Microscope Slides

Microscopes are often used with microscope slides. These are usually thin, flat pieces of glass typically measuring 75 × 26 mm and 1 mm thick [1]. Slides are used to hold samples in place for examination under a microscope and help to prevent the microscope and lenses from becoming contaminated. Plain slides can be used either way up, but some have a frosted area for labelling. When using these slides, the frosted side should be uppermost [1].

Cover Slips

Coverslips are flat pieces of glass measuring around 20 mm wide and fewer than 1 mm thick. Cover slips are placed over a specimen on a microscope slide to hold the specimen in place and to protect it from contamination. Cover slips also serve to protect the objective lenses from contamination and damage [1].

Microscope Care

Microscopes are expensive instruments that must be carefully stored and maintained. The following points should be considered [1, 2, 11]:

• The microscope must be turned off and covered when not in use.

  

  
  
• The microscope must be stored on a flat surface, away from water, excessive heat and vibrations.
  
• When moving the microscope, it should be lifted by the limb with one hand under the base.
  
• The stage should be cleaned after each use with disinfectant wipes.
  
• The eyepieces and the objective lenses should only be cleaned with lens tissue. Lens tissue is soft, lint‐free tissue paper that will not scratch or damage the lens.
  
• Safety checks should be carried out regularly, including electrical testing.
  
• Immersion oil should be used sparingly with the X100 lens.
  
• When oil has been used, the oil immersion lens should be cleaned immediately after use with lens tissue. This will prevent the oil from solidifying and damaging the lens.
  
• The light should be switched off when not in use to prolong the life of the bulb and stop it from overheating. If the light does need to be left on, the light intensity switch (rheostat) should be turned down to the lowest setting.

  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  
  

  Microscope component		Function
  1	Eyepiece	Monocular microscopes have one eyepiece; binocular microscopes have two. Eyepieces typically have a magnifying power of X10. They contain an ocular lens which magnifies the primary image for the operator to view
  2	Objective turret	The objective turret holds the objective lenses and rotates in a clockwise direction allowing the operator to move from a lower‐power objective lens, to a higher‐power objective lens
  3	Limb, body and base	The limb of the microscope connects the body and the base. A microscope should be carried by holding the limb with a second hand on the base of the microscope. The base includes the light source and the on/off switch
  4	Stage (including mechanical stage)	The stage is a flat platform with a hole in the middle, which allows light from the condenser to illuminate the specimen. The microscope slide is placed on the stage and is held in place by clips. Once a specimen is placed onto the stage, using the focusing mechanisms the stage can be moved up and down to assist with focusing. The mechanical stage, attached on top of the stage, can then be used to move the microscope slide horizontally, or vertically
  5	Objective lenses	The objective lenses are situated within the objective turret of the microscope. There are usually 4 objective lenses, starting at X4, which is the lowest power; X10, X40, which is the highest power for dry magnification; and X100, which would be used for oil immersion
  6	Vernier scale	The Vernier scale is found on the mechanical stage. It consists of a horizontal and a vertical scale, which allows for the precise location of the specimen to be recorded and also relocated when required
  7	Substage condenser	The substage condenser is situated below the stage and is made up of two lenses which condense the light from the light source and onto the specimen. The position of the condenser and the quantity of light passing through the specimen can be altered using the iris diaphragm
  8	Iris diaphragm	The iris diaphragm regulates the amount of light that can pass through the substage condenser. Closing the diaphragm has advantages when examining parasites as it increases the contrast, which helps with visibility. Opening the diaphragm will help with visibility when examining cytological samples
  9	Focusing mechanism (Fine and coarse)	Found on the side of a microscope, this mechanisms allows raising and lowering of the stage to help with the focus of the image. The larger wheel is the coarse focus; the smaller is the fine focus. To start with, the stage should be racked as high as possible (while watching the stage at all times) and then be lowered; this will prevent the objective lens from accidentally hitting the slide and potentially causing damage
  10	Rheostat	The rheostat allows the level of light produced by the light source to be altered
  11	On/off switch	The on/off switch is usually situated on the base of the microscope. Before turning the microscope on, the rheostat should be checked to make sure it is at its lowest setting; otherwise damage to the bulb could occur

  
  
• After use, the stage should be lowered, and the lowest power objective lens should be moved into position.

