Lab 5: Direct Stain and Indirect Stain - Biology

Lab 5: Direct Stain and Indirect Stain - Biology

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In our laboratory, bacterial morphology (form and structure) may be examined in two ways:

  • by observing living unstained organisms (wet mount), or
  • by observing killed stained organisms.

Since bacteria are almost colorless and therefore show little contrast with the broth in which they are suspended, they are difficult to observe when unstained. Staining microorganisms enables one to:

  • see greater contrast between the organism and the background,
  • differentiate various morphological types (by shape, arrangement, gram reaction, etc.),
  • observe certain structures (flagella, capsules, endospores, etc.).

Before staining bacteria, you must first understand how to "fix" the organisms to the glass slide. If the preparation is not fixed, the organisms will be washed off the slide during staining. A simple method is that of air drying and heat fixing. The organisms are heat fixed by passing an air-dried smear of the organisms through the flame of a gas burner or holding it in front of the opening of a microincinerator. The heat coagulates the organisms' proteins causing the bacteria to stick to the slide.

The procedure for heat fixation is as follows:

1. If the culture is taken from an agar medium:

a. Using the dropper bottle of deionized water found in your staining rack, place 1/2 of a normal sized drop of water on a clean slide by touching the dropper to the slide (see Fig. 2). Altenately, use your sterilized inoculating loop to place a drop of deionized water on the slide.

b. Using your sterile inoculating loop, aseptically remove a small amount of the culture from the agar surface and gently touch it 2 - 3 times to the drop of water until the water becomes visibly cloudy (see Fig. 3).

c. Incinerate the remaining bacteria on the inoculating loop. If too much culture is added to the water, you will not see stained individual bacteria.

d. After the inoculating loop cools, spread the suspension over approximately half of the slide to form a thin film (see Fig. 4).

e. Allow this thin suspension to completely air dry (see Fig. 5). The smear must be completely dry before the slide is heat fixed!

f. To heat-fix the bacteria to the slide, pick up the air-dried slide with coverslip forceps and hold the bottom of the slide opposite the smear near the opening of the microincinerator for 10 seconds (see Fig. 6) as demonstrated by your instructor. If the slide is not heated enough, all of the bacteria will wash off. If it is overheated, the bacteria structural integrity can be damaged.

2. If the organism is taken from a broth culture:

a. Using a sharpie, draw a circle about the size if a nickel on the bottom of your microscope slide (see Fig. 10)

b. Turn the slide over. Using your sterile inoculating loop, aseptically place 2 or 3 loops of the culture within this circle on the top of the slide (see Fig. 11). Do not use water.

c. Using the inoculating loop, spread the suspension over the area delineated by the circle to form a thin film.

d. Allow this thin suspension to completely air dry.

e.To heat-fix the bacteria to the slide, pick up the air-dried slide with coverslip forceps and hold the bottom of the slide opposite the smear near the opening of the microincinerator for 10 seconds (see Fig. If it is overheated, the bacteria structural integrity can be damaged.

In order to understand how staining works, it will be helpful to know a little about the physical and chemical nature of stains. Stains are generally salts in which one of the ions is colored. (A salt is a compound composed of a positively charged ion and a negatively charged ion.) For example, the dye methylene blue is actually the salt methylene blue chloride which will dissociate in water into a positively charged methylene blue ion which is blue in color and a negatively charged chloride ion which is colorless.

Dyes or stains may be divided into two groups: basic and acidic. If the color portion of the dye resides in the positive ion, as in the above case, it is called a basic dye (examples: methylene blue, crystal violet, safranin). If the color portion is in the negatively charged ion, it is called an acidic dye (examples: nigrosin, congo red).

Because of its chemical nature, the cytoplasm of all bacterial cells has a slight negative charge when growing in a medium of near neutral pH. Therefore, when using a basic dye, the positively charged color portion of the stain combines with the negatively charged bacterial cytoplasm (opposite charges attract) and the organism becomes directly stained (see Fig. 1). An acidic dye, due to its chemical nature, reacts differently. Since the color portion of the dye is on the negative ion, it will not readily combine with the negatively charged bacterial cytoplasm (like charges repel). Instead, it forms a deposit around the organism, leaving the organism itself colorless (see Fig. Since the organism is seen indirectly, this type of staining is called indirect or negative, and is used to get a more accurate view of bacterial size, shapes, and arrangements.

In today's lab, we will make both direct and indirect stains of several microorganisms.


In direct staining the positively charged color portion of the basic dye combines with the negatively charged bacterium and the organism becomes directly stained.


Your pure cultures of Staphylococcus epidermidis (coccus with staphylococcus arrangement) or Micrococcus luteus (coccus with a tetrad or a sarcina arrangement) and Escherichia coli (small bacillus) or Enterobacter aerogenes (small bacillus) from Lab 3.

PROCEDURE (to be done individually)

1. Escherichia coli or Enterobacter aerogenes

a. Heat-fix a smear of either Escherichia coli or Enterobacter aerogenes as follows:

1. Altenately, use your sterilized inoculating loop to place a drop of deionized water on the slide.

2. 3).

3. If too much culture is added to the water, you will not see stained individual bacteria.

4. 4).

5. The smear must be completely dry before the slide is heat fixed!

6. If it is overheated, the bacteria structural integrity can be damaged.

b. Place the slide on a staining tray and cover the entire film with safranin (see Fig. 7A). Stain for one minute.

c. Pick up the slide by one end and hold it at an angle over the staining tray. Using the wash bottle on the bench top, gently wash off the excess safranin from the slide (see Fig. 7B). Also wash off any stain that got on the bottom of the slide as well.

d. Use a book of blotting paper to blot the slide dry (see Fig. 8). Observe using oil immersion microscopy.

2. Micrococcus luteus or Staphylococcus epidermidis

a. Heat-fix a smear of either Micrococcus luteus or Staphylococcus epidermidisas follows:

1. Stain with crystal violet for one minute.

c. Wash off the excess crystal violet with water.

d. Blot dry and observe using oil immersion microscopy.

3. Prepare a third slide of the normal flora and cells of your mouth as follows:

a. Use a sterile cotton swab to vigorously scrape the inside of your mouth and gums.

b. Rub the swab over the slide (do not use water), air dry, and heat-fix.

c. Stain with crystal violet for 30 seconds.

d. Wash off the excess crystal violet with water.

e. Blot dry and observe. Find epithelial cells using your 10X objective, center them in the field, and switch to oil immersion to observe your normal flora bacteria on and around your epithelial cells.

4. Make sure you carefully pour the used dye in your staining tray into the waste dye collection container, not down the sink.

C. INDIRECT STAIN USING AN ACIDIC DYE In negative staining, the negatively charged color portion of the acidic dye is repelled by the negatively charged bacterial cell. Therefore the background will be stained and the cell will remain colorless.


Use the pure culture of Micrococcus luteus provided.

PROCEDURE (to be done individually)

1. Place a small drop of nigrosin towards one end of a clean microscope slide.

2. Using your sterile inoculating loop, aseptically add a small amount of Micrococcus luteus to the dye and mix gently with the loop.

3. Using the edge of another slide, spread the mixture with varying pressure across the slide so that there are alternating light and dark areas (see Fig. 9). Make sure the dye is not too thick or you will not see the bacteria!

4. Let the slide air dry completely on the slide. Do not heat fix and do not wash off the dye.

5. Make sure you carefully pour the used dye in your staining tray into the waste dye collection container, not down the sink.

