Chapter 4: Microscopic techniques and principles of ... · Staining techniques ... Specialized...

Applied Veterinary Bacteriology and Mycology: Bacteriological Techniques Chapter 4: Microscopic techniques and

principles of staining methods used in a diagnostic bacterial laboratory

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Applied Veterinary Bacteriology and Mycology: Bacteriological techniques

Chapter 4: Microscopic techniques and Principles of Staining Methods Used in a Diagnostic Bacteriology Laboratory

Author: Dr. J.A. Picard

Licensed under a Creative Commons Attribution license.

TABLE OF CONTENTS

INTRODUCTION .......................................................................................................................................... 2

COMPOUND LIGHT, BRIGHT-FIELD MICROSCOPY ............................................................................... 2

Adjustment of Köhler illumination ............................................................................................................ 3

Precautions when using a microscope .................................................................................................... 6

Determination of the size of objects viewed under the compound light microscope ............................... 6

Methods in light microscopy .................................................................................................................... 7

Fluorescence microscopy ........................................................................................................................ 8

Methods of Sample Preparation for Microscopic Examination ................................................................ 9

Staining techniques ............................................................................................................................... 10

REFERENCES ........................................................................................................................................... 13

APPENDIX ................................................................................................................................................. 14

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INTRODUCTION

In spite of the development of new identification techniques, microscopic examination of clinical material remains

one of the most effective means of judging specimen quality and detecting the presence of potential pathogens

in clinical material. It is also used to examine the morphology and differential staining characteristics of artificially

cultivated bacteria and fungi. Specialized techniques and stains when used with different microscopy techniques

will further aid in the identification of an organism.

COMPOUND LIGHT, BRIGHT-FIELD MICROSCOPY

This is the most commonly used of all microscopes, due to its versatility, ease of use and low cost of

maintenance.

A typical light microscope is illustrated in Figure 1. A light source, usually provided by a coiled tungsten filament

starts as a horizontal beam, which is transmitted through the condenser via a mirror.When reaching the

condenser lens it is focused just below the plane of the specimen (Figure 2). This prevents glare from the beam

as well as affects the resolution of the image. This type of light transmission is known as Köhler illumination and

is most efficient when properly centred in the light path of the specimen. This is accomplished by using the

condenser adjustment screws on the stage.



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Adjustment of Köhler illumination

(Do this for the most common magnification you use, as the setting varies according to magnification used).

1. Place a mounted specimen on the specimen stage (1)

2. Set the light intensity control dial (2)

3. Make sure that the selected condenser lens (3) is in the path of the light.

4. Using the condenser ratchet (4), set it to the highest position (nearest to the specimen)

5. Focus on the material at infinity, using the fixed eyepiece (5)

6. Close the field diaphragm (6) until a ring of light just fits into the field

7. Set the second eyepiece (7) so that the image is sharply focused

8. Focus the light by slowly lowering the condenser (4). There is a ring of light that changes from the blue to

red colour range. Adjust it so that this colour change just occurs.

Figure 1: A typical light, bright-field compound

microscope. (Use this diagram with the description

below on how to adjust so that ideal Köhler

illumination is obtained).

1. Stage 2. Light intensity adjustment 3. Substage condensor 4. Ratchet for adjusting substage condenser 5. Ocular objective adjustable to eyes 6. Field diaphragm: adjusts light field 7. Ocular objective – non-adjustable 8. Condenser adjustment screws 9. Objective 10. Condenser aperture adjustor 11. Placement of light bulb/light source 12. Fine focus 13. Focus 14. Substage diaphragm



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9. Centre the light in the field of vision using the two centering screws (8) in the condenser holder. If the light

path is not visible, lower the condenser first, find the light path and follow this while raising the condenser

again.

10. Open up the field diaphragm (6) until the edge of light path starts to disappear from view.

11. The diameter of the diaphragm must be two-thirds of the opening of the objective. Check by removing

eyepiece (5).

12. Contrast can be adjusted by setting the condenser opening (10).

13. If the light is not uniform, check the lamp (11). Do not touch the globe. The reflection of the filament image

must be the same size, when looking through the eyepiece (5) with ocular lenses removed.

