underlying mechanism for metachromasia is the presence of polyanions within the tissue. When these tissues are stained with a concentrated basic dye solution, such as toluidine blue, the dye molecules are close enough to form dimeric and polymeric aggregates. The absorption properties of these aggregations differ from those of the individual nonaggregated dye molecules. Cell and tissue structures that have high concentrations of ionized sulfate and phosphate groups—such as the ground substance of cartilage, heparin-containing granules of mast cells, and rough endoplasmic reticulum of plasma cells—exhibit metachromasia. Therefore, toluidine blue will appear purple to red when it stains these components. Aldehyde Groups and the Schiff Reagent


Histology_Ross

Histology_Ross. Aldehyde Groups and the Schiff Reagent The ability of bleached basic fuchsin (Schiff reagent) to react with aldehyde groups results in a distinctive red color and is the basis of the periodic acid–Schiff and Feulgen reactions. The periodic acid–Schiff (PAS) reaction stains carbohydrates and carbohydrate-rich macromolecules. It is used to demonstrate glycogen in cells, mucus in various cells and tissues, the basement membrane that underlies epithelia, and reticular fibers in connective tissue. The Feulgen reaction, which relies on a mild hydrochloric acid hydrolysis, is used to stain DNA.


Histology_Ross

Histology_Ross. The PAS reaction is based on the following facts:  Hexose rings of carbohydrates contain adjacent carbons, each of which bears a hydroxyl (–OH) group.  Hexosamines of glycosaminoglycans contain adjacent carbons, one of which bears an –OH group, whereas the other bears an amino (–NH2) group.  Periodic acid cleaves the bond between these adjacent carbon atoms and forms aldehyde groups. These aldehyde groups react with the Schiff reagent to give a distinctive magenta color. The PAS staining of basement membrane (Fig. 1.2) and reticular fibers is based on the content or association of proteoglycans (complex carbohydrates associated with a protein core). PAS staining is an alternative to silver-impregnation methods, which are also based on reaction with the sugar molecules in the proteoglycans.


Histology_Ross

Histology_Ross. The Feulgen reaction is based on the cleavage of purines from the deoxyribose of DNA by mild acid hydrolysis; the sugar ring then opens with the formation of aldehyde groups. Again, the newly formed aldehyde groups react with the FIGURE 1.2 • Photomicrograph of kidney tissue stained by the PAS method. This histochemical method demonstrates and localizes carbohydrates and carbohydrate-rich macromolecules. The basement membranes are PAS positive as evidenced by the magenta staining of these sites. The kidney tubules (T ) are sharply delineated by the stained basement membrane surrounding the tubules. The glomerular capillaries (C) and the epithelium of Bowman’s capsule (BC) also show PAS-positive basement membranes. 360.


Histology_Ross

Histology_Ross.  FOLDER 1.2 Functional Considerations: Feulgen Microspectrophotometry is a technique devel-sion. Currently, Feulgen microspectrophotometry is used to study changes in the DNA content in dividing cells undergoing differentiation. It is also used clinically to analyze abnormal chromosomal number (i.e., ploidy patterns) in malignant cells. Some malignant cells that have a largely diploid pattern are said to be well differentiated; tumors with these types of cells have a better prognosis than tumors with aneuploid (nonintegral multiples of the haploid amount of DNA) and tetraploid cells. Feulgen microspectrophotometry has been particularly useful in studies of specific adenocarcinomas (epithelial cancers), breast cancer, kidney cancer, colon and other gastrointestinal cancers, endometrial (uterine epithelium) cancer, and ovarian cancer. It is one of the most valuable tools for pathologists in evaluating the metastatic potential of these tumors and in making prognostic and treatment


Histology_Ross

Histology_Ross. (uterine epithelium) cancer, and ovarian cancer. It is one of the most valuable tools for pathologists in evaluating the metastatic potential of these tumors and in making prognostic and treatment decisions.


Histology_Ross

Histology_Ross. of reaction. In a typical reaction to display a hydrolytic enzyme, the tissue section is placed in a solution containing a substrate (AB) and a trapping agent (T) that precipitates one of the products as follows: where AT is the trapped end product and B is the hydrolyzed substrate. By using such methods, the lysosome, first identified in differential centrifugation studies of cells, was equated with a vacuolar component seen in electron micrographs. In lightly fixed tissues, the acid hydrolases and esterases contained in lysosomes react with an appropriate substrate. The reaction mixture also contains lead ions to precipitate (e.g., lead phosphate derived from the action of acid phosphatase). The precipitated reaction product can then be observed with both light and electron microscopy.


Histology_Ross

Histology_Ross. Similar light and electron microscopy histochemical procedures have been developed to demonstrate alkaline phosphatase, adenosine triphosphatases (ATPases) of many varieties (including the Na /K -ATPase that is the enzymatic basis of the sodium pump in cells and tissues), various esterases, and many respiratory enzymes (Fig. 1.3). The specificity of a reaction between an antigen and an antibody is the underlying basis of immunocytochemistry.


Histology_Ross

Histology_Ross. Antibodies, also known as immunoglobulins, are glyco proteins that are produced by specific cells of the immune system in response to a foreign protein, or antigen. In the laboratory, antibodies can be purified from the blood and conjugated (attached) to a fluorescent dye. In general, ﬂuorescent dyes (ﬂuorochromes) are chemicals that absorb oped to study DNA increases in developing cells and to analyze ploidy–that is, the number of times the normal DNA content of a cell is multiplied (a normal, nondividing cell is said to be diploid; a sperm or egg cell is haploid). Two techniques, static cytometry for tissue sections and ﬂow cytometry for isolated cells, are used to quantify the amount of nuclear DNA. The technique of static cytometry of Feulgen-stained sections of tumors uses microspectrophotometry coupled with a digitizing imaging system to measure the absorption of light emitted by cells and cell clusters at 560-nm wavelength. In contrast, the flow cytometry technique uses


Histology_Ross

Histology_Ross. coupled with a digitizing imaging system to measure the absorption of light emitted by cells and cell clusters at 560-nm wavelength. In contrast, the flow cytometry technique uses instrumentation able to scan only single cells flowing past a sensor in a liquid medium. This technique provides rapid, quantitative analysis of a single cell based on the measurement of fluorescent light emis-


Histology_Ross

Histology_Ross. Schiff reagent to give the distinctive magenta color. The reaction of the Schiff reagent with DNA is stoichiometric, meaning that the product of this reaction is measurable and proportional to the amount of DNA. It can be used, therefore, in spectrophotometric methods to quantify the amount of DNA in the nucleus of a cell. RNA does not stain with the Schiff reagent because it lacks deoxyribose. Enzyme digestion of a section adjacent to one stained for a specific component—such as glycogen, DNA, or RNA— can be used to confirm the identity of the stained material. Intracellular material that stains with the PAS reaction may be identified as glycogen by pretreatment of sections with diastase or amylase. Abolition of the staining after these treatments positively identifies the stained material as glycogen.


Histology_Ross

Histology_Ross. Similarly, pretreatment of tissue sections with deoxyribonuclease (DNAse) will abolish the Feulgen staining in those sections, and treatment of sections of protein secretory epithelia with ribonuclease (RNAse) will abolish the staining of the ergastoplasm with basic dyes. Histochemical methods are also used to identify and localize enzymes in cells and tissues. To localize enzymes in tissue sections, special care must be taken in fixation to preserve the enzyme activity. Usually, mild aldehyde fixation is the preferred method. In these procedures, the reaction product of the enzyme activity, rather than the enzyme itself, is visualized. In general, a capture reagent, either a dye or a heavy metal, is used to trap or bind the reaction product of the enzyme by precipitation at the site FIGURE 1.3 • Electron histochemical procedure for localization of membrane ATPase in epithelial cells of rabbit gallbladder.