6. Observe using oil immersion microscopy. Find an area that has neither too much nor too little dye (an area that appears light purple where the light comes through the slide). If the dye is too thick, not enough light will pass through; if the dye is too thin, the background will be too light for sufficient contrast.


Make drawings of your three direct stain preparations and your indirect stain preparation.

Tech Tip: Combined Direct and Indirect Immunofluorescence Using Primary Antibodies from the Same Host

Immunofluorescence can be performed in two ways, by direct or indirect detection. Direct immunofluorescence uses a fluorophore-conjugated antibody to stain the target protein. Indirect immunofluorescence involves first binding the primary antibody to the target, then detecting the primary antibody using a conjugated secondary antibody. Indirect immunofluorescence offers the advantage of higher sensitivity. Each conjugated antibody molecule usually bears 4-6 fluorescent dyes. With direct immunofluorescence using monoclonal antibodies, only one antibody may bind to its target antigen on the protein, tagging it with 4-6 dye molecules. With indirect immunofluorescence, multiple secondary antibodies bind to each primary antibody, greatly increasing the number of dyes associated with the target protein. However, direct immunofluorescence offers the advantage of being able to stain a sample with multiple primary antibodies from the same host species simultaneously.

Secondary antibodies will still recognize a dye conjugated primary antibody, therefore a directly labeled antibody cannot be combined with an unconjugated primary from the same species in a single staining step, because the labeled secondary antibody will detect both the labeled and unlabeled primary antibodies. However, staining can be performed sequentially to allow indirect detection of the first primary antibody using a secondary, followed by staining with a directly labeled primary antibody from the same species, without cross-reactivity with the secondary. The following protocol provides an example of sequential staining with two unconjugated primary antibodies from rabbit and mouse hosts, followed by one directly labeled mouse primary. It can be adapted for other combinations of primary antibody host species.

Protocol for combined direct and indirect immunofluorescence

1. Perform fixation, permeabilization, and blocking of your samples according to the preferred protocol for your primary antibodies.

Note: See our Protocols for Antibody-Based Detection for more information on fixation, permeabilization, and blocking.

2. Incubate samples with unlabeled mouse and rabbit primary antibodies diluted in blocking buffer at the manufacturer’s recommended concentration for 2 hours at room temperature or 4°C overnight.

3. Rinse samples three times with PBS, and then perform 3 x 5 minute washes with PBS.

4. Incubate samples with fluorescent anti-mouse and anti-rabbit secondary antibodies diluted in blocking buffer for 30 minutes to 2 hours at room temperature, protected from light. Typically, fluorescent secondary conjugates are used at a final concentration of 1-2 ug/mL.

Note: Be sure to use highly cross-adsorbed secondary antibodies with minimal cross-reactivity for all antibody host species. If staining human or animal tissue sections that may contain endogenous immunoglobulins, make sure the secondary antibodies also have minimal cross-reactivity to the tissue itself.

5. Rinse samples three times with PBS, then perform 3 x 5 minute washes with PBS, protected from light.

6. Incubate samples with directly labeled mouse primary antibody diluted in blocking buffer for 2 hours at room temperature. Nuclear counterstains or labeled phalloidin may be included at this step.

Note: You may need to use a higher concentration of directly labeled primary antibody than you would use for indirect immunofluorescence.

7. Rinse samples three times with PBS, then perform 3 x 5 minute washes with PBS, protected from light.

8. Mount samples in antifade mounting medium and image.

Example data

Combined direct and indirect immunofluorescence staining of rat retina cryosection with mouse anti-neurofilament H and CF™568 goat anti-mouse (min x rat) (neuronal processes, red), rabbit anti-GFAP and CF™488A goat anti-rabbit, highly cross-adsorbed (glial cells, green), followed by CF™640R Mix-n-Stain™ labeled mouse anti-ZO1 (tight junctions, magenta). Mounted in Everbrite™ Mounting Medium with DAPI (nuclei, blue). Combined direct and indirect immunofluorescence staining of rat testis with mouse anti-tubulin and CF™488A goat anti-mouse (min x rat) (microtubules, green), followed by CF™555 Mix-n-Stain labeled mouse anti-ZO1 (tight junctions, red) and CF™640R phalloidin (actin filaments, cyan).

Related Products

Biotium offers a wide selection of highly cross-adsorbed secondary antibodies conjugated to our bright and photostable CF™ dyes. We also offer more than 1000 primary antibodies, available in purified format or with your choice of 13 CF™ dye colors. In addition, our Mix-n-Stain™ antibody kits provide a fast and simple way for you to conjugate your own primary antibody with a wide selection of CF™ dyes. Also, be sure to see our accessory products for immunofluorescence, including TrueBlack™ Lipofuscin Autofluorescence Quencher, EverBrite™ antifade mounting media, and CoverGrip™ Coverslip Sealant.

Direct vs indirect immunofluorescence

​​Immunofluorescence (IF) or cell imaging techniques rely on the use of antibodies to label a specific target antigen with a fluorescent dye (also called fluorophores or fluorochromes ) such as fluorescein isothiocyanate ( FITC ). Antibodies that are chemically conjugated to fluorophores are commonly used in IF.

​ The fluorophore allows visualization of the target distribution in the sample under a fluorescent microscope ( eg epifluorescence and confocal microscopes). We distinguish between two IF methods depending on whether the fluorophore is conjugated to the primary or the secondary antibody:

  • Direct IF uses a single antibody directed against the target of interest. The primary antibody is directly conjugated to a fluorophore.
  • Indirect IF uses two antibodies. The primary antibody is unconjugated and a fluorophore-conjugated secondary antibody directed against the primary antibody is used for detection.

The diagram below represents both direct and indirect methods.

​​ Both methods have their advantages and disadvantages as shown in the table below.

Protocols for direct IF are usually shorter as they only require one labeling step.

Direct and indirect methods are not limited to immunofluorescence. They are also relevant to other techniques that rely on the use of fluorophore-conjugated antibodies such as flow cytometry, ELISA, western blot and immunohistochemistry.

Detection of low abundance proteins can be sometimes challenging even with indirect methods. Biotinylated antibodies offer an extra layer for increased signal amplification. Learn more about how methods based on the use of biotin-conjugated antibodies work here.

Negative Staining Procedure

  1. Take a clean, grease-free and dry glass slide.
  2. Put a minimal drop of nigrosin towards one end of the glass slide via a dropper.
  3. Then, take the inoculum from the culture plates or slant culture via a sterilized inoculating loop and mix it with a drop of nigrosin.
  4. After that, take another clean, grease-free and dry glass slide and place the one end of it towards the centre.
  5. Then, tilt the glass slide over the stain containing the test organism by making an acute angle.
  6. Slightly draw the tilted slide until it touches the drop of the culture organism, and drag it across the edge of the glass slide to make an even, broad and thin bacterial smear.
  7. Allow the glass slide to air dry (do not heat fix).
  8. At last, put a drop of oil immersion and observe the glass slide under the microscope for the appearance of a colourless bacterial cell with a grey background.


  • Through negative staining, clear unstained cells are easily observable against the black coloured stained background.
  • A negative staining method does not involve the heat-fixing of the specimen. As a result, the cell will not deform by the heat.
  • It can also stain heat-sensitive microorganisms like Spirochetes, Yeasts etc.
  • The negative staining technique also permits examining a transparent capsule around the cell wall of various microorganisms like Cryptococcus neoformans.
  • It is quite an easy and rapid method that makes the use of a single acidic stain only.