The contrast of the specimen may be enhanced by decreasing the condenser aperture and thus the amount of

oblique light waves reaching the specimen. This is useful when examining wet mounts or uniformly stained

specimens. It does result however, in decreased resolution of the image.

As the human eye is most sensitive to blue wave lengths, a blue filter over the field diaphragm enhances

visualization and causes less fatigue when multiple slides are viewed.

Lenses

This microscope uses at least two magnifying lenses, namely the ocular and objective lenses. The total

magnification of the microscope is the product of the magnification of the ocular and objective lenses.

The ocular lens is usually set at a magnification of 10X. Several objective lenses are used, they are

commonly:

4X scanning lens

10X or 20X intermediate lenses

40X high dry lens

100X immersion oil lens



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Figure 2: Illustration of condenser function showing focusing light paths at the plane of the specimen

Achromatic (corrected for light distortion in the red and blue ranges) objective lenses are important to use in

microscopy as they correct for chromatic aberration (splits the light into colours – prism effect) which is an

inherent property of most convex lenses. Apochromatic lenses are used in microphotography as they correct for

both chromatic and spherical aberrations (fuzzy images). Another factor which affects the viewing of the image

is the resolution, which is defined as the smallest distance between two objects which allows them to be viewed

as distinct objects. Immersion oils with a refractive index similar to glass are used with the 100X objective lens.

The 100X objective lens is usually close (0.2 mm) to allow the entry of more rays of light. This is further assisted

by the use of immersion oil. Note that the immersion oil used is specified by the manufacture and it is not

recommended that oils are mixed as it can decrease the resolution power of the lens. Also clean the lens with a

lens paper every time after use to prevent the oil hardening on the lens. Note that many solvents can cause the

glue holding the lens in its casing to dissolve. The resolution limit for the compound light microscope is 0.2m

that is obtained with the 100X oil immersion lens.



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Precautions when using a microscope

As the light supply and lens must meet each other in a straight line, it is critical that care is taken not to handle

them roughly or drop them. They must always be placed in a sturdy box with packaging to prevent any

movement and to protect them.

Determination of the size of objects viewed under the compound light microscope

An eyepiece micrometer is used to measure the size of objects. Before use, it should be calibrated for the

magnification you will be using.

This is done by placing a table micrometer on the specimen stage and focusing on the scale.

Turn the eyepiece containing the micrometer so that the two scales are parallel to each other (Figure 3).

Find the number of units on the eyepiece scale that exactly coincides with one or more units on the table

micrometer: each unit on the table micrometer represents 0.1mm. To calculate the value of one eyepiece unit

(X), the following calculation is done:

X = table micrometer units (mm)

Number of eyepiece micrometer units over the same distance

Figure 3: The positioning of the eyepiece micrometer to lie adjacent to the table micrometer when viewed through the microscope.



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Methods in light microscopy

Dark-field microscopy

This method of microscopy, using a darkfield condenser, excludes directly transmitted light and only

allows oblique or scattered light to be directed onto the specimen (Figure 4). This set-up allows finer

structures to be seen as the resolution improves to approximately 0,1 m. The background appears dark

whereas objects in the fluid such as bacteria appear as brightly luminous against a black background. It is

commonly used for visualization of the spirochaetes.

Figure 4: Comparison of bright-field microscopy (A) and dark-field microscopy (B)

Phase contrast microscopy

Due to the small nature of micro-organisms, it is not possible to discern internal structures with the use of

a normal microscope. Phase contrast microscopy increases the contrast of an object by converting slight

differences in refractive index and cell density into easily detected variations in light intensity (Figure 5). In

order to achieve this effect an annular diaphragm is placed at the lower focal plane (Figure 5). This

permits only a ring of light to pass through the condenser and objective lens which is then focused on the

phase ring just before reaching the eyepiece. This ring is there to change the phase of the light (not hitting

an object on the slide) reaching it, so that it is not a quarter of a wavelength different to the diffracted rays



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(those scattered by bouncing off an object on the slide). At some points these waves will come into

contact with each other either amplifying each other (bright) or nullifying (dark) each other. By using this

method of microscopy the background is illuminated and the unstained object is dark surrounded by a

halo of light. Internal structures of cells such as endospores and nuclear bodies will also show up in a

similar fashion.

