Microbiology

Capsule Staining

Some bacterial cells are surrounded by an extracellular slime layer called a capsule or glycocalyx. The capsule stain is a differential stain which selectively stains external capsules surrounding bacterial cells. The polymers which make up the capsule tend to be uncharged and they are not easily stained hence direct staining methods cannot be utilized.

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Negative Staining

Method

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  • Prepare a clean, grease-free slide; a small drop of nigrosine is mixed with a small drop from a broth culture or with a quantity of dry material.
  • The drop is spread across the slide using the edge of another slide as a spreader. Same procedure that is used for preparing blood smears. Allow the smear to air dry, don???t heat fix.
  • After air drying, the smear is observed using the high dry lens, or oil immersion if desired.
  • The smear will be denser where the nigrosine dye is deposited on the slide. The background should be blue-gray. Bacteria will be evident by the absence of any color.

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Anthony???s Capsule Staining

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Method

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  • Prepare a smear from a 12- to 18-hour culture grown in milk broth or litmus milk. This is to provide a proteinaceous background for contrast.

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  • Allow the smear to air dry. Do not heat fix because capsule is readily destroyed by heating
  • Cover the slide with 1% crystal violet for 2 minutes.

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  • Rinse gently with a 20% solution of copper sulfate.

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  • Air-dry the slide. Do not blot as blotting will remove the un-heat-fixed bacteria from the slide and/or cause disruption of the capsule.

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  • Examine the slide under an oil immersion lens. Bacterial cells and the proteinaceous background will appear purplish while the capsules will appear transparent.

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Maneval???s Capsule Staining

  • A few drops of Congo red are placed on a clean slide.
  • Mix a small amount of culture into it.
  • Air-dry the slide. Do not heat fix as it will destroy the protein capsules on the surface of the bacteria
  • Place the slide on a rack of the staining tray. Gently flood the smear with the Maneval solution and wait 5 minutes.
  • Lift the slide and gently discard excess stain.
  • Rinse with water with great care. As the sample is not heat fixed to the slide it could be washed off.
  • Air-dry the slide placing it on an absorbent paper. Do not blot
  • Examine the slide with an oil immersion lens at 1,000X magnification.

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Gin Staining

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  • Use a loop to mix a drop of water, a drop of india ink and a small amount of bacterial culture together at the end of a slide.
  • Use another slide to spread the smear like a blood smear. Allow the smear to air dry and do not heat fix.
  • Flood the smear with crystal violet, 1 minute. Wash with water, blot, dry, and observe.

Capsule Staining Anthonys Stain Method

Capsules are composed primarily of polysaccharides or glycoproteins and are gelatinous in texture. They are readily destroyed by heating and hence direct staining methods cannot be utilized.

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Anthony???s Capsule Staining

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Method

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  • Prepare a smear from a 12- to 18-hour culture grown in milk broth or litmus milk. This is to provide a proteinaceous background for contrast.

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  • Allow the smear to air dry. Do not heat fix because capsule is readily destroyed by heating
  • Cover the slide with 1% crystal violet for 2 minutes.

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  • Rinse gently with a 20% solution of copper sulfate.

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  • Air-dry the slide. Do not blot as blotting will remove the un-heat-fixed bacteria from the slide and/or cause disruption of the capsule.

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  • Examine the slide under an oil immersion lens. Bacterial cells and the proteinaceous background will appear purplish while the capsules will appear transparent.

Capsule Staining Negative Staining

Most bacteria have some kind of capsule. Most bacterial capsules are composed of polysaccharide however some genera produce polypeptide capsules. The polymers which make up the capsule tend to be uncharged and as such they are not easily stained. For this reason we use a negative stain to visualize them.

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Negative Staining

Method

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  • Prepare a clean, grease-free slide; a small drop of nigrosine is mixed with a small drop from a broth culture or with a quantity of dry material.
  • The drop is spread across the slide using the edge of another slide as a spreader. Same procedure that is used for preparing blood smears. Allow the smear to air dry, don???t heat fix.
  • After air drying, the smear is observed using the high dry lens, or oil immersion if desired.
  • The smear will be denser where the nigrosine dye is deposited on the slide. The background should be blue-gray. Bacteria will be evident by the absence of any color.

Endospore Staining

Endospore production is a very important characteristic of some bacteria, allowing them to resist adverse environmental conditions such as desiccation, chemical exposure, extreme heat, radiation, etc. Gram staining and simple staining techniques may or may not reveal the presence of endospores in a bacterial sample.??