Magnification and Focusing

Most binocular microscopes have four objective lenses; each one has a different function [1, 2, 11]:

• X4 objective lens: This is a very low‐power lens mainly used to obtain an initial focus on a sample. It can also be used for the examination of coat brushings and to scan a slide for an area of interest. When using this objective lens, the condenser and light should both be on a low setting.
  
• X10 objective lens: This is another low‐power objective lens. This lens can be used for the identification of parasites and the assessment of urine sediment. It can also be used for locating areas of interest before increasing the magnification. When using this objective lens, the condenser and light should be on a low setting.
  
• X40 objective lens: This is a high‐power objective lens. These lenses are mainly used to examine areas of interest in more detail, for example, when examining a pathological sample. When using this objective lens, the condenser and light should be on a medium setting.
  
• X100 objective lens: This is a high‐power lens used with oil. This lens uses light that gets refracted through the thin layer of oil to the lens. It is used for cytology and to examine blood smears and fine needle aspirates. When using this objective lens, the condenser and light should be on a high setting.

Examining a Specimen on a Microscope Slide [1]

1. Use PPE if required.
   
2. Remove the microscope dust cover and ensure the microscope is plugged into an electrical socket.
   
3. Adjust the light beam diaphragm control so that it is in the middle position (to do this, look from the side, not down the eyepiece) and ensure the rheostat is turned down to its lowest setting.
   
4. Turn the microscope on. The power switch is usually on the base of the microscope.
   
5. Move the stage down to its lowest setting and secure the slide on the mechanical stage using the spring arm clips.
   
6. Rotate the objective turret (clockwise) and click the X4 or X10 objective lens into place (depending on the microscope).
   
7. Looking at the stage directly, move it back up until it almost touches the objective lens. Ensure that the lens does not touch the slide.
   
8. Look down the eyepieces and adjust the distance between them; only one field should be viewed (a single image) when the eyepieces are in the correct place.
   
9. Adjust the rheostat to a medium setting and adjust the substage condenser to a couple of millimetres below the stage.
   
10. While looking down the eyepieces, slowly move the stage downwards using the course focus until the image comes into view.
    
11. Once the specimen is visualised, the fine focus should be used to sharpen the image.
    
12. Rotate through the objectives lenses if a larger magnification is needed. If it was in focus at a low power, it should remain in focus at the higher power. Every time a larger objective lens is used, it should be observed during movement to ensure it does not touch the slide on the mechanical stage.
    
13. The Vernier scale should be read and recorded as this can be used to identify the position of interest and enable colleagues to examine the same area on the slide.

Oil Immersion Technique

The oil immersion technique is used when a detailed examination is required, for example, when looking at a blood smear. It works by refracting light through a thin layer of oil to the lens. Oil immersion must only be used with the oil immersion objective lens (X100). The oil immersion technique should be used during the final stage of an examination to prevent contamination of the other objective lenses.

When using oil immersion, the rheostat should be set to the highest value, as this will facilitate better visibility. The objective lenses should be rotated slightly so none are fixed into position; this leaves a space to drop a small amount of oil immersion directly onto the slide over the spot of light. If a coverslip is present, this will need to be removed. The oil immersion objective lens should be moved into place, ensuring it does not come into contact with the slide. Watching the stage position from the side, the stage should be raised slowly until the oil drop touches the lens. If the specimen was in focus with a smaller magnification, any adjustments should be made using the fine focus [1, 11].

