CHAPTER FIVE:
ANIMAL MICRO-TECHNIQUES
Kabir Mohammed Adamu Department of Biology, IBB University, Lapai |
& | Kate Isioma Iloba Department of Animal and Environmental Biology, Delta State University, Abraka |
5.1 Animal Cell Culture
Animal cell culture is the processes by animal cells are grown in a favorable artificial environment. These cells are derived from multi-cellular eukaryotes with already established cell lines or established cell strains. In the mid-1900s, animal cell culture became a common laboratory technique, but the concept of maintaining live cell lines separated from their original tissue source was discovered in the 19th century. Animal cell culture is now one of the major tools used in the life sciences in areas of research that have a potential for economic value and commercialization.
The development of basic culture media has enabled scientists to work with a wide variety of cells under controlled conditions; this has played an important role in advancing our understanding of cell growth and differentiation, identification of growth factors, and understanding of mechanisms underlying the normal functions of various cell types. New technologies have also been applied to investigate high cell density bioreactor and culture conditions. Many products of biotechnology (such as viral vaccines) are fundamentally dependent on mass culturing of animal cell lines.
At present, cell culture research is aimed at investigating the influence of culture conditions on viability, productivity, and the constancy of posttranslational modifications such as glycosylation, which are important for the biological activity of recombinant proteins. Biological produced by recombinant DNA (rDNA) technology in animal cell cultures include anticancer agents, enzymes, immunobiological [interleukins, lymphokines, monoclonal antibodies (mABs)] and hormones.
Animal cell culture has found use in diverse areas, from basic to advanced research. It has provided a model system for various research efforts:
a. The study of basic cell biology, cell cycle mechanisms, specialized cell function, cell-cell and cell-matrix interactions.
b. Toxicity testing to study the effects of new drugs.
c. Gene therapy for replacing non-functional genes with functional gene-carrying cells.
d. The characterization of cancer cells, the role of various chemicals, viruses, and radiation in cancer cells.
e. Production of vaccines, mABs, and pharmaceutical drugs.
f. Production of viruses for use in vaccine production (e.g., chicken pox, polio, rabies, hepatitis B, and measles).
5.1.1 Methods of Obtaining Cell Culture
There are three methods commonly used to initiate a culture from animals thus;
5.1.1.1 Organ Culture
Organ culture refers to a three-dimensional culture of tissue retaining some or all of the histological features of the tissue in vivo. The whole organ or part of the organ is maintained in a way that allows differentiation and preservation of architecture, usually by culturing the tissue at the liquid-gas interface on a grid or gel. There are disadvantages to organ cultures. Organs cannot be propagated so each piece of tissue can only be used once, which makes it difficult to assess the reproducibility of a response. And, of course, the particular cells of interest may be very small in number in a given piece of tissue so the response produced may be difficult to detect and quantify. It may not be possible to supply adequate oxygen and nutrients throughout the tissue because of the absence of a functioning vascular system, so necrosis of some cells occurs fairly rapidly.
5.1.1.2 Explant/Organotypic Culture
In explant culture, small pieces of the tissue of interest are simply allowed to attach to an appropriate substrate, usually one that has been coated with collagen, and are cultured in a rich medium, usually one containing serum. Following attachment, cell migration is promoted in the plane of the solid substrate. Traditionally, explants have been maintained in Maximov chambers in which cells are grown on coverslips sealed over a depression in a thick glass slide, and this approach is still in use. More recently, it has become common to use regular culture dishes, which are much more convenient since they do not need to be disassembled and reassembled at each feeding. One of the principal advantages of this method is that some aspects of the tissue's architecture can be preserved within the explant.
5.1.1.3 Dissociated Cell Culture
Cell culture refers to cultures derived from dissociated cells taken from the original tissue ('primary cell culture'). Cells are dispersed (mechanically and/or enzymatically) into a cell suspension which may then be cultured as a monolayer on a solid substrate, or as a suspension in the culture medium. These cultures have lost their histotypic architecture and often some of the biochemical properties associated with it. Cell cultures can be characterized and a defined population can be preserved by freezing. The most obvious advantage of cell culture, and of dissociated cell culture in particular, is that it makes individual living cells accessible. All in all, primary dissociated cell cultures are particularly amenable to study using morphological and physiological techniques, which can be applied on a cell by cell basis. They are obviously less well suited to traditional biochemical approaches because the quantity of material obtainable from these cultures is usually limited and they contain a heterogeneous population of cells. One final drawback of working with primary cell cultures is that success is not automatic. Finding the conditions that permit good cell growth and maturation, getting culture to grow reproducibly, and documenting that you have accomplished all of this entails plenty of hard work.
5.1.2 History of cell culture media
Dawn of cultivation experiments (1882–1907): In 1882, Sydney Ringer developed Ringer’s solution, a balanced salt solution of a composition that is close to that of bodily fluids, and successfully kept frog hearts beating after dissection and removal from the body. This was said to be the first instance of in vitro cultivation of animal tissue. Balanced salt solutions were developed one after another in the wake of Ringer’s report. After the success of Ringer’s solution, researchers began to pay attention to cells in culture devices and tried to maintain the cells. Nonetheless, the cells usually did not survive and rarely showed mitotic. In 1907, however, Ross G. Harrison successfully monitored an apparent outgrowth of nerve fibers of a frog for several weeks in lymph fluid that had been freshly drawn from the lymph sacs of an adult frog. This experiment is considered to be the beginning of animal cell cultivation.