Histology_Ross

Histology_Ross. FIGURE 1.3 • Electron histochemical procedure for localization of membrane ATPase in epithelial cells of rabbit gallbladder. Dark areas visible on the electron micrograph show the location of the enzyme ATPase. This enzyme is detected in the plasma membrane at the lateral domains of epithelial cells, which correspond to the location of sodium pumps. These epithelial cells are involved in active transport of molecules across the plasma membrane. 26,000.


Histology_Ross

Histology_Ross. light of different wavelengths (e.g., ultraviolet light) and then emit visible light of a specific wavelength (e.g., green, yellow, red). Fluorescein, the most commonly used dye, absorbs ultraviolet light and emits green light. Antibodies conjugated with fluorescein can be applied to sections of lightly fixed or frozen tissues on glass slides to localize an antigen in cells and tissues. The reaction of antibody with antigen can then be examined and photographed with a fluorescence microscope or confocal microscope that produces a three-dimensional reconstruction of the examined tissue (Fig. 1.4). Two types of antibodies are used in immunocytochemistry: polyclonal antibodies that are produced by immunized animals and monoclonal antibodies that are produced by immortalized (continuously replicating) antibody-producing cell lines.


Histology_Ross

Histology_Ross. In a typical procedure, a specific protein, such as actin, is isolated from a muscle cell of one species, such as a rat, and injected into the circulation of another species, such as a rabbit. In the immunized rabbit, the rat’s actin molecules are recognized by the rabbit immune system as a foreign antigen. This recognition triggers a cascade of immunologic reactions involving multiple groups (clones) of immune cells called B lymphocytes. The cloning of B lymphocytes eventually leads to the production of anti-actin antibodies. Collectively, these polyclonal antibodies represent mixtures of different antibodies produced by many clones of B lymphocytes that


Histology_Ross

Histology_Ross. FIGURE 1.4 • Confocal microscopy image of a rat cardiac muscle cell. This image was obtained from the confocal microscope using the indirect immunofluorescence method. Two primary antibodies were used. The first primary antibody recognizes a specific lactate transporter (MCT1) and is detected with a secondary antibody conjugated with rhodamine (red). The second primary antibody is directed against the transmembrane protein CD147, which is tightly associated with MCT1. This antibody was detected by a secondary antibody labeled with fluorescein (green). The yellow color is visible at the point at which the two labeled secondary antibodies exactly co-localize within the cardiac muscle cell. This three-dimensional image shows that both proteins are distributed on the surface of the muscle cell, whereas the lactate transporter alone is visible deep to the plasma membrane. (Courtesy of Drs. Andrew P. Halestrap and Catherine Heddle.) each recognize different regions of the actin molecule.


Histology_Ross

Histology_Ross. whereas the lactate transporter alone is visible deep to the plasma membrane. (Courtesy of Drs. Andrew P. Halestrap and Catherine Heddle.) each recognize different regions of the actin molecule. The antibodies are then removed from the blood, purified, and conjugated with a fluorescent dye. They can now be used to locate actin molecules in rat tissues or cells. If actin is present in a cell or tissue, such as a fibroblast in connective tissue, then the fluorescein-labeled antibody binds to it and the reaction is visualized by fluorescence microscopy.


Histology_Ross

Histology_Ross. Monoclonal antibodies (Folder 1.3) are those produced by an antibody-producing cell line consisting of a single group (clone) of identical B lymphocytes. The single clone that becomes a cell line is obtained from an individual with multiple myeloma, a tumor derived from a single antibody-producing plasma cell. Individuals with multiple myelomas produce a large population of identical, homogeneous antibodies with an identical specificity against an antigen. To produce monoclonal antibodies against a specific antigen, a mouse or rat is immunized with that antigen. The activated B lymphocytes are then isolated from the lymphatic tissue (spleen or lymph nodes) of the animal and fused with the myeloma cell line. This fusion produces a hybridoma, an immortalized individual antibody-secreting cell line. To obtain monoclonal antibodies against rat actin molecules, for example, the B lymphocytes from the lymphatic organs of immunized rabbits must be fused with myeloma cells.


Histology_Ross

Histology_Ross. FOLDER 1.3 Clinical Correlation: Monoclonal Antibodies in Medicine Monoclonal antibodies are now widely used in im-munocytochemical techniques and also have many clini-cal applications. Monoclonal antibodies conjugated with radioactive compounds are used to detect and diagnose tumor metastasis in pathology, differentiate subtypes of tumors and stages of their differentiation, and in infec-tious disease diagnosis to identify microorganisms in blood and tissue fluids. In recent clinical studies, mono-clonal antibodies conjugated with immunotoxins, chemotherapy agents, or radioisotopes have been used to deliver therapeutic agents to specific tumor cells in the body. Both direct and indirect immunocytochemical methods are used to locate a target antigen in cells and tissues.


Histology_Ross

Histology_Ross. Both direct and indirect immunocytochemical methods are used to locate a target antigen in cells and tissues. The oldest immunocytochemistry technique used for identifying the distribution of an antigen within cells and tissues is known as direct immunoﬂuorescence. This technique uses a fluorochrome-labeled primary antibody (either polyclonal or monoclonal) that reacts with the antigen within the sample (Fig. 1.5a). As a one-step procedure, this method involves only a single labeled antibody. Visualization of structures is not ideal because of the low intensity of the signal emission. Direct immunofluorescence methods are now being replaced by the indirect method because of suboptimal sensitivity.


Histology_Ross

Histology_Ross. Indirect immunoﬂuorescence provides much greater sensitivity than direct methods and is often referred to as the “sandwich” or “double-layer technique.” Instead of conjugating a fluorochrome with a specific (primary) antibody directed against the antigen of interest (e.g., a rat actin molecule), the fluorochrome is conjugated with a secondary antibody directed against rat primary antibody (i.e., goat anti-rat antibody; Fig. 1.5b). Therefore, when the fluorescein is conjugated directly with the specific primary antibody, the method is direct; when fluorescein is conjugated with a secondary antibody, the method is indirect. The indirect method considerably enhances the fluorescence signal emission from the tissue. An additional advantage of the indirect labeling method is that a single secondary antibody can be used to localize the tissue-specific binding of several different primary antibodies (Fig. 1.6). For microscopic studies, the secondary antibody can be conjugated with different


Histology_Ross

Histology_Ross. antibody can be used to localize the tissue-specific binding of several different primary antibodies (Fig. 1.6). For microscopic studies, the secondary antibody can be conjugated with different fluorescent dyes so that multiple labels can be shown in the same tissue section (see Fig. 1.4). Drawbacks of indirect immunofluorescence are that it is expensive, labor intensive, and not easily adapted to automated procedures.


Histology_Ross

Histology_Ross. It is also possible to conjugate polyclonal or monoclonal antibodies with other substances, such as enzymes FIGURE 1.5 • Direct and indirect immunoﬂuorescence. a. In direct immunofluorescence, a fluorochrome-labeled primary antibody reacts with a specific antigen within the tissue sample. Labeled structures are then observed in the fluorescence microscope in which an excitation wavelength (usually ultraviolet light) triggers the emission of another wavelength. The length of this wavelength depends on the nature of the fluorochrome used for antibody labeling. b. The indirect method involves two processes. First, the specific primary antibodies react with the antigen of interest. Second, the secondary antibodies, which are fluorochrome labeled, react with the primary antibodies. The visualization of labeled structures within the tissue is the same in both methods and requires the fluorescence microscope.