  • Negative staining does not provide much information about the cell rather than the cell size, shape and arrangement.
  • By using this technique, we cannot examine a particular strain or a type of organism.


Therefore, we can conclude that a negative staining technique is a simple method to examine the microorganism using a single acidic stain. Negative staining results in an unstained or clear specimen with a dark coloured background.

Repulsion occurs between the negatively charged stain and the specimen, after which we could observe cells of different shapes and sizes as unstained outlines against a stained background.

Lab 5: Flatworm and Smaller Lophotrochozoans

Phylum Rotifera: rotifers
Phylum Acanthocephala: spiny-headed worms
Phylum Ectoprocta (Bryozoa) &ndash &ldquomoss animals&rdquo (Bugula, Plumatella)
Phylum Brachiopoda &ndash &ldquolampshells&rdquo (Lingula, Terebratella)

Phylum From this point on, all animals covered in the Zoo Lab website have primary bilateral symmetry and are triploblastic, that is, three true germ layers (the ectoderm, mesoderm and endoderm) are formed during gastrulation of the blastula stage of development. While radial symmetry may be well suited for sessile or slow-moving forms, animals that are active in seeking food, shelter and mates require a new body plan. Bilateral symmetry coupled with cephalization solves these problems. The anterior end moves forward and the posterior follows. The dorsal side is kept facing up and the ventral side is kept down and usually specialized for locomotion.

The bilateral grade of metazoans is further subdivided into two main divisions: the protostomes and deuterostomes, which are separated on the basis of a number of embryological differences. Evidence from sequence analysis of the small-subunit ribosomal gene suggests that some time after ancestral deuterostomes and protostomes diverged from one another during the Cambrian period, protostomes split into two large groups (superphyla), the Ecdysozoa and Lophotrochozoa. In this lab we will examine one acoelomate lophotrochozoan phylum and several smaller pseudocoelomate and eucoelomate lophotrochozoan phyla.

The Phylum Platyhelminthes contains over 20,000 free-living and parasitic species of acoelomate animals called flatworms. In flatworms, the body that is flattened dorsoventrally, with the mouth and genital pore usually located in a ventral position. The space between the gut and outside is filled with mesodermal muscle fibers and undifferentiated parenchyma. Although fluid-filled spaces in the parenchyma serve as a hydrostatic skeleton for support and to aid in internal transport, the animals lack a body cavity, which is why they are called acoelomate. Most free-living flatworms have a gastrovascular-type digestive system (a mouth is present but no anus), while parasitic forms generally have no digestive system.

Flatworms have a centralized nervous system consisting of pair of cerebral ganglia and longitudinal nerve cords connected to transverse nerves. The excretory system (absent in some forms) consists of two lateral canals with protonephridia bearing flame cells. Although many flatworms are free-living, the phylum includes some very important parasitic species as well.

In terms of reproduction, flatworms can reproduce sexually or asexually. Most species are monoecious but practice cross fertilization. Many freshwater turbellarians can reproduce asexually by fission in which the animal simply divides into two halves, each of which regenerates the other half. In some turbellarians (as it is in most other animals), the yolk that provides nutrition for the developing embryo is containing within the egg cell itself, a condition described as endolecithal. In the monogeneans, trematodes and cestodes (as well as in a few turbellarians), yolk is contributed by cells released from organs called yolk glands, and the eggs are therefore described as ectolecithal. Development may be direct or indirect.

The Class Turbellaria contains mostly free-living forms ranging in size from a few mm to 50 cm. Most species are bottom dwellers in marine and freshwater environments that crawl over rocks, sand or vegetation. Smaller forms can swim by means of ventral cilia, but more often they move by laying down a sheet of mucus that aids in adhesion and helps the cilia gain traction. Larger forms use powerful muscle contractions to crawl or swim. Unique to turbellarians are rod-shaped rhabdites found among the ventral epidermal cells of the body surface. These rhabdites secrete mucus that coats the animal's body, possibly for protection against predators or to prevent drying.

In terms of nutrition, most turbellarians are predators and scavengers. Epidermal mucous secretions trap and kill prey items. A muscular pharynx everted though the ventral mouth is used to secrete digestive enzymes into prey, which is then sucked into the branched intestine that forms a gastrovascular cavity. In addition to a simple nervous system, turbellarians have light-sensitive eye spots called ocelli that help orient the animal to the direction of light. Touch and chemical receptors in some forms like the planarian seen in lab are concentrated in lateral projections from the head called auricles that look like ear lobes. Reproduction in turbellarians can occur asexually through fission or sexually all forms are monoecious but practice cross-fertilization. Planarians are also known for their tremendous powers of regeneration, and a planarian that has been cut into three pieces will give rise to three new complete individuals!

The Class Monogenea contains animals called monogenetic flukes. Although most of species are ectoparasites on the skin or gills of fish, there a few forms found in the bladders of frogs and even one that lives in the eye of a hippopotamus! The life cycle of a monogenetic fluke is direct, with a single host. Since they must depend on a single host for both reproduction and transmission, monogenetic flukes have evolved mechanisms that usually ensure that the parasites do not endanger the lives of their hosts, but in crowded conditions (such as fish hatcheries), they can produce serious, damaging infestations.

The Class Trematoda contains about 8,000 species of leaf-like animals called digenetic flukes. The adults are endoparasites on vertebrates but many invertebrates serve as intermediate hosts, and many species of medical and economic importance! Development is trematodes indirect not only adults but larvae reproduce and all species have at least two hosts, one for transmission and the other for reproduction. The vast majority of flukes possess two large suckers that are used for attachment, an anterior one called an oral sucker, which surround the mouth and a posterior one called a ventral sucker, or acetabulum.

In trematodes, one egg leads to the production of many progeny! Eggs are typically deposited in water via the urine or feces of the definitive host. When they reach freshwater, the egg opens and a ciliated free-swimming larva called a miracidium swims out. The miracidium will then swim about until it finds a suitable intermediate host, which is usually an aquatic snail to which it is chemically attracted. When the miracidium finds snail, it penetrates it, loses its cilia and develops into a sporocyst, which produces asexually either more sporocysts or a number of rediae that also produce asexually either more rediae or tailed forms called cercariae. The cercariae emerge from the snail, swim around and penetrate a second intermediate host, the final (definitive) host or encyst on vegetation (in the case of the sheep liver fluke), where they are transformed into metacercariae, which are juvenile flukes the adult grows from the metacercariae when it is eaten by the definitive host.

Trematode Infection in Humans

Humans can be infected with a number of serious trematodes in a variety of ways. In the case of the Oriental liver fluke (Clonorchis sinensis), infection occurs by eating raw or poorly cooked fish (which serve as the second intermediate host of the parasite) containing the metacercariae of the trematode.

In the case of blood flukes (Schistosoma), infection can occur when tailed cercariae burrow through exposed skin of people bathing or working in waters containing the cercariae (such as Asian rice paddies). Schistosomiasis is a chronic illness that can damage internal organs and, in children, impair growth and cognitive development. The urinary form of schistosomiasis is associated with increased risks for bladder cancer in adults, and the disease is the second most socioeconomically devastating parasitic disease after malaria!

The sheep liver fluke (Fasciola hepatica) is a common parasite of sheep and cattle, which become infected by eating aquatic plants containing encysted metacercariae (juvenile flukes). Humans can acquire the parasite by eating raw watercress (which grows naturally at the edges of lakes and ponds and is cultivated in many countries in Asia and Europe) containing the metacercariae of the fluke.

