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Diagnostic Tools Used in Fish Disease Diagnosis

Arun Sudhagar S, Ezhil Nilavan S, Linga Prabu D, Rathi Bhuvaneswari G, Chandrasekar S and Rajesh kumar R.

Central Institute of Fisheries Education,




Diagnosis is a label given for a medical condition or disease identified by its signs, symptoms, and from the results of various diagnostic procedures. The term "diagnostic criteria" designates the combination of signs, symptoms, and test results that allows the clinician to ascertain the diagnosis of the respective disease. It can be definition as "the recognition of a disease or condition by its outward signs and symptoms," or "the analysis of the underlying physiological/biochemical cause(s) of a disease or condition." Disease diagnosis is refers to the various procedures and techniques used to identify the nature of disease and to precisely pinpoint the primary and secondary pathogens involved. Proper diagnosis leads to accurate therapies and avoid indiscriminate use of chemotherapeutics. Disease diagnosis is an integral part of aquatic health management. Proper diagnosis helps to adopt accurate therapy and avoid indiscriminate use of chemotherapeutics. Information on case history and clinical signs should be carefully used while examining samples for diagnosis.

Importance of case history

Case history information is very vital for proper disease diagnosis and for taking proper remedial steps. During a disease outbreak, careful scrutiny of the case history information will help to precisely pinpoint the circumstances under which the disease has developed. Some of the case history that are taken in to account are,

  • Water quality

  • Feeding percentage

  • Feed intake

  • Fertilization schedule

  • Time of last Algal bloom

  • Liming details

  • Treatment details

  • Source of seed, stocking details

Classification of disease diagnostic methods

The disease diagnostic methods can be classified in to following types

  • Microscopic diagnosis

  • Histological diagnosis

  • Microbiological diagnosis

  • Immunological diagnosis

  • Molecular diagnosis

Microscopic diagnosis

Without a microscope it is simply impossible to tell the difference between a water quality and a parasite problem. The microscope should be considered the most basic of tools in fish disease diagnosis. Microscopy can be done quickly, but accuracy depends on the experience of the microscopist and quality of equipment. Most specimens are treated with stains that color pathogens, causing them to stand out from the background, although wet mounts of unstained samples can be used to detect fungi, parasites (including helminth eggs and larvae) and motile organisms. Visibility of fungi can be increased by applying 10% potassium hydroxide (KOH) to dissolve surrounding tissues and nonfungal organisms.

The stain is based on the likely pathogens, but no stain is 100% specific. Most samples are treated with Gram stain and, if mycobacteria are suspected, an acid-fast stain. However, some pathogens are not easily visible using these stains; if these pathogens are suspected, different stains or other identification methods are required because microscopic detection usually requires a microbe concentration of about 1 x 105/ml.

The microscopes that are commonly used in fish disease diagnosis are

  1. Light microscope

  2. Electron microscope

Light microscope

The light microscope is the most commonly used microscope in disease diagnosis. In this microscope the scrapings from the fish can be directly visualized or a squash of organ can be prepared for the examination. It can also be used for interpreting histological slides and stained microorganisms.

Electron microscope

They are broadly classified in to two types

  1. Transmission Electron Microscopy

They are used for observing the changes in the tissue at higher magnification.

  1. Scanning Electron Microscopy

They are used for observing the surface level changes in the tissues at higher magnification.

Histological diagnosis

Histology is a branch of biology that involves the microscopic examination of thin, stained tissue sections in order to study their structure and function and, in the case of histopathology, to determine changes which may be due to pathogens and disease. Histology and have a central role in disease diagnosis.Stained fish tissue sections are prepared and examined by light microscopy. Tissue changes resulting from infectious or non-infectious disease are identified and described. Immunohistochemical methods are also used to detect specific pathogens in tissue sections.


Following sampling, fish tissues are placed in an aqueous fixative. This fixative preserves the morphology (structure and chemical constituents) of tissues and cells, so that they are capable of withstanding further preparatory steps without change. It is essential that tissues are fixed within a very short time after death to avoid disintegration of tissues or cells by the action of their own enzymes. Following fixation, tissues are gradually dehydrated to remove any tissue water, using a series of graded alcohols. The tissues are then 'cleared', which involves treatment with a substance that mixes completely with both the dehydrating fluid and the embedding agent. Next the tissues are embedded in molten paraffin wax and cooled to harden the wax so that thin sections can be cut using a microtome and then mounted onto glass microscope slides. The wax is removed from the sections before staining.


After clearing and rehydration, the tissue sections can be stained using biological stains or dyes. Haematoxylin and eosin (H&E) is the most widely used histological stain because of its ability to reveal a wide range of different tissue components. Gram's stain is a staining method for differentiating microorganisms. The technique is based on the capability of bacteria cell walls to retain the crystal violet dye in the Gram stain during solvent treatment. The cell walls for Gram positive micro-organisms retain the primary violet as they have a higher peptidoglycan (sugars) and a lower lipid content than Gram negative bacteria. The Periodic Acid-Schiff (PAS) reaction is used to demonstrate certain carbohydrates that are present in some tissues, and provide identification of infecting fungus in fish tissues. The PAS positive sites stain pink/red.


Immunohistochemical staining methods have been developed for the detection of viruses such as infectious pancreatic necrosis virus (IPNV), infectious salmon anaemia virus (ISAV) and nodavirus in paraffin-embedded tissue sections. Viral antigen is localised by an antibody raised against the virus and subsequent detection steps result in a coloured product that can be visualised by light microscopy

Histological techniques enable the description of tissue pathology and highlight the sequence of cellular changes and their progression caused by infectious and noninfectious diseases. By examining stained sections, viruses, bacteria, fungi and parasites can be identified, and using immunohistochemical techniques, certain infectious agents can be detected in tissue sections. The increased use of image analysis tools by FRS allows qualitative data to be generated to enhance disease diagnosis.

