1. Action: Place the microscope on an even, stable surface.

    Rationale: The slightest vibration will be magnified when looking down the eyepiece making viewing an object very difficult.

2. Action: Before use, clean the eyepieces (Fig. 5.1), objective lens and the condenser with special lens tissue.

    Rationale: Lens tissue is lint free so that bits are not left on the surfaces. Other types of tissue or cloth may cause scratches. Cleaning should be done after you finish using the microscope, but you may need to do it again.

3. Action: Clean the oil immersion lens with the proper cleaning fluid.

    Rationale: This lens should be cleaned after every use with isopropanol. Oil left on the lens may damage the lens and cause it to consolidate.

4. Action: Turn the light control to a minimum and turn on the instrument.

    Rationale: This prevents the sudden power surge breaking the microscope bulb.

5. Action: Adjust the eyepieces.

    Rationale: If using a binocular microscope arrange the eyepieces so that both fields converge as one. If you normally wear glasses remove them – you can alter the focus of the microscope to suit your eyes. Make sure that you are comfortable.

6. Action: Rotate the nosepiece in a clockwise direction so that the ×10 objective lens clicks into the viewing position.

    Rationale: Always start viewing a slide through the lowest magnification.

7. Action: Rack up the sub-stage condenser until its top surface is as high as possible.

    Rationale: This condenses the light source onto the specimen to make it bright and sharper.

8. Action: Looking from the side of the microscope, adjust the iris diaphragm control lever so that it is in its middle position.

    Rationale: This regulates the amount of light passing through the condenser. As the lens magnification is increased the aperture of the iris diaphragm should be increased.

9. Action: Place the slide on the stage over the hole in the centre.

    Rationale: The stage is made of black vulcanite with a hole in the centre through which light from the condenser illuminates the slide. There may be clips on the stage to hold the slide firmly in place.

10. Action: Position the area to be viewed over the light source by using the control knobs on the mechanical stage.

    Rationale: This allows the slide to be examined smoothly and accurately without touching the slide with your fingers.

11. Action: Focus using the coarse adjustment knob and then with the fine adjustment knob.

    Rationale: Always focus up from the slide to prevent the objective lens descending on to the slide and cracking it. Avoid using the fine focus to excess – if you have to do this it means you are not near enough to focus with the coarse focus.

12. Action: To increase magnification, turn the nosepiece clockwise until the next objective lens clicks into place.

    Rationale: There will usually be three objective lenses screwed into the nosepiece and one oil immersion lens. These provide different levels of magnification:

13. Action: After you have finished, remove the slide from the stage.

    Rationale: Slides should be stored in an appropriate box or, if not needed, should be disposed of in the sharps bin.

14. Action: After use, wipe the objective lenses with lens tissue and turn the nosepiece so that the lowest-powered lens is in position. Reduce the light and switch off and cover the microscope with its cover.

    Rationale: The microscope is now clean and ready to use the next time.

1. Action: Set up the microscope as described above and place the slide on the stage.

    Rationale: You are now ready to use oil immersion.

2. Action: Rotate the nosepiece so that neither the ×50 or ×100 lens is in position.

    Rationale: You need to have clear access to the slide when you place the drop of oil.

3. Action: Place a drop of oil on the slide (Fig. 5.2).

    Rationale: This oil is specifically designed for this purpose. Do not use any other type of oil.

4. Action: Rotate the nosepiece until the ×100 lens is above the slide and click into position.

    Rationale: This technique can only be done using the ×100 lens.

5. Action: Gently adjust the focus so that the lens descends to touch the drop of oil.

    Rationale: Always watch what you are doing. Do not look at the slide through the eyepiece as you will find it impossible to judge the distance and may smash through the slide. The lens must be lying in the oil to avoid distortion of the image and to achieve magnification.

6. Action: Looking through the eyepiece, adjust the focus using the fine control. At this point the oil may be seen to spread out (Fig. 5.2).

    Rationale: You may need to adjust the light to improve the view.

7. Action: When you have finished, remove the slide and clean the microscope as described above.

    Rationale: The microscope is now clean and ready to use the next time.