Histology_Ross

Histology_Ross. FIGURE 1.6 • Microtubules visualized by immunocytochemical methods. The behavior of microtubules (elements of the cell cytoskeleton) obtained from human breast tumor cells can be studied in vitro by measuring their nucleation activity, which is initiated by the centrosome. This image was photographed in the fluorescence microscope. By use of indirect immunofluorescence techniques, microtubules were labeled with a mixture of anti– -tubulin and anti– -tubulin monoclonal antibodies (primary antibodies) and visualized by secondary antibodies conjugated with fluorescein dye (fluorescein isothiocyanate–goat anti-mouse immunoglobulin G). The antigen–antibody reaction, performed directly on the glass coverslip, results in visualization of tubulin molecules responsible for the formation of more than 120 microtubules visible on this image. They originate from the centriole and extend outward approximately 20 to 25 m in a uniform radial array. 1,400. (Photomicrograph courtesy of Drs. Wilma L.


Histology_Ross

Histology_Ross. than 120 microtubules visible on this image. They originate from the centriole and extend outward approximately 20 to 25 m in a uniform radial array. 1,400. (Photomicrograph courtesy of Drs. Wilma L. Lingle and Vivian A. Negron.) (e.g., horseradish peroxidase), that convert colorless substrates into an insoluble product of a specific color that precipitates at the site of the enzymatic reaction. The staining that results from this immunoperoxidase method can be observed in the light microscope (Folder 1.4) with either direct or indirect immunocytochemical methods. In another variation, colloidal gold or ferritin (an iron-containing molecule) can be attached to the antibody molecule. These electron-dense markers can be visualized directly with the electron microscope.


Histology_Ross

Histology_Ross. Hybridization is a method of localizing messenger RNA (mRNA) or DNA by hybridizing the sequence of interest to a complementary strand of a nucleotide probe. In general, the term hybridization describes the ability of single-stranded RNA or DNA molecules to interact (hybridize) with complementary sequences. In the laboratory, hybridization requires the isolation of DNA or RNA, which is then mixed with a complementary nucleotide sequence (called a nucleotide probe). Hybrids are detected most often using a radioactive label attached to one component of the hybrid. Binding of the probe and sequence can take place in a solution or on a nitrocellulose membrane. In in situ hybridization, the binding of the nucleotide probe to the DNA or RNA sequence of interest is performed within cells or tissues, such as cultured cells or whole embryos. This technique allows the localization of specific nucleotide sequences as small as 10 to 20 copies of mRNA or DNA per cell.


Histology_Ross

Histology_Ross. Several nucleotide probes are used in in situ hybridization. Oligonucleotide probes can be as small as 20 to 40 base pairs. Single-or double-stranded DNA probes are much longer and can contain as many as 1,000 base pairs. For specific localization of mRNA, complementary RNA probes are used. These probes are labeled with radioactive isotopes (e.g., 32P, 35S, 3H), a specifically modified nucleotide (digoxigenin), or biotin (a commonly used covalent multipurpose label). Radioactive probes can be detected and visualized by autoradiography. Digoxigenin and biotin are detected by immunocytochemical and cytochemical methods, respectively.


Histology_Ross

Histology_Ross. The strength of the bonds between the probe and the complementary sequence depends on the type of nucleic acid in the two strands. The strongest bond is formed between a DNA probe and a complementary DNA strand and the weakest between an RNA probe and a complementary RNA strand. If a tissue specimen is expected to contain a minute amount of mRNA or a viral transcript, then polymerase chain reaction (PCR) amplification for DNA or reverse transcriptase-PCR (RT-PCR) for RNA can be used. The amplified transcripts obtained during these procedures are usually detected using labeled complementary nucleotide probes in standard in situ hybridization techniques. Recently, fluorescent dyes have been combined with nucleotide probes, making it possible to visualize multiple probes at the same time (Fig. 1.7). This technique, called


Histology_Ross

Histology_Ross. Recently, fluorescent dyes have been combined with nucleotide probes, making it possible to visualize multiple probes at the same time (Fig. 1.7). This technique, called FIGURE 1.7 • Example of the FISH technique used in a prenatal screening test. Interphase nuclei of cells obtained from amniotic fluid specimens were hybridized with two specific DNA probes. The orange probe (LSI 21) is locus specific for chromosome 21, and the green probe (LSI 13) is locus specific for chromosome 13. The right nucleus is from a normal amniotic fluid specimen and exhibits two green and two orange signals, which indicates two copies of chromosomes 13 and 21, respectively. The nucleus on the left has three orange signals, which indicate trisomy 21 (Down syndrome). DNA has been counterstained with a nonspecific blue stain (DAPI stain) to make the nucleus visible. 1,250. (Courtesy of Dr. Robert B. Jenkins.)


Histology_Ross

Histology_Ross. FOLDER 1.4 Proper Use of the Light Microscope continued next page This brief introduction to the proper use of the light micro-scope is directed to those students who will use the micro-scope for the routine examination of tissues. If the following comments appear elementary, it is only because most users of the microscope fail to use it to its fullest advan-tage. Despite the availability of today’s fine equipment, rel-atively little formal instruction is given on the correct use of the light microscope. Expensive and highly corrected optics perform optimally only when the illumination and observation beam paths are centered and properly adjusted. The use of proper settings and proper alignment of the optic pathway will contribute substantially to the recognition of minute details in the specimen and to the faithful display of color for the visual image and for photomicrography. Köhler illumination is one key to good microscopy and is incorporated in the design of practically all


Histology_Ross

Histology_Ross. the specimen and to the faithful display of color for the visual image and for photomicrography. Köhler illumination is one key to good microscopy and is incorporated in the design of practically all modern laboratory and research microscopes. Figure F1.4.1 shows the two light paths and all the controls for alignment on a modern laboratory microscope; it should be referred to in following the instructions given below to provide appropri-ate illumination in your microscope. The alignment steps necessary to achieve good Kler illumination are few and simple:  Focus the specimen.  Close the field diaphragm.  Focus the condenser by moving it up or down until the outline of its field diaphragm appears in sharp focus.  Center the field diaphragm with the centering controls on the (condenser) substage. Then open the field diaphragm until the light beam covers the full field observed.  Remove the eyepiece (or use a centering telescope or a phase telescope accessory if available) and


Histology_Ross

Histology_Ross. substage. Then open the field diaphragm until the light beam covers the full field observed.  Remove the eyepiece (or use a centering telescope or a phase telescope accessory if available) and observe the exit pupil of the objective. You will see an illuminated cir-cular field that has a radius directly proportional to the numeric aperture of the objective. As you close the con-denser diaphragm, its outline will appear in this circular field. For most stained materials, set the condenser di-aphragm to cover approximately two thirds of the objec-tive aperture. This setting results in the best compromise between resolution and contrast (contrast simply being the intensity difference between dark and light areas in the specimen). FIGURE F1.4.1 • Diagram of a typical light microscope. This drawing shows a cross-sectional view of the microscope, its operating components, and light path. (Courtesy of Carl Zeiss, Inc., Thornwood, NY.) eyepiece final image exit pupil (eyepoint) real


Histology_Ross

Histology_Ross. This drawing shows a cross-sectional view of the microscope, its operating components, and light path. (Courtesy of Carl Zeiss, Inc., Thornwood, NY.) eyepiece final image exit pupil (eyepoint) real interme-diate image exit pupil of objective specimen condenser diaphragm field diaphragm light source focusing control light source tube objective auxiliary condenser lens stage condenser diaphragm condenser stage control field diaphragm KÖHLER ILLUMINATION THROUGH THE MICROSCOPE IMAGING BEAM PATH ILLUMINATING BEAM PATH the ﬂuorescence in situ hybridization (FISH) procedure, is extensively used in the clinic for genetic testing. For example, a probe hybridized to metaphase chromosomes can be used to identify the chromosomal position of a gene.