Figure 5: The optics of a phase-contrast microscope

Nomarski interference

This type of microscopy is used for bigger objects than bacteria, such as vegetative cells and spores of fungi.

Normarski interference is similar to phase contrast microscopy, but the presence of a polarizer and special

prisms in the condenser result in the formation of a clearer image that can be obtained by phase contrast

microscopy. The object being examined has no halo and appears three dimensional. The resolution power is

0.1m compared to 0.2m of a phase contrast microscope.

Fluorescence microscopy

This technique has become commonplace in most laboratories. Fluorescence is dependent on the ability of

fluorophores (naturally fluorescent substances) or fluorochromes (fluorescent dyes) to absorb the energy of non-

visible UV and short visible wavelenghts become excited, and remit the energy in the form of longer visible

wavelenghts. Figure 6 gives a diagrammatic representation of illumination required for fluorescent microscopy.

A special adapter containing the light source (high pressure gas lamps of mercury, xenon or halogen), filters,

and a dichromatic mirror beam splitter can be attached to a compound bright-field microscope so that light



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passes through the objective lens. This option has greatly reduced the costs of fluorescence microscopy. This

adapter is usually adequate to detect most organisms or their antibodies using direct or indirect fluorescence

staining techniques. Its sensitivity is 84% and specificity 93% compared to that of a standard fluorescence

microscope when fluorescence (auramine stain) is used to detect acid-fast bacteria. Note that a special

ultraviolet, opaque filter is placed in the microscope tube to protect the eyes from the ultraviolet rays.

Figure 6: Fluorescence microscope incident illumination light path and microscope components

Methods of Sample Preparation for Microscopic Examination

Wet preparations and hanging mounts

These are unstained, wet preparations of the material, usually made to observe the viable microorganism

and examined using reduced light, phase contrast or dark field microscopy. On these smears one is able

to observe motility, which is indicative of flagellae or fimbriae, bacterial spores, intracellular granules and

spirochaetes. A plain wet mount is done by suspending bacteria in a drop of fluid on a microscope slide

and covering it with a thin cover slip. A ring of petroleum jelly (Vaseline) can be drawn around the drop

with a toothpick to prevent drying out.

A hanging drop method is used when free movement of the micro-organisms is necessary. Briefly pick

some bacteria (not too many – as it will result in overcrowding) from a colony on a culture plate or a

loopful from a broth and suspend in a drop of water or saline on a coverslip. Invert to coverslip position



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either over a slide with a hollow or over a thin ring made from plasticine (Prestik) (Figure 12). A hanging

drop method is not suitable for anaerobes, as air might inhibit their movement. It is best to observe them in

sealed capillary tubes containing growth media.

Staining techniques

Stains are used to determine bacterial morphology and to distinguish bacteria belonging to different groups or

species by their differential staining characteristics. Prior to staining all slides are fixed by heat (most common),

methyl alcohol, formalin, magnesium chloride or osmic acid. Fixation immobilises and kills vegetative bacteria

and thus renders them more permeable to staining. As a result of fixation, there is protoplasmic shrinkage, thus

a string of bacteria, will appear to have spaces between them, and some bacteria, such as the diphtheria

bacterium will be beaded and Pasteurella species will appear to be bipolar.

Different types of stains can be used and include:

Simple stains e.g. carbol fuschin stain.

Negative staining e.g. India ink.

Silver impregnation.

Differential stains e.g. Gram’s stain

Simple stains

The application of a basic dye, such as methylene blue, methyl violet, basic fuschin or carbol fuschin, will

show the presence of organisms and the nature of cellular contents in exudates. Sometimes a mordant is

added to these dyes to allow better penetration of the dye. A basic dye stains bacteria because coloured

positively charged particles combines firmly with the negatively charged group in the bacterial protoplasm,

especially with the phosphate group in nucleic acids. The excess stain is then washed off with water and

the combined stain remains. Very rarely are acid dyes used as they stain bacteria at a low pH. They are,

however, used for negative staining. Carbol fuschin is useful for visualising Campylobacter, Helicobacter,

spirochaetes and Fusobacterium in tissue smears. Mature Löffler’s methylene blue stain (see Appendix) is

used to stain Bacillus anthracis in blood and tissue smears.