Microbiological diagnosis

Bacterial diseases can be identified by various microbiology methods. The samples are taken from the fish under aseptic condition and they are first grown in non-selective media, then the various microbial diagnostic methods are used. The different microbiological diagnostic methods are

  1. Staining methods

  2. Motility test

  3. Culturing in selective medium

  4. Biochemical test

Staining method

Various staining methods are used to identify various bacterial pathogens among them gram staining is the most important and most commonly used method.

  • Gram stain: The Gram stain classifies bacteria according to whether they retain crystal violet stain (gram-positive — blue) or not (gram-negative — red) and highlights cell morphology (eg, bacilli, cocci) and cell arrangement (eg, clumps, chains, diploids). Such characteristics can direct antibiotic therapy pending definitive identification. To do a Gram stain, technicians heat-fix specimen material to a slide and stain it by sequential exposure to Gram's crystal violet, iodine, decolorizer, and counterstain (typically safranin).

  • Acid-fast and moderate (modified) acid-fast stains: These stains are used to identify acid-fast organisms (Mycobacterium sp) and moderately acid-fast organisms (primarily Nocardia sp). These stains are also useful for staining Rhodococcus and related genera, as well as oocysts of some parasites (eg, Cryptosporidium).

  • Fluorescent stains: These stains allow detection at lower concentrations (1 104 cells/mL). Examples are acridine orange (bacteria and fungi), auramine-rhodamine and auramine O (mycobacteria), and calcofluor white (fungi, especially dermatophytes). Coupling a fluorescent dye to an antibody directed at a pathogen (direct or indirect immunofluorescence) should theoretically increase sensitivity and specificity. However, these tests are difficult to read and interpret, and few (eg, Pneumocystis and Legionella direct fluorescent antibody tests) are commercially available and commonly used.

  • India ink (colloidal carbon) stain: This stain is used to detect mainly encapsulated fungi in a cell suspension. The background field, rather than the organism itself, is stained, which makes any capsule around the organism visible as a halo. Leukocytes may appear encapsulated.

  • Wright's stain and Giemsa stain: These stains are used for detection of parasites in blood, phagocytes and tissue cells, intracellular inclusions formed by viruses and some intracellular bacteria.

  • Trichrome stain (Gomori-Wheatley stain) and iron hematoxylin stain: These stains are used to detect intestinal protozoa. The Gomori-Wheatley stain is used to detect microsporidia. It may miss helminth eggs and larvae. The iron hematoxylin stain differentially stains cells, cell inclusions, and nuclei. Helminth eggs may stain too dark to permit identification.

Motility test

Histological techniques enable the description of tissue pathology and highlight the sequence of cellular changes and their progression caused by infectious and noninfectious diseases. By examining stained sections, viruses, bacteria, fungi and parasites can be identified, and using immunohistochemical techniques, certain infectious agents can be detected in tissue sections. The increased use of image analysis tools by FRS allows qualitative data to be generated to enhance disease diagnosis.

There are three methods to see the motility of the bacteria and they are

  1. Hanging Drop Method

  2. Semi Solid Agar Method (Craigie)

  3. Three Coverslip Method

Hanging Drop Method

  • Place a small drop of liquid bacterial culture in the center of a coverslip

  • Place a small drop of water at each corner of the coverslip

  • Invert a slide with a central depression over the coverslip

  • The coverslip will stick to the slide and when the slide is inverted the drop of bacterial culture will be suspended in the well

  • Examine microscopically (x400) for motile organisms

Note: If well slides are not available, a ring of Vaseline or plasticine may instead be made on an ordinary microscope slide

Positive result: A darting, zigzag, tumbling or other organised movement

Negative result: No movement or Brownian motion only.

Semi Solid Agar Method (Craigie)

  • Inoculate the test organism - the central glass tube

  • Incubate at the relevant temperature for 18-24 hours

  • Subculture from the outer section of the medium

Positive result: Organism can be recovered from the outer section of the medium
Negative result: Organism remains in the inner tube

Three coverslip method

  • Set microscope slide according to Figure 1

  • Place a small drop of bacterial culture in center of third coverslip.

  • Place a small drop of water at each corner of the coverslip.

  • Invert the prepared microscope slide over the third coverslip.

  • The coverslip should stick to the other coverslips on the slide and when inverted the drop of bacterial culture should be suspended in the gap between the microscope slide and third coverslip.

  • Examine microscopically (x400) for motile organism.

Positive result: A darting, zigzag, tumbling or other organised movement.

Negative result: No movement or only Brownian motion.

Culturing bacteria pathogen in selective medium

Certain bacteria have the capacity to grow in selective media. This property of bacteria is used to identify the bacterial pathogen. Only particular kind of bacteria can grow in selective media. Example for selective medium is TCBS for Vibrios.

Biochemical tests

Various biochemical test are used for the identification of bacteria based on the character of a particular bacteria to give its positive and negative result for a particular biochemical test.

  1. The Indole Test

The test organism is inoculated into tryptone broth, a rich source of the amino acid tryptophan. Indole positive bacteria such as Escherichia coli produce tryptophanase, an enzyme that cleaves tryptophan, producing indole and other products. When Kovac's reagent (p-dimethylaminobenzaldehyde) is added to a broth with indole in it, a dark pink color develops. The indole test must be read by 48 hours of incubation because the indole can be further degraded if prolonged incubation occurs. The acidic pH produced by Escherichia coli limits its growth.

  1. The Methyl Red and Voges-Proskauer Tests

The methyl red (MR) and Voges-Proskauer (VP) tests are read from a single inoculated tube of MR-VP broth. After 24-48 hours of incubation the MR-VP broth is split into two tubes. One tube is used for the MR test; the other is used for the VP test. Media contains glucose and peptone. All enterics oxidize glucose for energy; however the end products vary depending on bacterial enzymes. Both the MR and VP tests are used to determine what end products result when the test organism degrades glucose. E. coli is one of the bacteria that produces acids, causing the pH to drop below 4.4. When the pH indicator methyl red is added to this acidic broth it will be cherry red (a positive MR test). Klebsiella and Enterobacter produce more neutral products from glucose (e.g. ethyl alcohol, acetyl methyl carbinol). In this neutral pH the growth of the bacteria is not inhibited. The bacteria thus begin to attack the peptone in the broth, causing the pH to rise above 6.2. At this pH, methyl red indicator is a yellow color (a negative MR test).