The lung fluke (Paragonimus westermani) is a potential dangerous parasite found in Asia and South America that can cause death in human hosts. Their eggs are coughed up from the lungs of their host, swallowed and eliminated in feces Humans can become infected by eating raw or poorly cooked freshwater crabs (the second intermediate host of the parasite) containing the metacercariae of the fluke.

The Class Cestoda contains about 4,000 species of tapeworms, all of which are highly modified endoparasites that live in just about every known vertebrate species. The long, flattened body of a tapeworm (which is referred to as the strobila) is divided into segments called proglottids. Most forms have an organ called a scolex at the anterior end with suckers, hooks, etc. that attach to the wall of the gut and prevent them from being swept away.

Tapeworms lack a digestive system and feed by absorbing nutrients directly from the host. The entire body surface is covered with minute projections called microtriches that greatly increase the absorptive surface area of the tapeworm. Tapeworms also secrete substances that inhibit the digestive enzymes of their host as well as lowering the pH around them to a level that they but not the digestive enzymes of their host can function. In tapeworms, much of the strobila is given over to reproduction. Each proglottid is monoecious, and cross-fertilization or even self-fertilization is common. Proglottids can be filled with up to 100,000 eggs!

With few exceptions, all cestodes require at least two hosts, and the adult is the parasite in the digestive tract of vertebrates. Often one of the intermediate hosts is an invertebrate (most often an arthropod such as a flea, louse or copepod) that is eaten by the final host. The eggs within the proglottids are shed daily in the feces into the soil where they may lie dormant for quite some time. Sometimes the egg-bearing proglottids crawl out of the anus by themselves and can be found wriggling about on an infected dog, cat or child or on infected clothing and bedding. Once the eggs are released, they must be ingested by an intermediate host in order to hatch into hooked larvae called oncospheres, which bore through the intestinal wall and picked up by the circulatory system where they are transported to skeletal muscle, heart or even some other organ where they encyst as cysticerci (bladder worms). Each cysticercus is essentially an inside-out scolex that everts after the infected tissue (so-called &ldquomeasly meat&rdquo) of the intermediate host is eaten by the final host. The scolex then attaches to the lining of the intestine by means of suckers and/or hooks.

Tapeworm Infection in Humans

Humans can become infected with tapeworms by eating poorly cooked meat containing the cysticerci of the tapeworm. The most important tapeworms that infect humans are the beef tapeworm (Taenia saginata) and the pork tapeworm (Taenia solium).

Another species of cestode that can infect humans is the broad fish tapeworm (Diphyllobothrium latum), which is common in fish inhabiting the Great Lakes. Again, infection occurs by ingesting cysticerci in raw or poorly cooked fish. In most cases, tapeworms found in the gut do not cause much damage to their human hosts, but occasionally they migrate to other organs such as the eyes or even the brain, where they can cause serious neurological problems and even death from cerebral cysticercosis!

The dog tapeworm (Diplydium caninum) is common in dogs but can be picked up by humans (usually kids) who ingest infected fleas that serve as intermediate hosts of the parasite.

In contrast to radiate and acoelomate phyla in which the space between the body wall and the digestive tract is filled with mesoglea or with solid mesenchymal parenchyma, the remaining bilateral animals covered in the Zoo Lab website have a body cavity in which internal organs are located. In the pseudocoelomates, the embryonic blastocoel persists as a body cavity. Since it is not lined with mesodermal peritoneum (the lining of the coelom), it is called a &ldquofalse cavity&rdquo or pseudocoel.

The Phylum Acanthocephala contains about 1,000 species of parasitic animals called spiny-headed worms, all of which are endoparasites in the intestinal tracts of vertebrates (especially fishes). Two hosts are required to complete the life cycle, and the juveniles are parasites of crustaceans and insects. Most species are quite small (less than 40 mm). Spiny-headed worms have an eversible proboscis covered with recurved spines that provides a means of attachment in the host's intestine. Eggs pass out host and are eaten by certain insects or crustaceans where they hatch and go through several developmental stages. When the intermediate host is eaten by a bird, mammal or fish, the larva inside attaches to the intestinal wall with its spiny proboscis.

The Phylum Rotifera contains about 1,800 species of microscopic animals called rotifers that bear an anterior crown of cilia that give the appearance of a revolving wheel. Although cosmopolitan (widely distributed), most are found only in freshwater environments. The general body plan of a rotifer is divided into three regions: a head, which bears a ciliated organ called a corona (wheel organ), which creates currents that draw small planktonic forms into the mouth, which opens into a muscular pharynx called a mastax. The mastax is equipped with intricate jaws composed of seven hard pieces called trophi that are used for grasping and chewing the prey. The trunk contains the visceral organs, and the foot (when present) is segmented and ringed into joints that can shorten or telescope. Pedal glands on the foot secrete a sticky substance that anchors the animal to the substrate or allows it to creep along with leech like movements.

From this point on, all animals that will be studied in the Zoo Lab website are eucoelomate, that is, they have a true coelom (body cavity) that is lined with a thin layer of mesodermal tissue called the peritoneum. Note: The development of the coelom must be considered one of the most important steps in the evolution of larger and more complex forms for it provides plenty of space for organs that can be held in place by thin membranes called mesenteries!

The Phylum Ectoprocta (also called Bryozoa) contains about 4,000 species of small colonial forms called moss animals that are found in shallow freshwater and marine environments. Although bryozoans are also well represented in the fossil record, they are also quite abundant today. Modern marine forms exploit all kinds of firm substrates including shells, rocks, marine timbers, and ship bottoms. In fact, like barnacles, the ectoprocts are one of the most important groups of fouling organisms that need to be removed periodically from ship and boat hulls. Each member of a colony lives in a tiny chamber called a zoecium (&ldquoanimal house&rdquo), which is secreted by its epidermis.

Each individual (zooid) consists of a feeding part and a case-forming part. The zoecium can be gelatinous, chitinous or calcareous, and sometimes it is impregnated with sand grains. The feeding portion of the animal contains the lophophore (a ciliated feeding device that can also be used for gas exchange), digestive tract, muscles and nervous system. Each individual lives a kind of &ldquojack-in-the-box&rdquo existence, popping up to feed and then quickly withdrawing into its protective chamber that is often sealed with a tiny trapdoor (operculum).

The Phylum Brachiopoda (&ldquoarm foots&rdquo) contains animals that are known as lampshells. This is an ancient group that is well represented in the fossil record (with some 30,000 described species) but only about 300 living species. Brachiopods resemble bivalve molluscs, but unlike bivalves, they have shells that are located on the ventral and dorsal side rather than left and right.

Brachiopods are divided into two classes based on whether they have shells that are connected by a hinge with interlocking &ldquoteeth&rdquo or with shells of unequal size. Brachiopods in the latter group are called lampshells because the larger ventral valve resembles a Roman oil lamp. Some brachiopods attach themselves to the substrate by a pedicel on the ventral valve while others just cement the ventral valve to the substrate (like an oyster) or burrow into the sediment. Like bryozoans, brachiopods also have a lophophore surrounding the mouth that is used for feeding and gas exchange.

Lab-5 01

This slide contains two specimens of the free-living turbellarian flatworm Planaria. One specimen has been stained, while the other has been injected with carbon black to reveal the extent of the blind gastrovascular cavity, which is divided into three, many-branched trunks (one anterior and two posterior). Without an anus, food must first pass through the mouth into the gastrovascular cavity where it is digested after which waste products exit through the same opening. Note the large, eversible pharynx in each planarian that is used for feeding. In the head region are lateral projections called auricles (not well developed on the specimens shown) that contain touch and chemical receptors as well as light-sensitive ocelli (eye spots).

