Lab 4: Acid-Fast, Spores, and Capsule Stains - Biology

Lab 4: Acid-Fast, Spores, and Capsule Stains - Biology

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Acid-fast stain is a differential stain used to identify acid-fast organisms such as members of the genus Mycobacterium. Acid-fast organisms are characterized by wax-like, nearly impermeable cell walls; they contain mycolic acid and large amounts of fatty acids, waxes, and complex lipids. This type of cell wall is resistant to most compounds, therefore acid-fast organisms require a special staining technique.

The primary stain used in acid-fast staining, carbol fuchsin, is lipid-soluble and contains phenol, which helps the stain penetrate the cell wall. This is further assisted by the addition of heat in the form of heat (steam). Steam helps to loosen up the waxy layer and promotes entry of the primary stain inside the cell. The smear is then rinsed with a very strong decolorizer, which strips the stain from all non-acid-fast cells but does not permeate the cell wall of acid-fast organisms. The decolorized non-acid-fast cells then take up the counterstain, which in our case is methylene blue.

1. Working in pairs, prepare THREE slides as directed below (2 for back-up and one to stain).

2. Label each slide and draw a circle on the center of the slide with a wax pencil.

3. Prepare an emulsion on each slide with 4 loopfuls of Staphylococcus epidermidis from your broth culture onto the slide (these will be your acid-fast negative bacteria).

4. Then, add one loopful of Mycobacterium chelonae (these are your acid fast positive bacteria) and mix the two bacteria together.

5. Allow the slide(s) to air dry on the slide warmers (while these slides are drying, prepare your slides for the spore stain).

6. Once the liquid has completely evaporated, heat fix the bacteria by passing it through your flame three times.

7. Make sure the slide rack on top of your beaker is completely level. Then, bring your water to boil while the slides are drying.

You only need about 200 milliliters of water. If you add more, you will be waiting all lab period for your water to boil.

8. Once the water is boiling, place your slide on the slide rack above the boiling water.

9. Cover the area of your smear on the slide with a square piece of PRECUT paper towel. Make sure none of the paper is hanging off the slide.

10. Carefully apply the CARBOL FUCHSIN stain to the paper towel.

If a stain appears in the water you are boiling, please stop and discard the stained water in the liquid waste disposal. The fumes from carbol fuchsin can be toxic.

11. Steam with the stain on the slide for 7 minutes while continuously applying more stain so the paper square never dries out.

12. Gently remove the paper with forceps and discard it in the small waste paper cup that will be provided on your bench. Then, rinse the slide with water.

13. Put the slide on your staining basin and gently rinse with water.

14. Decolorize with 6 drops of acid alcohol (not ethanol from your gram stain kit), then rinse with water.

15. Counterstain with methylene blue for 2 minutes.

16. Rinse with water and blot dry with bibulous paper (do not use the slide warmer).

17. Examine under the 100X objective lens with oil immersion and record your results.

The colors of this image may be slightly off due to printing/copying.


The endospore stain is a differential stain used to visualize bacterial endospores. An endospore is a dormant form of a bacterium, which some species of bacteria produce under stressful conditions such as poor nutrition, high temperatures, or dry environments. The outer layer is composed of keratin which resists staining. The malachite-green stain is forced into the spore using steam. Spores can be central, terminal, and subterminal. This stain is commonly used to detect spores produced from the genera of Bacillus and Clostridium.

18. Prepare two slides with emulsions one from plate A and one from plate B.

19. Heat fix the slides and place them on the slide rack above the boiling water.

20. Cover the area of your smear on each slide with a square piece of PRECUT paper towel.

21. Carefully apply the MALACHITE GREEN stain to the paper towel.

22. Steam for 10 minutes and keep the paper soaked with the stain during this time.

23. Gently remove the paper with forceps, discard in the small waste paper cup that will be provided on your bench, and then rinse the slide with water.

24. Put the slides on your staining basin and gently rinse with water.

25. Counterstain with SAFRANIN for 1 minute and then rinse with water. Blot dry with bibulous paper.

26. Examine under the 100X objective lens with oil immersion and record your results.

The colors of this image may be slightly off due to printing/copying.


Most capsules are composed of polysaccharides or polypeptides which are a thick, detectable, and discrete layer outside the cell wall. Some capsules have well-defined boundaries while some have fuzzy, trailing edges. Capsules protect bacteria from the phagocytic action of immune cells and allow pathogens to invade the body. If a pathogen loses its ability to form capsules, it often ceases to be pathogenic.