1. Action: Place the slide on the microscope stage and fix it with the clips.

    Rationale: It is important that the slide does not move around as this will invalidate your scale references.

2. Action: Locate the object you wish to identify.

3. Action: Look at the scale on the vertical axis of the stage.

    Rationale: See Figure 5.3.

4. Action: Record the number where the zero mark on the Vernier plate meets the main scale.

    Rationale: In Figure 5.3 the zero mark falls between 31 and 32. If it falls between two divisions, record the lower number.

5. Action: Make a note of which of the marks on the Vernier plate is exactly opposite a division on the main scale.

    Rationale: In Figure 5.3, mark number 6 is exactly opposite a division on the main scale.

6. Action: Record this reading, placing it after the decimal point.

    Rationale: In Figure 5.3 this will give a reading of 31.6.

7. Action: Repeat steps 3–6 using the horizontal scale.

    Rationale: You will now have two readings – always write the horizontal scale first, e.g. 90.1 × 31.6.

8. Action: You now have a grid reference for that object on that slide provided that the slide is placed in the same position on the stage.

    Rationale: By tradition, slides are placed on the stage with the label to the right.

9. Action: You may now remove the slide, but can return to the same location using the grid reference.

    Rationale: This is useful if you wish to show someone else what you have found. You can locate it quickly and easily.

1. Action: Always ensure that the centrifuge is placed on an even, stable surface.

    Rationale: Slight vibration may occur when the centrifuge is switched on, and an uneven or unstable surface may cause the machine to move around and even fall to the ground.

2. Action: Use only the tubes recommended by the manufacturer.

    Rationale: These tubes, apart from microhaematocrit tubes, have conical bottoms and are designed to withstand centrifugal force.

3. Action: When placing the tubes in the machine, make sure that the top is not protruding above the top of the bucket.

    Rationale: This may affect the way that the machine works and make it unbalanced.

4. Action: Make sure that the sample tubes are balanced by placing in diametrically opposite buckets assessed by weight not volume.

    Rationale: If you do not do this then the machine will be unbalanced and may vibrate excessively. Stop the machine and check the tubes – this is the most common cause of vibration and instability.

5. Action: When using microhaematocrit tubes, ensure that the Plasticine® end is placed against the outer ring of the instrument.

    Rationale: This prevents the blood inside from escaping as it is subjected to centrifugal force.

6. Action: Vacutainer® tubes may be spun with their stoppers in place.

    Rationale: If the tube is opened or broken then aerosol contamination of the environment might occur.

7. Action: Lock the lid of the centrifuge securely.

    Rationale: If you do not the lid will fly open; however, most machines will not switch on if the lid is not secured properly.

8. Action: Set the spin speed as appropriate.

    Rationale: Different procedures require different speeds; for example, urine requires a lower speed than heparinized blood for biochemistry.

9. Action: Never attempt to open the centrifuge until the head has stopped rotating.

    Rationale: If you do this you may damage the machine and yourself.

10. Action: After use, turn off the power supply and clean the machine thoroughly using a mild disinfectant and a soft cloth.

    Rationale: To prevent contamination of the next samples.

11. Action: Wear disposable gloves to remove any spillages or broken glass.

    Rationale: To prevent any harm to yourself.

1. Action: Collect sample in an EDTA tube and rotate gently to mix the contents.

    Rationale: EDTA is an anticoagulant. Heparin tubes may also be used. Do not shake the tube as you may damage the blood cells.

2. Action: Remove the cap and tilt the sample so that a clear surface free of air bubbles can be seen.

    Rationale: If bubbles get into the capillary tube, they may cause an air-lock and slow the rate of filling.

3. Action: Place the end of a capillary tube into the blood sample, tilting the sample to at least 55°, and allow blood to run in until the tube is about full.

    Rationale: Blood will run into the capillary tube by capillary action. The microhaematocrit reader requires at least 5–7 cm of blood.

4. Action: Wipe the blood from the outside of the tube with a piece of tissue.

    Rationale: This reduces the risk of spread of infection to you.