Lab-5 02

  1. Buccal cavity
  2. Gastrodermis
  3. Gastrovascular cavity
  4. Epidermis
  5. Pharynx
  6. Parenchyma

This slide contains a cross section through the pharyngeal (middle) region of the free-living flatworm Planaria. Note the large muscular pharynx that lies within a space called the buccal cavity. During feeding, the pharynx can be everted through the mouth and used to suck up fluids and soft tissue from captured prey. Two branches of the extensive gastrovascular cavity can also be seen. This cavity is lined with large, vacuolated cells that comprise the gastrodermis. On the outside of the flatworm is a ciliated epidermis that contains many gland cells as well as dark-staining rod-shaped bodies called rhabdites that can discharge their contents to form a protective mucous layer around the body. Lacking a body cavity, the space between the gut and epidermis in these acoelomates is filled with a meshwork of mesodermal parenchyma as well as muscle fibers that run circularly, longitudinally and diagonally.

Lab-5 03

  1. Gastrovascular cavity
  2. Gastrodermis
  3. Parenchyma
  4. Rhabdites on epidermis

Lab-5 04

This slide shows a stained whole mount of the Oriental liver fluke (Clonorchis sinensis), an important trematode parasite of the humans in many regions of Asia, especially China, Southeast Asia and Japan. Humans are infected by eating raw or poorly cooked fish containing the encysted metacercariae. After being ingested, these cysts dissolve in the intestine, releasing the young flukes which then migrate to the bile duct and liver.

Anterior section

Lab-5 05

  1. Mouth and oral sucker
  2. Pharynx
  3. Esophagus
  4. Intestinal cecum
  5. Genital pore
  6. Ventral sucker (acetabulum)

Middle section

Lab-5 06

  1. Uterus
  2. Intestinal ceca
  3. Yolk glands
  4. Yolk duct
  5. Ovary
  6. Seminal receptacle
  7. Testes
  8. Excretory bladder

Posterior section

Lab-5 07

Lab-5 08

This slide shows the redia larva of a trematode parasite. This larval stage normally develops in the tissues of an aquatic snail. The redia contains groups of cells called "germ balls" that eventually develop into the tailed cercaria larvae, which emerge from the snail and penetrate a second intermediate host or encyst on vegetation to become a metacercaria.

Lab-5 09

This slide shows the tailed-cercaria larva of a trematode parasite. This larval stage, which normally develops in the tissue of an aquatic snail, will emerge from its intermediate host and penetrate a second intermediate host or encyst on vegetation to become a metacercaria.

Lab-5 10

This slide contains stained sections of the dog tapeworm Diplydium caninum taken from four different regions. The anterior most portion contains the scolex, a specialized attachment organ that often contains hooks and/or suckers. The rest of the body is divided into a linear series of segments called proglottids, each of which contains a complete set of reproductive organs. The youngest proglottids in the first part of the strobila (body) of the tapeworm are immature, while those in the middle are mature. The oldest terminal proglottids are gravid, which means they are filled with eggs. Dogs and cats can become infected by eating adult fleas (the intermediate hosts) containing cysticercoid larvae.

Scolex (close-up)

Lab-5 11

Lab-5 12

  1. Testes
  2. Vas deferens
  3. Vagina
  4. Ovary
  5. Yolk gland
  6. Genital pore

This slide shows a mature proglottid from the dog tapeworm Diplydium caninum. Note that there are two complete sets of male and female reproductive structures that include testes, vasa deferentia (the plural of vas deferens), ovaries, yolk glands, vaginas and genital pores. Dogs and cats become infected by eating adult fleas (the intermediate hosts) containing cysticercoid larvae.

Lab-5 13

  1. Excretory canal
  2. Testes
  3. Uterus
  4. Genital pore
  5. Vas deferens
  6. Vagina
  7. Ovaries
  8. Yolk glands

This slide shows a mature proglottid from the tapeworm Taenia pisiformis, a species commonly found in the small intestines of dogs and cats. Note that each segment contains a complete set of reproductive structures including testes, vas deferens (sperm duct), ovary, yolk gland, vagina and genital pore.

Lab-5 14

This slide shows a scolex from the anterior most region of the tapeworm Taenia pisiformis. Note the series of hooks on a raised portion of the scolex called a rostellum as well as the four lateral suckers. These hooks and suckers enable to tapeworm to remain attached to the intestinal wall of its host.

Hooks on rostellum

Lab-5 15

This slide shows a magnified view of the raised tip of the scolex (rostellum) from the dog and cat tapeworm Taenia pisiformis. Note the formidable array of hooks that are used by the tapeworm to hang on to the intestinal tract of its host.

Lab-5 16

This slide shows the cysticercus larva of the tapeworm Taenia pisiformis. Note the invaginated scolex on the right end of this "bladder worm". After infected tissue of the intermediate host is eaten by the definitive host, the scolex everts and attaches to the lining of the intestine by means of hooks and suckers.

Lab-5 17

This slide shows a stained specimen of an adult spiny-headed worm belonging to the Phylum Acanthocephala. Although human infections have been recorded, adult worms normally parasitize the digestive tracts of fish, birds and domestic and wild mammals. The larvae of spiny-headed worms develop in various species of crustaceans or insects. Note the everted proboscis containing numerous recurved spines that give the organism its name. These spines (which permit the worms to remain attached to the digestive tract) can cause massive and sometimes painful destruction of the intestinal mucosa.

Acanthocephalan proboscis (close-up)

Lab-5 18

Lab-5 19

This slides shows two stained rotifers. These pseudocoelomate animals derive their name from a distinctive ciliated crown (corona) that, when beating, gives the impression of a rotating wheel. The movement of these cilia creates water currents that draw food items into the mouth of the organism. Once inside, food is chewed and ground up in a muscular portion of the pharynx called a mastax that is equipped with small hard jaws called trophi. Although there are a few marine species, most rotifers are found in freshwater habitats throughout the world.

Photographs of living rotifers

Lab-5 20

This microscope image shows two live specimens of the common rotifer Philodina. Note the lateral extension of the body wall in the head region of the specimen on the right (pointed to by the red arrow). This structure (which is called an antenna) contains many, tiny sensory bristles. The corona ("wheel organ") containing two large ciliated trochal discs and foot with its two toes (the spurs pointed to by the blue arrow) can be seen on the specimen on the left. Pedal glands (which open by ducts at the tips of the toes) produce an adhesive substance used for temporary attachment to the substrate.

Lab-5 21

This microscope image shows a magnified view of the freshwater rotifer Philodina. Note the conspicuous corona (wheel organ) with its cilia and the centrally-located mastax (pointed to by the red arrow), a muscular portion of the pharynx equipped with chitinous jaws (trophi) that grind and shred ingested food.

Lab-5 22

This microscope image shows another species of rotifer in the genus Monostyla. This common freshwater species has a rigid, chitin-like covering called a lorica.

Lab-5 23

This slide shows several zooids of the freshwater ectoproct Plumatella. Note the conspicuous lophophores. These feeding devices consist of masses of ciliated tentacles borne on ridges surrounding the mouth. In addition to reproducing by budding, freshwater bryozoans reproduce asexually by means of special resistant bodies called statoblasts (not visible on this slide). These dark, disc-shaped structures (which are similar to the gemmules of freshwater sponges) are produced during the summer and fall, and can remain dormant until environmental conditions improve in the spring.