27. Label and prepare a slide with an emulsion of Klebsiella pneumoniae.


29. Stain with 1% crystal violet for 2 minutes (DO NOT USE CRYSTAL VIOLET FROM GRAM STAIN).

30. VERY GENTLY rinse with 6 drops of copper sulfate.

31. Let it air dry on the counter (do not use slide warmer).

32. Examine under the 100X objective lens with oil immersion and record your results.

Capsule Staining- Principle, Reagents, Procedure and Result

The main purpose of capsule stain is to distinguish capsular material from the bacterial cell. A capsule is a gelatinous outer layer secreted by bacterial cell and that surrounds and adheres to the cell wall. Most capsules are composed of polysaccharides, but some are composed of polypeptides. The capsule differs from the slime layer that most bacterial cells produce in that it is a thick, detectable, discrete layer outside the cell wall. The capsule stain employs an acidic stain and a basic stain to detect capsule production.

Capsule Stain

Capsule stain is a type of differential stain which involves the use of two stains primary stain and the counterstain. Here, it's worth noting that for most part, capsules are non-ionic.

As a result, acidic and basic stains will often fail to adhere on to them. For this reason, it becomes necessary to stain the background using an acidic stain while the cell is stained using a basic stain.

Two of the most commonly used methods used in capsule staining include:


Not only are most bacteria very small, they are also very clear and difficult to view under a microscope without first staining. You must firmly attach your bacteria to a glass slide before you can stain them. There are two important things to consider when preparing a slide for staining:

  1. The bacteria must be evenly and lightly dispersed. If there are too many bacteria on the slide they will form a big glob and you will not be able to see the morphology of the individual cells. Large blobs of cells also do not stain properly and could yield erroneous results from the improper staining.
  2. The bacteria need to be firmly attached to the slide so they are not washed off during the staining procedures. All procedures that attach the bacteria to the slide result in some morphological changes. The cells typically shrink in size and will exhibit some changes in shape and extra-cellular matrixes.

You will be preparing slides for staining from both broth and agar surfaces. While the goals are the same for both, evenly and lightly dispersed cells firmly adhered to the slide surface, the techniques are slightly different. Staining is as much art as science. It will undoubtedly take you several tries before you are successful.


  • Clean glass slides
  • Inoculating loops or needles
  • Sterile water
  • Marking pen
  • Assorted broth and plate cultures

Safety Considerations

Be careful of aerosols when transferring bacteria from your loop to the slide. The loop is very flexible and it is easy to zing off a loop-full of organisms. Do not assume your organism is dead. Heat or methanol fixation is not guaranteed to kill the organism. Dispose of your completed slides in the disinfectant bucket at your bench.

General Considerations

You are striving for a light suspension of cells that will leave a faint cloudy deposit on your slide. You have lots of room on your slide use it! It helps to initially draw a circle on the bottom of the slide so you know where to look for your smear. It is very easy to get confused which side of the slide your smear is on. Be sure to label the far edge of the slide. Do this consistently on the same end of the slide to help orient your slide.

Be patient and take the time to let your slide air dry before proceeding with adhering it to the slide. If your slide is wet and you heat fix it, the bacteria will boil and the cellular morphology will be lost. If your slide is wet and fix it in methanol, it will most likely wash off the slide. Smears that are too thick will most likely wash off the slide regardless of the fixation method.

Smear from Broth

Broth cultures are usually easier to work with because the cells are already diluted in the broth. Be sure to carefully mix the culture tube to suspend the bacteria in the broth.

  1. Label your slide. Aseptically transfer a loop-full of organism onto the center of your slide.
  2. Use the flat part of the loop to smear the broth drop around the slide. Use a spiraling, circular motion to spread out the drop. Because the broth is full of protein, the smear will usually stay spread out and not bead up on the surface of the slide.
  3. Set the slide aside to air dry. This will take several minutes at least. Do not rush this step.

Smear from Plate

You can scoop a lot of organisms off with your loop. You may want to use an inoculating needle to transfer your organism to the slide. Be sure to use sterile water to dilute your samples. Regular tap water or the de-ionized water in your rinse bottles are often contaminated with bacteria.

  1. Label your slide. Aseptically transfer a loop-full of sterile water to the center of the slide.
    • This serves to both dilute your bacteria and give you something to spread around.
  2. Pick a well-isolated colony.
  3. Prick it with your sterile needle, or slightly scoop the edge of the colony with your sterile loop.
  4. Place your needle/loop in the center of the drop and with a spiraling circular motion spread the bacteria on the slide.
  5. Set the slide aside to air dry. This will take several minutes at least. Do not rush this step.


The fixation procedure is the same regardless of smear source, plate or broth. There are two methods of adhering your bacteria to the slide, heat fixation or methanol fixation. Heat fixing is only used with BSL1 organisms. The organisms we will be working with are BSL2, so you will need to use the methanol fixation technique. Heat fixing the slide can create aerosols and with BSL2 organisms, we need to prevent this as much as possible. Methanol fixation causes fewer changes in cellular morphology and creates no aerosols.