Use of natural media (1907-): Alexis Carrel was a French surgeon and biologist who received the Nobel Prize in Physiology and Medicine in 1912 for his research on the vascular suture and transplantation of blood vessels and organs. He contributed greatly to tissue culture technology by devising a prototype of the cell culture flask that is used widely today and by establishing the aseptic manipulation technique. The first success of animal cell culture by Harrison inspired Carrel to send Montrose T. Burrows to work under Harrison’s supervision in 1909. In 1912, Carrel demonstrated that the long-term cultivation of the cells that have been obtained from the connective tissues of chick fetuses is possible (for several months) with a periodic exchange of the medium. In 1913, he discovered that adding embryonic extract to blood plasma can dramatically increase cellular proliferation and extend the culture period of fibroblasts from the chick embryo heart.
Endeavors toward synthetic media (1911-): Margaret R. Lewis and Warren H. Lewis (1911) demonstrated that the Locke–Lewis solution—which is modified Locke’s solution that additionally contains amino acids, bouillon, and glucose (or maltose)—is more effective for chick embryo cell cultivation than simple balanced salt solutions. They reported that glucose is especially important: if the concentration of glucose is not sufficient in the medium, the chick embryo cells show vacuolar degeneration and die within a few days. In contrast, in their search for the active ingredients in embryonic extract, it was ascertained that the active substance is in the protein fraction and that the partially hydrolyzed proteins effectively promote the cell growth of chick embryo fibroblasts. They also confirmed the growth-promoting activity of amino acids and glutathione for chick embryo fibroblasts. They hypothesized that glutathione is required for the control of the redox environment during cell cultivation. Carrel’s medium was supplemented with several natural products, such as casein digests (thymus-derived) nucleic acids, liver ash, and hemoglobin.
Birth of established cell lines (1940-): It is rare for healthy somatic cells that are derived from animals to acquire unlimited proliferative capacity during cultivation. They typically stop growing after a certain number of divisions (i.e. the Hayflick limit). In 1940, Wilton R. Earle et al. used carcinogens to successfully create immortal mouse fibroblasts (L cells), revealing that proliferation from a single cell is possible. In 1951, George O. Gey and his coworkers created an infinitely proliferating human cell line from a tissue of a patient with uterine cervical cancer (HeLa cells). Due to the emergence of these established cell lines, the sampling of cells from the tissues of animals in each experiment became unnecessary, enabling researchers worldwide to perform assays by using the same homogenous population of cells. This state of affairs made it easier to examine and to precisely quantify the subtle differences in the effects of culture media on cells. Thus, the development of culture media advanced rapidly as a result.
Establishment of basal media and research into protein-free media (1946-): Baker’s medium and the other media that had been developed up to this point contained naturally derived components of unknown composition, including plasma, serum, bouillon, peptone, and tissue extracts. In order to find the crucial components in those natural materials and to develop defined media that are comparably efficient in the cultivation of cells, relative to the media containing natural ingredients, two main strategies were undertaken. The first strategy was to use dialyzed serum for the support of cells at minimum levels and to add defined components to maximize the proliferation of cells. The second strategy was not dependent on serum, or even proteins at all, and involved the formulation of media exclusively from definitive components. Fischer was a pioneer of the first strategy. He dialyzed blood plasma to remove the low-molecular- weight fraction. Culture media that were supplemented with dialyzed blood plasma could sustain cells only for a short period, indicating that the low-molecular-weight fraction was essential for the survival of cells. Then, he discovered that the amino acids are the key substance in the low-molecular- weight fraction.
Identification of serum substitutes and the development of serum-free media tailored to a cell type (1970-): Insulin was discovered earlier by Frederick Banting and Charles Best (1921), but full-scale research into this peptide as a supplement for culture media began in the 1960s. Initially, the effectiveness of insulin alone was found to be inferior to that of serum, but the use of insulin in combination with low-concentration serum yielded a higher level of efficacy of baby hamster kidney (BHK) cell growth. This finding led researchers to conclude that insulin acts in a coordinated manner with serum components. Growth factors were discovered one after another during this era: nerve growth factor, epidermal growth factor, insulin-like growth factor, fibroblast growth factor (FGF), platelet-derived growth factor, and transforming growth factor (TGF). The addition of these growth factors to a culture medium increased cellular proliferation. Nevertheless, their effect on cell proliferation, as with insulin, was found to be almost always inferior to the effect of serum. Under these circumstances, in 1976, three key reports were published that accelerated the development of serum-free media. Ham’s group discovered that a trace element of selenite is necessary for the serum-free cultivation of human diploid cells. Larry J. Guilbert and Iscove showed that, besides selenite, a combination of transferring and albumin is a good serum substitute. Izumi Hayashi and Gordon H. Sato discovered that a combination of several hormones and growth factors is an effective serum substitute. Prompted by their discoveries, attempts at serum-free culture by using serum substitutes (e.g. several hormones and growth factors, transferrin, and selenite) grew in number and a variety of serum-free media was developed, with each medium tailored to researchers’ cell type of interest.