The reagents used for the VP test are Barritt's A (alpha-napthol) and Barritt's B (potassium hydroxide). When these reagents are added to a broth in which acetyl methyl carbinol is present, they turn a pink-burgundy color (a positive VP test). This color may take 20 to 30 minutes to develop.

  1. Catalase Test

Catalase is the enzyme that breaks hydrogen peroxide (H2O2) into H2O and O2. Hydrogen peroxide is often used as a topical disinfectant in wounds, and the bubbling that is seen is due to the evolution of O2 gas. H2O2 is a potent oxidizing agent that can wreak havoc in a cell; because of this, any cell that uses O2 or can live in the presence of O2 must have a way to get rid of the peroxide. One of those ways is to make catalase.

  1. The Citrate Test

The citrate test utilizes Simmon's citrate media to determine if a bacterium can grow utilizing citrate as its sole carbon and energy source. Simmon's media contains bromthymol blue, a pH indicator with a range of 6.0 to 7.6. Bromthymol blue is yellow at acidic pH's (around 6), and gradually changes to blue at more alkaline pH's (around 7.6). Uninoculated Simmon's citrate agar has a pH of 6.9, so it is an intermediate green color. Growth of bacteria in the media leads to development of a Prussian blue color (positive citrate). Enterobacter and Klebsiella are citrate positive while E.coli is negative.

  1. Oxidase Test

The oxidase test identifies organisms that produce the enzyme cytochrome oxidase.  Cytochrome oxidase participates in the electron transport chain by transferring electrons from a donor molecule to oxygen. The oxidase reagent contains a chromogenic reducing agent, which is a compound that changes color when it becomes oxidized.  If the test organism produces cytochrome oxidase, the oxidase reagent will turn blue or purple within 15 seconds.

  1. Nitrate Reduction Test

Nitrate broth is used to determine the ability of an organism to reduce nitrate (NO3) to nitrite (NO2) using the enzyme nitrate reductase. It also tests the ability of organisms to perform nitrification on nitrate and nitrite to produce molecular nitrogen. Nitrate broth contains nutrients and potassium nitrate as a source of nitrate. After incubating the nitrate broth, add a dropperful of sulfanilic acid and a-naphthylamine. If the organism has reduced nitrate to nitrite, the nitrites in the medium will form nitrous acid. When sulfanilic acid is added, it will react with the nitrous acid to produce diazotized sulfanilic acid. This reacts with the a-naphthylamine to form a red-colored compound. Therefore, if the medium turns red after the addition of the nitrate reagents, it is considered a positive result for nitrate reduction.If the medium does not turn red after the addition of the reagents, it can mean that the organism was unable to reduce the nitrate, or it could mean that the organism was able to denitrify the nitrate or nitrite to produce ammonia or molecular nitrogen. Therefore, another step is needed in the test. If the medium does not turn red after the addition of the nitrate reagents, add a small amount of powdered zinc. Be careful, as powdered zinc is hazardous! If the tube turns red after the addition of the zinc, it means that unreduced nitrate was present. Therefore, a red color on the second step is a negative result. The addition of the zinc reduced the nitrate to nitrite, and the nitrite in the medium formed nitrous acid, which reacted with sulfanilic acid. The diazotized sulfanilic acid that was thereby produced reacted with the a-naphthylamine to create the red complex. If the medium does not turn red after the addition of the zinc powder, then the result is called a positive complete. If no red color forms, there was no nitrate to reduce. Since there was no nitrite present in the medium, either, that means that denitrification took place and ammonia or molecular nitrogen were formed.

  1. Urease Test

Urease broth is a differential medium that tests the ability of an organism to produce an exoenzyme, called urease that hydrolyzes urea to ammonia and carbon dioxide.  The broth contains two pH buffers, urea, a very small amount of nutrients for the bacteria, and the pH indicator phenol red.  Phenol red turns yellow in an acidic environment and fuchsia in an alkaline environment.  If the urea in the broth is degraded and ammonia is produced, an alkaline environment is created, and the media turns pink. Many enterics can hydrolyze urea; however, only a few can degrade urea rapidly.  These are known as "rapid urease-positive" organisms.  Members of the genus Proteus are included among these organisms. Urea broth is formulated to test for rapid urease-positive organisms.  The restrictive amount of nutrients coupled with the use of pH buffers prevent all but rapid urease-positive organisms from producing enough ammonia to turn the phenol red pink.

  1. Phenol Red Broth

Phenol Red Broth is a general-purpose differential test medium typically used to differentiate gram negative enteric bacteria.  It contains peptone, phenol red (a pH indicator), a Durham tube, and one carbohydrate.  We use three different kinds of phenol red broths. One contains glucose; one contains lactose, and the last contains sucrose.  The objective of the exercise is to determine which organisms can utilize each sugar. Phenol red is a pH indicator which turns yellow below a pH of 6.8 and fuchsia above a pH of 7.4. If the organism is able to utilize the carbohydrate, an acid by-product is created, which turns the media yellow.  If the organism is unable to utilize the carbohydrate but does use the peptone, the by-product is ammonia, which raises the pH of the media and turns it fuchsia. When the organism is able to use the carbohydrate, a gas by-product may be produced. If it is, an air bubble will be trapped inside the Durham tube.  If the organism is unable to utilize the carbohydrate, gas will not be produced, and no air bubble will be formed.