The Battlement Technique

The battlement technique is used for a quantitative microscopic examination of blood when determining the number of cells seen. This differs from a qualitative examination used when assessing cellular morphology. When examining blood smears, a logical examination must occur to reduce the possibility of counting the same blood cell twice, which would give an inaccurate result. The battlement technique should begin on the left‐hand side of a microscope slide and involves moving two fields to the right, two fields up, two fields to the right and then two fields down (see Figure 8.2). This pattern should be repeated across the slide until 100 cells have been counted. All cells in each field should be counted. For the most accurate count, the edge and the middle of the body of the smear should be examined. This is to compensate for the maldistribution of cells between the centre and the edge of the blood smear. The battlement technique is most commonly used when carrying out a differential white blood cell count where the percentages of the different cell types are determined [1–3].

Vernier Scale Readings

Vernier scale readings can be used to record the exact location of a specimen or a particular part of an image. Examples include an ectoparasite from a skin scrape or a white blood cell on a blood smear. There are two scales: the vertical Vernier scale (Y‐axis, which runs from top to bottom) and the horizontal Vernier scale (X‐axis, running from left to right). The main scale is marked on the stage, which gives half of the reading; the other half comes from the smaller Vernier scale. Both the vertical and horizontal Vernier scales must be read, and the readings must be recorded accurately.

The Method to Read the Vernier Scale is as Follows [1]:

1. The stage can be lowered if required to facilitate an accurate reading. The stage should not be touched again.
   
2. For each direction (vertical or horizontal), there is a main scale (marked on the stage) and a smaller, Vernier scale located next to the main stage.
   
3. For the main scale reading (which gives the first part of the scale), observe where the 0 on the Vernier scale aligns with the main scale. In Figure 8.3, it is 18. If this falls between two divisions, the lower number should be used.
   
4. For the Vernier scale reading, locate the number on the Vernier scale that is in closest coincidence with the main scale. In Figure 8.3, it is 4.
   
5. This will give the first complete reading of 18.4.
   
6. This procedure should then be repeated for the other axis.
   
7. Once both readings have been identified, they can be used to relocate an area of interest on a microscope slide.
   
8. All results should be recorded.

Haematology Analysers

Haematology analysers are used to count and characterise blood cells for the detection of diseases and monitoring purposes. They can carry out tests such as cell counts and coagulation tests, but in equine practice, they are most often used for their differential counts of both white and red blood cells. Most of these analysers work on either a coulter electrical impedance method or flow cytometry; both methods rely on the cells being passed rapidly through a laser beam (or a small aperture) to allow each cell to be counted and differentiated.

Haematology analysers provide more accurate cell counts than manual counts; for example, some of the analysers perform the differential white blood cell count on 10,000 rather than 100 cells. They are also much quicker; some analysers can perform 120 tests per hour. But for this to be accurate and timely, they need to be used correctly and serviced regularly. Haematology machines can help in the diagnosis of conditions such as anaemia, dehydration and different types of infections [1–3, 12, 13].

Biochemistry Analysers

Biochemistry analysers are used to measure the levels of various biochemical substances found within the blood, for example, glucose and total protein (TP) (Figure 8.4). Results from biochemistry machines are compared to a reference range for each parameter; the results give information on the patient’s clinical condition, and this facilitates a more accurate diagnosis and treatment plan. Biochemistry can help to diagnose conditions such as hepatopathy, bacterial infections and metabolic abnormalities. Often, biochemistry is initially used alongside haematology to get a full picture in relation to the health status of a patient.

There are two types of biochemistry machines [1–3, 14]:

• Dry chemistry analysers: These are the most commonly used analysers. The sample is dropped onto a series of chemical‐impregnated slides, initiating a colour change. This colour change is then read and interpreted by the machine. The colour change reflects the amount of substance being tested.
  
• Wet chemistry analysers: These use wells of fluid rather than slides. Chemical reactions between the fluid and the blood create a colour change, which determines the biochemical levels in the individual sample.