Improvements to basal media (1970-): In addition to being a source of hormones, growth factors, carrier proteins, and lipids, serum increases the levels of various low-molecular- weight compounds in basal media. As a result, traditional basal media from which serum is excluded were sometimes unable to adequately support cell growth. The report that the performance of DMEM, MEM, and Ham’s F-12 as basal media in serum-free cultures is inadequate, but that the DMEM/F-12 medium, in which Ham’s F-12 and DMEM are combined in a 1:1 ratio, shows better performance occasionally when used for certain types of cells. The reason seems to be the large number of constituents in Ham’s F-12 and the high concentration of several nutritional constituents in DMEM: mixing the two allows each to complement the weaknesses of the other. Moreover, Murakami et al. reported that the RDF medium - a 2:1:1 mixture of RPMI 1640, DMEM, and Ham’s F-12— yields more effective cell growth of hybridomas than does the DMEM/F12 medium. Mixed media, however, do not always show a level of performance that is better than that of a single medium. For example, the ferrous sulfate that is contained in Ham’s F-12 is toxic to nerve cells and nerve cells proliferate more readily at a reduced osmolarity. Thus, DMEM alone is more effective than DMEM/F-12 in this case. Naturally, the composition of a basal medium that is used for serum-free culture should be optimized for each cell type. In addition, it seems that the optimization also depends on the scale of the culture and its method.
Medical and industrial applications of animal-cell culture technology (1978-): Inspired by the 1982 clinical application of recombinant human insulin expressed in Escherichia coli, researchers actively proceeded to produce growth hormones, interferon α, and other substances by using E. coli or yeast as a host. With E. coli and yeast, however, it was impossible to produce proteins with glycosylation. Animal cells thus started to be used for the production of recombinant proteins, like tissue plasminogen activator, erythropoietin, interferon β, and monoclonal antibodies. The host cells that have been used in the manufacture of biopharmaceutical products include CHO cells, mouse myeloma NS0 cells. BHK cells, human embryonic kidney 293 cells, and human retinal cells. Among these, the CHO and NS0 cells have become especially popular in the field of biopharmaceutical manufacturing for the following reasons:
ü technological advances in mass-culture methods for these two cell lines;
ü sufficient knowledge about the safety of viruses that these two cell lines contain; and
ü remarkable advances in high-expression sub-lines that were derived from these two cell lines.
5.1.3 Applications of Animal Cell Culture
a) Model system: Cell cultures are used as model system to study the basic cell biology and biochemistry. To study the interaction between cell and disease-causing agents like bacteria. To study the effect of drugs. To study the process of aging and used to study triggers of aging. Nutritional study.
b) Cancer research: The difference between the normal and cancer cells can be studied using animal cell culture techniques. Normal cell can be converted into cancer cells by using radiation. Chemicals and viruses. The mechanisms and causes of cancer cells can be studied. Cell culture can be used to determine the effective drugs for selectively destroy only cancer cells.
c) Virology: Animal cell cultures are used to replicate the viruses instead of animals for vaccine production. Cell culture can be used to detect and isolate viruses. It can be used to study growth and development cycle of viruses. It can also be used to study the mode of infection.
d) Toxicity testing: Animal cell culture is used to study the effects of new drugs, cosmetics and chemicals on the survival and growth of a number of types of cell especially liver and kidney cells. Used to determine the maximum permissible dosage of new drugs.
e) Vaccine production: Used in the production of viruses and these viruses are used to produce vaccines. Vaccines such as polio, rabies, chicken pox, measles etc. are produced using animal cell culture.
f) Genetically engineered protein: Animal cell cultures are used to produce commercially important genetically engineered protein such as monoclonal antibodies, insulin, hormones etc.
g) Replacement tissue/organ: Animal cell culture can be used as replacement tissue or organs. E.g. artificial skin can be produced using this technique to treat patients with burns and ulcer.
h) Genetic counseling: Fetal cell culture from pregnant women can be used to study or examine the abnormalities of chromosomes, genes using karyotyping and these findings can be used in early detection of fetal disorders.
i) Genetic engineering: Culture animal cells can be used to introduce new genetic materials like DNA or RNA into the cell. It is used to study the expression of new genes and its effect on the health of the cell. Insect cells are used to produce commercially important proteins by infecting them with genetically altered baculoviruses.
j) Gene therapy: Culture animal cells can be genetically altered and used in gene therapy techniques or use of viral vector.
k) Drug screening and development: They are used to study cytotoxicity of new drugs. Used to find out effective and safe dosage of new drug. Cell –based assay plays an important role in pharmaceutical industry.
5.1.4 Basic Equipment and Facilities in Animal Cell Culture
1) Sterile Work Area: Where possible, a separate room is made available for clean cell culture work. The room should be free of through traffic and, if possible, equipped with an air flow cabinet which supplies filtered air around the work surface. A HEPA (High Efficiency Particle Air Filter) filtered air supply is desirable but not always affordable. Primary animal tissue and micro-organisms must not be cultured in or near the cell culture laboratory and the laboratory must be specifically designated for clean cell culture work. Clean laboratory coats should be kept at the entrance and should not be worn outside of this laboratory and brought back in. If strict sterility is needed, a laminar flow hood offers the best sterile protection available. If a hazardous chemical is to be handled a Class II Biohazard Cabinet which has a vertical laminar flow should be used. However, for primary cultures and also if no laminar flow hood or sterile room is available, an area for sterile work should be set aside, where there is no thoroughfare. If aseptic techniques are adhered to and the area kept clean and tidy, sterility can be easily maintained. All work surfaces, benches and shelves and the base of the airflow cabinets must be kept clean by frequent swabbing with 70% ethanol or an alternative disinfectant. If an airflow cabinet cannot be provided, the culture work may be done on a clean bench using a Bunsen burner to create a sterile 'umbrella' under which the work can be done.