  1. Casease Test

Skim milk agar is a differential medium that tests the ability of an organism to produce an exoenzyme, called casease, that hydrolyzes casein. Casein forms an opaque suspension in milk that makes the milk appear white. Casease allows the organisms that produce it to break down casein  into smaller polypeptides, peptides, and amino acids that can cross the cell membrane and be utilized by the organism.  When casein is broken down into these component molecules, it is no longer white.  If an organism can break down casein, a clear halo will appear around the areas where the organism has grown.

  1. Gelatinase Test

Nutrient gelatin is a differential medium that tests the ability of an organism to produce an exoenzyme, called gelatinase, that hydrolyzes gelatin. Gelatin is commonly known as a component of gelled salads and some desserts, but it's actually a protein derived from connective tissue. When gelatin is at a temperature below 32°C (or within a few degrees thereof), it is a semisolid material.  At temperatures above 32°C, it is a viscous liquid. Gelatinase allows the organisms that produce it to break down gelatin into smaller polypeptides, peptides, and amino acids that can cross the cell membrane and be utilized by the organism. When gelatin is broken down, it can no longer solidify.  If an organism can break down gelatin, the areas where the organism has grown will remain liquid even if the gelatin is refrigerated.

  1. Lipase Test

Tributyrin agar is a differential medium that tests the ability of an organism to produce an exoenzyme, called lipase, that hydrolyzes tributyrin oil.  Lipases break down lipids (fats). Tributyrin oil is a type of lipid called a triglyceride. Other lipase tests use different fat sources such as corn oil, olive oil, peanut oil, egg yolk, and soybean oil. Lipase allows the organisms that produce it to break down lipids into smaller fragments. Triglycerides are composed of glycerol and three fatty acids.  These get broken apart and may be converted into a variety of end-products that can be used by the cell in energy production or other processes. Tributyrin oil forms an opaque suspension in the agar. When an organism produces lipase and breaks down the tributyrin, a clear halo surrounds the areas where the lipase-producing organism has grown.

  1. Starch Hydrolysis

Starch agar is a differential medium that tests the ability of an organism to produce certain exoenzymes, including a-amylase and oligo-1,6-glucosidase, that hydrolyze starch.  Starch molecules are too large to enter the bacterial cell, so some bacteria secrete exoenzymes to degrade starch into subunits that can then be utilized by the organism.  Starch agar is a simple nutritive medium with starch added.  Since no color change occurs in the medium when organisms hydrolyze starch, we add iodine to the plate after incubation. Iodine turns blue, purple, or black (depending on the concentration of iodine) in the presence of starch. A clearing around the bacterial growth indicates that the organism has hydrolyzed starch.

  1. Triple Sugar Iron Agar

Triple sugar iron agar (TSI) is a differential medium that contains lactose, sucrose, a small amount of glucose (dextrose), ferrous sulfate, and the pH indicator phenol red.  It is used to differentiate enterics based on the ability to reduce sulfur and ferment carbohydrates.  As with the phenol red fermentation broths, if an organism can ferment any of the three sugars present in the medium, the medium will turn yellow.  If an organism can only ferment dextrose, the small amount of dextrose in the medium is used by the organism within the first ten hours of incubation. After that time, the reaction that produced acid reverts in the aerobic areas of the slant, and the medium in those areas turns red, indicating alkaline conditions. The anaerobic areas of the slant, such as the butt, will not revert to an alkaline state, and they will remain yellow. This happens with Salmonella and Shigella

  1. Decarboxylation Test

Decarboxylase broth tests for the production of the enzyme decarboxylase, which removes the carboxyl group from an amino acid. Decarboxylase broth contains nutrients, dextrose (a fermentable carbohydrate), pyridoxal (an enzyme cofactor for decarboxylase), and the pH indicators bromcresol purple and cresol red. Bromcresol purple turns purple at an alkaline pH and turns yellow at an acidic pH. We also add a single amino acid to each batch of decarboxylase broth.   The three amino acids we test in our decarboxylase media are arginine, lysine, and ornithine. The decarboxylase test is useful for differentiating the Enterobacteriaceae.Each decarboxylase enzyme produced by an organism is specific to the amino acid on which it acts. Therefore, we test the ability of organisms to produce arginine decarboxylase, lysine decarboxylase, and ornithine decarboxylase using three different but very similar media. If the organism is unable to ferment dextrose, there will be no color change in the medium.  If an organism is able to ferment the dextrose, acidic byproducts are formed, and the media turns yellow.  As the organisms ferment the dextrose, the media initially turns yellow, even when it has been inoculated with a decarboxylase-positive organism.  The low pH and the presence of the amino acid will cause the organism to begin decarboxylation. If an organism is able to decarboxylate the amino acid present in the medium, alkaline byproducts are then produced.  Arginine is hydrolyzed to ornithine and is then decarboxylated.  Ornithine decarboxylation yields putrescine.  Lysine decarboxylation results in cadaverine. These byproducts are sufficient to raise the pH of the media so that the broth turns purple.  If the inoculated medium is yellow, or if there is no color change, the organism is decarboxylase-negative for that amino acid.  If the medium turns purple, the organism is decarboxylase-positive for that amino acid.

  1. Coagulase Test

The coagulase test identifies whether an organism produces the exoenzyme coagulase, which causes the fibrin of blood plasma to clot. Organisms that produce catalase can form protective barriers of fibrin around themselves, making themselves highly resistant to phagocytosis, other immune responses, and some other antimicrobial agents. The coagulase slide test is used to identify the presence of bound coagulase or clumping factor, which is attached to the cell walls of the bacteria. Bound coagulase reacts with the fibrinogen in plasma, causing the fibrinogen to precipitate. This causes the cells to agglutinate, or clump together, which creates the "lumpy" look of a positive coagulase slide test.  You may need to place the slide over a light box to observe the clumping of cells in the plasma.The coagulase tube test has been set up as a demo for you to observe in class. This version of the coagulase test is used to identify the presence of either bound coagulase or free coagulase, which is an extracellular enzyme. Free coagulase reacts with a component of plasma called coagulase-reacting factor. The result is to cause the plasma to coagulate.  In the demo, the coagulase plasma has been inoculated with Staphylococcus aureus and Staphylococcus epidermidis and allowed to incubate at 37°C for 24 hours.  Staphylococcus aureus produces free coagulase; Staphylococcus epidermidis does not. The coagulase test is useful for differentiating potentially pathogenic Staphylococci such as Staphylococcus aureus from other Gram positive, catalase-positive cocci.

