Principle:

• Soil is the prime storage of the microorganisms producing antibiotics which are able to inhibit the growth of other microorganisms. Antibiotics have been implemented in one or other forms for centuries. The screening of wild isolates from the soil has yielded the broad majority of new antibiotics. Even if the purification of several hundred naturally produced antibiotics has been performed, only a few have been proved to be successful to be used in medical practice. Those which are presently of greatest use were derived from a comparatively small group of microorganisms belonging to the genera Penicillium, Streptomyces, Cephalosporium, Micomonospora and Bacillus. In this era, the continuous efforts to develop new antibiotics are the emerging trends.
• Even if soils from various parts of the world are continually screened in industrial laboratories in order to isolate new antibiotic-producing microorganisms, industrial microbiology is directing its efforts toward chemical modification of existing antibiotic substances. This is completed by adding or replacing chemical side chains, reorganizing intramolecular bonding, or producing mutant microbial strains able to excrete a more potent form of the antibiotic. The establishment of chemical congeners accounts for the overcoming of antibiotic resistance, reducing adverse side effects in the host and increasing the effective spectrum of a given antibiotic.
• I: We will use the crowded-plate technique for the isolation of antibiotic producing microorganisms from two soil samples, one of which is seeded with Streptomyces griseus to serve as a positive control.
• II: For the determination of anti-microbial spectrum of isolates, isolates manifesting antibiotic activity will be screened against several different microorganisms to establish their effectiveness.

I. Isolation of antibiotic producing microorganisms

Requirements:

1.  Soil Suspensions:
   • – 1:500 dilution of soil sample suspension (0.1 g of soil per 50 ml of tap water) to serve as an unknown
   • – 1:500 dilution of soil sample seeded with S. griseus (0.1 g of soil per 50 ml of tap water) to serve as a positive control.
2. Media:
   • Six 15-ml Trypticase soy agar deep tubes, and two Trypticase soy agar slants.
3. Equipment:
   – 500-ml beaker
   – test tubes
   – test tube rack
   – sterile Petri dishes
   – inoculating needle
   – hot plate
   – thermometer
   – 1-ml and 5-ml pipettes
   – mechanical pipetting device
   – magnifying hand lens.

Procedure for Isolation of antibiotic producing microorganisms

• Label two sets of three sterile Petri dishes with the types of soil samples being used and dilutions (1:1000, 1:2000, and 1:4000).
• Place six Trypticase soy agar deep tubes into a beaker of water and bring to 100°C on a hot plate. Once agar is liquefied, add cool water to the water bath. Cool to 45°C, checking the temperature with a thermometer.
• Prepare a serial dilution of the unknown and positive control 1:500 soil samples as follows:
  – Label three test tubes 1, 2, and 3. With a pipette, add 5 ml of tap water to each tube.
  – Shake the provided 1:500 soil sample thoroughly for 5 minutes to effect a uniform soil-water suspension.
  – Using a 5-ml pipette, transfer 5 ml from the 1:500 dilution to Tube 1 and mix. The final dilution is 1:1000.
  – Using another pipette, transfer 5 ml from Tube 1 to Tube 2 and mix. The final dilution is 1:2000.
  – Using another pipette, transfer 5 ml from Tube 2 to Tube 3 and mix. The final dilution is 1:4000.
  – Using separate 1-ml pipettes, transfer 1 ml of the 1:1000, 1:2000, and 1:4000 dilutions to their appropriately labelled Petri dishes.
  – Pour one tube of molten Trypticase soy agar, cooled to 45°C, into each plate and mix by gentle rotation.
  – Allow all plates to solidify.
• Incubate all plates in an inverted position for 2 to 4 days at 25°C.
• Examine all crowded-plate dilutions for colonies exhibiting zones of growth inhibition. Use a hand magnifying lens if necessary. Record in the Lab Report the number of colonies showing zones of inhibition.
• Aseptically isolate one colony showing a zone 34 of growth inhibition from each soil culture
  with an inoculating needle and streak onto
  Trypticase soy agar slants labelled with the soil sample from which the isolate was obtained

• Incubate the slants for 2 to 4 days at 25°C. These will serve as stock cultures of antibiotic-producing isolates to be used in Part B.