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Negative staining

A stain such as India ink or nigrosin, stains the background dark, so that bacteria or fungi are visualised

as clear transparent objects. It is a good method to show shape, size and arrangement of bacteria and

fungi. For example, it is the best stain to visualize the heavily capsulated yeast, Cryptococcus neoformans

in tissue smears. Some bacilli, such as those of the coliform and haemophilic groups, also have a central

dark staining portion in their cells resembling a nucleus.

Silver impregnation

This method aids in the visualization of fine, morphological distinct microorganisms such as spirochaetes

and Gram-negative curved bacteria e.g. Campylobacter in tissue sections.

Differential stains

Note that the methods for performing the stains are described in the Appendix.

Gram’s stain

This is the stain most commonly used in diagnostic bacteriology and is used to place bacteria into one of

two groups: Gram-negative and Gram-positive, as well as to examine the morphology of bacteria. The wall

of Gram-positive bacteria is able to retain basic dyes, such as crystal violet, at a higher hydrogen ion

concentration and is more permeable to these dyes. Crystal violet and iodine form a complex within the

cell wall which is impermeable to water, but moderately soluble and dissociable in alcohol or acetone.

Thus on decolourization by acetone-alcohol, a thinner wall (as is possessed by Gram-negative bacteria)

will allow easier leaching of dye. Thus, Gram-positive bacteria will stain purple and Gram-negative

bacteria, being decolourized, will stain with the pink counterstain (Safranin). Old or damaged Gram-

positive bacteria e.g. Bacillus spp. and Streptococcus spp. will, however, stain Gram-negative. Note too

that bacteria cultured in acidic media will also stain Gram-negative. Some bacteria such as mycobacteria

which have a highly impermeable cell wall do not stain well with Gram’s stain.

Acid-fast or Ziehl-Neelsen stain

Certain bacteria such as mycobacteria are relatively impermeable to most stains, but do stain with a

strong reagent such as hot carbol fuschin in 5% phenol. Once stained these bacteria resist

decolourisation by strong acids e.g. sulphuric acid. The smear is then counterstained with either

methylene blue or malachite green. Acid-fast bacteria stain pink and any cellular material or other bacteria

stain blue or green, dependent on the counterstain used. Mycobacteria are acid-fast as their cell wall is

rich in lipids, fatty acids and the higher alcohols.

Partial acid-fast stains or Stamp’s stain



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Certain bacteria cannot withstand decolourisation by a strong acid, but do if a weaker acid such as 0,5%

acetic acid is used. Brucella, Chlamydia, Coxiella and Nocardia species stain with this method. Some

bacterial endospores are also partially acid-fast.

Giemsa and Diff Quik (CAM’s Quick, Rapid-Diff) stains

These stains are very useful in the staining of certain bacteria and to visualize cellular morphology in

tissue smears. They are used to demonstrate the capsule of Bacillus anthracis and the spirochaete

Borrelia in blood and tissue smears. Bacteria usually stain well (deep purple) as the relationship of

bacteria to tissue cells can be well demonstrated. Yeasts also stain well with these stains. However,

moulds and mycobacteria tend to stain poorly or not at all.

Staining for spores

Spores are usually easy to observe as they stain negatively, whereas bacteria stain positively when

normal bacterial stains are used. It is, however, possible to stain spores using acid-fast staining

techniques. At times, the ideal growth conditions of artificial culture media can inhibit the production of

spores. Therefore, before a spore stain is attempted, it might be necessary to culture the bacteria on a

starch or trace element constrained medium.