Immunological Diagnosis

Antigen — Antibody reactions are highly specific and sensitive. This forms the basis of immunodiagnostics. These tools are used for the qualitative and quantitative estimation of the pathogens and/or the protective antibody. These tests can be used at the farm level without the aid of the instruments. It is apparent that conventional diagnostic methods rely solely on the microscopic examination and its visual recognition, which requires a great deal of experience with the organisms that often change so dramatically in their morphology during the course of their development. Therefore, in the evolution of disease diagnostic procedures in aquaculture, antibody-based (protein-based) immunodiagnosis plays a crucial role. This method has the advantage over other traditional method in that it can detect sub-clinical/latent/carrier sate of infection and can also discriminate the antigenic differences. This technique is relatively rapid and more specific and sensitive. Further refinement of conventional immunodiagnostic techniques has resulted in the development of monoclonal antibody-based techniques and this has increased the accuracy of detection and has allowed studying the pathogenesis of diseases. Nevertheless, the specificity of antibodies also limits their usefulness because major antigens are not conserved among life stages of certain pathogens (Bartholomew et al., 1995). Although there is an array of polyclonal and monoclonal antibody-based diagnostics available for various aquatic animal pathogens.

  1. Agar Gel Precipitation Test,

  2. Agglutination Test,

  3. ELISA,

  4. Dot ELISA,

  5. Latex Agglutination Test and

  6. Fluorescent Antibody Test.

  1. Agar Gel Precipitation Test

In this test, antibody and possible antigens are placed in wells in agar plates and allowed to diffuse toward one another. The antibody is placed in a center well and antigens (specific or nonspecific) are placed in surrounding wells. When an antibody and its specific antigen meet one another and are at the proper concentrations, the precipitate will form a visible white line between the two wells. This line is called a precipitin line.

  1. Agglutination Test

This test is used to identify unknown antigens; blood with the unknown antigen is mixed with a known antibody and whether or not agglutination occurs helps to identify the antigen; used in tissue matching and blood grouping and diagnose. In the direct agglutination test, serum is added to a suspension of cells that have the surface self-Ag to be tested. If the individual's serum contains the specific auto-Abs, Ig will bind and, at the appropriate Ab concentration, the cells will become cross linked. This will cause agglutination, and the cells will form a mat at the bottom of the test well. Auto-Abs attached to a patient's cells can be detected by the addition of a second Ab and observed for agglutination. Selective soluble self-Ags can also be used to assay auto-Abs by attaching them to the surface of red blood cells. This latter type of agglutination test is called passive or indirect hemagglutination.

  1. ELISA

The purpose of an ELISA is to determine if a particular protein is present in a sample and if so, how much. There are two main variations on this method: you can determine how much antibody is in a sample, or you can determine how much protein is bound by an antibody. The distinction is whether you are trying to quantify an antibody or some other protein. In this example, we will use an ELISA to determine how much of a particular antibody is present in an individual's blood. ELISAs are performed in 96-well plates which permits high throughput results. The bottom of each well is coated with a protein to which will bind the antibody you want to measure. Whole blood is allowed to clot and the cells are centrifuged out to obtain the clear serum with antibodies (called primary antibodies). The serum is incubated in a well, and each well contains a different serum. A positive control serum and a negative control serum would be included among the 96 samples being tested. After some time, the serum is removed and weakly adherent antibodies are washed off with a series of buffer rinses. To detect the bound antibodies, a secondary antibody is added to each well. The secondary antibody would bind to all human antibodies and is typically produced in a rodent. Attached to the secondary antibody is an enzyme such as peroxidase or alkaline phosphatase. These enzymes can metabolize colorless substrates (sometimes called chromagens) into colored products. After an incubation period, the secondary antibody solution is removed and loosely adherent ones are washed off as before. The final step is the addition the enzyme substrate and the production of colored product in wells with secondary antibodies bound.When the enzyme reaction is complete, the entire plate is placed into a plate reader and the optical density (i.e. the amount of colored product) is determined for each well. The amount of color produced is proportional to the amount of primary antibody bound to the proteins on the bottom of the wells.


Dot-ELISA (Enzyme Linked Immunosorbent Assay) is an extensively used immunological tool in research as well as analytical/diagnostic laboratories. In sandwich Dot-ELISA, the antigen is sandwiched directly between two antibodies which react with two different epitopes on the same antigen. Here one of the antibodies is immobilized onto a solid support and the second antibody is linked to an enzyme. Antigen in the test sample first reacts with the immobilized antibody and then with the second enzyme-linked antibody. The amount of enzyme linked antibody bound is assayed by incubating the strip with an appropriate chromogenic substrate, which is converted to a coloured, insoluble product. The latter precipitates onto the strip in the area of enzyme activity, hence the name Dot-ELISA. The enzyme activity is indicated by intensity of the spot, which is directly proportional to the antigen concentration.

  1. Latex Agglutination Test

The latex agglutination test is a laboratory method to check for certain antibodies or antigens in a variety of bodily fluids including saliva, urine, cerebrospinal fluid or blood. In latex agglutination, an antibody (or antigen) is coated on the surface of latex particles which is known as sensitized latex. When a sample containing the specific antigen (or antibody) is mixed with the milky-appearing sensitized latex, it causes visible agglutination

  1. Fluorescent Antibody Test

It is a laboratory test that uses antibodies tagged with fluorescent dye that can be used to detect the presence of microorganisms. This method offers straight-forward detection of antigens using fluorescently labeled antigen-specific antibodies. Because detection of the antigen in a substrate of patient sample (cellular smear, fluid or patient- inoculated culture medium) is the goal, DFA is seldom quantitative.