II. Determination of antimicrobial spectrum of isolates

Requirements:

1. Cultures:
   • – 24-hour Trypticase soy broth cultures of Escherichia coli, Staphylococcus aureus, Mycobacterium smegmatis, and Pseudomonas aeruginosa.
2. Media:
   Two Trypticase soy agar plates.
3. Equipment:
   – Bunsen burner
   – inoculating loop
   – glassware marking pencil.

Procedure  for determination of antimicrobial spectrum of isolates.

• Label the Trypticase soy agar plates with the soil sample source of the isolate.
• Using aseptic technique, make a single-line streak inoculation of each isolate on the surface of an agar plate so as to divide the plate in half
• Incubate the plates in an inverted position for 3 to 5 days at 25°C.
• Following incubation, on the bottom of each plate draw four lines perpendicular to the growth of the antibiotic-producing isolate
• Aseptically make a single-line streak inoculation of each of the four test cultures following the inoculation template on each plate. Start close to, but not touching, the growth of the antibiotic-producing isolate and streak toward the edge of the plate.
• Incubate the plates in an inverted position for 24 hours at 37°C.
• Examine all plates for inhibition of test organisms, and record your observations in the Lab Report.

Observations and Results interpretations:

• I: Isolation of Antibiotic-producing micro-organisms.
  -Number of colonies showing zone of inhibition in different serial dilutions were noted and were further cultured to obtain pure cultures.
• II: Determination of anti-microbial spectrum of isolates.
  • Draw a representation of the observed antibiotic activity against the test organisms.
  • Based on your observations, record in the chart the presence (+) or absence (−) of antibiotic activity against each of the test organisms and the spectrum of antimicrobial activity (broad or narrow).

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Principle:

• Members of the anaerobic genera Clostridium and Desulfotomaculum and the aerobic genus Bacillus are examples of organisms that have the capacity to exist either as metabolically active vegetative cells or as highly resistant, metabolically inactive cell types called spores. When environmental conditions become unfavorable for continuing vegetative cellular activities, particularly with the exhaustion of a nutritional carbon source, these cells have the capacity to undergo sporogenesis and give rise to a new intracellular structure called the endospore, which is surrounded by impervious layers called spore coats. As conditions continue to worsen, the endospore is released from the degenerating vegetative cell and becomes an independent cell called a free spore. Because of the chemical composition of spore layers, the spore is resistant to the damaging effects of excessive heat, freezing, radiation, desiccation, and chemical agents, as well as to the commonly employed microbiological stains. With the return of favorable environmental conditions, the free spore may revert to a metabolically active
  and less resistant vegetative cell through germination. It should be emphasized that sporogenesis and germination are not means of reproduction but merely mechanisms that ensure cell survival under all environmental conditions.

In practice, the spore stain uses two different stains and decolorizing agents:

1. Primary Stain (Malachite Green):

• Unlike most vegetative cell types that stain by common procedures, the free spore, because of its impervious coats, will not accept the primary stain easily. For further penetration, the application of heat is required. After the primary stain is applied and the smear is heated, both the vegetative cell and spore will appear green.

2. Decolorizing Agent (Water):

• Once the spore accepts the malachite green, it cannot be decolorized by tap water, which removes only the excess primary stain. The spore remains green. On the other hand, the stain does not demonstrate a strong affinity for vegetative cell components; the water removes it, and these cells will be colorless.

3. Counter stain (Safranin):

• This contrasting red stain is used as the second reagent to color the decolorized vegetative cells, which will absorb the counterstain and appear red. The spores retain the green of the primary stain.

Requirements

• i). Bacterial Culture: 48-72 hrs nutrient agar slant culture of Bacillus cereus and thioglycollate culture of Clostridium sporogenes.
• ii. Reagents: Malachite green and safranin.
• iii. Equipment: Microincinerator or Bunsen burnerhot plate, staining tray, inoculating loop, glass slides, bibulous paper, lens paper, and microscope.