Please be careful when working with the methanol, if you forget you have fixed it with methanol and your slide isn’t totally dry, the remaining methanol will catch on fire.

Methanol Fixing (BSL2)

  1. Be sure your slide is totally dry. Set it on the staining rack over the sink.
  2. Carefully flood the slide with 95% methanol. Let it sit for two minutes.
  3. Tilt the slide and pour off the methanol.
    • Touch the edge of the slide to a paper towel to wick off the excess methanol.
  4. Set the slide aside to air dry before staining.

Simple Stain

Simple stains are just that - add one stain to a fixed smear slide, let it sit, rinse it off, let it dry, and view. It is a quick procedure for determining the presence and morphology of bacteria in clinical samples such as stool and discharges.

Methylene blue is used to determine the morphology of fusiform and spirochetes in oral infections. It is also the stain of choice for identifying the metachromatic granules in Corynebacterium diphtheriae. The granules will stain a distinctly deeper blue than the surrounding blue bacteria. Other species of Corynebacterium do not have the metachromatic granules. Any basic dyes, such as methylene blue, crystal violet, malachite green, or safranin work well.

Basic (cationic or positively charged) dyes bind to negatively charged components in the cell membrane and cytoplasm.


  • Methylene blue
  • Safranin
  • Crystal violet
  • Malachite Green
  • Staining racks
  • Micro tool boxes
  • Prepared smear slides

General Considerations

Staining is part art and part science. There are no hard and fast rules for staining and rinsing times. The times listed are suggestions that usually work well. You will need to experiment with what works for the bacteria you have and the techniques you use.

It is essential that you record exactly what you do and the results you observe in your lab book. You will be repeating these stains later in the semester and you don’t want to waste your time re-inventing your successful staining procedure. It would be useful for each lab bench member to pick a different stain so you can see what they all look like.

Simple Stain Procedure

  1. Place your carefully prepared fixed smear slides on the stain rack over the sink.
    • Do one slide at a time.
    • Cover the smear with any of the basic dyes available to you.
    • You only need enough dye to cover the smear. The stain should not drip off the slide.
  2. Let the stain sit for 1-5 minutes.
  3. Using the clothespin, grab the long end of the slide, tilt the slide over the sink and rinse the stain off with a stream of water from the wash bottle.
    • Be sure to spray above the smear and let it dribble down.
    • If you spray directly on the smear you are liable to wash the smear off the slide.
    • Rinse till the water runs clear or is only slightly colored.
  4. Touch the edge of the slide to a paper towel to remove excess water. You can now let it air dry. Alternately you can dry it with blotting paper by placing it in the blotting paper book and pressing lightly. While this method is quicker, you can also blot off a poorly adhered smear.
  5. View your slide under oil immersion and record your observations in your lab book.
  6. Discard your used stained slides in the disinfectant bucket in the sink.

Negative Stain

Negative stains are even simpler than simple stains because you do not have to make a smear. A drop of cells is spread on a slide and viewed without fixation. The stain is a suspension of carbon, found in India ink or nigrosin. The carbon particles are negatively-charged, as is the cell membrane. The background looks black or sepia colored and the cells remain clear, since they repel the dye.

Some positively charged inclusion bodies, such as sulfur, may stain. This stain gives accurate information on cell morphology and capsule presence because the cells are not fixed. Cell size appears slightly larger because any extracellular coatings or secretions on the outside of the cell membrane also do not stain. Negative stains are useful for rapid determination of the presence of Cryptococcus neformans, the causative agent of cryptococcisis, in cerebral spinal fluid. This technique is also used when you stain for endospores and capsules.


General Considerations

Just as in preparing a smear, you only need a small amount of organism. If you have too many organisms, you won’t be able to see the morphology of individual cells. It is also important not use too much nigrosin. If it is too thick, the background will have a cracked appearance similar to mud puddles drying in the sun. You want to get a light film. Your instructor will demonstrate this technique for you.

Negative Staining Procedure

  1. Label your slide. If you are working from a broth culture, place a loop-full of organisms about three fourths of the way on the left side of the slide. If you are working from a plate culture, add a drop of sterile water to the slide and dilute your organism in the drop without spreading the drop.
  2. Put one or two drops of nigrosin on another slide. Use your sterilized loop to pick up a loop-full of nigrosin. Carefully mix it in with the drop of cells, without spreading the drop too much.