Lab-5 24

This slide shows a portion of a branching colony of the marine bryozoan (ectoproct) Bugula. Branching within the colony is produced by repeated asexual budding of individuals called zooids. Note the tentacles of the lophophores (ciliated feeding devices surrounding the mouth that can also be used for gas exchange). Like many colonial cnidarians, ectoproct colonies are polymorphic, with most of the zooids functioning as feeding individuals. Defensive zooids called avicularia protect the colony against small organisms, including settling larvae and crawling tube-building polychaete worms and arthropods. Each avicularium resembles the head of a bird complete with powerful musculature and a sharp beak-like structure (rostrum) that is used to seize the appendages of trespassing organisms.

Avicularium (close-up)

Lab-5 25

This slide shows a magnified view of an avicularium from the marine colonial bryozoan Bugula. Note the mandible, bird-like beak (which is called a rostrum) and musculature. Avicularia protect the colony from small organisms, including settling larvae and crawling tube-building polychaete worms and arthropods.

Lab-5 26

This is a slide of a monogenetic fluke taken from the gills of an Atlantic stingray. Unlike the digenetic trematodes, monogenetic species have a direct life cycle in which ciliated larvae called oncomiracidia develop on or within a single host. Although a few species are found in the urinary bladders of frogs and turtles, most such flukes cling to the gills and external surfaces of fish by means of a posterior attachment organ called an opisthaptor that is equipped with hooks.

Lab-5 27

This slide shows a stained whole mount of the sheep liver fluke (Fasciola hepatica). This large trematode is a common parasite of sheep and cattle, which become infected by eating aquatic plants containing the encysted metacercariae (juvenile flukes). Once ingested, the cyst walls are digested and the larvae burrow through the intestinal wall to the body cavity and eventually to the liver.

Lab-5 28

This slide shows an adult specimen of the lung fluke Paragonimus westermani. Found in east Asia, southwest Pacific and some parts of South America, the fluke parasitizes a number of wild carnivores, pigs, rodents and humans. Infection with lung flukes causes respiratory symptoms, with breathing difficulties and chronic cough, and fatalities are common! Humans get infected with lung flukes by eating raw or poorly-cooked freshwater crabs containing the fluke metacercariae.

Lab-5 29

This slide shows a pair of adult blood flukes in copulation. Blood flukes differ from most other flukes by being dioecious (i.e., having separate sexes). Males are larger and have a large, ventral groove called a gynecophoric canal posterior to the ventral sucker that holds the smaller (more darkly stained) female during copulation, which is continuous. Schistosoma mansoni is one of the three species of blood flukes responsible for the disease in humans called schistosomiasis. Humans get infected when the tailed cercaria larvae (which escape from freshwater snails that serve as their intermediate hosts) burrow into the exposed skin of individuals bathing, swimming or working in such habitats.

Lab-5 35

This model includes several views of a free-living turbellarian flatworm. The image on the left shows the nervous system (painted white), which consists of a pair of cerebral ganglia with two ventral nerve cords that are connected by a series of transverse nerves called commissures, giving it a ladder-like appearance. Other sensory structures include simple, light-sensitive eyes (ocelli) and chemical receptors that are concentrated in lateral projections of the head called auricles (because they look like ear lobes). Although reproduction in planarians can occur asexually through fission, all forms are monoecious with both male and female reproductive organs. Several features of the reproductive system (shown in blue and yellow) are also seen on the model on the left.

The model on the right shows the many branched gastrovascular cavity (shown in red) that exits through a single ventral opening at the end of a muscular, eversible pharynx (shown in off-white on both models as well as on the small upper planarian
model). Also seen on the model is a portion of the excretory/osmoregulatory system (shown in green) that is made up of protonephridia that collect and secrete some wastes as well as the excess water that enters freshwater forms by osmosis. Protonephridia consist of excretory tubules that are closed internally and open to the outside by a series of collecting ducts that lead to a posterior opening called a nephridiopore. The internal ends of each of these tubules terminate in so-called flame cells (one of which is shown on the small, lower model), which have tufts of cilia that flicker like the flame of a candle. The beating of these cilia pulls water through a mesh-like cup, producing a filtrate of water and small molecules.

Anterior sections

Lab-5 36

  1. Cerebral ganglion
  2. Ventral nerve cord
  3. Ocellus
  4. Auricles
  5. Testes
  6. Ovary
  7. Oviduct
  8. Circular muscle layer
  9. Longitudinal muscle layer

Posterior sections

Lab-5 37

  1. Gastrovascular cavity
  2. Pharynx
  3. Excretory system
  4. Ventral nerve cord
  5. Testes
  6. Seminal vesicle
  7. Oviduct
  8. Parenchyma

Flame cell close-up

Lab-5 38

Lab-5 30

This image shows a model of the Oriental liver fluke (Clonorchis sinensis), an important trematode parasite of the humans in many regions of Asia. Humans are infected by eating raw or under cooked fish containing the encysted metacercariae. After being ingested, these cysts dissolve in the intestine, releasing the young flukes which then migrate to the bile duct and liver. For close-up views of labeled structures found in different sections of the liver fluke, click on the links below.


Lab-5 32

1. Pharynx 2. Cerebral ganglia 3. Intestinal ceca 4. Seminal vesicle 5. Uterus


Lab-5 31

1. Mouth 2. Oral sucker 3. Esophagus 4. Intestinal ceca 5. Ventral sucker 6. Genital pore 7. Uterus 8. Yolk glands


Lab-5 34

1. Seminal vesicle 2. Uterus 3. Intestinal cecum 4. Seminal receptacle 5. Excretory bladder 6. Excretory pore 7. Yolk ducts 8. Mehlis' gland


Lab-5 33

1. Uterus 2. Yolk glands 3. Mehlis' gland 4. Ovary 5. Seminal receptacle 6. Testes 7. Excretory bladder 8. Intestinal cecum

Definition of Simple Staining

Simple staining is defined as one of the ordinaries yet the popular method used to elucidate the bacterial size, shape and arrangement to differentiate the various bacteria groups. It stains the bacterial cell uniformly and thus increases the visibility of an organism. A term simple staining sometimes interchangeable with the terms like direct, positive or monochrome staining. Now let us understand why simple staining is called by such alternative names.

  • Direct staining: Because it is a direct method that directly stains the bacterial cell with a colourless background.
  • Positive staining: Because it uses positively charged basic dyes that bind with the negatively charged bacterial cell.
  • Monochrome staining: Because it adds contrast to the specimen by using a single stain only.

Simple Stains

Simple stains can be defined as the basic dyes, which are the alcoholic or aqueous solutions (diluted up to 1-2%). These can easily release OH – and accepts H + ion, due to which the simple stains are positively charged. As the simple stains are positively charged, they usually termed as positive or cationic dyes.

It is commonly used to colour most of the bacteria. As the simple stain carry a positive charge, it firmly adheres to a negative bacterial cell and makes the organism coloured by leaving a background colourless. Examples of simple stain include safranin, methylene blue, crystal violet etc.

The basic stains have different exposure time to penetrate and stain the bacterial cell.

Basic stainsExposure time to stain the bacteria
Methylene blue1-2 minutes
Crystal violet20-60 seconds
Carbol fuschin15-30 seconds
Safranin30-60 seconds


Its principle is based on producing a marked contrast between the organism and its surroundings by using basic stain. A basic dye consists of a positive chromophore, which strongly attracts to the negative cell components and charged molecules like nucleic acids and proteins. Thus, a simple staining technique results in a coloured bacterial cell against a colourless background.