Procedure for Spore staining:

1. Obtain two clean glass slides.
2. Make individual smears in the usual manner using aseptic technique.
3. Allow smear to air-dry, and heat fix in the usual manner.
4. Flood smears with malachite green and place on top of a beaker of water sitting on a warm hot plate, allowing the preparation to steam for 2 to 3 minutes. Note: Do not allow stain to evaporate; replenish stain as needed. Prevent the stain from boiling by adjusting the hot
   plate temperature.
5. Remove slides from hot plate, cool, and wash under running tap water.
6. Counterstain with safranin for 30 seconds.
7. Wash with tap water.
8. Blot dry with bibulous paper and examine under oil immersion.
9. In the chart provided in the Lab Report, complete the following:
   • Draw a representative microscopic field of each preparation.
   • Describe the location of the endospore within the vegetative cell as central, sub-terminal, or terminal on each preparation.
   • Indicate the color of the spore and vegetative cell on each preparation

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• It is possible to describe a cell culture contaminant as any elements in the culture system that are unacceptable because of their possible adverse effects on either the system or its usage.
• Contamination cannot be entirely removed, but both the level of occurrence and the severity of its effects can be controlled to decrease.

Consequences of contamination in cell culture lab:

• Loss of time, cash and commitment.
• Unfavorable impacts on cultures.
• Inaccurate or false scientific observations.
• Personal awkwardness.
• Loss of valuable items

Contamination issues in cell culture:

• The contamination issues in cell culture can be classified into three classes:

1. Minor annoyances: When several plates and flasks are lost to contamination.
2. Severe problems: When the frequency of contamination rises and whole experiments or cell cultures are lost.
3.  Major disasters: Contaminants that are usually other cell lines or mycoplasma that bring into question the validity of your past and present work

Types of Contamination:

1. Biological contamination
2. Chemical contamination

I. Chemical contamination:

Types and Sources of chemical contamination:

• The involvement of any nonliving material that results in adverse effects on the culture system is best explained as chemical contamination.
• Media: Most chemical contaminants are present in the media for cell culture and come either from the reagents and water used to make them, or from the additives used to supplement them, such as sera.
• Sera: Cause of both biological and chemical contaminants, variance of hormone and growth factors.
• Water: A common cause of chemical contamination is water used for the manufacture of media and cleaning glassware and needs careful precautions to ensure its quality.
• Endotoxins: lipopolysaccharide-containing gram-negative bacteria by-products typically present in water, serums, and some culture additives (especially those manufactured using microbial fermentation)
• Storage vessels: Another cause of contaminants may be media stored in glass or plastic bottles that have already contained heavy metal solutions or organic compounds, such as electron microscopy stains, solvents, and pesticides.
  • – During storage of the initial solution, the contaminants can be adsorbed on the surface of the container or its cap (or ingested into the bottle if it is plastic).
  • Fluorescent lights:
  • – the application of media containing HEPES (N-[2-hydroxyl ethyl] piperazine-N’-[2-ethanesulfonic acid])-an organic buffer widely used to supplement bicarbonate-based buffers), riboflavin or tryptophan to regular fluorescent lighting is a significant yet sometimes underestimated cause of chemical contamination.- These media components are likely to be photoactivated resulting hydrogen peroxides and free radicals that are harmful to cells, the longer the exposure, the more toxic it is.
  • Incubators:
  • – The incubator may also be a source of chemical contamination, often viewed as a significant source of biological contamination.
  • – Gas mixture perfused through certain incubators (usually including carbon dioxide to help control media H) which contain toxic contaminants, especially oils or other gases such as carbon monoxide that may have previously been used in the storage cylinder or tank.

II. Biological contamination:

• Centered on the difficulties of detecting them in cultures, biological contaminants can be subdivided into two groups:
• 1. Those that are easy to detect: Bacteria, molds and yeast.
• 2. Those which are more difficult to detect and cause more severe problems in culture: Viruses, protozoa, insects, mycoplasmas, and cross-contamination from other cell lines.