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Staining solutions

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Malachite green stain (0.5% (wt/vol) aqueous solution)

0.5 g of malachite green

100 ml of distilled water

Safranin counterstain

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Stock solution (2.5% (wt/vol) alcoholic solution)

2.5 g of Safranin O

100 ml of 95% ethanol

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Working solution of Safranin

10 ml of stock solution

90 ml of distilled water

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Method

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  • Smear the organism and heat fix to a slide
  • Place the slide over a steam bath and cover with Malachite Green
  • Keep the stain over the bath for 3 – 5 minutes, recovering the slide with Malachite Green if some evaporates.
  • Pour the Malachite Green off and allow it to cool
  • Rinse the slide with water to remove excess stain
  • Cover the smear with Safranin for two minutes
  • Rinse the slide with water to remove excess stain
  • Blot dry the stain and view under a microscope.
  • Once completely dried, the vegetative cells are red/pink and the spores are green.

Acid Fast Staining

The acid-fast stain is performed on samples to demonstrate the characteristic of acid fastness in certain bacteria and the cysts of Cryptosporidium and Isospora. Clinically, the most important application is to detect??Mycobacterium tuberculosis??in sputum samples to confirm or rule out a diagnosis of tuberculosis in patients.

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Ziehl-Neelsen method for acid-fast staining

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Staining solution:

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Stock Solution A??(stable for 6 months)

????????????????????????????????????????????????Ingredients?????????????????????????????????????????????? ??????????????????????????????????????????????Amount

?????????????????????????????????????????????? L.O.C. High Suds (Amway) ?????????????????? ?? ?? ?? ?? ?? ?? ?? ????0.6 ml

?????????????????????????????????????????????? Distilled water ??????????????????????????????????????????????????????????????????????????????????????100 ml

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Stock Solution B

???????????? ??????????????????????????????????Ingredients???? ?????????????????????????????????????????????????????????????????? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ??Amount

?????????????????????????????????????????????? Basic fuchsin?? ??????????????????????????????????????????????????????????????????????????????????????1 g

?????????????????????????????????????????????? Absolute ethyl alcohol?????????????????????????????????? ????????????????????????????10 ml

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?????????????? The two solutions can be kept as stock solution and mixed before use.

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Working Solution??(stable for 1 month)

?????????????????????????????????????????????? Mix 50ml of A with 5 ml of B.

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Acid alcohol??- 3% hydrochloric acid in 95% ethyl alcohol

????????????????????????????????????????????????Ingredients???????????????????????????????????????? ???????????????????????????????????????????? ?? ?? ?? ?? ?? ?? ?? ??Amount????????

?????????????????????????????????????????????? Absolute ethyl alcohol???????????????????????????????????????????????? ????????????95ml

?????????????????????????????????????????????? Distilled water???????????????????????????????????????????????????????????????????????? ??????????2 ml

?????????????????????????????????????????????? Concentrated hydrochloric acid?? ???????????????? ?? ?? ?? ????3 ml

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Make up the alcohol solution then add the concentrated acid. Use extreme care when handling concentrated acid.

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0.25% methylene blue in 1% acetic acid

?????????????????????????????????????????????? Ingredients???? ???????????????????????????????????????????????????????????????????? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ?? ????Amount

?????????????????????????????????????????????? Methylene blue ????????????????????????????????????????????????????????????????????????????????????0.25 g

?????????????????????????????????????????????? Distilled water ????????????????????????????????????????????????????????????????????????????????????????99 ml

?????????????????????????????????????????????? Acetic acid ??????????????????????????????????????????????????????????????????????????????????????????????????1 ml

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Method

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  • Place a strip of blotting paper over the slide.
  • Place the covered slide over a screened water bath and then saturate blotting paper with primary stain Ziehl???s carbol fuchsin.
  • Allow the slide to sit over water bath for 3 ??? 5 minutes, reapplying stain if it begins to dry out.
  • Remove blotting paper and rinse slide until water runs clear.
  • Flood slide with decolorizer, Acid Alcohol, for 10 ??? 15 seconds and then rinse.
  • Flood slide with counterstain, Crystal Violet, for one minute and then rinse.
  • Gently blot the slide dry. It is now ready to be viewed under oil immersion (1000x TM) with a bright-field compound microscope.
  • Bacteria described as acid fast will appear red while examining specimens using bright-field microscopy. Non-acid-fast cells and field debris will appear blue.

Spread Plate Technique

A pure culture theoretically contains a single bacterial species. One of the most important and simpler methods for isolation of a pure culture is spread plate technique.