  1. Hold the right end of the slide in your right-hand with your left–hand take another slide at a 45° or less angle to the first slide, just past your nigrosin/cell drop.
  2. Scoot the angled slide back along the surface of the first slide till it just touches the drop of nigrosin and cells. Wait for capillary action to draw the liquid along the leading edge of the angled slide.
  3. Push the angled slide across the surface of the flat slide. Most of the nigrosin should still be left on the original spot. Discard the slide in the disinfectant bucket.
  4. Set the stained slide aside to air dry before observing it under oil immersion. Be sure to start examining your slide in the area with the faintest gray background.
  5. Record your observations in your lab book.
  6. Discard your used stained slide in the disinfectant bucket.

Special Note

Nigrosin comes off the slide and onto your oil immersion lens very easily. Be sure to thoroughly clean your oil lens when you are finished. Then clean it again. Once it dries on the lens it is very difficult to remove and will impair your ability (and the other micro students using that scope) to see clearly out of the lens.

Gram Stain

The Gram stain is the most common differential stain used in microbiology. Differential stains use more than one dye. The unique cellular components of the bacteria will determine how they will react to the different dyes. The Gram stain procedure has been basically unchanged since it was first developed in 1884. Almost all bacteria can be divided into two groups, Gram negative or Gram positive. A few bacteria are gram variable. Trichomonas, Strongyloides, some fungi, and some protozoa cysts also have a Gram reaction. Very small bacteria or bacteria without a cell wall, such as Treponema, Mycoplasma, Chlamydia, or Rickettsia do not have a gram reaction. The characterization of any new bacteria must include their gram reaction.

Typically a differential stain has four components the primary stain, a mordant that sets the stain, a decolorizing agent to remove the primary stain, and a counter stain. In the Gram stain, the primary stain is crystal violet. This gives the cell an intense purple color. The mordant, iodine, forms a complex with the crystal violet inside the cell wall. The cell is then washed with either Gram’s de-colorizer or 95% ethanol. Gram positive cells will retain the dye complex and remain purple. The dye rinses out in gram negative cells. The counter stain, safranin, is used to color the cells that lost the primary stain, other wise they would remain colorless and you wouldn’t be able to see them.

The large iodine-crystal violet complex is retained within the cell walls of gram positive cells because of the molecular structure of the many layers of peptidoglycan in the cell wall. There are lots of cross-linked teichoic acids and the iodine-dye complex cannot physically get out. There is also less lipid in the membrane and the decolorizing agent cannot get to it as well. Gram negative cells have an outer membrane and only one layer of peptidoglycan, with more lipid. The crystal violet dye is easily washed out.

General Considerations

The accuracy of the Gram stain is dependent on the integrity of the bacterial cell wall. There are a variety of things that can influence the cell wall integrity old cells (i.e. cultures over 24 hours old), the sample is from someone treated with antibiotics that target that cell wall such as penicillin, the cells have been roughly handled or you over heat fixed them. Under these conditions, gram positive cells will come out as gram-negative. If you de-colorize too long, Gram-positive cells will look like Gram-negative cells. Conversely, if you do not decolorize enough, Gram-negatives will look like Gram-positives. The only way you can trust your results it to always run a known Gram-positive and a known Gram-negative on the same slide. If they stain as predicted you can be pretty sure the result of your unknown sample is reliable.

The Gram staining takes practice to get right. Do not expect to get a good Gram stain on your first try. It is a good idea to hold your slide with a clothespin your gloves will get pretty psychedelic as will everything you touch!

Gram Stain Procedure

  1. Label your slide. Prepare your smears on a slide with a Gram negative on the left, your unknown in the middle, and a Gram positive on the right.
    • Don't forget to methanol fix your slide!
  2. Laying your slide on the staining rack, cover the smears with crystal violet for 1 minute.
    • Use just enough stain to fully cover the smear but not so much that it runs or drips off the slide.
  3. Tilt the slide and pour the crystal violet off and briefly rinse with water from your wash bottle.
    • Remember not to spray directly on the smears or you will wash them off.
  4. Flood the slide with iodine mordant for 1 minute.
  5. Tilt the slide, pour off the excess iodine and gently decolorize with Gram’s de-colorizer until it just begins to run clear.
    • This is the tricky part!
  6. Tilt your slide on some paper towels to remove excess de-colorizer.
  7. Flood your slide with the safranin for 1 minute.
  8. Tilt the slide to pour off the excess safranin and gently rinse with water until it runs clear.
  9. Let the slide air dry
  10. Observe under oil immersion.
    • The Gram positive control should be purple and the Gram negative control should be pink.
  11. Discard your used slide in the disinfectant bucket.

Congo Red Capsule Stain

The Congo Red Capsule stain is a modification of the nigrosin negative stain you may have done previously. The bacteria take up the congo red dye and the background is stained then with acid fuchsin dye. The capsule or slime layers, highly hydrated polymers, exclude both dyes. The background will appear blue, the bacterial cells will appear pink, and the clear halos are the capsules.