Sources of biological contamination:

• Accidents and failures.
• Contact with nonsterile supplies, media or solution.
• Particulate or aerosol fallout during manipulation, transportation, or incubation.
• Swimming, crawling, or growing into culture vessels

1. Bacteria, yeasts and molds:

• Found almost anywhere and able to colonize and grow rapidly in the environment provided by cell culture.
• In the absence of antibiotics, microbes can be identified either by direct microscopic analysis or by their effects on culture in a culture within a few days (pH shifts, turbidity and cell destruction).
• Resistant species may establish low-level infections that are very difficult to detect by direct visual examination when antibiotics are regularly used.
• a. Bacterial contaminants:
  • -common in nature
  • – maybe mistaken as cellular debris, usually at lower levels.
  • – Search for:
    • Signs of motility
    • acidic pH
    • Size and shape uniformity
• b. Fungal and yeast contaminants:
  • – also common\- formation of clumps or mats
  • – Easier identification because of large size .
  • – Fungi will form small colonies on medium initially.
  • – Yeast will normally show signs of budding.

2. Viruses:

• Viruses are the most challenging cell culture pathogens to detect in culture due to their small size.
• Their small size also makes it difficult to extract them from biological media, solutions, and other solutions.
• While viruses may be more widespread in cell cultures than many researchers know, unless they have cytopathic or other adverse effects on cultures, they are typically not considered a serious issue.
• Their effect on cultures is not a major concern for the use of virally infected cultures, but they pose possible health risks for laboratory staff.
• Viral contaminants:
  • – Not very common:
  • identified by their adverse affects on  cultures
  • Many unknown consequences are wrongly blamed on viruses
  • Fetal bovine sera consists of bovine viruses
  • – No reliable way to completely eliminate viral contamination
  • -Detection:
    • Immunostaining
    • ELISA
    • PCR

3. Protozoa:

• Single-celled protozoa, such as amoebas, have sometimes been classified as cell culture contaminants, both parasitic and free-living.
• Amebas, typically of soil origin, can form spores and are readily isolated from the air, sometimes from tissues, as well as from laboratory personnel’s throat and nose swabs.
• Cytopathic symptoms similar to viral damage can be induced and a culture can be completely killed within ten days.
• Amoebas are very hard to detect in culture due to their sluggish growth and morphological similarity to cultured cells.
• Fortunately, contaminants of this kind are uncommon, but it is important to be aware of the likelihood of their occurrence.

4. Invertebrates:

• Insects and arachnids are commonly found in laboratory environments, and both cultures and sterile supplies may be infected by flies, ants, cockroaches and mites.

5. Mycoplasmas:

• Mycoplasmas were first observed by Robinson and coworkers in 1956 in cell cultures.
• They attempted to research the effects of PPLO (pleuropneumonia-like species, the original name for mycoplasma) on HeLa cells, when they found that the control HeLa cultures were already contaminated by PPLO.
• The most popular form of contaminant in today’s cell culture!
• There are approximately 180 different species.
• The most popular transmission method is from other cultures that have been contaminated.
• The smallest (0.2-0.3μm) free living organisms.
• No cell wall; it cannot be seen under phase contrast microscopy.
• Effects of Mycoplasma:
  • Effects on almost every aspect of cell behavior & growth, including microarrays.
  • A medium of up to 108 mycoplasma/mL without turbidity
  • Intervention with screening assays.
  • This triggers chromosome breakage.
  • This leads to inaccurate or incorrect results.

How to reduce contamination of cultures ?

• The basic step is autoclaving, it reduces the contamination.
• Commercial cleanup kits: It can help with temporary reduction of contamination between detectable levels. It may alter characteristics or kill cells.
• In case of detection of mycoplasma:
  • – Tylosin
  • – BM Cycline 
  • MRA; mycoplasma removal  agent 
  • – Ciprofloxacin
  • Non- Antibiotic: Mynox Mycoplasma Elimination  Reagent 