5. Action: Holding the tube between your finger and thumb, insert the opposite end (from the blood) into the Plasticine® or Cristaseal® block. Twist two or three times and take it out of the Plasticine®.

    Rationale: This creates a plug, which prevents the blood from coming out of the end of the tube. If you use the wrong end of the tube you will contaminate the Plasticine®. The tube can be heat sealed in the flame of a Bunsen burner.

6. Action: Hold the tube vertically, sealed end down, and allow the blood column to run down.

    Rationale: Make sure that there is no evidence of a leak.

7. Action: Place the tube into one of the grooves of the microhaematocrit with the Plasticine® plug facing outwards towards the rim.

    Rationale: Centrifugal forces will cause the cells and fluid to spin outwards. The sealed end will prevent blood escaping.

8. Action: Place a similar tube on the opposite side of the centrifuge.

    Rationale: This balances the centrifuge and reduces vibration, although it is not essential in this type. If you are doing several tubes from a variety of patients then make a note of the number of each groove.

9. Action: Screw the safety plate over the tubes and close the lid.

    Rationale: The safety plate holds the tubes in place while they are spun. If you do not do this you will regret it!

10. Action: Set the timer for 6 minutes.

    Rationale: A microhaematocrit centrifuge spins at 10 000 r.p.m. and it takes 6 minutes to pack the erythrocytes properly.

11. Action: After 6 minutes allow the machine to stop naturally.

    Rationale: Avoid using the brake as this can damage the machine. Never attempt to open the machine while it is still running.

12. Action: Remove the capillary tube and check the colour and the thickness of the buffy coat. Write down your observations.

    Rationale: Check the plasma for evidence of haemolysis, jaundice and lipaemia. The buffy coat is made up of the leucocytes.

13. Action: Place the tube into the groove on the microhaematocrit reader (Fig. 5.4).

    Rationale: The blood will have separated into three layers (Fig. 5.4A):

14. Action: Adjust the tube vertically so that the bottom of the red cell layer is aligned with the 0%.

    Rationale: Make sure that you do not include the Plasticine® plug in your measurement.

15. Action: Slide the perspex plate so that the top of the plasma aligns with the 100% mark (Fig. 5.4B).

    Rationale: Use the bottom of the plasma meniscus as your measuring point.

16. Action: Adjust the reader handle on the left so that the line passes through the buffy coat / red cell junction.

    Rationale: This can be quite thick and pinpoint accuracy may be difficult.

17. Action: Read the measurement from the scale on the right hand side.

    Rationale: The scale on the reader runs from 0 to 100 and this is expressed as a percentage.

18. Action: PCV can also be calculated without the use of the reader by measuring the total length of the blood column (B) and the length occupied by the red blood cells (A) (Fig. 5.4A).

    Rationale: Do the following calculation:

1. Action: Select a new clean glass microscope slide and wipe it with lint-free tissue. If it has been soaked in methanol, rinse and dry it.

    Rationale: The slide must be as clean as possible to avoid the inclusion of dirt in the smear. Methanol removes grease and stops gaps appearing in the smear.

2. Action: Prepare a spreader by chipping the corner off another glass slide (Fig. 5.5). You may use a glass cutter to do this.

    Rationale: The use of the spreader prevents the smear overlapping the sides of the slide.

3. Action: Using a chinagraph pencil or an indelible felt pen, label the slide on the underside.

    Rationale: This identifies the slide. Marking it on the underside prevents the label being removed during staining.

4. Action: Take the EDTA tube containing the blood sample and gently rotate it between your finger and thumb. The tube must be at room temperature.

    Rationale: This resuspends the cells evenly within the plasma. Do not be overvigorous as this will damage the cells.

5. Action: Place the slide flat on the bench, preferably on a white background, with the long edges parallel to the edge of the bench.

    Rationale: The white background will help to show up the blood smear.

6. Action: Dip a capillary tube into the blood sample and allow it to collect a small amount of blood.

    Rationale: Blood will move up the capillary tube by capillary action.

7. Action: Place a small drop of blood on the right-hand end of the slide about 1 cm from its edge.

    Rationale: If you are left handed place the drop on the left-hand end. Too large a drop of blood will make the smear too thick; too small a drop gives too short a smear and / or ‘hesitation lines’.