2) Incubation Facilities: In addition to an airflow cabinet and benching which can be easily cleaned, the cell culture laboratory will need to be furnished with an incubator or hot room to maintain the cells at 30-40 °C. The incubation temperature will depend on the type of cells being cultivated. Insect cells will grow best at around 30 °C while mammalian cells require a temperature of 37 °C. It may be necessary to use an incubator which has been designed to allow CO2 to be supplied from a main supply or gas cylinder so that an atmosphere of between 2-5% CO2 is maintained in the incubator. In general, many cell lines can be maintained in an atmosphere of 5% CO2:95% air at 99% relative humidity. The concentration of CO2 is kept in equilibrium with sodium bicarbonate in the medium. Different media have differing buffering capacity. If a CO2 controlled incubator is not available, or cultures must be kept sealed in flasks (i.e., after treatment with some volatile substances), then cells may be maintained in flasks sealed after gassing with 5% CO2:95% air, or vessels kept in boxes gassed and then sealed with pressure sensitive tape. In the case of boxes, the humidity must be maintained with a dish of water. Various media may be used so that a controlled CO2 atmosphere is not required and in this case a CO2 incubator is not necessary. Hepatocytes in primary culture are often maintained in Leibovitz L-15 medium which does not require a CO2 atmosphere; however, flasks must not be sealed (as the hepatocytes require a high O2 tension which is reduced with time in sealed ungassed vessels). Most cell lines are maintained at 36.5 °C, although some cultures, such as skin cultures may require lower temperatures. Cultured cells can generally survive lower temperatures, but rarely survive temperatures greater than 2 °C above normal, and therefore the incubator should be set to cut out at approximately 38.5 °C to prevent cell death. Incubators are designed to regulate an even temperature and this is more important than accuracy, i.e., temperature should be ±0.5°
3) Refrigerators and Freezer (-20°C): Both items are very important for storage of liquid media at 4°C and for enzymes (e.g., trypsin) and some media components (e.g., glutamine and serum) at -20°C. A refrigerator or cold room is required to store medium and buffers. A freezer will be needed for keeping pre-aliquoted stocks of serum, nutrients and antibiotics. Reagents may be stored at a temperature of -20°C but if cells are to be preserved it may be necessary to provide liquid nitrogen or a -70°C freezer.
4) Microscopes: A simple inverted microscope is essential so that cultures can be examined in flasks and dishes. It is vital to be able to recognize morphological changes in cultures since these may be the first indication of deterioration of a culture. A very simple light microscope with x100 magnification will suffice for routine cell counts with a hemocytometer, although a microscope of much better quality will be required for chromosome analysis or autoradiography work. A microscope with normal Kohler illumination will be needed for cell counting. An inverted microscope will also be needed for examining flasks and multi-well dishes from underneath. Both microscopes should be equipped with a x10 and a x20 objective and it may be useful to provide a x40 and a x100 objective for the normal microscope. Additional features such as a camera, CCD video camera, adapter and attachments and UV facility may also be required for some purposes.
5) Tissue Culture Ware: A variety of tissue culture plasticware is available, the most common being specially treated polystyrene. Although all tissue culture plasticware should support cell growth adequately, it is essential when using a new supplier or type of dish to ensure that cultures grow happily in it. The tests to ensure this, such as growth curves and time of reaching a confluent monolayer, are similar, to those used to ensure that serum batches are satisfactory. Cells can be maintained in Petri dishes or flasks (25 cm2 or 75 cm2) which have the added advantage that the flasks can be gassed and then sealed so that a CO2 incubator need not be used. This is particularly useful if incubators fail. Tissue culture ware is always chosen to match the procedure. Sometimes it may be necessary to condition a surface by pretreatment with 'spent' medium which has been used with another culture (conditioned medium). The choice of vessel depends on several factors: whether the culture is in suspension or grows as a monolayer; the cell yield; whether it needs CO2 or not; and what form of sampling is to be taken place. Cost can also be a limiting factor. Cell yield is proportional to available surface areas. It is important to ensure that an even monolayer can grow, especially in the currently popular multi-well dishes (24, 48 and 96 wells). For adherent cells to which histological stains may be applied, cover slips fitted into multi-well dishes which can be removed and treated with various organic solvent in staining are required. Commercially available multi-well slide-chamber dishes are also a convenient, but costly, alternative. Normal tissue culture ware is not resistant to organic solvents.
6) Washing Up and Sterilizing Facilities: Availability of a wide range of plastic tissue culture reduces the amount of necessary washing up. However, glassware such as pipettes should be soaked in a suitable detergent, then passed through a stringent washing procedure with thorough soaking in distilled water prior to drying and sterilizing. Pipettes are often plugged with nonabsorbent cotton wool before putting into containers for sterilizing. Glassware, such as pipettes, conical flasks, beakers (covered with aluminum foil) are sterilized in a hot air oven at 160 °C for one hour. All other equipment, such as automatic pipette tips and bottles (lids loosely attached) are autoclaved at 121 °C for 20 min. Sterilizing indicators such as sterile test strip is necessary for each sterilizing batch to ensure that the machine is operating effectively. Autoclave bags are available for loose items. Aluminum foil also makes good packaging material.