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• Lactose is a disaccharide of glucose and galactose which is connected by a β-galactoside bond.
• Lactose fermenting bacteria consist of both the permease and beta-galactosidase enzymes that are responsible for acid production during lactose fermentation.
• Permease permits the lactose to enter into the bacterial cell wall.
• Then, beta-galactosidase breaks down the lactose into glucose and galactose.
• But in some organisms, permease is absent and hence appear as late or non-lactose fermenters.
• O-Nitrophenyl-β-D-galactopyranoside (ONPG) resembles to lactose in structure, except that glucose is substituted by orthonitrophenyl group.
• In contrast to lactose, the substrate O-nitrophenyl-beta-D galactopyranoside (ONPG) can penetrate the bacterial cell wall even in absence of permease.
• ONPG test is a very sensitive test for lactose-fermentation.
• In this test, O-Nitrophenyl-β-D-galactopyranoside (ONPG) (artificial in nature) acts as a substrate for beta-galactosidase to ascertain the specific enzyme activity that serves in the identification and differentiation of organisms.
• If the organism consists of beta-galactosidase, then it will hydrolyse ONPG to yield galactose and o-nitrophenol.
• The positive test is confirmed by the yellow colour, which is given by ortho-nitrophenol.
• A positive ONPG test is mainly focused on providing the rapid identification of delayed lactose fermentation.

Requirements:

1. Sodium phosphate buffer, 1 M, pH 7.0
2. O-Nitrophenyl-β-D-galactopyranoside (ONPG), 0.75 M
3. Physiologic saline
4. Toluene
5. ONPG broth: Ingredients per 1000 mL
   • Na2HPO4 —————- (9.46 g)
   • phenylalanine ——— (4 g)
   • ONPG ———————– (2 g)
   • KH2PO4 ——————–(0.907 g)
   • pH 8.0
6. ONPG disk: 
   • ONPG Differentiation Disk is made by impregnating controlled concentrations of ONPG onto a 0.25 inch diameter filter paper disk.

Procedure of ONPG test:

• For ONPG disk method:
  • Keep an ONPG disk into a sterile tube.
  • Add 0.2 mL saline to it.
  • Use a loopful of test isolate to heavily inoculate the tube.
  • Incubate at 35-37°C for up to 4 hours.
  • Observe for colour change of the disk.
• For broth method:
  • Keep the test medium to room temperature.
  • Use heavy inoculum from a pure 18-24 hr culture to inoculate the test medium.
  • Incubate aerobically, at 35- 37ºC, with caps loosened.
  • Observe for a yellow colour development at 1 hour.
  • If no colour change is observed in the tube even after 1 hour of incubation, continue incubation for up to 24 hours.

Results interpretations:

• Positive result:
  • The positive test is indicated by the yellow colour of ortho-nitrophenol resulted by the hydrolysis of ONPG.
• Negative result:
  • The negative test is suggested by no change in colour, that means the absence of enzyme.

Limitations of ONPG test:

• Further biochemical, molecular, immunological, or mass spectrometry testing on colonies from pure culture is advised for complete identification.
• Cultures which naturally yield yellow colour cannot be tested in this media.
• Inoculation of test organism should be done only from lactose- containing medium.

O-Nitrophenyl-β-D-Galactopyranoside (ONPG) test: Principle, Procedure and Results

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• L-Pyrrolidonyl Arylamidase (PYR) test is one of the rapid tests employed for the identification of group a beta-hemolytic Streptococci and Enterococci on the basis of activity of the enzyme pyrolidonyl arylamidase.
• It is also termed as pyrrolidonyl aminopeptidase.
•  PYR is a bacterial enzyme
• The test disk is impregnated with L-pyroglutamic acid-β-naphthylamide that is hydrolysed by PYR hence serves as a substrate for the detection of PYR
• Hydrolysis of the L-pyroglutamic acid-β-naphthylamide yields beta-naphthylamine which in combination with the PYR Reagent (p-dimethylamino-cinnamaldehyde)  forms a bright pink to cherry red colour.
• Either broth assay method or rapid disk method is used for the test.