Procedure of Simple Staining

It involves the following three steps:

Smear Preparation

Bacterial smear appears as a thin film of bacterial culture. For the smear preparation, we need to perform the following steps:

  1. Take a clean, grease-free glass slide.
  2. Add a drop of distilled water at the centre of the glass slide.
  3. Then, add inoculum from the bacterial culture with sterilized inoculating loop on the glass slide.
  4. After that, mix the inoculum with a drop of distilled water to make a thin film by uniformly rotating the inoculating loop until the formation of a thin bacterial film.

Heat Fixing

After smear preparation, move the prepared slides over Bunsen burner’s flame for at least three times. Then, allow the slide to air dry. There are many reasons to perform heat fixing, and it can not be skipped because:

  • Heat fixing helps in the fixation of a specimen to the glass slide.
  • Heat fixing helps the stain to penetrate the smear.

Staining of Bacteria

It is the last and the most crucial step, in which one can identify the morphological characteristics of the bacteria through microscopic examination, once the cells get stained. This stage involves the following steps, which are as follows:

  1. Add stain to the heat fixed smear.
  2. Allow the stain to stand for at least 1 minute so that it can penetrate between the cells.
  3. Wash off the glass slide carefully.
  4. Blot dry the slide with absorbent paper (do not wipe the slide).
  5. Examine the glass slide under the microscope from low to high power to get a magnified view of the specimen. One can also add a drop of oil immersion over the glass slide’s stained area to observe it under 100X objective.



  • Simple staining is a very simple method to perform, which stains the organism by using a single reagent.
  • It is a rapid method which reduces the performance time by taking only 3-5 minutes.
  • Simple staining helps to examine or elucidate the bacterial shape, size and arrangement.
  • It also helps us to differentiate the bacterial cells from the non-living structures.
  • Simple staining can be useful in the preliminary study of the bacteria’s morphological characteristics.


  • It does not give much information about the cell apart from the bacteria’s morphological characteristics.
  • Through simple staining, we cannot classify a particular type of organism.


Therefore, we can conclude that a simple staining method is the easiest way to colour the microscopic object as it uses a single basic stain. The results of simple staining are based on the type of basic stain that has been used.

The colour of a stain will decide the colour of a specimen that has to be identified. For example, when the bacteria retain the safranin colour, they appear pink-red, and the same goes with the other stains.

There is an attraction between the positive stain and the negative bacterial cell in simple staining, which results in the observation of coloured bacteria with a bright background.

Observations on living bacteria

Sometimes assay results are compromised because a contaminating organism grows in the medium instead of the intended bacterial isolate. For a quick check to verify that cell morphology is consistent with the culture from which the inoculum was taken, a wet mount can be prepared and examined in dark field and/or phase contrast. If present, endospores are often evident in phase contrast, allowing one to avoid having to do a spore stain.

Very often, identification of an unknown organism requires knowledge of its motility, that is, its capability for translational movement. The results of motility agar incubations can be difficult to interpret, partically for aerobic bacteria that don't grow well deep into the agar. A good quick check for motility is to examine a very young culture using the hanging drop method. A young culture would be a broth culture inoculated the night before, or a broth culture that was diluted 10 fold or so in the morning, incubated, and examined in the afternoon. A hanging drop culture is prepared by placing a very small drop of medium on a coverslip, then inverting the coverslip over a depression slide so that the bottom of the drop does not make contact with the slide itself. Vaseline can be used if necessary, to make a sealed chamber.

Hanging drops can be examined using all objective lenses, although to be able to look throughout the depth of the drop the limit may be 100x. The curved depression slide will distort the effects of phase contrast, but dark field may work and will be sufficient to detect movement. All live bacteria move by Brownian (molecular) motion, at a vibration rate that is inversely proportional to the size of the cell. Rapid Brownian movement is a common characteristic of non-motile cocci such as Staphylococcus, Streptococcus, or Micrococcus. However some bacteria are flagellated, and exhibit translational movement as well. Truly motile organisms will zip across the microscope field. Look for definite directional motion, tumbling, and movement against currents.


    Obtain a uniform suspension of cells: Follow the typsinization/trypsin neutralization protocol for the specific cell type. Place the cell suspension in a suitably-sized conical centrifuge tube. For an accurate cell count to be obtained, a uniform suspension containing single cells is necessary. Pipette the cell suspension up and down in the tube 5-7 times using a pipette with a small bore (5 ml or 10 ml pipette). For cells thawed from cryopreservation (in 1ml cryopreservation medium), pipette up and down 7-10 times using a one ml pipette.

For an accurate determination, the total number of cells overlying one 1 mm 2 should be between 15 and 50. If the number of cells per 1 mm 2 exceeds 50, dilute the sample and count again. If the number of cells per 1 mm 2 is less than 15, use a less diluted sample. If less dilute samples are not available, count cells on both sides of the hemocytometer (8 x 1 mm 2 areas).

Keep a separate count of viable and non-viable cells. If more than 25% of cells are non-viable, the culture is not being maintained on the appropriate amount of media. Reincubate the culture and adjust the volume of media according to the confluency of the cells and the appearance of the media. Include cells on top and left touching middle line. The cells touching middle line at bottom and right are not counted.

i. Trypan Blue is the "vital stain" excluded from live cells.
ii. Live cells appear colourless and bright (refractile) under phase contrast.
iii. Dead cells stain blue and are non-refractile.

  • %Cell Viability = [Total Viable cells (Unstained) / Total cells (Viable +Dead)] X 100.
  • Viable Cells/ml = Average viable cell count per square x Dilution Factor x 10 4 /
  • Average viable cell count per square = Total number of viable cells in 4 squares / 4.
  • Dilution Factor = Total Volume (Volume of sample + Volume of diluting liquid) / Volume of sample.
  • Total viable cells/Sample = Viable Cells/ml x The original volume of fluid from which the cell sample was removed.
  • Volume of media needed = (Number of cells needed/Total number of viable cells) x 1000.

Immunohistochemical Methods

The Enzymatic method for immunohistochemistry uses reagents like Calcium Chloride, Sodium Hydroxide, Hydrochloric Acid solutions, Xylenes for dewaxing, and Methanol. Immunohistochemistry use different staining procedures such as one step direct method, ABC methods, two-step indirect method and Tyramide signal amplification.

Direct Method

Direct method is one step staining method, and involves a labeled antibody (i.e. FITC conjugated antiserum) reacting directly with the antigen in tissue sections. This technique utilizes only one antibody and the procedure is short and quick. However, it is insensitive due to little signal amplification and rarely used since the introduction of indirect method.

Indirect Method

Indirect method involves an unlabeled primary antibody (first layer) which react with tissue antigen, and a labeled secondary antibody (second layer) react with primary antibody (Note: The secondary antibody must be against the IgG of the animal species in which the primary antibody has been raised). This method is more sensitive due to signal amplification through several secondary antibody reactions with different antigenic sites on the primary antibody. In addition, it is also economy since one labeled second layer antibody can be used with many first layer antibodies (raised from the same animal species) to different antigens. The second layer antibody can be labeled with a fluorescent dye such as FITC, rhodamine or Texas red, and this is called indirect immunofluorescence method. The second layer antibody may be labeled with an enzyme such as peroxidase, alkaline phosphatase or glucose oxidase, and this is called indirect immunoenzyme method.