Histology_Ross

Histology_Ross. The FISH procedure is used to simultaneously examine chromosomes, gene expression, and the distribution of gene products such as pathologic or abnormal proteins. Many specific ﬂuorescent probes are now commercially available and are used clinically in screening procedures for cervical cancer or for the detection of HIV-infected cells. The FISH procedure can also be used to examine chromosomes from the lymphocytes of astronauts to estimate the radiation dose absorbed by them during their stay in space. The frequency of chromosome translocations in lymphocytes is proportional to the absorbed radiation dose. Autoradiography makes use of a photographic emulsion placed over a tissue section to localize radioactive material within tissues.


Histology_Ross

Histology_Ross. Autoradiography makes use of a photographic emulsion placed over a tissue section to localize radioactive material within tissues. Many small molecular precursors of larger molecules, such as the amino acids that make up proteins and the nucleotides that make up nucleic acids, may be tagged by incorporating a radioactive atom or atoms into their molecular structure. The radioactivity is then traced to localize the larger molecules in cells and tissues. Labeled precursor molecules can be injected into animals or introduced into cell or organ cultures. In this way, synthesis of DNA and subsequent cell division, synthesis and secretion of proteins by cells, and localization of synthetic products within cells and in the extracellular matrix have been studied.


Histology_Ross

Histology_Ross. FIGURE 1.8 • Examples of autoradiography used in light and electron microscopy. a. Photomicrograph of a lymph node section from an animal injected with tritiated [3H]thymidine. Some of the cells exhibit aggregates of metallic silver grains, which appear as small black particles (arrows). These cells synthesized DNA in preparation for cell division and have incorporated the [3H]thymidine into newly formed DNA. Over time, the low-energy radioactive particles emitted from the [3H]thymidine strike silver halide crystals in a photographic emulsion covering the specimen (exposure) and create a latent image (much like light striking photographic film in a camera). During photographic development of the slide with its covering emulsion, the latent image, actually the activated silver halide in the emulsion, is reduced to the metallic silver, which then appears as black grains in the microscope. 1,200. (Original slide specimen courtesy of Dr. Ernst Kallenbach.) b. Electron microscopic


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Histology_Ross. in the emulsion, is reduced to the metallic silver, which then appears as black grains in the microscope. 1,200. (Original slide specimen courtesy of Dr. Ernst Kallenbach.) b. Electron microscopic autoradiograph of the apical region of an intestinal absorptive cell. In this specimen, 125I bound to nerve growth factor (NGF) was injected into the animal, and the tissue was removed 1 hour later. The specimen was prepared in a manner similar to that for light microscopy. The relatively small size of the silver grains aids precise localization of the 125I–NGF complexes. Note that the silver grains are concentrated over apical invaginations (inv) and early endosomal tubular profiles (tub). 32,000. (Electron micrograph courtesy of Dr. Marian R. Neutra.)


Histology_Ross

Histology_Ross. Sections of specimens that have incorporated radioactive material are mounted on slides. In the dark, the slide is usually dipped in a melted photographic emulsion, thus producing a thin photographic film on the surface of the slide. After appropriate exposure in a light-tight box, usually for days to weeks, the exposed emulsion on the slide is developed by standard photographic techniques and permanently mounted with a coverslip. The slides may be stained either before or after exposure and development. The silver grains in the emulsion over the radioactively labeled molecules are exposed and developed by this procedure and appear as dark grains overlying the site of the radioactive emission when examined with the light microscope (Fig. 1.8a).


Histology_Ross

Histology_Ross. These grains may be used simply to indicate the location of a substance, or they may be counted to provide semiquantitative information about the amount of a given substance in a specific location. For instance, after injection of an animal with tritiated thymidine, cells that have incorporated this nucleotide into their DNA before they divide will have approximately twice as many silver grains overlying their nuclei as will cells that have divided after incorporating the labeled nucleotide. Autoradiography can also be carried out by using thin plastic sections for examination with the EM. Essentially the same procedures are used, but as with all TEM preparation techniques, the processes are much more delicate and difficult; however, they also yield much greater resolution and more precise localization (Fig. 1.8b).


Histology_Ross

Histology_Ross. A microscope, whether simple (one lens) or compound (multiple lenses), is an instrument that magnifies an image and allows visualization of greater detail than is possible with the unaided eye. The simplest microscope is a magnifying glass or a pair of reading glasses. TABLE Eye Versus Instrument Resolution1.3 Distance Between Resolvable Points Human eye 0.2 mm Bright-field microscope 0.2 m SEM 2.5 nm TEM Theoretical 0.05 nm Tissue section 1.0 nm Atomic force microscopy 50.0 pm The resolving power of the human eye—that is, the distance by which two objects must be separated to be seen as two objects (0.2 mm)—is determined by the spacing of the photoreceptor cells in the retina. The role of a microscope is to magnify an image to a level at which the retina can resolve the information that would otherwise be below its limit of resolution. Table 1.3 compares the resolution of the eye with that of various instruments.


Histology_Ross

Histology_Ross. Resolving power is the ability of a microscope lens or optical system to produce separate images of closely positioned objects. Resolution depends not only on the optical system but also on the wavelength of the light source and other factors such as specimen thickness, quality of fixation, and staining intensity. With light of wavelength 540 nm (see Table 1.1), a green-filtered light to which the eye is extremely sensitive, and with appropriate objective and condenser lenses, the greatest attainable resolving power of a bright-field microscope would be about 0.2 m (see Folder 1.4, page 12 for method of calculation). This is the theoretical resolution and, as mentioned, depends on all conditions being optimal. The ocular or eyepiece lens magnifies the image produced by the objective lens, but it cannot increase resolution.


Histology_Ross

Histology_Ross. Various light microscopes are available for general and specialized use in modern biologic research. Their differences are based largely on such factors as the wavelength of specimen illumination, physical alteration of the light coming to or leaving the specimen, and specific analytic processes that can be applied to the final image. These instruments and their applications are described briefly in this section. The microscope used by most students and researchers is the bright-field microscope.


Histology_Ross

Histology_Ross. The microscope used by most students and researchers is the bright-field microscope. The bright-field microscope is the direct descendant of the microscopes that became widely available in the 1800s and opened the first major era of histologic research. The bright-field microscope (Fig. 1.9) essentially consists of:  a light source for illumination of the specimen (e.g., a substage lamp), a condenser lens to focus the beam of light at the level of the specimen, a stage on which the slide or other specimen is placed, an objective lens to gather the light that has passed through the specimen, and an ocular lens (or a pair of ocular lenses in the more commonly used binocular microscopes) through which the image formed by the objective lens may be examined directly. cussed earlier are used. Examination of a Histologic Slide Preparation in the Light Microscope Organs are three-dimensional, whereas histologic sections are only two-dimensional.


Histology_Ross

Histology_Ross. cussed earlier are used. Examination of a Histologic Slide Preparation in the Light Microscope Organs are three-dimensional, whereas histologic sections are only two-dimensional. As discussed in the earlier “Tissue Preparation” section, every tissue sample prepared for light microscopic examination must be sliced into thin sections. Thus, two-dimensional sections are obtained from an original three-dimensional sample of tissue. One of the most challenging aspects for students using the microscope to study histology is the ability to mentally reconstruct the “missing” third dimension.


Histology_Ross

Histology_Ross. For example, slices in different planes through an orange are shown in Figure 1.10. Note that each cut surface (indicated by the dotted line) of the whole orange reveals different sizes and surface patterns, depending on the orientation of the cut. Thus, it is important when observing a given section cut through the orange to be able to mentally reconstruct the organization of the structure and its component parts. An example of a histologic structure—in this case, a kidney renal corpuscle—is shown as it would appear in different sectional planes (Fig. 1.10). Note the marked difference in each section of the renal corpuscle. By examining a number of such two-dimensional sections, it is possible to create the three-dimensional configuration of the examined structure. Artifacts in histologic slides can be generated in all stages of tissue preparation.