Clinically, the capsules of some highly pathogenic bacteria (i.e.: pneumococci, Haemophilis influenzae, and meningococci), can be distinguished with the use of antisera specific for that type of capsule. The bacteria are suspended in the antisera and then mixed with methlyene blue. In the antisera staining procedure, the bacteria will appear blue surrounded by a clear halo and then surrounded by a thin blue line where the antisera have attached to the capsule.


  • Congo Red stain
  • Acid fuchsin stain
  • Acid alcohol
  • Klebsiella pneumoniae culture
  • Enterobacter aerogenes culture

Congo Red Capsule Stain Procedure

  1. Place a loop-full of Congo Red on a slide
  2. Mix a small amount of your organism into the drop of Congo Red.
    • Spread the organism/dye suspension well on the slide
  3. Let the slide thoroughly air dry.
    • Do not methanol fix!
  4. Fix the dried slide with acid alcohol for 15 seconds.
  5. Rinse with distilled water and cover the slide with acid fuchsin for 1-5 minutes.
  6. Rinse with water and allow to air dry.
  7. Examine the slide under oil immersion.
    • Cells stain red/pink, and the capsules appear as colorless halos against a dark blue background.

Wirtz's Endospore Stain

Endospore formation is characteristic of Clostridum and Bacillus spp. The ability to concentrate and coat their protoplasm allows them to survive the adverse environmental conditions they experience in their soil habitat. This also allows the spores to resist staining. The “live” organisms are easily visualized with simple stains and Gram’s stains.

Endospores are typically highly refractile, light striking them is deflected. Many Bacillus species have inclusion bodies that are highly refractile. These inclusion bodies may look like endospores with regular staining. The presence of endospores must be confirmed with endospore specific stains. The presence, and characteristic shape and position of endospores require special procedures to permeate the endospore coat. Most endospore stains involve heating the slides while keeping them continually moist with the dye. While quicker, it produces volatile chemicals and is just a big mess. The same results can be obtained by letting the dye sit on the slide for 30 minutes. It is a good idea to start this slide first and work on another stain while you are waiting for the dye to permeate the endospore.

General Considerations

You will be using a Bacillus species for the endospore stain. The shape and position of B. cereus spores are very similar to those of B. anthracis. Bacillus does not start forming spores until it runs out of food. If the cultures are too young, you will mostly see just the pink rods of the bacteria. If the cultures are too old, you will mostly see just the small green ovals of the endospores. Ideally, you should see the green oval bodies of the endospore surrounded by the pink vegetative bacterial cell. Select a sample from the middle of a colony with the straight inoculating needle for the best results. The edge of the colony is still actively growing and will have few endospores.


A capsule in bacteria is the result of amorphous viscid secretion released by the bacteria. When this secretion remains loose and un-demarcated it is called a slime layer and when it is organized into a sharply defined structure it is called as Capsule.

Most of the capsules are made up of polysaccharides but there are many which are composed of Polypeptides (Proteins).

Capsules are usually fragile and can easily be destroyed or distorted by heating so the capsule staining is designed in such a way that maintains the overall morphology of bacteria as well as preserved the Capsule structure so that we can easily identify it.

Also, to get the better results and to enhance the size of the bacterial capsule a drop of serum can be used while preparing a smear which makes it more convenient to observe bacterial capsule with a typical compound/light microscopes.

There is no use of Complex processes and multiple strong reagents or stains in Capsule staining process and simplest methods like India ink method gives the satisfactory results so that we can easily determine the presence or absence of capsule in the bacterial cell.

Heat fixing is also avoided as it will destroy or distort the capsule which makes it more convenient and moreover the newbie’s to microbiology can easily do it and explore more in the microbial world….

Briefly, in Capsule staining technique, the background and the bacterial cell body is stained whereas capsule remains colorless. However, the result varies as per the methodology used for capsule staining.


Firstly, let’s discuss the bacterial cell structures which are involved in this staining technique and later on we’ll move to the principles involved in capsule staining technique.

A capsulated bacterium acquires a capsule around its body which appears as a clear halo. A Bacterial capsule is non-ionic in nature, so there is no or very minute possibility that the acidic or basic stains will adhere to their surfaces so it remains colorless.

The excellent way to demonstrate the capsule is to stain the bacterial cell body and the background leaving the bacterial capsule as colorless.

The Bacterial cell body is stained best with the Basic stains like Crystal violet, Safranine, Methylene blue etc. whereas the background is stained best with the Acidic stains which include the India ink, Nigrosin, Eosin, Congo red etc.