8. Action: Place the spreader to the left of the blood at an angle of 45° to the horizontal and draw backwards to ‘pick up’ the blood (Fig. 5.5).

    Rationale: The blood will run along the edge of the spreader as soon as it makes contact with the blood. The angle of the spreader helps to determine the thickness of the smear.

9. Action: Move the spreader forwards towards the left-hand end of the slide in a single smooth movement so that the blood is smeared along the slide.

    Rationale: The blood is drawn along behind the spreader. If you were to push the blood along the slide without drawing back, as in step 6, you would damage the blood cells. The sides of the smear should be parallel and there should be a feathery ‘tail’. It should take up about of the slide.

10. Action: Dry the slide in the air by holding the sides between your finger and thumb and waving it gently.

    Rationale: The use of heat to dry the slide will damage the cells. Instant drying will preserve cell morphology.

11. Action: Make another blood smear.

    Rationale: It is always a good idea to make more than one smear in case one is unsatisfactory.

12. Action: Assess the quality of your smears.

    Rationale: The quality of the smear affects its use in diagnosis:

13. Action: Stain the most satisfactory smears.

    Rationale: Use an appropriate stain depending on what you need to know from the slide.

1. Action: Place a prepared blood smear with the smear uppermost, on a staining rack across a sink or staining bath.

    Rationale: This is a messy process and you must allow the stain to drain off into a sink or staining bath.

2. Action: Pour neat Leishman’s stain through a piece of filter paper on to the slide. Leave for 2 minutes.

    Rationale: There must be enough stain to cover the entire slide. Filtering the stain removes sediment, which could be mistaken for blood parasites.

3. Action: Do not wash off the stain, but add twice the volume of buffered distilled water (pH 6.8) and mix well. Leave for 15 minutes. A gold film may be seen within a few minutes.

    Rationale: This indicates that the pH is correct. The slide may be placed, using forceps, in a vertical position, in a Coplin jar – this prevents the build-up of sediment.

4. Action: Tip the stain off the slide and wash well, both front and back, with buffered water.

    Rationale: Do not use tap water as the changed pH will alter the staining characteristics of the cells.

5. Action: Wipe the underside of the slide with tissue and air dry.

    Rationale: Avoid touching the smear as you will wipe it off.

6. Action: Examine under the microscope using ×100 oil immersion.

    Rationale: This stain is useful for differential white cell counts as it provides good cellular definition.

1. Action: Place a prepared blood smear with the smear uppermost, on a staining rack across a sink or staining bath.

    Rationale: This is a messy process and you must allow the stain to drain off into a sink or staining bath.

2. Action: Using a pipette, flood the smear with methanol. Leave for 3–5 minutes.

    Rationale: This fixes the smear.

3. Action: Drain off the methanol and allow the slide to air dry.

    Rationale: This is essential before using the stain.

4. Action: Flood the slide with neat Giemsa stain pouring it through a piece of filter paper. Leave for 30 seconds.

    Rationale: Filtering the stain removes sediment, which may be mistaken for blood parasites.

5. Action: Using forceps transfer the slide to a Coplin jar containing Giemsa stain diluted 1 : 3 with buffered distilled water pH 6.8. Leave for 15–20 minutes.

    Rationale: The slide is held in a vertical position in the Coplin jar, which prevents the build-up of sediment on the slide.

6. Action: Remove the slide from the Coplin jar and rinse on both sides with buffered distilled water.

    Rationale: Do not use tap water as the changed pH will alter the staining characteristics of the cells.

7. Action: Wipe the underside of the slide and allow to air dry. Do not touch the top of the slide.

    Rationale: You may remove the stained smear.

8. Action: Examine under the microscope using ×100 oil immersion.

    Rationale: This stain is used to identify blood parasites and can also be used for differential white cell counts.

1. Action: Place a prepared blood smear stained in either Leishman’s or Giemsa under a light microscope.

    Rationale: Both Leishman’s and Giemsa provide good cellular definition for this procedure.