Histology_Ross

Histology_Ross. Artifacts in histologic slides can be generated in all stages of tissue preparation. The preparation of a histologic slide requires a series of steps beginning with the collection of the specimen and ending with the placement of the coverslip. During each step, an artifact (an error in the preparation process) may be introduced. In general, artifacts that appear on the finished glass slide are linked to methodology, equipment, or reagents used during preparation. The inferior purity of chemicals and reagents used in the process (fixatives, reagents, and stains), imperfections in the execution of the methodology (too short or too long intervals of fixation, dehydration, embedding, staining, or careless mounting and placement of the coverslip), or improper equipment (e.g., a microtome with a


Histology_Ross

Histology_Ross. FIGURE 1.9 • Diagram comparing the optical paths in different types of microscopes. For better comparison between all three types of microscopes, the light microscope (left) is shown as if it were turned upside down; the TEM (middle); and the SEM (right). Note that in both the TEM and the SEM, specimens need to be inserted into the high vacuum (10 4 to 10 7 Pa) environment. image in electron eye detector with CCD camera defective blade) can produce artifacts in the final preparation. It is important for students to recognize that not every slide in their slide collection is perfect and that they should be familiar with the most common artifacts found on their slides.


Histology_Ross

Histology_Ross. Besides bright-field microscopy, which is commonly used for routine examination of histologic slides, other optical systems (described below) are used in clinical and research laboratories. Some of them are used to enhance the contrast without staining (such as phase contrast microscope), whereas others are designed to visualize structures using specific techniques such as immunofluorescence (fluorescence and confocal microscopes). The phase contrast microscope enables examination of unstained cells and tissues and is especially useful for living cells.


Histology_Ross

Histology_Ross. The phase contrast microscope takes advantage of small differences in the refractive index in different parts of a cell or tissue sample. Light passing through areas of relatively high refractive index (denser areas) is deflected and becomes out of phase with the rest of the beam of light that has passed through the specimen. The phase contrast microscope adds other induced, out-of-phase wavelengths through a series of optical rings in the condenser and objective lenses, essentially abolishing the amplitude of the initially deflected portion of the beam and producing contrast in the image. Dark portions of the image correspond to dense portions of the specimen; light portions of the image correspond to less dense portions of the specimen. The phase contrast microscope is therefore used to examine living cells and tissues (such as cells in tissue culture) and is used extensively to examine unstained semithin (approximately 0.5-m) sections of plastic-embedded tissue.


Histology_Ross

Histology_Ross. Two modifications of the phase contrast microscope are the interference microscope, which also allows quantification of tissue mass, and the differential interference microscope (using Nomarski optics), which is especially useful for assessing surface properties of cells and other biologic objects. In dark-field microscopy, no direct light from the light source is gathered by the objective lens. In dark-field microscopy, only light that has been scattered or diffracted by structures in the specimen reaches the objective. The dark-field microscope is equipped with a special condenser that illuminates the specimen with strong, oblique


Histology_Ross

Histology_Ross. FIGURE 1.10 • Example of sections from an orange and a kidney renal corpuscle. The dotted lines drawn on the intact orange indicate the plane of section that correlates with each cut surface. Similarly, different sections through a kidney renal corpuscle, which is also a spherical structure, show differences in appearance. The size and internal structural appearance are reflected in the plane of section. light. Thus, the field of view appears as a dark background on which small particles in the specimen that reflect some light into the objective appear bright. The effect is similar to that of dust particles seen in the light beam emanating from a slide projector in a darkened room. The light reflected off the dust particles reaches the retina of the eye, thus making the particles visible.


Histology_Ross

Histology_Ross. The resolution of the dark-field microscope cannot be better than that of the bright-field microscope, using, as it does, the same wavelength source. Smaller individual particles can be detected in dark-field images, however, because of the enhanced contrast that is created. The dark-field microscope is useful in examining autoradiographs, in which the developed silver grains appear white in a dark background. Clinically, dark-field microscopy is useful in examining urine for crystals, such as those of uric acid and oxalate, and in demonstrating specific bacteria such as spirochetes, particularly Treponema pallidum, the microorganism that causes syphilis, a sexually transmitted disease. The fluorescence microscope makes use of the ability of certain molecules to fluoresce under ultraviolet light.


Histology_Ross

Histology_Ross. The fluorescence microscope makes use of the ability of certain molecules to fluoresce under ultraviolet light. A molecule that fluoresces emits light of wavelengths in the visible range when exposed to an ultraviolet (UV) source. The ﬂuorescence microscope is used to display naturally occurring fluorescent (autofluorescent) molecules such as vitamin A and some neurotransmitters. Because autofluorescent molecules are not numerous, however, the microscope’s most widespread application is the display of introduced fluores cence, as in the detection of antigens or antibodies in immunocytochemical staining procedures (see Fig. 1.6). Specific fluorescent molecules can also be injected into an animal or directly into cells and used as tracers. Such methods have been useful in studying intercellular (gap) junctions, in tracing the pathway of nerve fibers in neurobiology, and in detecting fluorescent growth markers of mineralized tissues.


Histology_Ross

Histology_Ross. Various filters are inserted between the UV light source and the specimen to produce monochromatic or near-monochromatic (single-wavelength or narrow-band–wavelength) light. A second set of filters inserted between the specimen and the objective allows only the narrow band of wavelength of the fluorescence to reach the eye or to reach a photographic emulsion or other analytic processor. The confocal scanning microscope combines components of a light optical microscope with a scanning system to dissect a specimen optically.


Histology_Ross

Histology_Ross. The confocal scanning microscope combines components of a light optical microscope with a scanning system to dissect a specimen optically. The confocal scanning microscope allows visualization of a biologic specimen in three dimensions. The two lenses in the confocal microscope (objective and phototube lens) are perfectly aligned to focus light from the focal point of one lens to the focal point of the other lens. The major difference between a conventional and a confocal microscope is the addition of a detector aperture (pinhole) that is conjugate with the focal point of the lens; therefore, it is confocal. This precisely positioned pinhole allows only “in-focus” light to pass into a photomultiplier (detector) device, whereas the “out-of-focus” light is blocked from entering the detector (Fig. 1.11). This system


Histology_Ross

Histology_Ross. FIGURE 1.11 • Diagram of the in-focus and out-of-focus emitted light in the confocal microscope. a. This diagram shows the path of the laser beam and emitted light when the imaging structure is directly at the focus of the lens. The screen with a pinhole at the other side of the optical system of the confocal microscope allows the light from the structure in focus to pass through the pinhole. The light is then translated into an image by computer software. Because the focal point of the objective lens of the microscope forms a sharp image at the level at which the pinhole is located, these two points are referred to as confocal points. b. This diagram shows the path of the laser beam and the emitted light, which is out of focus in relation to the pinhole. Thus, the light from the specimen that gets blocked by the pinhole is never detected.


Histology_Ross

Histology_Ross. FIGURE 1.12 • Structure of the confocal microscope and diagram of the beam path. The light source for the confocal microscope comes from a laser. The laser beam (red line) travels to the tissue sample via a dichroic beam splitter and then to two movable scanning mirrors; these mirrors scan the laser beam across the sample in both x and y directions. Finally, the laser beam enters the fluorescence microscope and travels through its optical system to illuminate an examined tissue sample. The emitted light by the illuminated tissue sample (blue line) travels back through the optical system of the microscope, through both scanning mirrors, passes through the beam splitter, and is focused onto the pinhole. The light that passes through the pinhole is received and registered by the detector attached to a computer that builds the image one pixel at a time.