Molecular diagnosis

Molecular techniques are potentially faster and more sensitive than culture, serology, and histology methods that are traditionally used to identify fish pathogens. During the last 15 years or so, molecular techniques have been increasingly employed to diagnose fish diseases. These techniques include polymerase chain reaction (PCR), restriction enzyme digestion, probe hybridization, in situ hybridization, and microarray. Pathogens can be detected from asymptomatic fish by molecular diagnostic techniques so disease outbreak could be prevented. Thus antibiotic treatment can be reduced so that creation of antibiotic resistant bacteria may be eliminated. In this paper molecular techniques for detection of fish pathogens are reviewed and the potential for their application are discussed. The application of new techniques as a routine tool in a diagnostic laboratory is an area where relevant literature is scarce and this may contribute to the reticence of some to adopt these methods.

  1. Polymerase Chain Reaction

Polymerase chain reaction is a technique for amplifying a specific region of DNA, defined by a set of two "primers" at which DNA synthesis is initiated by a thermostable DNA polymerase. Usually, at least a million-fold increase of a specific section of a DNA molecule can be realized and the PCR product can be detected by gel electrophoresis. The regions amplified are usually between 150-3,000 base pairs (bp) in length. Primer design is important to obtain greatest possible sensitivity and specificity. Therefore, the primers should be sufficiently long to allow a high annealing temperature and reduce the opportunity for nonspecific primer annealing, but primers that are too long may facilitate nonspecific annealing even to regions of DNA that are not perfectly complementary to the primer sequence. The reaction includes template DNA that may be in various forms, from a simple tissue lysate to purified DNA, primers, polymerase enzyme to catalyze creation of new copies of DNA, and nucleotides to form the new copies. During each round of the thermocycling reaction, the template DNA is denatured, primers anneal to their complementary regions and polymerase enzyme catalyses the addition of nucleotides to the end of each primer, thus creating new copies of the target region in each round. Theoretically, the increase in amount of product after each round will be geometric. Reverse transcriptase polymerase chain reaction (RT-PCR) is used to detect specific mRNA and determine levels of gene expression. Compared to the two other commonly used techniques for quantifying mRNA levels, Northern blot analysis and RNase protection assay, RT-PCR can be used to quantify mRNA levels from much smaller samples. In fact, this technique is sensitive enough to enable quantitation of RNA from a single cell. Over the last several years, the development of novel chemistries and instrumentation platforms enabling detection of PCR products on a real-time basis has led to widespread adoption of real-time RTPCR as the method of choice for quantitative changes in gene expression. Furthermore, real-time RT-PCR has become preferred method for validating results obtained from array analyses and other techniques that evaluate gene expression changes on a global scale. The sensitivity and specificity achieved in a well-designed RT-PCR make it an ideal tool for use in the surveillance and monitoring of covert infections. As in the eukaryotes, the prokaryotic rRNA genes contain highly conserved sequences. The potential utility of conserved regions to identify or amplify the rRNA genes, followed by exploitation of more variable regions of the genes or spacers to detect or identify bacteria that may be difficult or even impossible to culture has long been recognized.

  1. Multiplex PCR

New developments such as design of PCR conditions that can detect several pathogens at one time in a multiplex reaction will improve time and cost-efficiency of this methodology, countering one of the major arguments against the adoption of these techniques as routine. In multiplex PCR more than one target sequence can be amplified by including more than one pair of primers in the reaction. Multiplex PCR has the potential to produce considerable savings of time and effort within the laboratory without compromising test utility. Since its introduction,multiplex PCR has been successfully applied in many areas of nucleic acid diagnostics, including gene deletion analysis, quantitative and RNA detection. In the field of infectious diseases, the technique has been shown to be a valuable method for identification of viruses, bacteria, fungi and parasites.

  1. Labeling and detection of nucleic acids

A variety of labeling and detection systems exist for nucleic acid probes. Radioisotopes were once the norm but, in the interests of researcher safety, other methods are becoming increasingly popular. The variety of labels and detection methods now available can provide a system suitable for any application, from dot blots to in situ hybridization. These include labeling with a variety of haptens such as biotin or digoxygenin and detection by antibody binding coupled with fluorescent, chemiluminescent or colorimetric detection methods

  1. Restriction enzyme digestion

Restriction enzymes (or restriction endonucleases) cleave DNA in a very specific fashion. Type II restriction enzymes, most commonly used for DNA analysis and genetic engineering, each have a unique nucleotide sequence at which it cuts a DNA molecule. A particular restriction enzyme will cleave DNA at that only recognition sequence that is often a six base pair palindromic sequence, but others recognize four or even eight base pair sequences. A common use for restriction enzymes is to generate a "fingerprint" of a particular DNA molecule. Because of the sequence specificity of restriction enzymes, these enzymes can cut DNA into discrete fragments which can be resolved by gel electrophoresis. This pattern of DNA fragments generates a "DNA fingerprint" and each DNA molecule has its own fingerprint. Other restriction enzymes can be used to further characterization of a particular DNA molecule. The location of these restriction enzyme cleavage sites on the DNA molecule can be compiled to create a restriction enzyme map. These maps are very useful for identifying and characterizing a particular DNA plasmid or region. Restriction enzymes recognize specific short sequences of DNA and cleave the DNA at that site. Single nucleotide changes can result in the gain or loss of a restriction site, thus altering the number of fragments produced following digestion of DNA. These RFLP can be visualized following gel electrophoresis of the digested DNA to separate the fragments according to size. Differences in the RFLP profiles have revolutionized criminal investigations and have become powerful tools in the identification of individuals in paternity and maternity cases, population genetics, and in the diagnosis of a variety of diseases.