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Spread plate technique

  • Select and prepare an agar medium based upon the type of bacteria to be enumerated or selected.??
  • After autoclaving, cool the agar to between 45 to 50??C prior pouring into the plates.
  • The thickness of the agar should be roughly 0.3 cm, which can be achieved by pouring??15 to 20 ml of media per 100 x 15 mm plate.??
  • Freshly prepared plates don???t work as well as dry plates as it takes longer for the inoculum to absorb into the agar.
  • Plates may be dried by keeping them at room temperature for roughly 24 hours.
  • A reusable glass or metal spreader should be flame sterilized by dipping in alcohol (such as 70% isopropyl or ethanol), shaking off the excess alcohol, and igniting the residue. The spreader is then allowed to cool.
  • Transfer 0.1 mm of culture on the plate with agar. The spreader is placed in contact with the inoculum on the surface of the plate and positioned to allow the inoculum to run evenly along the length of the spreader.
  • Even pressure is applied to the spreader and the plate is spun, on a turntable or by hand. Alternatively the spreader may be rotated over the agar surface as well.
  • Since some bacteria rapidly attach to the agar surface, the inoculum should be spread soon after it is applied.??
  • Avoid spreading the inoculum all the way to the edge of the agar.??
  • The goal is to evenly distribute the inoculum and to allow it to be absorbed into the agar.
  • Avoid disturbing plates for 10 to 20 minutes after spreading. Drying time varies with the room temperature and humidity.??
  • The plates are inverted and incubated at 37??C for 24 hours to 48 hours.
  • Observe the plates before the colonies have had time to fully develop. Continue the incubation as necessary.??

Incubation in closed humidified containers will help avoid problems with plates drying out when working with slow-growing colonies.

Pour Plate Technique

A pure culture theoretically contains a single bacterial species. One of the most important and simpler methods for isolation of a pure culture is pour plate technique.

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Pour plate technique

  • Label the bottom of the sterile Petri dish with the source of the culture and turn the plate
  • Obtain two tubes of liquefied agar by heating the nutrient agar.
  • Agar is cooled to 60??C and held in the water bath to maintain the temperature as higher temperature would kill the bacteria when introduced into it.
  • Agar should not be cooled for a long time as it would solidify at 42??C. Inoculum from the mixed bacterial culture is aseptically transferred into the tubes with agar with the help of a sterile inoculation loop.
  • Mix the tube by rolling the tube between your hands. Pour the inoculated liquid agar into the sterile Petri dish that is labeled initially and cover the dish.
  • Allow the agar to solidify for 5 minutes or so, turn the dish upside down once the agar is solidified.
  • Incubate the plate at 37 ??C for 24 to 48 hours.
  • Visible growth of colonies of organisms can be seen on the plate from which desired organisms can be isolated and maintained.

Pure Culture Techniques

A pure culture theoretically contains a single bacterial species. Simpler methods for isolation of a pure culture include: (i) Pour plating (ii) Spread plating (iii) Streak plating with a loop.

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Pour plate technique

  • Label the bottom of the sterile Petri dish with the source of the culture and turn the plate
  • Obtain two tubes of liquefied agar by heating the nutrient agar.
  • Agar is cooled to 60??C and held in the water bath to maintain the temperature as higher temperature would kill the bacteria when introduced into it.
  • Agar should not be cooled for a long time as it would solidify at 42??C. Inoculum from the mixed bacterial culture is aseptically transferred into the tubes with agar with the help of a sterile inoculation loop.
  • Mix the tube by rolling the tube between your hands. Pour the inoculated liquid agar into the sterile Petri dish that is labeled initially and cover the dish.
  • Allow the agar to solidify for 5 minutes or so, turn the dish upside down once the agar is solidified.
  • Incubate the plate at 37 ??C for 24 to 48 hours.
  • Visible growth of colonies of organisms can be seen on the plate from which desired organisms can be isolated and maintained.