Histology_Ross

Histology_Ross. has the capability to obtain exceptional resolution (0.2 to 0.5 m) and clarity from a thin section of a biologic sample simply by rejecting out-of-focus light. The confocal microscope uses an illuminating laser light system that is strongly convergent and therefore produces a high-intensity excitation light in the form of a shallow scanning spot. A mirror system is used to move the laser beam across the specimen, illuminating a single spot at a time (Fig. 1.12). Many single spots in the same focal plane are scanned, and a computer software program reconstructs the image from the data recorded during scanning. In this aspect, confocal microscopy resembles the imaging process in a computed axial tomography (CAT) scan.


Histology_Ross

Histology_Ross. Furthermore, by using only the narrow depth of the in-focus image, it is possible to create multiple images at varying depths within the specimen. Thus, one can literally dissect layer by layer through the thickness of the specimen. It is also possible to use the computer to make three-dimensional reconstructions of a series of these images. Because each individual image located at a specific depth within the specimen is extremely sharp, the resulting assembled three-dimensional image is equally sharp. Moreover, once the computer has assembled each sectioned image, the reconstructed three-dimensional image can be rotated and viewed from any orientation desired (see Fig. 1.4). The ultraviolet microscope uses quartz lenses with an ultraviolet light source.


Histology_Ross

Histology_Ross. The ultraviolet microscope uses quartz lenses with an ultraviolet light source. The image in the ultraviolet (UV) microscope depends on the absorption of UV light by molecules in the specimen. The UV source has a wavelength of approximately 200 nm. Thus, the UV microscope may achieve a resolution of 0.1 m. In principle, UV microscopy resembles the workings of a spectrophotometer; the results are usually recorded photographically. The specimen cannot be inspected directly through an ocular because the UV light is not visible and is injurious to the eye.


Histology_Ross

Histology_Ross. The method is useful in detecting nucleic acids, specifically the purine and pyrimidine bases of the nucleotides. It is also useful for detecting proteins that contain certain amino acids. Using specific illuminating wavelengths, UV spectrophotometric measurements are commonly made through the UV microscope to determine quantitatively the amount of DNA and RNA in individual cells. As described in the Folder 1.2 on page 7, Feulgen microspectrophoto metry is used clinically to evaluate the degree of ploidy (multiples of normal DNA quantity) in sections of tumors. The polarizing microscope uses the fact that highly ordered molecules or arrays of molecules can rotate the angle of the plane of polarized light. The polarizing microscope is a simple modification of the light microscope in which a polarizing filter (the polarizer) is located between the light source and the specimen, and a second polarizer (the analyzer) is located between the objective lens and the viewer.


Histology_Ross

Histology_Ross. Both the polarizer and the analyzer can be rotated; the difference between their angles of rotation is used to determine the degree by which a structure affects the beam of polarized light. The ability of a crystal or paracrystalline array to rotate the plane of polarized light is called birefringence (double refraction). Striated muscle and the crystalloid inclusions in the testicular interstitial cells (Leydig cells), among other common structures, exhibit birefringence. Two kinds of EMs can provide morphologic and analytic data on cells and tissues: the transmission electron microscope and the scanning electron microscope. The primary improvement in the EM versus the light microscope is that the wavelength of the EM beam is approximately 1/2,000 that of the light microscope beam, thereby increasing resolution by a factor of 103. The TEM uses the interaction of a beam of electrons with a specimen to produce an image.


Histology_Ross

Histology_Ross. The TEM uses the interaction of a beam of electrons with a specimen to produce an image. The optics of the TEM are, in principle, similar to those of the light microscope (see Fig. 1.9), except that the TEM uses a beam of electrons rather than a beam of light. The principle of the microscope is as follows:  An electron source (cathode, electron gun), such as a heated tungsten filament, emits electrons.  The electrons are attracted toward an anode.  An electrical difference between the cathode cover and the anode imparts an accelerating voltage of between 20,000 and 200,000 volts to the electrons, creating the electron beam.  The beam then passes through a series of electromag netic lenses that serve the same function as the glass lenses of a light microscope.


Histology_Ross

Histology_Ross. The condenser lens shapes and changes the diameter of the electron beam that reaches the specimen plane. The beam that has passed through the specimen is then focused and magnified by an objective lens and then further magnified by one or more projector lenses. The final image is viewed on a phosphor-coated ﬂuorescent screen or captured on a photographic plate. Portions of the specimen through which electrons have passed appear bright; dark portions of the specimen have absorbed or scattered electrons because of their inherent density or because of heavy metals added during specimen preparation. Often, an electron detector with a light-sensitive sensor such as a charge-coupled device (CCD) are placed above or below the viewing screen to observe the image in real time on a monitor. This allows for uncomplicated procedures of archiving images or videos in digital format on computers.


Histology_Ross

Histology_Ross. Specimen preparation for transmission electron microscopy is similar to that for light microscopy except that it requires finer methods. The principles used in the preparation of sections for viewing with the TEM are essentially the same as those used in light microscopy, with the added constraint that at every step one must work with specimens three to four orders of magnitude smaller or thinner than those used for light microscopy. The TEM, which has an electron beam wavelength of approximately 0.1 nm, has a theoretical resolution of 0.05 nm. Because of the exceptional resolution of the TEM, the quality of fixation—that is, the degree of preservation of sub-cellular structure—must be the best achievable. Routine preparation of specimens for transmission electron microscopy begins with glutaraldehyde fixation followed by a buffer rinse and fixation with osmium tetroxide.


Histology_Ross

Histology_Ross. Routine preparation of specimens for transmission electron microscopy begins with glutaraldehyde fixation followed by a buffer rinse and fixation with osmium tetroxide. Glutaraldehyde, a dialdehyde, preserves protein constituents by cross-linking them; the osmium tetroxide reacts with lipids, particularly phospholipids. The osmium also imparts electron density to cell and tissue structures because it is a heavy metal, thus enhancing subsequent image formation in the TEM. Ideally, tissues should be perfused with buffered glutaraldehyde before excision from the animal. More commonly, tissue pieces no more than 1 mm3 are fixed for the TEM (compared with light microscope specimens, which may be measured in centimeters). The dehydration process is identical to that used in light microscopy, and the tissue is infiltrated with a monomeric resin, usually an epoxy resin, that is subsequently polymerized.


Histology_Ross

Histology_Ross. The plastic-embedded tissue is sectioned on specially designed microtomes using diamond knives. Because of the limited penetrating power of electrons, sections for routine transmission electron microscopy range from 50 nm to no more than 150 nm. Also, for the reason that abrasives used to sharpen steel knives leave unacceptable scratches on sections viewed in the TEM, diamond knives with a nearly perfect cutting edge are used. Sections cut by the diamond knife are much too thin to handle; they are floated away from the knife edge on the surface of a fluid-filled trough and picked up from the surface onto plastic-coated copper mesh grids. The grids have 50 to 400 holes/inch or special slots for viewing serial sections. The beam passes through the holes in the copper grid and then through the specimen, and the image is then focused on the viewing screen, CCD, or photographic film.


Histology_Ross

Histology_Ross. Routine staining of transmission electron microscopy sections is necessary to increase the inherent contrast so that the details of cell structure are readily visible and photographable. In general, transmission electron microscopy sections are stained by adding materials of great density, such as ions of heavy metals, to the specimen. Heavy-metal ions may be bound to the tissues during fixation or dehydration or by soaking the sections in solutions of such ions after sectioning. Osmium tetroxide, routinely used in the fixative, binds to the phospholipid components of membranes, imparting additional density to the membranes. Uranyl nitrate is often added to the alcohol solutions used in dehydration to increase the density of components of cell junctions and other sites. Sequential soaking in solutions of uranyl acetate and lead citrate is routinely used to stain sections before viewing with the TEM to provide high-resolution, high-contrast electron micrographs.