  1. Restriction Fragment Length Polymorphism (RFLP)

RFLP is a technique in which organisms may be differentiated by analysis of patterns derived from cleavage of their DNA. If two organisms differ in the distance between sites of cleavage of a particular restriction endonuclease, the length of the fragments produced will differ when the DNA is digested with a restriction enzyme. The similarity of the patterns generated can be used to differentiate species (and even strains) from one another. Isolation of sufficient DNA for RFLP analysis is time-consuming and labor intensive. However, PCR can be used to amplify very small amounts of DNA, usually in 2-3 hours, to the levels required for RFLP analysis. Therefore, more samples can be analyzed in a shorter time.

  1. Amplified Fragment Length Polymorphism (AFLP)

A rapid PCR-based technique, AFLP can be used for typing prokaryotes and eukaryotes. The method is based on the selective PCR amplification of genomic restriction fragments of the whole genome and has been shown to be rapid, reproducible, and highly discriminatory. Selected markers are amplified in a PCR, which makes AFLP an easy and fast tool for strain identification in agriculture, botany, microbiology, and animal breeding. The AFLP method used was essentially that described by Valsangiacomo et al., (1995). AFLP analysis belongs to the category of selective restriction fragment amplification techniques, which are based on the ligation of adapters to genomic restriction fragments followed by a PCR-based amplification with adapter-specific primers. For AFLP analysis, only a small amount of purified genomic DNA is needed; this is digested with two restriction enzymes, one with an average cutting frequency (like EcoRI) and a second one with a higher cutting frequency (like MseI or TaqI). Double-stranded oligonucleotide adapters are designed in such a way that the initial restriction site is not restored after ligation, which allows simultaneous restriction and ligation, while religated fragments are cleaved again. An aliquot is then subjected to two subsequent PCR amplifications under highly stringent conditions with adapter specific primers that have at their 3' ends an extension of one to three nucleotides running into the unknown chromosomal restriction fragment. Alternative AFLP typing procedures are based on one enzyme with a single adapter and analysis by agarose gel electrophoresis. A major improvement has been obtained by switching from radioactive to fluorescently labeled primers for detection of fragments in an automatic sequence. In addition, it has been shown that for small bacterial and fungal genomes a single PCR amplification with one and two selective nucleotides, respectively, on both primers are sufficient.

  1. Random Amplified Polymorphic DNA (RAPD)

The technically demanding method of RAPD has been applied to the study of crayfish plague fungus, Astacus astaci . RAPD uses a single primer in low-stringency polymerase chain reactions. Random binding of primers results in different sizes of fragments from samples with nonidentical DNA. Application of the RAPD technique grouped different isolates of the fungus and provides the means to carry out epidemiological investigations (Lilley et al., 1997; Oidtmann et al., 1999). The method has also been used to examine another Aphanomyces species that has resulted in serious losses in both farmed and wild fish in Asia (Lilley et al., 1997). Other fish pathogens have been studied using RAPD, but problems with reproducibility and risks of contamination render the method unsuitable as a stand-alone method of diagnosis. However, RAPD can be a useful technique as a first step in the development of specific primers or probes and has been used in such a way in the study of bacteria.

  1. In Situ Hybridization

In situ PCR has become a powerful molecular tool in research as well as clinical practice. This technique has resulted in an increased understanding of infectious and neoplastic diseases and improvements in diagnosis of disease. In situ RT-PCR gives more detailed information by allowing for highly sensitive detection of low abundance gene expression in a given cell while providing anatomical information. The usefulness of these techniques has been hampered by low detection sensitivity, poor reproducibility, and high backgrounds. Moreover, many of the methods used to visualize the results of PCR amplification within cells and tissues employ radioactive tracers, making performance of the techniques cumbersome and costly. Researchers have developed a method for specific fluorescent detection of gene expression using in situ RT-PCR. This method enables the researcher or clinician to detect low levels of gene expression within tissues with very low background interference while addressing many of the other existing drawbacks to using in situ RT-PCR. Potential Areas of Application are detection and diagnosis of viruses and other infectious agents in specific cell types within tissues, detection, and characterization of tumor cells within a tissue, detection, and diagnosis of genetic mutations in inherited diseases, and detection of genes and gene expression in tissue. Fluorescence in situ hybridization, or FISH, is a method used to label cells or chromosomes according to the sequences of nucleic acids contained within them. In microbiology, the nucleic acid that is labelled as RNA or DNA of the ribosomes and the target is usually whole cells. The process works by taking fluorescently labelled pieces of DNA or RNA called probes that are around 20 nucleotides in length. The probes are incubated in the presence of cells munder appropriate conditions to permit specific hybridization of probe to target nucleic acid. Cells types that contain ribosomes with complementary RNA sequences become labeled by the binding of the fluorescent probe in situ. These labelled cells can then be visualized by flow cytometric or fluorescence microscopy.

  1. DNA microarrays

There are a number of ways of using DNA microarrays for the detection of unique DNA (or RNA) sequences. One method is to fluorescently label all the DNA sequences in the test sample. The sample DNA that hybridizes to a specific location on the microarray can be detected by fluorescent array detection and the data analyzed by computer programs. Often more practical is to use competitive hybridization in which the test sample competes for hybridization to the tethered oligonucleotide, on the chip, with a fluorescent labeled competitor oligonucleotide. When the test DNA is perfectly complimentary to the tethered oligonucleotide, it will hybridize to the chip. When the test DNA is not perfectly complementary to the tethered oligonucleotide, the fluorescent labeled competitor oligonucleotide will bind to the tethered oligonucleotide on the chip and displace the test DNA. A fluorescent microarray detector and computer program can then analyze the fluorescent array for the presence or absence of the species/strain specific DNA sequence. Compared with traditional nucleic acid hybridization with membranes, microarrays offer the additional advantages of high density, high sensitivity, rapid detection, lower cost, automation, and low background levels. Microarrays may provide a better option for largescale diagnostic testing and can survey a sample for a multitude of sequences simultaneously. Since most of the pathogens genetic sequences are available in the GenBank, oligonucleotide probes complementary to all pathogens can be made and inserted into microarray so that variety of microbes could be detected by a single microarray chip. As a result, microarray-based technology is potentially well suited for identifying fish pathogens in fish populations. The microarray techniques does not require such sequence conservation, however, because all of the diverse gene sequences from different populations of the same functional group can be fabricated on arrays and used as probes to monitor contagious fish disease especially during the asymptomatic period of the diseases. Microarrays are already proving valuable for assaying gene expression. A large number of probes on an array can reveal which genes are expressed or are present in the sample. This type of array would be particularly useful in studies of pathogens, where the presence of certain genes or gene products indicate whether the organism is pathogenic or not. Set up cost for the use of DNA microarrays is high. However, once the equipment is available and microarrays have been prepared, cost per unit of sample analyzed will be low. Furthermore, analysis time is extremely short. DNA microarray technology will be used in the future for fish diseases diagnosis especially during the asymptomatic period of diseases.