Staining of capsules

The capsules of bacteria present in pathological material are often clearly stained with standard bacterial

stains, such as basic fuschin, methylene blue, Giemsa or Diff Quik stains. Gram’s, Giemsa and Diff Quik

stains colours them pink. However, capsules of bacteria cultured artificially usually do not stain well. Thus

negative or relief staining techniques should be used. The best method to use is a wet-film India ink stain,

as there is no protoplasmic shrinkage which could cause a false positive result. Slime produced by

bacteria appears as irregular masses of pink amorphous material lying between the bacteria and outside

the capsule of capsulated bacteria. Some bacteria lose their capsules when cultured on artificial media

and special media and growth conditions may be required e.g. capsules will only be produced by Bacillus

anthracis if grown in 5% CO2 on bicarbonate rich agar medium.

Staining of flagella

Because of the small size of flagella, they are difficult to visualize with light microscopy. Thus techniques

are used to thicken them to ten times their normal size. A modified Leifson’s method is used, making use

of basic fuschin with tannic acid, which is deposited on the bacteria from an evaporating alcohol solution.

This stain will both swell and stain the flagella. The protoplasm of the bacteria is then stained with

methylene blue.

Staining of fungal hyphae and yeast cells



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Fungi often stain with difficulty when bacterial or cytological stains are used. Thus special stains have

been developed to observe them. In cultured material the vital stain lactophenol cotton blue is preferred.

Stains for the observation of fungi in clinical material include the Periodic-Acid-Schiff (PAS), and calcofluor

white. An Indian ink wet mount is used to observe Cryptococcus neoformans blastospores and 10-20%

potassium hydroxide with a vital dye to examine fungal infected hairs.

The PAS stain is a two-step procedure, in that the periodic acid hydrolyzes the cell wall aldehydes, which

then are able to combine with the modified Schiff reagent colouring the cell wall carbohydrates a bright

pink magenta.

Calcofluor is a nonspecific fluorochrome that binds to the 1,3-linked polysaccharides, specifically

cellulose and chitin in cell walls of fungi. This stain can be mixed with KOH to clear the specimens.

REFERENCES

1. Veterinary Microbiology and Microbial Disease, (2011). Quinn, P.J., Markey, B.K., Leonard, F.C., FitzPatrick,

E.S., Fanning, S., Hartigan, P.J. Wiley-Blackwell. ISBN 978-1-4051-5823-7



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APPENDIX

Stains used in bacteriology

Included are formulae and preparation methods

of commonly used stains in a bacteriology

laboratory. Although formulas for stains are

given, ready to use solutions are available

commercially. For other staining procedures it is

best to refer to bacteriology textbooks.

Points to be considered when making stains.

a) All dyes should be weighed accurately on an

analytical balance.

b) The dyes should be ground in a mortar with

the diluent, adding a small amount of the

diluent at a time until all has been added.

c) All staining solutions should be filtered

before use, but not before they have been

allowed to stand for at least 24 hours.

d) No solutions should be used after

precipitation has occurred. When

precipitation occurs, discard the solutions

from the bottom of the bottle after the stain

has been used or filter and use the filtrate.

e) Diluted solutions should be made in small

quantities. The stock solutions keep for a

much longer period of time than the diluted

solutions.

f) All solutions should show on the labels the

concentration of the dye present and the

date of preparation. Use stain-resistant

labels.

g) Keep all staining solutions out of direct

sunlight.

h) Keep all staining solutions in glass-

stoppered or sealed bottles.

Preparation of smears from pure cultures

a) Slides should be fat-free and free from lint or

any other foreign substance. New slides can

be cleaned by soaking in 95% ethyl alcohol,

then wiping dry with clean gauze.

b) Label slide with a diamond tipped pen,

graphite pencil or wax marker.

c) Place a loopful of distilled water in the centre

of the slide.

d) Flame a platinum wire

e) Lift cover of Petri dish, or lid from container

holding the culture. If necessary flame

mouth of container. Please note that if a

broth culture is used, solids may interfere

with the stain.

f) Cool the loop inside the tube or in a clean

portion of the agar, pick up a small portion of

the material to be smeared. If too much it

will be difficult to study the morphology of

individual bacteria and to observe motility.

g) Replace lid of container.

h) Holding slide in one hand, gently emulsify

material on platinum loop in water or saline

on the slide.

i) Flame loop.

j) Allow material on the slide to dry.

k) Fix smear by passing it three times through

the blue portion of the flame,

l) Allow to cool before staining. Hot slides

cause artefacts.

m) If fixation by alcohol instead of dry heat is

desired, the slide is placed in a Coplin jar of

methanol or ethanol.

n) Stain slide according to individual staining

instructions.

o) Always clean the back of the slide after

staining.

p) Allow the slides to air dry or if hasty use a

hand held hair dryer.