2. Action: Examine under ×10 objective lens.

    Rationale: This enables you to select an area that is one-third of the way from the tail end of the smear on the side edge.

3. Action: Scan over the slide and assess the distribution of the white cells.

    Rationale: If the distribution is uneven or clumped then it may be better to make another smear.

4. Action: Select an area near to the tail of the smear and apply a drop of immersion oil and move the ×100 objective into the oil.

    Rationale: See Figure 5.2. This increases the magnification.

5. Action: Adjust the condenser as described previously.

    Rationale: This provides maximum illumination.

6. Action: When focusing use the fine focus knob.

    Rationale: If you use the coarse focus knob the incremental movements mean your view moves out of focus very quickly.

7. Action: Move the slide so that it follows the line of a ‘battlement’ (Fig. 5.6) as follows: scan two fields along the edge of the smear, two fields down into the smear, two fields along, two fields back up towards the edge and so on.

    Rationale: This movement describes the battlement of a castle. By doing this you cover a wide area of the smear and it prevents biased cell distribution on the slide.

8. Action: While you are examining the fields count 200 white blood cells.

    Rationale: White blood cells have nuclei, which are stained in varying shades of pink or purple depending on their type (Fig. 5.7).

9. Action: Identify each type of white blood cell and record it (Fig. 5.7).

    Rationale: This will form the basis of your blood count. You may record your results manually on paper or by using some form of commercial counter.

10. Action: You can also scan the slide by travelling in a straight line along the length of the smear starting at the tail.

    Rationale: The tail is the thinnest part of the smear and the cells may be more clearly seen in this area. This method achieves the same final result as the battlement method.

11. Action: Using the records of each type of white blood cell, calculate the percentage of each type. The more cells you count in total the more accurate your results will be.

    Rationale: For example if you have 144 neutrophils in a sample of 200 then the percentage is

12. Action: Compare your results with the normal ranges shown in Table 5.2.

1. Action: Using a pipette, dispense 2 ml of methylene blue stain into a centrifuge tube.

    Rationale: Ensure that the stain is new methylene blue. McFadyean’s methylene blue is used to stain and identify Bacillus anthracis, the causal organism of anthrax.

2. Action: Using a pipette, add 4–5 drops from a well-mixed sample of blood in an EDTA tube.

    Rationale: Do not use heparin for this procedure.

3. Action: Mix the two components gently in the tube and place in an incubator at 37°C for 30 minutes.

    Rationale: If no incubator is available, the sample can be incubated at room temperature.

4. Action: Remove from the incubator and place in a centrifuge.

    Rationale: Make sure that you balance the centrifuge with a tube of similar weight as described previously.

5. Action: Spin at 1000 r.p.m. for 1 minute.

    Rationale: If you spin for longer or faster you will damage the cells.

6. Action: Remove and discard the supernatant.

    Rationale: This is not needed for the procedure.

7. Action: Gently flick the tube with your fingers and use this remaining liquid to make a smear as previously described; allow to air dry.

    Rationale: This resuspends the cells in any remaining liquid.

8. Action: Place the slide under the microscope and examine under ×10 and ×40 objective lenses. Look for red blood cells with dark-blue-stained strands in them (Fig. 5.8).

    Rationale: The dark-blue strands are the remnants of the endoplasmic reticulum within the cell and they are known as Howell-Joly bodies (Fig. 5.8).

9. Action: Select an area where individual cells can be easily seen and count a total of 500 cells within the smear. Make a note of how many reticulocytes you see.

    Rationale: If individual cells can be seen clearly this indicates that the smear is one cell thick.

10. Action: Using this information, calculate the percentage of reticulocytes.

    Rationale: To calculate:

11. Action: A correction factor is now applied.

    Rationale: To measure accurately the responsiveness of the bone marrow, the PCV of the patient is used.

12. Action: The final figure provides an indication of how the bone marrow is responding to the clinical situation.

    Rationale: To calculate:

1. Action: Place the dog in a sitting position or in sternal recumbency on an examination table covered in a non-slip mat.

    Rationale: If the dog feels secure and comfortable it will be less inclined to move or to try and escape.