There are numerous methods available for the demonstration of the capsule in bacterial cells. The result of the staining varies as per the Method followed but commonly all the methods have one thing in common that it stains the bacterial cell, capsule and/or Background. The Two most commonly used methods are as follows:

India ink method / Nigrosin method

This one is the easiest method of capsule staining. In this technique, two dyes are used that is Crystal violet and the India ink / Nigrosin.

The Capsule of the bacterial cell appears as a clear halo around the stained bacterial cell body and the Dark background (color depends upon the dye used – India ink or Nigrosin).

Anthony’sstain method

This one is another most commonly used method of Capsule staining. In this technique, the crystal violet stain is used as the primary stain.

Besides that, a 20% solution of copper sulfate is used which plays an important role in staining procedure by acting as a decolorizing agent as well as Counterstain.

20% CuSO4 decolorizes the bacterial capsules by removing the Crystal violet that sticks on it but has no such effect on bacterial cell body and it also stains the capsule which later appears as faint blue color zone or halo around purple bacterial cell body.

One important thing to note here is that there is no mordant and no strong decolorizers are used.


  • Microscopic glass slide.
  • Inoculating loop.
  • Spirit Lamp.
  • Staining Rack.
  • Wash bottle.
  • Microscope (with 100X objective lens).

For India ink / Nigrosin method

For Anthony’s Stain method


The procedure of capsule staining using India ink / Nigrosin method

Take a clean, Dry, Scratch and Grease free Microscopic glass slide and place a drop of India ink or Nigrosin on it at one end near the edge.

Take a small portion of the bacterial colony or a loopful of broth culture with the help of sterilized Inoculating loop.

Mix well the culture with dye taken on the Glass slide.

Now, take another Microscopic Glass slide, place it near to the specimen-dye mixture at an angle of about 30° – 45°.

Move the slide toward the drop of the specimen-dye mixture until the contact is made with the drop at the specific angle. Then move the spreader slide smoothly and rapidly forward over the specimen slide, drawing the dye mixture behind it into a thin film.

Allow the smear to Air dry.

NOTE: Do not Heat as heat will melt the capsule and distorts the actual shape of the bacterial cell.

Now, Flood the smear slide with crystal violet solution for 1 minute and rinse carefully and gently with water.

NOTE: Be cautious while doing this step as water may remove the capsule from the cell as well as it may wash out the smear

Allow the smear slide to air dry.

NOTE: Do not Blot dry the slide as it may distort the capsule as well as the smear.

Observe under the microscope at High power objective (45X) and oil immersion (100X) objectives. To easily visualize the capsule make sure to decrease the amount of light while observing under the microscope.

The procedure of capsule staining using Anthony’s stain method

The procedure of Anthony’s stain method is quite similar to that of India ink / Nigrosin method initially which is as follows:

Take a clean, Dry, Scratch and Grease free Microscopic glass slide and place a drop of Crystal violet on it at one end near the edge.

Take a small portion of the bacterial colony or a loopful of broth culture with the help of sterilized Inoculating loop.

Mix well the culture with dye taken on the Glass slide.

Now, take another Microscopic Glass slide, place it near to the specimen-dye mixture at an angle of about 30° – 45°.

Move the slide toward the drop of the specimen-dye mixture until the contact is made with the drop at the specific angle. Then move the spreader slide smoothly and rapidly forward over the specimen slide, drawing the dye mixture behind it into a thin film.

Allow the smear to Air dry.

NOTE: Do not Heat as heat will melt the capsule and distorts the actual shape of the bacterial cell.

Now, Tilt the smear slide and rinse it with the 20% copper sulfate (CuSO4) solution.

NOTE: Do not Blot dry the slide as it may distort the capsule as well as the smear.

Observe under the microscope at High power objective (45X) and oil immersion (100X) objectives. To easily visualize the capsule make sure to decrease the amount of light while observing under the microscope.


Results of Capsule staining using India ink / Nigrosin Method

The capsule appears as the Clear halo or clear zone around the purple colored bacterial cell body on a dark background. The Background color is due to the India ink or Nigrosin whatever you’ll use.


Results of Capsule staining using Anthony’s stain Method

The capsule appears as the Faint blue halo around the purple bacterial cell body on a transparent background as there is no background stain is used in this technique.



By using India ink / Nigrosin technique, we stain the background and bacterial cell body, not the bacterial capsules actually.

By Anthony’s Stain method we stain the Bacterial cell body as well as the Capsule but not the Background.

Further Reading:

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Capsule Stain:

Capsules are the gelatinous outer layer of the bacterial cells and these structures cannot retain the color of the staining agents. The capsules can be visualized by means of two methods.