7) Liquid N2/Deep Freezer: Invariably for continuous and finite cell lines, samples of cultures will need to be frozen down for storage. It is important to maintain continuity in cells to prevent genetic drift and to guard against loss of the cell line through contamination and other disasters. The procedure for freezing cells is general for all cells in culture. They should be frozen in exponential phase of growth with a suitable preservative, usually dimethylsulfoxide (DMSO). The cells are frozen slowly at 1 °C/min to -50°C and then kept either at -196°C immersed in liquid N2 (in sealed glass ampoules) or above the liquid surface in the gas phase (screw top ampoules). Deterioration of frozen cells has been observed at -70°C, therefore, -196°C (liquid N2) seems to be necessary. To achieve slow freezing rates a programmable freezer or an adjustable neck plug or freezing tray for use in a narrow-necked liquid nitrogen freezer can be used. Alternatively, ampoules may be frozen in a polystyrene box with 1" thick walls. This will insulate the ampoules to slow the freezing process to 1 °C/min in a -70°C freezer.
5.2 Preparation of Histological Sections
In order to prepare thin sections for examination by microscopy, it is necessary to preserve the tissues (fixation) and embed them in a supporting medium (such as paraffin wax or resins) prior to sectioning. Sections are usually stained in order to provide contrast.
5.2.1 Fixation
The process through which cell structure is preserved is called fixation. Since cells rapidly deteriorate after a tissue has been removed from the body, achieving adequate fixation is often the most difficult task confronting a histologist. "Artifacts" are changes to the original structure of cells and tissues that arise from tissue deterioration and from the fixation process itself. Thus, a skilled histologist employs techniques that minimize the formation of artifacts in different types of tissues, and has is the ability to distinguish artifacts from normal cell structures.
Fixation of tissue is done for several reasons. One reason is to kill the tissue so that postmortem decay (autolysis and putrefaction) is prevented. Fixation preserves a sample of biological material (tissue or cells) as close to its natural state as possible in the process of preparing tissue for examination. The fixative needs to preserve the tissues as close as possible to the living state. These fixatives have the ability to either stabilize or denature proteins.
The primary function of a fixative is to preserve the cellular structure of the tissue. Fixation is necessary to protect and harden the tissue against the deleterious effects of later procedures which otherwise would disrupt cellular structure beyond recognition. Furthermore, fixation minimizes a process called autolysis. Autolysis is the degradation of cellular structure which results from the release of degradative enzymes from the excised tissue itself.
There are varying types of fixatives such as chemical fixatives. A widely used example is formaldehyde, which has the advantage of being cheap and penetrates tissues rapidly. For better fixation, it is necessary to use pH buffers in the fixative. The most common fixative for light microscopy is 10% neutral buffered formalin (4% formaldehyde in phosphate buffered saline). For electron microscopy, the most commonly used fixative is glutaraldehyde, usually at a 2.5% solution in phosphate buffered saline. These fixatives preserve tissues or cells mainly by irreversibly cross-linking proteins. The main action of these aldehyde fixatives is to cross-link amino groups in proteins through the formation of CH2 (methylene) linkage, in the case of formaldehyde, or by a C5H10 cross-links in the case of glutaraldehyde. Other fixatives often used for electron microscopy are osmium tetroxide or uranyl acetate. Formalin fixation leads to degradation of mRNA, mRNA and DNA in tissues.
Frozen section fixation: this is a rapid way to fix and mount histology sections. It is used in surgical removal of tumors, and allow rapid determination of margin (that the tumor has been completely removed). It is done using a refrigeration device called a cryostat. The frozen tissue is sliced using a microtome, and the frozen slices are mounted on a glass slide and stained the same way as other methods. It is a necessary way to fix tissue for certain stain such as antibody linked immunofluorescence staining. It can also be used to determine if a tumor is malignant when it is found incidentally during surgery on a patient.
5.2.1.1 Types of Fixations
There are generally three types of fixation process:
a. Heat fixation: After a smear has dried at room temperature, the slide is gripped by tongs or a clothespin and passed through the flame of a Bunsen burner several times to heat-kill and adhere the organism to the slide. This generally preserves overall morphology but not internal structures because it denatures the proteolytic enzyme and prevent autolysis.
b. Perfusion: forceful flooding of tissue. The fixative is injected into the heart with the injection volume matching cardiac output. The fixative spreads through the entire body, and the tissue doesn't die until it is fixed. This has the advantage of preserving perfect morphology, but the disadvantages that the subject dies and the cost is high (because of the volume of fixative needed for larger organisms)
c. Immersion: In this the sample of tissue is immersed in fixative of volume at a minimum of 20 times greater than the volume of the tissue to be fixed. The fixative must diffuse through the tissue to fix, so tissue size and density, as well as type of fixative must be considered. Using a larger sample means it takes longer for the fixative to reach the deeper tissue. Best in a slight vacuum.
5.2.2 Dehydration
Tissues fixed in aqueous solutions will maintain a high-water content, a condition that can be a hindrance to later processing. Except in special cases (freezing method, water-soluble waxes, and special cell contents), the tissue must be dehydrated (water removed) before certain steps in this processing can be successful. Dehydration is achieved using an ascending series of alcohols (70%, 95%, 100%).