Procedure:

• Broth Method
  • Inoculate PYR broth with 2-5 colonies from overnight (18-24 hours) pure culture.
  • Incubate the tube aerobically at 35-37°C for 4 hours.
  • Add 2-3 drop of PYR reagent and observe for colour change.
  • Observe for the red colour development within 1-2 minutes.
• Disk Method (Rapid)
  • With 10 µl sterile distilled water or deionized water, wet the PYR test disc on the strip.
  • Note: Do not flood the disk.
  • From 18-24 hrs culture, place 5-10 colonies of the tested strain on the surface of the disc with a loop and smear them lightly on it.
  • Incubate the disc for 1-2 minutes at room temperature.
  • Add 1 drop of N, N-dimethylaminocinnamaldehyde after incubation.
  • Observe for red colour development within 1-2 minutes.

Results interpretations:

• Positive test:
  • The positive test is indicated by the appearance of bright pink or cherry red color within 1-2 mins. Examples: Citrobacter, Klebsiella etc.
• Negative test:
  • The negative test is suggested by no any color change or a blue color because of indole reaction. Examples: Streptococcus bovis, S. equinus

Limitations:

• PYR is only for the presumptive identification of group A Streptococci and group D enterococci from other streptococci thus other tests are recommended for complete identification.
• If the disk or filter paper are too moist, a false-negative test can result.
• Few isolates of lactococci and aerococci maybe PYRase positive.
• If reactions are read after 20 seconds, non-specific color reactions may occur.

L-Pyrrolidonyl Arylamidase (PYR) test: Principle, Requirements, Procedure and Results

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• Decarboxylases are a group of enzymes which acts by hydrolyzing an amino acid to form an amine.
• Decarboxylase test is mainly performed to differentiate decarboxylase producing Enterobacteriaceae from other gram-negative rods.
• The decarboxylation of the amino acid yields in an alkaline pH and a change in color of pH indicators bromocresol and cresol red from orange to purple is observed.

Requirements:

1. Media: Brain-heart infusion broth, Blood agar, Decarboxylase broth
2. Test-tubes
3. Incubator
4. Sterile mineral oil

composition of Decarboxylase broth

1. Composition———————————–g/ltr
   • Peptic digest of animal tissue——– (5 g)
   • beef extract ———————————-(5 g)
   • bromocresol purple ———————-(0.1 g)
   • cresol red————————————- (0.005 g)
   • dextrose —————————————(0.5 g)
   • pyridoxal ————————————(0.005 g)
   • amino acid ———————————–(10 g)
2. pH-6.0

Procedure of Decarboxylase test:

1. Glucose-Nonfermenting Organisms:
   • Prepare a suspension (≥McFarland No. 5 turbidity standard) in brain-heart infusion broth from 18-24hr old culture on 5% sheep blood agar.
   • With 4 drops of broth, inoculate each of the three decarboxylase broths (arginine, lysine, and ornithine) and the control broth (no amino acid).
   • To each tube, add a 4-mm layer of sterile mineral oil.
   • Incubate the cultures at 35°-37°C in ambient air.
   • Examine the tubes at 24, 48, 72, and 96 hours.
2. Glucose-Fermenting Organisms:
   • With 1 drop of an 18- to 24-hour brain-heart infusion broth culture, inoculate each tubes.
   • To each tube, add a 4-mm layer of sterile mineral oil.
   • Incubate the cultures for 4 days at 35°-37°C in ambient air.
   • Observe and examine the tubes at 24, 48, 72, and 96 hours.

Results Interpretations:

• Positive test:
  • It is indicated by the alkaline pH and change in coloration from orange to purple.
• Negative test:
  • It is suggested by no change in colour in both test and control tubes and growth is visible in the control tube.

Decarboxylase test-Principle, Procedure and Results

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• Leucine Amino Peptidase (LAP) test is one of the rapid tests which is employed for the detection of the enzyme leucine amino peptidase.
• The LAP test is generally performed for the preliminary characterization of catalase- negative, gram-positive cocci, particularly non-beta hemolytic cocci.
• Leucine- -naphthalamide impregnated disk acts as a substrate.
• The LAP enzyme is responsible for hydrolysis of substrate that yields -naphthalamine.
• The product on addition to p-methyl (an aminocinnamaldehyde reagent) forms a highly visible red colored Schiff’s base.