Electrolyte and Blood Gas Analysers

These analysers read electrolyte levels in plasma and can give rapid results for tests such as sodium, chloride and calcium. They can be used for blood gas analysis in critical care patients and to monitor patients undergoing general anaesthesia. Handheld analysers can be used to measure glucose and lactate in blood samples.

PPE For Sample Collection

When setting up for sample collection, the need for PPE should always be considered. The handling of pathological specimens and some preservatives, such as formal saline, is classed as hazardous and precautions should be taken to ensure good health and safety standards. Some cases that require sampling could be classed as infectious or being treated as such until the sample results are known. Some infectious diseases may also be zoonotic, such as salmonella or dermatophytosis, so strict biosecurity measures will be required to prevent the spread of infection to staff and other patients. PPE should include an apron or laboratory coat and gloves. Depending on the collection method used and the patient’s disease state, PPE could also include a mask, hair cover, disposable boiler suit, foot covers and goggles [1–3].

Blood Cell Counts

Blood cell counts are a quantitative examination of blood as they determine the number of blood cells seen. Within most practices, quantitative blood analysis is often carried out using a haematology machine. Such machines produce results within a quicker timeframe than manual blood cell counts and are often more accurate if used correctly. White blood cell counts are carried out on blood smears using the X10, X40 and then X100 oil immersion objective lenses. These are often performed to determine the quantities of each type of white blood cell, known as a differential white blood cell count (see section on blood smears). Red blood cell counts are commonly known as a packed cell volume (PCV) and are discussed below.

Packed Cell Volume (PCV)

PCV is a rapid test that can be performed in‐house to determine a patient’s hydration status, and the level of blood loss in patients who are or have been haemorrhaging. PCV is a measurement of the proportion of blood that is made up of cells. This value is expressed as a percentage. The PCV reference ranges in horses differ with age, fitness and breed. The normal reference ranges are as follows [3]:

• Neonates: 40–52%
  
• Six months of age: 29–41%
  
• Adults: 32–53%

To measure the PCV of a patient, blood is drawn into a microhematocrit or capillary tube from a blood sample. Microhematocrit tubes are thin tubes which are most commonly made from soda lime glass. There are two types of microhematocrit tubes [1]:

• Red‐tipped tubes: Contain sodium heparin to prevent the sample from clotting. These tubes are used with plain blood samples.
  
• Blue‐tipped tubes: Do not contain sodium heparin and so are used with blood samples that have been mixed with an anticoagulant.

The correct microhaematocrit tube should be selected for the sample being tested.

The method for preparing a PCV sample is displayed in Table 8.5.

When the microhaematocrit tube is removed from the centrifuge after spinning, the sample will have spun into layers. The red blood cells will be concentrated in a layer at the bottom of the tube next to the clay plug. Above this is the buffy coat containing white blood cells and platelets. Above this is the plasma (see Figure 8.11). A Hawksley haematocrit reader is most commonly used to read the %PCV of a sample (see Figure 8.12).

The Hawksley haematocrit reader is used as follows:

1. Place the capillary tube into the slot in the reader with the clay seal facing downwards (the capillary tube holder) (see Figure 8.13).
   
2. Align the top of the clay seal with the zero line on the reader, then move the capillary tube holder along until the very top of the plasma is aligned with the 100% line which is on the reader.
   
3. There is an adjustable reading line on the left side of the reader. This reading line should be moved until it is directly over the intersection between the red blood cells and the buffy coat layer.

   

   

   

   

   
   
4. Once the adjustable reading line is in place, read the percentage it is lined up with on the right‐hand side of the reader; this gives the PCV reading (see Figure 8.14).

If a Hawksley reader is unavailable, the following simple calculation can be used to manually identify the % PCV:

The length of the column of red blood cells divided by the length of the red blood cells, buffy coat and plasma combined X100. So, as an example, if the height of the red blood cells is 2.4 cm, this is divided by the height of all three layers, which is 4.8 cm [1–3]:

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