2. Action: Ask the owner to restrain the dog appropriately.

    Rationale: This is not really painful, but the dog should be prevented from jumping off the table during the procedure. Local anaesthetic must not be used.

3. Action: Clip the hair from the area of skin to be used. Include a reasonable area around the site.

    Rationale: In dogs the most common site is the ear pinna.

4. Action: Gently clean the area with surgical spirit and allow to dry.

    Rationale: Take care to avoid causing hyperaemia of the area by overvigorous rubbing.

5. Action: Using a sterile scalpel blade, make an incision about 1 cm long and 2 mm deep and start the stopwatch.

    Rationale: This provides a base from which to measure the clotting time.

6. Action: After 15 seconds, touch a piece of filter paper to the blood so that it absorbs into the paper.

    Rationale: Avoid touching the skin.

7. Action: Repeat 15 seconds later using a fresh area of filter paper.

    Rationale: If using a circular piece of filter paper, move it around 1 cm at a time every time you touch the blood.

8. Action: Repeat the action until no more blood oozes from the incision. Stop the stopwatch.

    Rationale: The time on the stopwatch is the bleeding time and it is normally less than 5 minutes.

1. Action: The patient will normally be under a general anaesthetic.

    Rationale: This depends on the site and on the degree of pain – if the lesion is superficial the patient may be sedated, or fully conscious and well restrained.

2. Action: Preparation of the area must be as aseptic as possible.

    Rationale: To prevent the spread of infection.

3. Action: Make a cut across the surface of the tissue.

    Rationale: This will expose fresh uncontaminated cells from deep within the lesion.

4. Action: Blot until dry with a piece of paper towel.

    Rationale: If the tissue is too wet the cells may not stick and may run off the glass slide.

5. Action: Take a precleaned glass slide and roll the tissue against it.

    Rationale: All equipment must be clean and / or sterile to prevent the spread of infection. Rolling the tissue against the slide preserves the shape of the cells.

6. Action: Make several imprints on each slide.

    Rationale: This ensures that you obtain the maximum chance of understanding what is happening within the lesion.

7. Action: Allow the slides to air dry and stain.

1. Action: Place the patient in a comfortable position on the table with the area to be sampled uppermost and accessible.

    Rationale: If the technique is to be performed on a superficial mass then the patient may be either conscious or sedated and firmly restrained. If the area to be sampled is an abdominal mass then the patient may need to be sedated or under general anaesthesia.

2. Action: Clip the hair from the selected site and prepare the area aseptically.

    Rationale: To reduce the chance of introducing infection.

3. Action: For superficial masses, fix the mass firmly with one hand and insert the needle with the other hand.

    Rationale: Use 21–23G (–3″) needles depending on the depth of tissue to be sampled.

4. Action: If internal organs such as the liver, spleen and kidney or intra-thoracic or intra-abdominal organs are to be sampled, guidance of the needle by ultrasound may be necessary

    Rationale: It is impossible to fix these organs with one hand and there is a risk of penetrating another organ. Use 21–23G (–3″) needles as appropriate.

5. Action: If an internal organ such as the liver, spleen or kidney is to be sampled, haemorrhage may be a problem. In such cases use a Tru-cut® needle (Fig. 5.9).

    Rationale: A Tru-cut® needle is used to collect tiny samples. The needle has a small notch cut out of its barrel and a cover that slides down trapping the sample. This can then be removed, fixed and examined by cytology or histopathology.

6. Action: For a true FNA, when the needle is inserted attach a 5–10 ml syringe and pull back on the plunger several times while moving the needle to and fro within the lesion. Do not withdraw the needle.

    Rationale: This action helps to exfoliate cells from around the needle so that they can be drawn up into the needle. There is a risk of damaging fragile cells and increasing the amount of blood in the sample.

7. Action: When you are satisfied that you have collected enough cells, withdraw the needle and syringe from the site.

    Rationale: This keeps the harvested cells within the needle. Creating too much negative pressure by withdrawing the needle and syringe with the plunger half-way up would cause the cells to rush into the barrel of the syringe as the needle was withdrawn from the lesion. If there were relatively few cells in the sample, you might then lose them within the syringe.