Procedure:

1. Place a LAP disk in a sterile petri dish, and leave the disk to warm to room temperature.
2. Lowly dampen the LAP disk either with reagent grade water or with a little sterile distilled water.
3. Use a wooden applicator stick to rub a small amount of several colonies of an 18 to 24 hour pure culture onto a small area of the LAP disk.
4. Incubate at room temperature for 5 minutes.
5. Add 1 drop of cinnamaldehyde reagent after the incubation period, and read within one minute.

Results interpretation:

• Positive result:
  • The positive test is indicated by the development of a red/pink color.
• Negative result:
  • The negative test is suggested by no change or slight yellow color.

Limitations:

• The confirmation of test organism to be gram-positive coccus and catalase negative is mandatory for the LAP test.
• There might be chances of false negative if inadequate columns are taken.

Leucine Amino Peptidase (LAP) test

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• Microdase test is also termed as modified oxidase test.
• It is one of the rapid tests employed for the differentiation between Staphylococcus and Micrococcus which are Gram positive cocci having catalase enzyme.
• The test is based on the detection of oxidase enzyme.
• Filter paper disks impregnated with tetramethyl-p-phenylenediamine in dimethyl sulfoxide (DMSO) is used in order to detect the oxidase enzyme.
• 1% (w/v) tetramethyl-p-phenylenediamine in certified grade dimethyl sulfoxide is the preparation for modified oxidase reagent.
• DMSO serves for the permeability of cells to the reagent along with providing solubility and stability against auto-oxidation.
• When exposed to atmospheric oxygen, the oxidase enzyme reacts with the oxidase reagent and cytochrome C to yield the coloured compound, indophenol indicated as blue or purplish blue coloration on the disc after the bacterial colony is introduced on the disc.

Requirements:

1. Media: Blood agar
2. Oxidase Disc
3. Filter paper disks impregnated with tetramethyl-p-phenylenediamine dihydrochloride in DMSO.
4. Incubator
5. Forceps
6. Petri dish
7. Glass slide
8. Fresh pure colonies of test organisms

Procedure:

1. Use forceps to place the disk in an empty petri dish or on a clean glass slide.
2. Use a wooden applicator stick and rub a small amount of various colonies of an 18- to 24-hour pure culture grown on blood agar onto a small area of the microdase disk.
   Note: Do not rehydrate the disk before use.
3. Incubate at room temperature for 2 minutes.
4. Observe for a blue colour development.

Results interpretations:

• Positive result:
  • The positive test is suggested by the development of blue to purple colour within the 2 mins.
• Negative result:
  • The negative test is indicated by no any change in color.

Limitations:

• Neither too young nor too old cultures are used as they might yield inaccurate results.
• Microdase test is recommended only for the Gram-positive, catalase positive cocci.
• Microdase is limited only for the routine testing of oxidase activity of Staphylococcus and Micrococcus.

Microdase (Modified Oxidase) test: Principle, Procedure and Results interpretation

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• After E. coli, Staphylococcus saprophyticus is the second most causative organism for the Urinary tract infections (UTIs) in sexually active young women.
• Laboratory identification of Staphylococcus saprophyticus is performed on the basis of haemolysis, coagulase and resistance to novobiocin.
• Thus, after isolation of coagulase negative staphylococcus, laboratory should further identify the isolate and find out if the isolate is sensitive to novobiocin or not.
• Novobiocin is the antibiotic produced by the actinomycete Streptomyces nivens.
• Sensitivity to novobiocin is detected by placing a novobiocin impregnated paper disk on a agar plate seeded with the organism under identification.
• If the bacteria are sensitive to novobiocin, the formation of visible zone of inhibition  is seen around the disk.
• The zone of inhibition represents the area where the growth of organism was inhibited by the antibiotic concentration.
• No zone of inhibition suggests the resistance of organism to the antibiotic.