PAP Method

(peroxidase anti-peroxidase method): PAP method is a further development of the indirect technique and it involves a third layer which is a rabbit antibody to peroxidase, coupled with peroxidase to make a very stable peroxidase anti-peroxidase complex. The complex, composed of rabbit gaba-globulin and peroxidase, acts as a third layer antigen and becomes bound to the unconjugated goat anti-rabbit gaba-globulin of the second layer. The sensitivity is about 100 to 1000 times higher since the peroxidase molecule is not chemically conjugated to the anti IgG but immunologically bound, and loses none of its enzyme activity. It also allows for much higher dilution of the primary antibody, thus eliminating many of the unwanted antibodies and reducing non-specific background staining.

Avidin-Biotin Complex (ABC) Method

ABC method is standard IHC method and one of widely used technique for immunhistochemical staining. Avidin, a large glycoprotein, can be labeled with peroxidase or fluorescein and has a very high affinity for biotin. Biotin, a low molecular weight vitamin, can be conjugated to a variety of biological molecules such as antibodies. The technique involves three layers. The first layer is unlabeled primary antibody. The second layer is biotinylated secondary antibody. The third layer is a complex of avidin-biotin peroxidase. The peroxidase is then developed by the DAB or other substrate to produce different colorimetric end products.

Labeled StreptAvidin Biotin (LSAB) Method

Streptavidin, derived from streptococcus avidini, is a recent innovation for substitution of avidin. The streptavidin molecule is uncharged relative to animal tissue, unlike avidin which has an isoelectric point of 10, and therefore electrostatic binding to tissue is eliminated. In addition, streptavidin does not contain carbohydrate groups which might bind to tissue lectins, resulting in some background staining. LSAB is technically similar to standard ABC method. The first layer is unlabeled primary antibody. The second layer is biotinylated secondary antibody. The third layer is Enzyme-Streptavidin conjugates (HRP-Streptavidin or AP-Streptavidin) to replace the complex of avidin-biotin peroxidase. The enzyme is then visualized by application of the substrate chromogen solutions to produce different colorimetric end products. The third layer can also be Fluorescent dye-Streptavidin such as FITC-Streptavidin if fluorescence labeling is preferred. A recent report suggests that LSAB method is about 5 to 10 times more sensitive than standard ABC method.

Guidelines for Indirect Staining in Flow Cytometry Using Secondary Detection Reagents

Fig. 2. Staining Strategies. Illustration of the different types of staining protocols that may need to be employed to have successful indirect and direct staining in flow cytometry.

1. Simple Single Primary Antibody Staining

When staining samples with a directly conjugated primary antibody, after the initial incubation and a few simple washes, the sample is ready to be acquired (Figure 2A). Indirect staining using an unconjugated primary antibody, detected with a secondary antibody, involves extra steps: incubate with the primary antibody, wash, and then incubate with the fluorescently-labeled secondary antibody that recognizes your primary antibody. After more washes, the sample is ready to be acquired (Figure 2B).

However if you have a combination of conjugated and unconjugated primary antibodies, which require secondary antibodies in your panel, staining can be more complicated. To avoid nonspecific binding, there are staining protocols, blocking and washing steps, and antibody choices to consider in addition to the usual flow cytometry controls.

2. Combining Conjugated and Unconjugated Antibodies

A common problem when combining unconjugated and conjugated antibodies is inappropriate detection of the conjugated primary antibody/antibodies by the secondary antibody, leading to false positives. This problem is illustrated in a video which demonstrates practically how using Bio-Rad&rsquos mouse IgG isotype specific secondary antibodies can deliver results you can trust in imaging multiplexing, without the risk of false positives, and in only two simple steps. To avoid this problem, it is necessary to be able to distinguish between the unconjugated and conjugated primary antibodies by means of species, class, or isotype. The primary antibodies can be incubated together, washed and then labeled with secondary antibodies, which only detect the species, class, or isotype of the unconjugated primary antibodies (Figure 2C). To optimize your staining, ideally both the primary and secondary antibodies should be titrated to ensure minimum background with a maximal specific signal. It is also recommended to include a secondary antibody-only control and a control containing all antibodies except the unlabeled primary antibody to check for nonspecific staining. To get help with your staining, view our detailed staining protocols.

3. Blocking and Washing

Incubation with an appropriate serum, before adding primary antibodies, can reduce background by blocking Fc receptors. This should be of the same species as the cell type being analyzed, never the same species as the host of the primary antibody, as the secondary antibody will recognize the blocking serum AND the primary antibody. If you are using bovine serum albumin, it may be contaminated with bovine IgG which can cross-react with closely related species such as goat and sheep. Include adequate washing steps to remove excess primary and secondary antibodies. Intracellular staining protocols may require more and longer washing steps to remove all the excess antibody. Furthermore, the choice of permeabilization and wash buffer may affect staining and should be optimized for your experiment.

4. Antibody Choice

The antibody choice for your primary and secondary antibodies will influence your experimental design. Unwanted species cross-reactivity can be removed by using cross-adsorbed or isotype specific secondary antibodies. This will ensure the signal from the secondary antibody is specific to the primary target. If you are staining immune tissue, which contains a lot of Fc receptors, you may want to consider a F(ab) fragment as a secondary antibody in addition to blocking with serum to reduce nonspecific binding. When multiplexing, if there are no primary antibodies raised in different species, different classes of immunoglobulin may be available, for example IgG and IgM, allowing class specific secondary antibodies to be used, from the same species. Many monoclonal primary antibodies are mouse or rat IgG1, IgG2a, or IgG2b. Isotype specific secondary antibodies allow multiplexing of purified antibodies from the same species. You may be able to increase your staining options by mixing and matching these methods, however always ensure your secondary antibodies do not recognize multiple targets and the correct controls are performed.

5. Compensation

When using labeled secondary antibodies in a multicolor panel, you should still perform the correct compensation control for the secondary fluorophore. The easiest way is to carry out the single staining compensation control using the unconjugated primary antibody and the associated secondary antibody. If this proves problematical, due to low expression for example, then compensation beads can be used. Depending upon the antibody-binding capability of the beads, it may be possible to use these directly with the secondary antibody, or with the primary plus secondary antibodies. A third option is to perform compensation using a primary antibody directly conjugated to the same fluorophore that will be used on the secondary antibody. However, the use of tandem fluorophores with this method should be done with extreme caution due to inherent differences between tandem fluorophore batches.

6. Streptavidin as a Secondary Reagent

Sometimes only a biotinylated primary antibody is available, in this case, the use of fluorescently labeled streptavidin with such an antibody will make it suitable in flow cytometry without needing secondary antibodies. This option removes the need to check for cross-reactivity of secondary antibodies with other primary antibodies and can expand co-staining options. However, it is still necessary to perform the correct negative and compensation controls.

7. General Flow Controls/Tips

Specific controls are required for flow cytometry such as unstained, isotype, viability, compensation, and FMO controls. Building multicolor panels can be tricky and following guidelines related to instrument and fluorophore choice are equally important when performing indirect staining. In addition to these standard flow cytometry controls and guidelines, use of secondary antibodies may require additional controls and optimization of both antibodies and staining protocol. The nonspecific binding of the secondary antibody can be assessed by staining with the secondary antibody without the specific primary antibody, either alone, or in combination with primary antibodies that are not the intended target. Alternatively a primary antibody, which the secondary antibody binds to, and known to be negative in your sample, can be used. This will give you an idea of the binding to both the sample and other antibodies in the panel, allowing you to assess the nonspecific background staining.

For more information on secondary antibodies and their use in various applications, visit our dedicated secondary antibodies webpage.

Watch the video: Lab 5 Special stain (January 2023).