5.2.3 Embedding
This is a process where dehydrated and cleared tissue are infiltrated with the embedding material. During this process the tissue samples are placed into molds along with liquid embedding material (such as agar, gelatin, or wax) which is then hardened. The most commonly used embedding or support medium is paraffin wax, with a melting point of about 56°C. The tissue is then transferred to molten paraffin wax (in an embedding oven) for a couple of hours. The tissue is then placed in a square or rectangular mold, and orientated in the required position, prior to adding hot wax to form a wax block. Embedding can also be accomplished using frozen, non-fixed tissue in a water-based medium. Pre-frozen tissues are placed into molds with the liquid embedding material, usually a water-based glycol, Cryogen, or resin, which is then frozen to form hardened blocks.
5.2.4 Microtomy
Sections of the tissue embedded in the wax block are cut on a machine, known as a microtome, using special knives (nowadays these are disposable). Typically, series or ribbons of sections are cut at a thickness of 6-8μm. The sections are transferred to the surface of a hot water-bath (where the sections flatten and lose any wrinkles). Sections are collected on glass microscope slides (standard dimensions of 3 x 1 inches). In order for the sections to adhere to the slides they are dried for up to 24 hours in a drying oven (at a temperature of about 40°C). This prevents sections falling off the slides in the later stages of preparation.
5.2.5 Staining
The most common staining technique is known as Hematoxylin and Eosin (H&E) staining. In order to stain the sections, the wax needs to be removed. This is done using a wax solvent such as xylene. The slide is then hydrated using a series of descending alcohols (100%, 95%, 70%) and then water. The slide is then immersed in Hematoxylin stain, rinsed in running water (preferably alkaline), followed by staining with Eosin, and rinsing in water.
Staining is an auxiliary technique used in microscopy to enhance contrast in the microscopic image. Stains and dyes are frequently used in biology and medicine to highlight structures in biological tissues for viewing, often with the aid of different microscopes. Stains may be used to define and examine bulk tissues (highlighting, for example, muscle fibers or connective tissue), cell populations (classifying different blood cells, for instance), or organelles within individual cells. Biological staining is also used to mark cells in flow cytometry, and to flag proteins or nucleic acids in gel electrophoresis. In vivo staining is the process of dyeing living tissues—in vivo means "in life" (compare with in vitro staining). By causing certain cells or structures to take on contrasting colour(s), their form (morphology) or position within a cell or tissue can be readily seen and studied. The usual purpose is to reveal cytological details that might otherwise not be apparent; however, staining can also reveal where certain chemicals or specific chemical reactions are taking place within cells or tissues.
In vitro staining involves colouring cells or structures that are no longer living. Certain stains are often combined to reveal more details and features than a single stain alone. Combined with specific protocols for fixation and sample preparation, scientists and physicians can use these standard techniques as consistent, repeatable diagnostic tools. A counterstain is stain that makes cells or structures more visible, when not completely visible with the principal stain.
The different types of staining techniques are:
i. Haematoxylin and eosin staining protocol is used frequently in histology to examine thin sections of tissue. Haematoxylin stains cell nuclei blue, while eosin stains cytoplasm, connective tissue and other extracellular substances pink or red. Eosin is strongly absorbed by red blood cells, colouring them bright red. In a skillfully made H & E preparation the red blood cells are almost orange, and collagen and cytoplasm (especially muscle) acquire different shades of pink. When the staining is done by a machine, the subtle differences in eosinophilia are often lost. Hematoxylin stains the cell nucleus and other acidic structures (such as RNA-rich portions of the cytoplasm and the matrix of hyaline cartilage) blue. In contrast, eosin stains the cytoplasm and collagen pink. The Hematoxylin is a basic dye that stains acidic components of cells a blue color. This characteristic is known as basophilia. Hematoxylin stains the nuclei of cells, and the RER of the cytoplasm. Eosin is an acidic dye that stains the basic components of the cells a reddish-pink color. This characteristic is known as acidophilia. Most of the cytoplasm of cells is stained by eosin. Bone matrix is also stained by eosin.
ii. Papanicolaou staining, or Pap staining, is a frequently used method for examining cell samples from various bodily secretions. It is frequently used to stain Pap smear specimens. It uses a combination of haematoxylin, Orange G, eosin Y, Light Green SF yellowish, and sometimes Bismarck Brown Y.
iii. PAS staining - Periodic acid-Schiff staining is used to mark carbohydrates (glycogen, glycoprotein, proteoglycans). It is used to distinguish different types of glycogen storage diseases. PAS is a widely used staining technique that stains the neutral sugars of glycosaminoglycans a pink color. Common components stained positively with PAS include mucus, the basal lamina and glycogen.
iv. Masson's trichrome is (as the name implies) a three-colour staining protocol. The recipe has evolved from Masson's original technique for different specific applications, but all are well-suited to distinguish cells from surrounding connective tissue. Most recipes will produce red keratin and muscle fibers, blue or green staining of collagen and bone, light red or pink staining of cytoplasm, and black cell nuclei.
v. The Romanowsky stains are all based on a combination of eosinate (chemically reduced eosin) and methylene blue (sometimes with its oxidation products azure A and azure B). Common variants include Wright's stain, Jenner's stain, Leishman stain and Giemsa stain. All are used to examine blood or bone marrow samples. They are preferred over H&E for inspection of blood cells because different types of leukocytes (white blood cells) can be readily distinguished. All are also suited to examination of blood to detect blood-borne parasites like malaria.