Requirements:

1. Isolated colonies of an aerobic, catalase positive, coagulase negative gram positive cocci.
2. Mueller Hinton agar
3. Blood agar/ Tryptic Soy agar
4. Incubator
5. Novobiocin disk
6. Distilled water
7. Sliding calipers/ metric ruler

Procedure of Novobiocin susceptibility test

1. The test isolate taken is of 18-72 hours and in pure culture.
2. Prepare a suspension of the test isolate in tryptic soy broth equal to a McFarland 0.5 standard or equivalent.
3. Dip a sterile swab into the suspension and rotate it against the side of the tube above the fluid level in order to remove excess inoculum.
4. Inoculate a blood agar or Mueller Hinton agar plate by streaking the expressed swab over the entire agar surface and repeat in 2 planes.
5. Allow the agar surface to dry for exactly15 minutes before applying a Novobiocin Disk.
6. Prepare a lawn of growth over the entire plate by use of sterile swab, swabbing over the entire plate in 3 directions and around the edge of the plate.
7. Using alcohol-dipped and flamed forceps, aseptically apply a novobiocin antibiotic disc to the surface of each inoculated plate.
8. Use sterile forceps to gently press the discs down to make sure that they adhere to the agar surface.
9. Incubate plate aerobically for 18 to 24 hours at 35 to 37°C.
10. Use sliding calipers or a metric ruler to measure the diameter of the zone of inhibition.

Results interpretation:

• Positive result:
  • The positive test is indicated by the zone of inhibition greater than 16mm suggesting the sensitivity of the organism.
• Negative result:
  • Negative test is indicated by the zone of inhibition less than 16mm which suggests the novobiocin resistance of the organism.

Limitations:

• Only isolated colonies of aerobic, catalase positive, coagulase negative gram positive cocci are to be tested.
• Biochemical, immunological tests are further recommended for the complete identification of the organism.
• Novobiocin disks can mislead the results if the test is performed for isolates other than urinary specimens.

Novobiocin susceptibility test: Principle, Procedure and Results interpretations

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• Acetate agar is employed to test an organism’s ability to utilize acetate. The medium consists of sodium acetate as the sole carbon source and inorganic ammonium salts as the sole source of nitrogen. Growth of organisms suggests the positive test for acetate utilization. During the metabolism of acetate by the bacteria , the ammonium salts are broken down to ammonia, which elevates alkalinity. The shift in pH turns the bromothymol blue indicator in the medium from green to blue. This medium is usually used for differentiating Shigella spp from Escherichia coli. Shigella spp are not able to metabolize actetate whereas approximately 94% Escherichia coli utilize acetate.

Requirements:

1. Media: Composition of Sodium acetate agar
   • Ingredients                                         gm/l
   • sodium chloride …………………………………….5.0 g
   • magnesium sulfate ………………………………..0.1 g
   • ammonium phosphate, monobasic …….1.0 g
   • potassium phosphate, dibasic …………….1.0 g
   • sodium acetate …………………………………….2.0 g
   • agar ………………………..………………………….20.0 g
   • bromothymol blue ……………………………. 0.08g
2. Sterile inoculating loops or sticks
3. Sterile pipette
4.  Incubator at 35°C
5.  Sterile saline

Procedure of Acetate utilization test:

•  Using an 18- to 24-h culture from a non-inhibitory culture plate, prepare a turbid saline suspension.
• Inoculate the slant with 1 drop of the suspension.
•  Alternatively, streak the slant back and forth with a light inoculum picked from the centre of a well-isolated colony.
• Place cap loosely on tube.
•  Incubate aerobically at 35 to 37°C for up to 5 days for Enterobacteriaceae; incubate at 30􏰀C for nonfermenting, gram-negative rods for up to 7 days.
• Observe a colour change from green to blue along the slant.

Results interpretations:

• Positive test:
  • It is indicated by the growth of organisms and conversion of colour from green to intense blue along the slant.
• Negative test:
  • No colour change indicates no growth and is suggestive of negative test.

Limitations:

• Stab on slant should be avoided as the test requires aerobic environment.
• Inoculation from broth cultures is avoided due to carry over of media.
• Light inoculum is preferred to avoid false positive results.

Acetate utilization test: Principle, media composition, Procedure and Results

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