Positive Capsule Staining

Since capsule is water soluble in nature, it is too difficult to stain the capsule with normal staining methods. The positive capsule staining method (Anthony Method) uses two reagents to stain the capsular material. The primary stain Crystal violet is applied over a non heat fixed bacterial smear so that both the bacterial cells and capsular material take up the color of the primary stain. The ionic nature of the bacterial cell binds the crystal violet stain more strongly while the non-ionic nature of the capsule get adhere with the crystal violet stain. When the decolorizing agent copper sulfate is added over the bacterial smear, the loosely adhered crystal violet stain is washed off from the capsular material without removing the tightly bound crystal violet from the cell wall. The capsular material absorbs the light blue color of the copper sulfate in contrast to the purple bacterial cell.

Negative Capsule Staining

Another simple method to visualize the bacterial capsules is by using negative staining Technique. During staining the non heat fixed bacterial smear with the acidic stains such as Nigrosin will not penetrate the bacterial cells (since both acidic stain and bacterial surface has negative charge). Instead the acidic stain deposits around the bacterial cells and create a dark back ground and the bacteria appear as unstained with a clear area around them, capsule.

Note: If you heat fix the bacterial smear for capsule staining, the cells will shrink creating a hallow zone around the bacterial cell and will be mistaken for the capsule.

What are Acid Fast Bacteria

The acid fast bacteria are a type of bacteria that resist decolorizing by acid after staining. Acid fastness is a physical property of bacteria, which rely on the structure of the bacterial cell wall. Typically, the cell wall of bacteria is made up of proteins, carbohydrates, and lipids. Acid fast bacteria comprise a thin layer of peptidoglycans. The mycolic acid is a long chain of fatty acids, attached to the peptidoglycans. Once the primary stain, carbol-fuchsin is added to the slide containing bacteria, the mycolic acid is attached to carbol-fuchsin. This makes the acid fast bacteria to stain in pink even after decolorization.

Figure 1: Acid Fast Bacteria

Mycobacteria is a common type of bacteria, which grows slowly hence, they are composed of a thick layer of mycolic acid. In addition to mycolic acid, acid fast bacteria consist of a large amount of fatty acids, complex lipids, and waxes. Due to the presence of thick bacterial cell wall, acid fast bacteria are highly resistant to disinfectants as well as dry conditions. The cell wall structure and the staining of acid fast bacteria are shown in figure 1.

Biochemical properties (continued)

Another test is whether or not your unknown has an hemolytic reaction. Most bacteria are gamma-hemolytic, which means that they do not have an hemolytic reaction. This test is mostly used on streptococci species: it differentiates non pathogenic streptococci from pathogenic streptococci. This is tested on a blood agar plate: a beta-hemolysis creates a white discoloration around the colony whereas an alpha-hemolysis has a brownish green zone around the colony. Streptococcus pyogenes is not a pathogen and therefore is beta-hemolytic whereas Streptococcus pneumoniae or Streptococcus salivarius are alpha-hemolytic.

Another biochemical property is the production of H2S from the oxidation of sulfur containing compounds like cysteine or the reduction of inorganic compounds like thiosulfates, sulfates or sulfites. The media used is peptone-iron agar. The peptone has sulfur containing amino acids which are used by the bacteria to produce H2S and the iron detects the H2S by forming a black residue along the stab line. Proteus vulgaris for example produces H2S.

The following test is the coagulase test which shows if bacteria are capable of coagulating oxolated plasma. It is an indication of pathogenicity since if a bacteria can coagulate the blood, it can wall off from the immune system. Staphylococcus aureus can coagulate oxolated plasma and therefore blood. It is also capable of secreting gelatinase which is the enzyme that hydrolyzes gelatine into polypeptides and amino acids.

The following series of tests is called IMVIC which stands for Indole, Methyl red, Voges-Proskauer and Citrate.

  • The indole production test shows if the bacterial strain is capable of breaking down tryptophan by tryptophanophase into indole, ammonia and pyruvate. We can detect this reaction by using Kovac&aposs reagent which is contained in amyl alcohol (not miscible in water). Kovac&aposs reagent reacts with indole to form Rosindol dye, forming a red color that will rise to the top of the broth culture. This test is positive for Escherichia coli and Proteus vulgaris but negative for Enterobacter aerogenes for example.
  • The methyl red test tests for glucose fermentors. It turns red when the pH is inferior to 4,3. It is positive for E. coli but negative for E. aerogenes.
  • The Voge-Proskauer tests shows the production of acetoin. The reagent used is potassium hydroxide, a creatine solution. The medium turns red if the test is positive for E. aerogenes for example. It is negative for E. coli.
  • Finally, the citrate test is used to differentiate enterics. It tests if the bacterium has the permease required to take up the citrate and use it as the sole carbon source. The indicator used is bromothymol blue: the black medium turns blue if the citrate is used. E. aerogenes has the permease however E. coli doesn&apost.