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Supplies

1. Absolute methanol

2. Analytical balance

3. Bunsen burner or alcohol flamer

4. Coplin jars (dependent on staining

technique)

5. Culture to be tested or specimen for direct

smears

6. Distilled water

7. Ethanol (95%)

8. Frosted or dark glass bottles

9. Glass microscope slides, frosted at one end

10. Gloves

11. Graphite marker (pencil)/ wax marker/

diamond tipped pen.

12. Bacteriological loop

13. Standard light compound bright-field

microscope

14. Blotting paper

15. Stains (see list of suppliers) according to

requirements of laboratory.

16. Non-absorbant paper and marking pen

17. Tripod for Bunsen burner

18. Water bath.

1. CAMSQuik/Rapid Diff/ DiffQuick

Use stain according to manufacturer’s

instructions. In samples where there is a heavy

bacterial load, it is best to use staining racks, to

prevent contamination of the stains. This stain

tends to overstain, particularly when new.

Procedure

1. Fix smear for 30s in methanol (fixative)

2. Stain in solution one (pink stain) for 8

seconds or eight dips or until stain clings to

smear.

3. Stain in solution two (purple) for 20 seconds.

4. Wash under running tap water, air dry and

examine.

Interpretation

Cell nuclei, protozoa, bacteria and yeast stain a

dark blue to purple. Cell cytoplasm, fibrin and

debris stain a light blue to pink.

1. Gram’s Stain (Hucker’s Modification)

Stock crystal violet

Crystal violet 10g

Ethanol (95%) 100ml

Stock oxalate solution

Ammonium oxalate 1g

Distilled water 100ml

Crystal violet working solution: Mix 20ml of stock

crystal violet with 80 ml stock oxalate solution.

Gram’s iodine solution.

Iodine crystals 1g

Potassium iodide 2g

Dissolve completely in 10ml of distilled water,

and then add distilled water to make 200ml

Store in an amber bottle.

Decolourizer

Ethanol (95%) 75ml

Acetone 25ml

Counterstain

Saffranin 2.5g



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Ethanol (95%) 100ml

Dissolve this stock solution 1:4 with distilled

water.

An alternative counterstain is:

Basic fuschin 3g

Ethanol (95%) 100ml

Procedure

Best performed on young cultures, because

older cultures decolourize too rapidly.

1. Make a thin smear of clinical material or thin

emulsion of colony. Air dry smear. Fix the

film by passing through a Bunsen burner

flame three times.

2. Flood the slide with crystal violet stain.

Leave for 60s.

3. Pour off the stain, and wash with water.

4. Flood the slide with Gram’s iodine. Leave for

60s.

5. Wash off iodine with water and shake off

excess water.

6. Decolourize with acetone-alcohol until

decolourizer flows off colourless from the

slide, but for a maximum of 20 seconds.

7. Counterstain with safranin for 30 s and wash

off with water. Weak carbol fuschin at a

dilution of 1:10 can also be used.

Interpretation

Gram-positive: Dark blue

Gram-negative: Pink

Note that old cultures of gram-positive bacteria

may stain Gram-negative.

2. Giemsa stain

Stock solution

Giemsa powder 0,3g

Glycerin 25ml

Absolute acetone-free methanol 25ml

(Available commercially)

If the stain does not go into complete solution, it

should be filtered.

Giemsa buffer

Sodium phosphate Na2HPO4 (anhydrous) M/15

9,47 g/l or Na2HPO4.2H2O M/15 11.87 g/l

61.1 ml

Potassium phosphate KH2PO4 (anhydrous) M/15

9,08 g/l 38.9 ml

Distilled/deionised water 900 ml

One volume of stock solution is diluted with 9

volumes of Giemsa buffer.