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Spread plate technique

  • Select and prepare an agar medium based upon the type of bacteria to be enumerated or selected.??
  • After autoclaving, cool the agar to between 45 to 50??C prior pouring into the plates.
  • The thickness of the agar should be roughly 0.3 cm, which can be achieved by pouring??15 to 20 ml of media per 100 x 15 mm plate.??
  • Freshly prepared plates don’t work as well as dry plates as it takes longer for the inoculum to absorb into the agar.
  • Plates may be dried by keeping them at room temperature for roughly 24 hours.
  • A reusable glass or metal spreader should be flame sterilized by dipping in alcohol (such as 70% isopropyl or ethanol), shaking off the excess alcohol, and igniting the residue. The spreader is then allowed to cool.
  • Transfer 0.1 mm of culture on the plate with agar. The spreader is placed in contact with the inoculum on the surface of the plate and positioned to allow the inoculum to run evenly along the length of the spreader.
  • Even pressure is applied to the spreader and the plate is spun, on a turntable or by hand. Alternatively the spreader may be rotated over the agar surface as well.
  • Since some bacteria rapidly attach to the agar surface, the inoculum should be spread soon after it is applied.??
  • Avoid spreading the inoculum all the way to the edge of the agar.??
  • The goal is to evenly distribute the inoculum and to allow it to be absorbed into the agar.
  • Avoid disturbing plates for 10 to 20 minutes after spreading. Drying time varies with the room temperature and humidity.??
  • The plates are inverted and incubated at 37??C for 24 hours to 48 hours.
  • Observe the plates before the colonies have had time to fully develop. Continue the incubation as necessary.??
  • Incubation in closed humidified containers will help avoid problems with plates drying out when working with slow-growing colonies.??

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Streak plate technique

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??????????????????Prepare lab bench by removing extraneous items and cleaning surface with table disinfectant.

??????????????????Label the bottom surface of the sterile agar plates.

??????????????????Use the T streak or quadrant streak or continuous streak, whichever is required.

??????????????????Obtain mix culture and shake gently to suspend organisms.??

??????????????????Perform desired streaking method to obtain pure culture of organisms after incubation.

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T-streak

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  • Outline the sections on the bottom of the agar plate in the shape of a ‘T’. Do the initial inoculation then flame the loop, let it cool for about 15 seconds.
  • Use the sterile cooled loop and draw over the agar surface in the first section around the sleeping line of T and flame and cool.
  • Use the sterile cooled loop and draw on the agar surface in the second section.
  • Go back and forth touching the first section few times and rest without touching them. Flame the loop and allow it to cool.
  • Use the sterile cooled loop and draw on the agar surface in the third section.
  • Again go back and forth touching the second section alone few times and rest without touching them. Now flame the loop and set aside.
  • Place the plate in the incubator, inverted at 30?? C for 2 days.

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Quadrant streak

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  • Prepare agar plates based on the type of bacteria to be enumerated or selected.??
  • Flame the loop and cool it for 5 seconds by touching an unused part of the agar surface close to the periphery of the plate, and then drag it rapidly several times across the surface of the first quadrant. Remove the loop and close the Petri dish.
  • Reflame and cool the loop, and turn the Petri dish to 90?? then touch the loop to a corner of the culture in the first quadrant and drag it several times across the agar in second quadrant, hitting the original streak a few times. The loop should never enter quadrant one again. Remove the loop and close the Petri dish.
  • Reflame and cool the loop and again turn the dish to 90?? anticlockwise. Streak third quadrant in the same manner as quadrant two, hitting last area several times. Remove the loop and close the Petri dish.
  • Flame the loop, again turn the dish to 90?? and then drag the culture from a corner of a third quadrant across the forth quadrant, contacting quadrant three several times and drag out the culture using a wider streak in a zigzag fashion.
  • Do not let the loop touch any of the previously streaked areas. The flaming of the loop is to effect the dilution of the culture so that fewer organisms are streaked in each area, resulting in the final desired separation. Remove the loop and close the Petri dish.
  • Incubate the plate in an inverted position in an incubator at 35??C for 24-48 hours.

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Continuous streak

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  • Mark the agar plate into two half-circles.??
  • Flame the loop and remove one colony from the mixed culture and streak one of the half-circle in a continuous back and forth line.??
  • Rotate the agar plate to 180?? and do the same thing in the second half circle without flaming the loop.??
  • Flame the loop and set aside. Cover the agar plate, flip it, and label it.
  • Place the inoculated agar plates in the incubator for 48 hours.

Hydrogen Sulphide Production Test

Hydrogen Sulphide Production testdetermines whether the microbe reduces sulfur-containing compounds to sulfides during the process of metabolism. Sulfide-Indole-Motility (SIM) medium is a nutrient medium allowing the detection of three different traits in bacteria which has

  • sulfates??to serve as the substrate for detecting??sulfide production
  • abundant tryptophan as a substrate for indole production
  • 0.5% agar is sufficient to allow bacterial motility, thereby allowing detection of motility.