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Histology_Ross. Sometimes, special staining is required to visualize results of histocytochemical or immunocytochemical reactions with the TEM. The phosphatase and esterase procedures are used for this purpose (see Fig. 1.3). Substitution of a heavy metal–containing compound for the fluorescent dye that has been conjugated with an antibody allows the adaptation of immunocytochemical methods to transmission electron microscopy. Similarly, routine EM autoradiography techniques have been refined for use with transmission electron microscopy (see Fig. 1.8b). These methods have been particularly useful in elucidating the cellular sources and intracellular pathways of certain secretory products, the location on the cell surface of specific receptors, and the intracellular location of ingested drugs and substrates. Freeze fracture is a special method of sample preparation for transmission electron microscopy; it is especially important in the study of membranes.


Histology_Ross

Histology_Ross. Freeze fracture is a special method of sample preparation for transmission electron microscopy; it is especially important in the study of membranes. The tissue to be examined may be fixed or unfixed; if it has been fixed, then the fixative is washed out of the tissue before proceeding. A cryoprotectant such as glycerol is allowed to infiltrate the tissue, and the tissue is then rapidly frozen to about 160 C. Ice crystal formation is prevented by the use of cryoprotectants, rapid freezing, and extremely small tissue samples. The frozen tissue is then placed in a vacuum in the freeze fracture apparatus and struck with a knife edge or razor blade. The fracture plane passes preferentially through the hydrophobic portion of the plasma membrane, exposing the interior of the plasma membrane.


Histology_Ross

Histology_Ross. The fracture plane passes preferentially through the hydrophobic portion of the plasma membrane, exposing the interior of the plasma membrane. The resulting fracture of the plasma membrane produces two new surfaces. The surface of the membrane that is backed by extracellular space is called the E-face; the face backed by the protoplasm (cytoplasm) is called the P-face. The specimen is then coated, typically with evaporated platinum, to create a replica of the fracture surface. The tissue is then dissolved, and the surface replica, not the tissue itself, is picked up on grids to be examined with the TEM. Such a replica displays details at the macromolecular level (see Fig. 2.5, page 30). In scanning electron microscopy, the electron beam does not pass through the specimen but is scanned across its surface.


Histology_Ross

Histology_Ross. In scanning electron microscopy, the electron beam does not pass through the specimen but is scanned across its surface. In many ways, the images obtained from SEM more closely resemble those seen on a television screen than on the TEM monitor. They are three-dimensional in appearance and portray the surface structure of an examined sample. For the examination of most tissues, the sample is fixed, dehydrated by critical point drying, coated with an evaporated gold–carbon film, mounted on an aluminum stub, and placed in the specimen chamber of the SEM. For mineralized tissues, it is possible to remove all the soft tissues with bleach and then examine the structural features of the mineral.


Histology_Ross

Histology_Ross. Scanning is accomplished by the same type of raster that scans the electron beam across the face of a television tube. Electrons reflected from the surface (backscattered electrons) and electrons forced out of the surface (secondary electrons) are collected by one or more detectors and reprocessed to form a high-resolution three-dimensional image of a sample surface. In earlier models of microscopes, images were captured on high-resolution cathode ray tube (CRT) or photographic plate; modern instruments, however, capture digital images using sensitive detectors and CCD for display on a high-resolution computer monitor. Other detectors can be used to measure X-rays emitted from the surface, cathodoluminescence of molecules in the tissue below the surface, and Auger electrons emitted at the surface. The scanning-transmission electron microscope (STEM) combines features of the TEM and SEM to allow electron-probe X-ray microanalysis.


Histology_Ross

Histology_Ross. The scanning-transmission electron microscope (STEM) combines features of the TEM and SEM to allow electron-probe X-ray microanalysis. The SEM configuration can be used to produce a transmission image by inserting a grid holder at the specimen level, collecting the transmitted electrons with a detector, and reconstructing the image on a CRT. This latter configuration of an SEM or scanning-transmission electron microscope (STEM) facilitates the use of the instrument for electron-probe X-ray microanalysis.


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Histology_Ross. Detectors can be fitted to the microscope to collect the X-rays emitted as the beam bombards the section; with appropriate analyzers, a map can be constructed that shows the distribution in the sections of elements with an atomic number above 12 and a concentration sufficient to produce enough X-rays to analyze. Semiquantitative data can also be derived for elements in sufficient concentration. Thus, both the TEM and the SEM can be converted into sophisticated analytical tools in addition to being used as “optical” instruments. The atomic force microscope has emerged as one of the most powerful tools for studying the surface topography at molecular and atomic resolution.


Histology_Ross

Histology_Ross. The atomic force microscope has emerged as one of the most powerful tools for studying the surface topography at molecular and atomic resolution. One newer microscope that has proved most useful for biologic studies is the atomic force microscope (AFM). It is a nonoptical microscope that works in the same way as a fingertip, which touches and feels the skin of our face when we cannot see it. The sensation from the fingertip is processed by our brain, which is able to deduce surface topography of the face while touching it.


Histology_Ross

Histology_Ross. In the AFM, an ultrasharp, pointed probe, approaching the size of a single atom at the tip, scans the specimen following parallel lines along the x-axis, repeating the scan at small intervals along the y-axis. The sharp tip is mounted at the end of a highly flexible cantilever so that the tip deflects the cantilever as it encounters the “atomic force” on the surface of the specimen (Fig. 1.13). The upper surface of the cantilever is reflective, and a laser beam is directed off the cantilever to a diode. This arrangement acts as an “optical lever” because extremely small deflections of the cantilever are greatly magnified on the diode. The AFM can work with the tip of the cantilever touching the sample (contact mode), or the tip can tap across the surface (tapping mode) much like the cane of a blind person (Fig. 1.13 insets).


Histology_Ross

Histology_Ross. As the tip moves up and down in the z-axis as it traverses the specimen, the movements are recorded on the diode as movements of the reflected laser beam. A piezoelectric device under the specimen is activated in a sensitive feedback loop with the diode to move the specimen up and down so that the laser beam is centered on the diode. As the tip dips down into a depression, the piezoelectric device moves the specimen up to compensate, and when the tip moves up over an elevation, the device compensates by lowering the specimen. The current to the piezoelectric device is interpreted as the z-axis, which along with the x-and y-axes renders the topography of the specimen at a molecular, and sometimes an atomic, resolution (Fig. 1.14).


Histology_Ross

Histology_Ross. A major advantage of the AFM for examining biologic specimens is that, unlike high-resolution optical instruments (i.e., TEM or SEM), the specimen does not have to be in a vacuum; it can even be in water. Thus, it is feasible to image living cells and their surrounding environments.


Histology_Ross

Histology_Ross. FIGURE 1.13 • Diagram of the atomic force microscope (AFM). An extremely sharp tip on a cantilever is moved over the surface of a biologic specimen. The feedback mechanism provided by the piezoelectric scanners enables the tip to be maintained at a constant force above the sample surface. The tip extends down from the end of a laser-reflective cantilever. A laser beam is focused onto the cantilever. As the tip scans the surface of the sample, moving up and down with the contour of the surface, the laser beam is deflected off the cantilever into a photodiode. The photodiode measures the changes in laser beam intensities and then converts this information into electrical current. Feedback from the photodiode is processed by a computer as a surface image and also regulates the piezoelectric scanner. In contact mode (left inset), the electrostatic or surface tension forces drag the scanning tip over the surface of the sample. In the tapping mode (right inset), the tip of the cantilever


Histology_Ross

Histology_Ross. scanner. In contact mode (left inset), the electrostatic or surface tension forces drag the scanning tip over the surface of the sample. In the tapping mode (right inset), the tip of the cantilever oscillates. The latter mode allows visualization of soft and fragile samples while achieving a high resolution.