  1. Loop Mediated Isothermal Amplification (LAMP)

Loop-mediated isothermal amplification (LAMP) is a novel nucleic acid amplification method that amplifies DNA with high specificity, efficiency and rapidity under isothermal conditions (Natomi et al, 2000). When combined with reverse transcription, this method can also amplify RNA sequences with high efficiency. The method relies on auto-cycling strand displacement DNA synthesis using a DNA polymerase with a high strand displacement activity and a set of four specially designed primers. These four primers, termed as inner and outer primers, recognize six distinct sequences of the target DNA, which improves the specificity of the reaction. The reaction is carried out at isothermal condition, as the denaturation of strands takes place by strand displacement.

  1. LAMP Reaction: In the initial stages of LAMP reaction all four primers are involved, however, in the later cycling reaction only the inner primers are used for strand displacement DNA synthesis. The LAMP reaction is initiated by an inner primer containing sequences of sense and anti-sense strands of the target DNA. This is followed by the release of a single-stranded DNA through the priming by an outer primer. This single-stranded DNA will serve as a template for DNA synthesis primed by the second inner and outer primers that can hybridize at the other end of the target. This process will result in the formation of a stem-loop DNA structure. In the subsequent step of LAMP cycling, one inner primer will hybridize to the loop on the product and initiate strand displacement DNA synthesis which will result in the original stem-loop DNA and a new stem-loop. Cycling continues for a period of approximately 1 h and results in the accumulation of 109 copies the target. The final products of the reaction are stem-loop DNA with several inverted repeats of the target and cauliflower-like structures with multiple loops.

  2. Visualization of Amplified Products: Several methods can be employed to visualize the end products of LAMP reaction. The most common method of visualization is by agarose gel electrophoresis. The agarose gel is stained with intercalating dyes such as ethidium bromide or SYBR Green I. Since the end products of LAMP consist of stem-loop DNA and cauliflower-like structures with multiple loops of various lengths, the agarose gel electrophoresis will reveal the products from the minimum length of the target DNA to the loading well, which appears as smear at the top and bands at the base of the gel. Since, one of the characteristics of the LAMP reaction is its ability to synthesize extremely large amount of DNA, addition of intercalating dye, SYBR Green I, into the reaction tube itself would help in visualizing the product under a UV- transilluminator. This method is useful in the field-level application, where gel electrohoresis will be a limiting factor. Another method is also based on the accumulation large amount of byproduct of the reaction. In the LAMP reaction large amount of byproduct, pyrophosphate ion is produced, which will yield white precipitate of magnesium pyrophosphate in the reaction mixture. Hence, detection of presence or absence of white precipitate will provide an easy distinction of whether the target DNA is amplified during the reaction. Further, since increase in the turbidity of the reaction mixture according to the production of the precipitate correlates to the amount of target DNA synthesized, a colorimetric estimation of the turbidity in real-time is also being used as an efficient method of visualizing the amplified product.

  3. Advantages of LAMP: LAMP amplifies the target DNA under isothermal amplification with high efficiency; The detection limit of LAMP is a few copies and comparable to PCR; No significant influence of the co-presence of non-target DNA; LAMP allows simple, easy and selective detection; Lamp is highly specific for the target sequence, as it employs four primers targeting multiple sequences; LAMP is simple and easy to perform, as it requires (after appropriate primers are prepared) only a regular laboratory water bath or heat block for the reaction; BY incorporating reverse transcription, LAMP can be used for amplifying RNA as well.

  4. Disadvantages: Because amplification of the target DNA is so high at the final stage it is vulnerable to contamination in subsequent amplifications; Multiplexing is not possible with LAMP.

Advantages of Molecular Methods

Apart from the sensitivity and rapidity of diagnosis, principal advantage of molecular diagnostic methods is in the detection of non-culturable agents; DNA amplification can assist in detecting the pathogens that are present in low numbers and also in handling a tiny volume of specimen; Can be sued to detect latent infection and thereby identify the reservoir hosts of infection that is significant in epizootiology; Can be used to differentiate antigenically similar pathogens.

Disadvantages of Molecular methods

These methods are cost-intensive procedures; These tests cannot detect unsuspected samples; Molecular methods will have difficulty in detecting new pathogens as the exclusive use of these would overlook such infections.


Diagnostic methods for aquatic animal pathogens have advanced from microscopic characterization and morphological descriptions to molecular characterization and probe-based diagnosis. Molecular tools are increasingly relevant to fish diseases. The sequencing of the complete genomes of pathogens is allowing great advances in studying the biology, and improving diagnosis and control of pathogens. Using nucleic acid as targets, and new methods of analyzing polymorphism in this nucleic acid, can improve specificity, sensitivity, and speed of diagnosis and offer means of examining the relationships between genotype and phenotype of various pathogens. Progress in techniques aids epidemiological studies as well as identifying causes of disease outbreaks or the presence of pathogens. Therefore, molecular biology can be a routine tool in the search for improved methods of diagnosis and control of fish pathogens and the epidemiology of infectious fish diseases.


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