Preparing Specimens for Electron Microscopy

Samples to be analyzed using a TEM must have very thin sections. But cells are too soft to cut thinly, even with diamond knives. To cut cells without damage, the cells must be embedded in plastic resin and then dehydrated through a series of soaks in ethanol solutions (50%, 60%, 70%, and so on). The ethanol replaces the water in the cells, and the resin dissolves in ethanol and enters the cell, where it solidifies. Next, thin sections are cut using a specialized device called an ultramicrotome (Figure 9). Finally, samples are fixed to fine copper wire or carbon-fiber grids and stained—not with colored dyes, but with substances like uranyl acetate or osmium tetroxide, which contain electron-dense heavy metal atoms.

Figure 9. (a) An ultramicrotome used to prepare specimens for a TEM. (b) A technician uses an ultramicrotome to slice a specimen into thin sections. (credit a: modification of work by “Frost Museum”/Flickr credit b: modification of work by U.S. Fish and Wildlife Service Northeast Region)

When samples are prepared for viewing using an SEM, they must also be dehydrated using an ethanol series. However, they must be even drier than is necessary for a TEM. Critical point drying with inert liquid carbon dioxide under pressure is used to displace the water from the specimen. After drying, the specimens are sputter-coated with metal by knocking atoms off of a palladium target, with energetic particles. Sputter-coating prevents specimens from becoming charged by the SEM’s electron beam.

Think about It

  • Why is it important to dehydrate cells before examining them under an electron microscope?
  • Name the device that is used to create thin sections of specimens for electron microscopy.

Using Microscopy to Diagnose Syphilis

The causative agent of syphilis is Treponema pallidum, a flexible, spiral cell (spirochete) that can be very thin (<0.15 μm) and match the refractive index of the medium, making it difficult to view using brightfield microscopy. Additionally, this species has not been successfully cultured in the laboratory on an artificial medium therefore, diagnosis depends upon successful identification using microscopic techniques and serology (analysis of body fluids, often looking for antibodies to a pathogen). Since fixation and staining would kill the cells, darkfield microscopy is typically used for observing live specimens and viewing their movements. However, other approaches can also be used. For example, the cells can be thickened with silver particles (in tissue sections) and observed using a light microscope. It is also possible to use fluorescence or electron microscopy to view Treponema (Figure 10).

Figure 10. (a) Living, unstained Treponema pallidum spirochetes can be viewed under a darkfield microscope. (b) In this brightfield image, a modified Steiner silver stain is used to visualized T. pallidum spirochetes. Though the stain kills the cells, it increases the contrast to make them more visible. (c) While not used for standard diagnostic testing, T. pallidum can also be examined using scanning electron microscopy. (credit a: modification of work by Centers for Disease Control and Prevention credit b: modification of work by Centers for Disease Control and Prevention credit c: modification of work by Centers for Disease Control and Prevention)

In clinical settings, indirect immunofluorescence is often used to identify Treponema. A primary, unstained antibody attaches directly to the pathogen surface, and secondary antibodies “tagged” with a fluorescent stain attach to the primary antibody. Multiple secondary antibodies can attach to each primary antibody, amplifying the amount of stain attached to each Treponema cell, making them easier to spot (Figure 11).

Figure 11. Indirect immunofluorescence can be used to identify T. pallidum, the causative agent of syphilis, in a specimen.

Lab 4: Acid-Fast, Spores, and Capsule Stains - Biology

Before staining, the specimen must be mounted and fixed on the slides, as previously done in the simple staining technique. Because of the 2 dyes used in the procedure–crystal violet and safrinin—as well as the decolorizer acetone-alcohol, bacteria will fall into 2 groups based on their gram reactivity. Gram positive bacteria retain the crystal violet even through the decolorizor step: gram negative bacteria do not retain the crystal violet, are decolorized, and then pick up the safrinin dye. Both gram + and – bind to the crystal violet: the key step to their differentiation is the decolorization.

Take a look at the accompanying diagram of the stain procedure and its effects on the bacterial color. During the crystal violet-iodine step, the bound molecules within the peptidoglycan of the gram + cell wall and within the membrane are held tightly. The acetone-alcohol actually causes the peptidoglycan molecules (arranged in a latticework) to shrink, thereby holding the crystal violetiodine even tighter. In the gram – cell, the outer lipopolysaccharide layer of the wall is dissolved by the decolorizer agents, and because the peptidoglycan layer is so thin in that group of bacteria, the crystal violet is leached out of the wall.

Although there is a standard routine and set reagents used in this stain, each person has to find a particular method that works best for them. The many variables that can affect this stain are age of the culture, amount of decolorizer used, the time of decolorization, the type of organism (acid-fast bacteria and spores do not stain well), thickness of the smear, and the general care of the stainer.


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