Staining procedure

1. Fix smear in methanol for 3-5 mins.

2. Dry in air

3. Immerse in diluted stain for 5 min (5%

Giemsa) or for 30 mins (10% Giemsa).

4. Wash with distilled water or buffer.

5. Allow to air dry.

Interpretation

Bacteria, fungi, protozoa and nuclei will stain a

dark blue. Cytoplasm and fibrin will stain a light

pink/blue. The capsule of Bacillus anthracis

stains magenta red.

3. Loeffler’s Methylene Blue Stain



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This is a simple basic stain. After the stain

ripens or is oxidized (12 months), it is useful for

staining the capsules of Bacillus anthracis.

(McFadyean’s reaction).

Methylene blue (1% in 95% ethanol) 30ml

Potassium hydroxide (0,001%

aqueous solution)

1ml


Procedure

1. The fixed smear is stained with the above

solution for 1 min.

2. Wash in running tap water and allow to air

dry.

Interpretation

Bacteria stain blue, with endospores appearing

as unstained bodies within the cells. The

beading and granules of corynebacteria may

also be seen. The capsule of Bacillus anthracis

appears as amorphous purplish material around

the bacteria (McFaydean’s reaction).

5. Schaeffer and Fulton spore stain

5% aqueous solution of malachite green

0,5% aqueous solution of safranin

Staining procedure

1. Prepare smear and heat fix as for Gram’s

stain.

2. Flood with malachite green.

3. Steam gently over a flame for 30 sec (don’t

allow to boil). The stain should be just

steaming. Placing the glass smear over a

jar with boiling water in the area of the

steam will have the same effect.

4. Wash with water and stain with safranin for

30 sec.

5. Wash under running tap water, allow drying

and examine.

Interpretation

Endospores stain a bright refractive green and

bacteria protoplasm pink.

6. Stamp’s stain (modified acid-fast for

Brucella)

a. Stock carbol fuschin (weak)

Basic fuschin 1g

Absolute methanol 10ml

Phenol (5%) 90ml

b. Decolourizer

Acetic acid 0,5ml

Distilled water 95,5ml

For Nocardia 0,05% aqueous sulphuric acid

4. Counterstain

Loeffler’s methylene blue (see Ziehl Neelsen

method)

Staining procedure

1. Make smear and heat fix.

2. Stain smears in a 1:10 solution of the stock

carbol fuschin for 15 mins.

3. Wash in running tap water to remove

excess stain.

4. Decolourize with acetic acid for 20 - 30

secs.

5. Wash in running tap water.

6. Counterstain with Loeffler’s methylene blue

for 30sec.

7. Wash and air dry. Examine.



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Interpretation

Red: Brucella, Nocardia, Chlamydia, Coxiella

Blue: Other organisms and background

7. Ziehl-Neelsen Acid-Fast Stain (rapid

method)

Solutions:

1. Carbolfuchsin stain

Basic fuchsin

Ethanol, 95%

2. This solution is mixed with

5% Phenol

3. Acid alcohol

Hydorchloric acid

(concentrated)

Ethanol, 95%

4. Counterstain

Malachite

green/methylene blue

Distilled water

0.3 g

100 ml

45 g/ 900

ml water

32 ml

970 ml

2.0 g

1000 ml

Procedure

1. Heat fix impression smears

2. Cover slides with carbolfuchsin stain and

heat steam for 5 minutes

3. Rinse with tap water

4. Decolorise by rinsing with acid alcohol for 2

minutes

5. Rinse with tap water

6. Counterstain with malachite green for 0.5 –

1 minute

7. Rinse with tap water and leave to air dry

Interpretation

Red: Mycobacteria

Blue (or green); Other organisms and

background material

8. Modified acid-fast stain for Nocardia

species.

Staining procedure

1. Make a smear of the organism from growth

media and heat fix.

2. Flood the slide with Kinyoun carbol fuchsin

for 5 minutes.

3. Pour off excess stain.

4. Decolourize with 1% aqueous sulphuric

acid.