vi. Silver staining is the use of silver to stain histologic sections. This kind of staining is important especially to show proteins (for example type III collagen) and DNA. It is used to show both substances inside and outside cells. Silver staining is also used in temperature gradient gel electrophoresis. Some cells are argentaffin. These reduce silver solution to metallic silver after formalin fixation. This method was discovered by Italian Camillo Golgi, by using a reaction between silver nitrate and potassium dichromate, thus precipitating silver chromate in some cells (see Golgi's method). Other cells are argyrophilic. These reduce silver solution to metallic silver after being exposed to the stain that contains a reductant, for example hydroquinone or formalin.
vii. Sudan staining is the use of Sudan dyes to stain sudanophilic substances, usually lipids. Sudan III, Sudan IV, Oil Red O, and Sudan Black B are often used. Sudan staining is often used to determine the level of fecal fat to diagnose steatorrhea.
viii. Orcein staining is used to stain elastic fibers a dark brown-purple color. This is used, for example, to show the elastic components in the walls of arteries, or in the matrix of elastic cartilage.
ix. Oil Red O is used to stain lipids a red-orange color in unfixed frozen sections.
a. Toluidine blue is a so-called metachromatic stain. It is a blue stain that stains specific components of tissues a purple color. This change in staining color is known as metachromasia. Metachromasia is seen in the matrix of hyaline cartilage, or in the granules of mast cells.
b. Impregnation is a staining technique in which blocks of tissue are processed in solutions containing metals such as silver or gold, which attach to specific
c. components in tissues. The silver or gold are then further processed (reduced) and develop into dark metallic deposits. The stained blocks are only then sectioned. Silver impregnation is widely used in neurohistology to stain neurons and their processes. Silver impregnation techniques are also widely used to demonstrate reticular fibers.
5.2.6 Paraffin Method of Section Preparation
This method is most and widely employed. Although this technique is not universally applicable, e.g. it does not work well with hard tissues such as bones from animals, it does present many advantages over alternative methods. The necessary reagents are inexpensive, readily available, and much less toxic to humans than those used in most other techniques. The steps for the method are;
a. Tissue Resection & Fixation
This occurs when an animal is sacrificed to remove (resect) certain organs (such as lungs, kidneys and liver). The ways in which for instance rodents can be sacrificed ethically includes by inhalation of carbon dioxide, methoxyflurane, or halothane. If exposure to chemicals contradicts the objectives of the investigation, cervical dislocation (rupturing of the spinal column in the neck) can be performed. Or the use of a rodent guillotine to decapitate sedated mice. This is instantaneous and painless, and allows bleeding out of the blood, which otherwise fills the body cavity during dissection. It also allows the organs to drain of blood that might interfere with an analysis of organ-specific proteins. The fixation process must start as soon as possible after resection of the sample by
1. Labeling the tissue cassettes in pencil as “lungs”
2. Fill a vial about 2/3 full with the fixative.
3. Remove the organ from the rodent (o.5g may be appropriate) by placing on a Petri plate chilled in ice bucket. Then transfer it into cryostorage vial and store on ice until transferring it to the -80OC freezer.
b. Dehydration
After fixation, the water must be removed from the tissue block, a process called dehydration. Isopropyl alcohol (IPA) is a favored reagent because it is miscible in paraffin. The tissue must not be dehydrated rapidly because this will cause distortion of the tissue. Rather, dehydration is carried out in a slow, step-wise manner by passing the tissue block through a series of solutions of increasing IPA concentration. In this way the water is fully leached out and replaced with IPA.
a. 70% IPA for 1 hr
b. 70% IPA for 1 hr
c. 85% IPA for 1 hr
d. 95% IPA for 1 hr
e. 100% IPA for 1 hr
f. 100% IPA for 1 hr
c. Infiltration and Embedding in Paraffin
Prior to sectioning, the tissue block must be infiltrated with a material that acts as a support during the sectioning process. For the method described here, paraffin serves this purpose. During infiltration, the paraffin will equilibrate within the tissue block, eventually occupying all of the space in the tissue that originally held by IPA. After infiltration, the tissue is allowed to solidify in a mold, embedded within a small cube of paraffin. Infiltration involves;
a. Discarding the 100% IPA from the last dehydration step, and fill the vial about 3/4 full with melted paraffin.
b. Allow the tissue to equilibrate for 1 hour in an incubator set at 58OC. Equilibration means to allow a solution to reach a stable concentration within a tissue. Thus, for example, after 1 hour the IPA will have reached 70% within the tissue block.
c. Pour the paraffin into the container labeled for paraffin disposal.
d. Repeat step ‘a’ using fresh melted paraffin.
Embedding involves;
a. Placing a base-pieces for two embedding molds in a plastic Petri plate – label the plate along the edge appropriately
b. Decant the paraffin from the second infiltration step into the waste container
c. Working quickly but carefully, use forceps to transfer the tissue blocks to the well of separate base mold, snap the base of tissue cassette into the base mold and then fill the mold with paraffin.
d. Allow the paraffin to solidify at room temperature (if the paraffin begins to solidify homogeneously around the tissue block, allow the paraffin in the base mold to melt in the incubator, and then allow it to solidify).