Histology_Ross

Histology_Ross. FIGURE 1.14 • Atomic force microscopic image of a single DNA molecule. This image was obtained in the contact mode in which the sharp scanning tip “bumps” up and down as it is moved back and forth over the surface of the sample. The sample lies on an ultrasmooth mica surface. An individual molecule of DNA easily produces enough of a bump to be detected. Thickenings along the DNA molecule are produced by proteins bound to the molecule, and these thickenings produce an even larger movement of the scanning tip. The scan field measures 540 nm by 540 nm. The length of the DNA molecule ranges from 0 to 40 nm. 185,000. (Courtesy of Dr. Gabriela Bagordo, JPK Instruments AG, Berlin, Germany.)


Histology_Ross

Histology_Ross. OVERVIEW OF THE CELL AND CYTOPLASM / 22 MEMBRANOUS ORGANELLES / 25 Plasma Membrane / 25 Endosomes / 35 Lysosomes / 38 Autophagy / 41 Proteasome-Mediated Degradation / 44 Rough-Surfaced Endoplasmic Reticulum / 45 Smooth-Surfaced Endoplasmic Reticulum / 49 Golgi Apparatus / 50 Mitochondria / 53 Peroxisomes (Microbodies) / 56 NONMEMBRANOUS ORGANELLES / 56 Microtubules / 56 Actin Filaments / 59 Intermediate Filaments / 62 Centrioles and Microtubule-Organizing Centers / 65 Basal Bodies / 71 INCLUSIONS / 71 CYTOPLASMIC MATRIX / 73 Folder 2.1 Clinical Correlation: Lysosomal Storage Diseases / 42 Folder 2.2 Clinical Correlation: Abnormalities in Microtubules and Filaments / 68 Folder 2.3 Clinical Correlation: Abnormal Duplication of Centrioles and Cancer / 72 Cells are the basic structural and functional units of all multicellular organisms.


Histology_Ross

Histology_Ross. Cells are the basic structural and functional units of all multicellular organisms. The processes we normally associate with the daily activities of organisms—protection, ingestion, digestion, absorption of metabolites, elimination of wastes, movement, reproduction, and even death—are all reﬂections of similar processes occurring within each of the billions of cells that constitute the human body. To a very large extent, cells of different types use similar mechanisms to synthesize protein, transform energy, and move essential substances into the cell. They use the same kinds of molecules to engage in contraction, and they duplicate their genetic material in the same manner. Specific functions are identified with specific structural components and domains within the cell.


Histology_Ross

Histology_Ross. Specific functions are identified with specific structural components and domains within the cell. Some cells develop one or more of these functions to such a degree of specialization that they are identified by the function and the cell structures associated with them. For example, although all cells contain contractile filamentous proteins, some cells such as muscle cells, contain large amounts of these proteins in specific arrays. This allows them to carry out their specialized function of contraction at both the cellular and tissue level. The specialized activity or function of a cell may be reﬂected not only by the presence of a larger amount of the specific structural component performing the activity but also by the shape of the cell, its organization with respect to other similar cells, and its products (Fig. 2.1). Cells can be divided into two major compartments: the cytoplasm and the nucleus.


Histology_Ross

Histology_Ross. Cells can be divided into two major compartments: the cytoplasm and the nucleus. In general, the cytoplasm is the part of the cell located outside the nucleus. The cytoplasm contains organelles (“little organs”) and inclusions in an aqueous gel called the cytoplasmic matrix. The matrix consists of a variety of solutes, including inorganic ions (Na , K , Ca2 ) and organic molecules such as intermediate metabolites, carbohydrates, lipids, proteins, and RNAs. The cell controls the concentration


Histology_Ross

Histology_Ross. FIGURE 2.1 • Histologic features of different cell types. These three photomicrographs show different types of cells in three different organs of the body. The distinguishing features include size, shape, orientation, and cytoplasmic contents that can be related to each cell’s specialized activity or function. a. Epithelial cells in the kidney. Note several shapes of epithelial cells: columnar cells with well-defned borders in the collecting duct (CD), squamous cells in the thin segment (TS ) of the nephron, and even more-fattened cells lining blood vessels, the vasa recta in the kidney (VR). 380. b. Dorsal root ganglion cells. Note the large size of these nerve cell bodies and the large, pale (euchromatic) nuclei (N) with distinct nucleoli. Each ganglion cell is surrounded by fattened satellite cells (S ). The size of the ganglion cell and the presence of a euchromatic nucleus, prominent nucleolus, and Nissl bodies (rough-surfaced endoplasmic reticulum visible as darker granules


Histology_Ross

Histology_Ross. satellite cells (S ). The size of the ganglion cell and the presence of a euchromatic nucleus, prominent nucleolus, and Nissl bodies (rough-surfaced endoplasmic reticulum visible as darker granules within the cytoplasm) refect the extensive synthetic activity required to maintain the exceedingly long processes (axons) of these cells. 380. C. Smooth muscle cells of the small intestine. Note that these cells are typically elongated, fusiform shaped, and organized in a parallel array. The nuclei are also elongated to conform to the general shape of the cell. 380.


Histology_Ross

Histology_Ross. of solutes within the matrix, which inﬂuences the rate of metabolic activity within the cytoplasmic compartment. The nucleus is the largest organelle within the cell and contains the genome along with the enzymes necessary for DNA replication and RNA transcription. The cytoplasm and nucleus play distinct functional roles but also work in concert to maintain the cell’s viability. The structure and function of the nucleus is discussed in Chapter 3. Organelles are described as membranous (membranelimited) or nonmembranous.


Histology_Ross

Histology_Ross. Organelles are described as membranous (membranelimited) or nonmembranous. Organelles include the membrane systems of the cell and the membrane-limited compartments that perform the metabolic, synthetic, energy-requiring, and energy-generating functions of the cell, as well as nonmembranous structural components. All cells have the same basic set of intracellular organelles, which can be classified into two groups: (1) membranous organelles with plasma membranes that separate the internal environment of the organelle from the cytoplasm, and (2) nonmembranous organelles without plasma membranes.


Histology_Ross

Histology_Ross. The membranes of membranous organelles form vesicular, tubular, and other structural patterns within the cytoplasm that may be convoluted (as in smooth-surfaced endoplasmic reticulum) or plicated (as in the inner mitochondrial membrane). These membrane configurations greatly increase the surface area on which essential physiologic and biochemical reactions take place. The spaces enclosed by the organelles’ membranes constitute the intracellular microcompartments in which substrates, products, and other substances are segregated or concentrated. In addition, each type of organelle contains a set of unique proteins; in membranous organelles, these proteins are either incorporated into their membranes or sequestered within their lumens. For example, the enzymes of lysosomes are separated by a specific enzyme-resistant membrane from the cytoplasmic matrix because their hydrolytic activity would be detrimental to the cell. In nonmembranous organelles, the unique proteins usually


Histology_Ross

Histology_Ross. by a specific enzyme-resistant membrane from the cytoplasmic matrix because their hydrolytic activity would be detrimental to the cell. In nonmembranous organelles, the unique proteins usually self-assemble into polymers that form the structural elements of the cytoskeleton.


Histology_Ross

Histology_Ross. Besides organelles, the cytoplasm contains inclusions, structures that are not usually surrounded by a plasma membrane. They consist of such diverse materials as crystals, pigment granules, lipids, glycogen, and other stored waste products (for details, see page 71).