5. Wash with tap water.

6. Counterstain with methylene blue for 1

minute.

7. Rinse with water and dry.

8. Examine with the 100X oil immersion optics.

Acid-fastness can be enhanced by growth on

Middlebrook 7H11 agar.

Interpretation

Acid-fast (red): Most Nocardia spp.

Negative (blue): Other actinomycetes.

1. Diene’s stain for mycoplasma cultures

Methylene blue 2.5g

Azure II 1.25g

Maltose 10g

Sodium carbonate 0.25g




19 | P a g e

Staining procedure

1. Place a small amount of the stain next to a

suspected colony (with a loop or cotton

swab).

2. The stain will diffuse and the colony will

become blue if it is mycoplasma. Live

bacteria decolourize the stain within a few

minutes.

Staining procedures for fungi

India ink wet mount

Clinical material or organisms from cultures are

mixed on a slide in a drop made up of loopfuls of

distilled water and India ink or permanent black

pen ink. Experience will indicate the amount of

India ink to use. A coverslip is added, and the

preparation examined. The large capsules of

Cryptococcus neoformans show up. An

alternative method is to suspend the material in

distilled water under a coverslip and then add

the India ink to the edge of the coverslip. The ink

will seep under the coverslip and provide areas

of variable density.

1. Lactophenol cotton blue

Lactophenol blue is used both as a mounting

fluid and a stain. Lactic acid acts as a clearing

agent and aids in the preservation of fungal

structures, phenol acts as a killing agent,

glycerol prevents drying and cotton blue gives

colour to the structures.

Stain preparation

Lactic acid 20ml

Phenol crystals (conc. Phenol) 20g

Glycerol or glycerine 40ml


Cotton blue

or 1% aqueous solution

0,05g

2ml

Dissolve phenol in the lactic acid, glycerol, and

water by gently heating (if crystals were used).

Then add cotton blue (Poirrier’s blue and aniline

blue are analogous to cotton blue). Mix well.

Staining procedure

1. Place a drop of the stain on a clean

microscope slide.

2. Pick up some fungal hyphae by using

either a dissecting needle or cellophane

tape (for methods refer to Chapter 25).

3. Place in or on the drop of stain.

4. Place a coverslip on top and examine

under a light microscope, with reduced

condenser aperture.

Interpretation

Young fungal hyphae stain blue.

3. Periodic-acid-Schiff (PAS)

Procedure

1. Fix the smear with formalin-ethanol for 1

min.

2. Drain alcohol and place in 5% periodic acid

for 5 mins.

3. Wash in running water for 2 mins.

4. Place in basic fuschin zinc (or sodium)

hydrosulphite (Schiff reagent) for 2 mins.

5. Wash for 2 mins. Under running water.

6. Immerse slide in sodium metabisulphate for

2 to 5 mins.

7. Wash for 5 mins under running water.

8. Counterstain with picric acid or light green

for 5 seconds.



20 | P a g e

9. Wash for 5-10 seconds.

10. Dip slide for 5 second intervals in 85%,

95% and absolute alcohol consecutively.

11. Dip in xylene, add mounting medium and

cover with a coverslip.

Interpretation

Fungal elements stain a bright pink-magenta or

purple against an orange background if picric

acid is used or green background if light green is

used.

4. Calcofluor white

This is a very rapid and useful method to

examine fungi in specimens.

To make up the stain

Calcofluor white M2R* 100mg

Evans Blue 50mg


*Fluorescent brightener 28 Product no F3397.

Sigma

Mix well and store at room temperature in a dark

bottle.

Method

1. Use I drop of calcofluor white

2. Add I drop of 10% KOH (for clearing)

3. Add a coverslip, allow to sit at room

temperature for approximately 3 minutes

and examine under a fluorescent

microscope with an exciter that transmits

wavelengths between 300 and 412nm.

4. If the slide is to be saved, remove the

coverslip, rinse the slide briefly with distilled

water and allow to air dry. The smear then

can be stained with a permanent stain.

Interpretation

Fungal structures are seen as a brilliant apple-

green or ghostly white, dependent on the

wavelength of light they are examined under.

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