Applied Veterinary Bacteriology and Mycology: Bacteriological Techniques  Chapter 4: Microscopic techniques and principles of staining methods used in a diagnostic bacterial laboratory

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 aCreative Commons Attribution license.

TABLE OF CONTENTS

INTRODUCTION

COMPOUND LIGHT, BRIGHT-FIELD MICROSCOPY

Adjustment of Köhler illumination

Precautions when using a microscope

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

Methods in light microscopy

Fluorescence microscopy

Methods of Sample Preparation for Microscopic Examination

Staining techniques

REFERENCES

APPENDIX

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.

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.
  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

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 0.2m that is obtained with the 100X oil immersion lens.

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.

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 (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 0.1m compared to 0.2m 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 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 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.

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

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.