5.2.7 Sectioning with a Microtome
This is accomplished by using a cutting apparatus called a microtome. The microtome will drive a knife across the surface of the paraffin cube and produce a series of thin sections of very precise thickness. The objective is to produce a continuous "ribbon" of sections adhering to one another by their leading and trailing edges. The thickness of the sections can be preset, and a thickness between 5 - 10 μm is optimal for viewing with a light microscope. The sections can then be mounted on individual microscope slides. Preparation and mounting of the embedded tissue block on the microtome is very important to successful sectioning. The paraffin surrounding the tissue block must be first trimmed, and then secured to a holder which is then mounted on the microtome.
5.2.8 Mounting on Microscope Slides
This is permanently attaching the section to microscope slides. If "serial" sections are desired, (i.e., sections that reveal sequential layers of the tissue structure) then sectioning must be performed carefully and systematically. Label the microscope slides appropriately. Wash the microscope slides with soap and water, and rinse free of soap with tap water. Place the slides in a coplin jar and rinse several times with roH2O. Handling the slides only by their edges, place the slides in your slide storage box, and allow to dry.
However, note that during sectioning the sections are not perfectly flat, but rather slightly crinkled. This is normal, and the sections will become flattened by floating them on water held at 45OC. The solution also contains an adhesive, Surgipath/Leica, which causes the tissue section to bind to the slide. Carefully transfer the sections to a solution held in a 45OC water bath. Within a few seconds sections flatten and the wrinkles disappear. Dip a clean microscope slide into the adhesive solution, and slowly pull it upward, out of the solution, allowing sections to adhere to the surface. Make sure that the slide is oriented with the label facing upward. Dry the bottom of the slide and carefully blot excess adhesive from around the sections (be careful not to touch the sections themselves). Allow the slides to dry overnight in the storage box.
5.2.9 Clearing and Staining
Before a section can be stained the paraffin must be removed, a process called clearing. After clearing, only the tissue remains adhering to the slide. Clearing is accomplished by passing the mounted sections through the solvent Clearene (Surgipath/Leica) that dissolves the paraffin. Staining of histological sections allows observation of features otherwise not
distinguishable. For routine histological work, it is customary to use two dyes, one that stains certain components a bright color and the other, called the counterstain that stains other cellular structures a contrasting color. While literally hundreds of staining techniques have been developed, the two stains most widely used for routine work are hematoxylin and Eosin Y. Hematoxylin stains negatively charged structures, such as DNA, a blue color. Eosin imparts a red color to most of the other cell components. To produce permanent staining with hematoxylin, the dye must be oxidized to "hematein", which is achieved by treating the tissue sections with Scott's solution. Clear and stain your slides with the following schedule of solutions held in Coplin jars:
a. Clearing and Rehydration: Clearing agent 1 for 3minutes; Clearing agent 2 for 2minutes and Clearing agent 3 for 1minute. 100% IPA for 30seconds; 85% IPA for 30seconds; 70% IPA for 30 seconds and Tap water for 30 seconds
b. Staining: Hematoxylin for 2 minutes; Tap water for 30 seconds; Scott’s solution for 1 minute; Tap water for 30 seconds; Buffer for 1 minute; Tap water for 30 seconds; 70% IPA for 1 minute; 95% IPA for 1 minute; Eosin Y for 1 minute.
c. Rinsing, Rehydration & Mounting Prep 95% IPA for 2-3 minutes; 100% IPA for 2-3 minutes; Clearing Agent for 1 minute; Clearing Agent for 1 minute and Clearing Agent for 1 minute
5.2.10 Preparation of permanent mounts
This is the final step in this procedure where the mount section is under a coverslip. This is accomplished by covering the section in a medium that will harden and produce a clear binder between the slide and cover slip. The ideal mounting medium should not distort the stain color, or yellow and become brittle with age. Thus, the use of mounting resin called Permount (Fisher Scientific). The procedure is;
a. Place 2-3 drops of resin over the section.
b. To avoid entrapping air bubbles, lower the cover slip slowly from one side of the droplet.
c. Place the slide on the slide warmer and carefully place a lead weight on top of the cover slip. There should be enough mounting medium to completely cover the bottom of the cover slide, and budge slightly around the edges.
d. Leave slides on the warmer for at least 24 hours; excess medium can then be cut from edges of cover slip with a razor blade.
e. Do not allow slides to dry before mounting under cover slides.
5.3 Preparation of Histological Sections of Bone
Because bone tissue is hard and calcified, special histological techniques are used to prepare sections.
i. Decalcification: The most common techniques involve calcium removal from the tissue (decalcification) after fixation and prior to wax embedding. Acids, such as formic acid or nitric acid, can be used as decalcifying agents. After decalcification the tissue is soft and can be embedded and processed as in standard histology. It is also possible to use chelating agents, such as EDTA, which specifically bind calcium. These chelating agents are less damaging to the tissue than acids, but the decalcification process may be quite long (several weeks or more).
ii. Ground sections: It is possible to grind the bone until the sample is sufficiently thin for histological observation. The cells and organic tissue are destroyed in such preparations, though the canaliculi and cell lacunae are well seen. (Similar techniques are used by geologists to prepare thin sections of rock samples).
iii. Sections of non-decalcified bone: It is possible to embed bone tissue in a hard resin and section it with special knives (tungsten-carbide). Small samples for electron microscopy can be cut with diamond knives.
References
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