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Sulfide-Indole-Motility (SIM) medium

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Formula / Liter

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Peptone?????????????????????? ???????????????????????????????????????????????? 30.0 g

Beef extract?????????? ?? ??????????????????????????????????????????????3.0 g

Ferrous ammonium sulfate???????????????? 0.2 g

Sodium thiosulfate???????????????? ?? ??????????????????????0.025 g

Agar???????? ?? ??????????????????????????????????????????????????????????????????????3.0 g

Final pH:?? 7.3 ?? 0.2 at 25??C

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Method

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  • Dissolve all the ingredients in one liter of purified water.
  • Heat the ingredients with frequent agitation to completely dissolve the medium. Autoclave at 121??C for 15 minutes at 15 lbs.
  • Prepare agar deeps with 5 ml of medium in sterile test tubes.
  • To test for motility, use a sterile inoculation needle to pick a well-isolated colony and stab the medium to within 1 cm of the bottom of the tube.
  • Be sure to keep the needle in the same line as it is inserted and??removed from the medium.
  • Incubate at 37??C for 18 hours or until growth is evident.
  • A positive motility test is indicated by a diffuse cloud of growth away from the line of inoculation.??
  • H2S production is shown by a blackening along the stab line.

Indole production is seen as the production of a red color after the addition of Kovac???s Reagent. Indole is produced from the tryptophan present in the medium.

Urease Test

The urease test identifies those organisms that are capable of hydrolyzing urea to produce ammonia and carbon dioxide. It is primarily used to distinguish urease-positive Protease??from other??Enterobacteriaceae.

Media Preparation

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Christensen???s urea agar is used to detect urease activity in a variety of microorganisms. Stuart???s urea broth is used primarily for the differentiation of??Proteus??species.

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Christensen???s Urea Agar

Ingredient?????????????????????????????????????????????????????????????????????????? Amount??

Peptone???????????????????????????????????????????????????????????????????????????????????? ????1g

Dextrose?????????????????????????????????????????????????????????????????????????????????? ????1g

Sodium chloride?????????????????????????????????????????????????????????????? 5 g

Potassium phosphate, monobasic ?????? 2 g

Urea?? ???? ????????????????????????????????????????????????????????????????????????????????????????????20 g

Phenol red?????????????? ?? ????????????????????????????????????????????????????????????????0.012 g

Agar ??????????????????????????????????????????????????????????????????????????????????????????????????15 to 20 g

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Preparation

  • To prepare the urea base, dissolve all the ingredients except agar in 100 ml of distilled water and filter sterilize (0.45-mm pore size).
  • Suspend the agar in 900 ml of distilled water, boil to dissolve completely, and autoclave at 121 degree C and 15 lbs for 15 minutes. Cool the agar to 50 to 55 degree C.
  • Aseptically add 100 ml of filter-sterilized urea base to the cooled agar solution and mix thoroughly. Distribute 4 to 5 ml per sterile test tube and slant the tubes during cooling until solidified. It is desirable to have a long slant and short butt. Prepared media will have a yellow-orange color.
  • Once prepared, do not reheat the medium as the urea will decompose. Inoculate the entire surface of the urea slant (slope) with the provided growth from the tryptic broth culture using the inoculating loop (do not stab the butt).
  • The slant of the medium is inoculated by streaking the surface of the agar in a zigzag manner. Incubate at 37??C for 24-48 hours.
  • After incubation observe the color change where the entire slant changes into pink color indicating positive result and yellow color indicating negative result, the control remains unchanged.

Stuart???s Urea Broth

Ingredient?????????????????????????????????????????????????????????????????????????? Amount??

Yeast extract?????? ?????????????????????????????????????????????????????????????????????? 0.1 g

Potassium phosphate, monobasic?????????????????? 9.1 g

Potassium phosphate, dibasic?????? ??????????????????????9.5 g

Urea???????? ?????????????????????????? ????????????????????????????????????????????????????????????????????20 g

Phenol red?????????????? ??????????????????????????????????????????????????????????????????????0.01 g

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Preparation

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  • Dissolve all ingredients in 1 liter of distilled water and filter sterilize (0.45-mm pore size).
  • Distribute 3 ml of prepared broth per sterile test tube. Prepared media will have a yellow-orange color. Do not heat the medium as the urea will decompose.
  • Inoculate the urea broth with the inoculation loop containing the organism from the tryptic soy broth culture.
  • Incubate for 24-48 hours at 37??C.
  • After incubation observe the color change where the entire broth changes into pink color indicating positive result and yellow color indicating negative result and the control remains unchanged .