8. Action: Detach the syringe from the needle and attach a clean air-filled syringe.

    Rationale: This will ensure that only the contents of the needle are expelled.

9. Action: Depress the plunger and expel the contents of the needle onto one or more clean microscope slides.

    Rationale: Ensure that all the cells are expelled.

10. Action: Prepare smears as described below and examine under a microscope.

1. Action: Place the animal in a sitting position on a stable examination table covered in a non-slip mat.

    Rationale: If the animal feels secure it will be less likely to try and escape.

2. Action: Ask the owner or an assistant to restrain the animal so that you have access to the ear.

    Rationale: See Chapter 1. This technique is not painful, but if an animal has an inflamed ear it may resent the procedure.

3. Action: Using a clean cotton bud soaked in a little sterile saline, insert it into the external ear canal and move it gently over the surface of the canal.

    Rationale: The aim is to pick up lining cells without further traumatizing the delicate epithelium.

4. Action: Roll the cotton bud over the surface of a clean microscope slide.

    Rationale: This deposits the cells harvested from the ear canal on the slide.

5. Action: If you have picked up a great deal of material you may have to spread or tease it out.

    Rationale: See the starfish technique below. The aim is to produce a thin layer of cells that can be easily stained and identified.

6. Action: Allow the slide to air dry and stain using Diff-Quik®.

    Rationale: Read the manufacturer’s instructions.

7. Action: Place a drop of microscope oil on the slide and then place a cover slip on top.

    Rationale: This helps to protect and spread the layer of cells.

8. Action: Place the slide under the microscope and examine under ×100 oil immersion lens.

    This technique can be used to diagnose the presence of Malassezia pachydermatis, although you may also find bacteria and Otodectes mites, which will more easily be seen under ×4 or ×10 magnification. Gram stain is used to differentiate between Gram negative and Gram positive bacteria.

1. Action: Expel the cells within the needle onto the centre of a clean glass microscope slide.

    Rationale: Ensure that all the cells are expelled.

2. Action: Place another clean glass slide (the spreader) at right angles to the first and apply gentle pressure (Fig. 5.10).

    Rationale: This aids spreading, but too much pressure may rupture the cells.

3. Action: Draw the spreader quickly and smoothly across the bottom slide.

    Rationale: The underside of this slide will pick up the cells making a useable smear.

4. Action: Allow the smear on the spreader to air dry.

    Rationale: Do not use heat because the cells will be damaged.

5. Action: Stain appropriately.

    Rationale: Choice of stain depends on what you are looking for.

1. Action: Expel the aspirate at one end of a clean glass microscope slide.

    Rationale: This gives maximum space on the slide to make the smear.

2. Action: Using another glass slide, break off one corner using a glass cutter – this creates a spreader.

    Rationale: The use of this shaped spreader to prepare the smear avoids spreading the cells over the edge of the slide.

3. Action: Place the spreader in front of the aspirate at an angle of approx. 30° and draw it backwards until the spreader picks up the aspirate (see Fig. 5.5).

    Rationale: This is the same technique as has been described for preparing a blood smear. The aspirate will flow along the spreader.

4. Action: Push the spreader quickly and smoothly along of the slide.

    Rationale: The spreader will draw the aspirate, smearing the cells along the slide.

5. Action: At this point, abruptly lift the spreader upwards.

    Rationale: This creates a line in which the cells within the smear are concentrated. This is different to the true blood smear technique.

6. Action: Allow to air dry and stain appropriately.

    Rationale: Choice of stain depends on what you are looking for.

1. Action: Expel the aspirate onto a clean glass microscope slide.

    Rationale: Ensure that all the cells are expelled.

2. Action: Using the point of the needle, drag the aspirate in several directions.

    Rationale: This method tends not to damage the cells. The aim is to draw out the cells to create a thin layer.

3. Action: Continue until the shape resembles that of a starfish with many projections (Fig. 5.11).

    Rationale: The thick layer of fluid around the cells may prevent evaluation of the cellular detail.

4. Action: Allow to air dry and stain appropriately.

    Rationale: Choice of stain depends on what you are looking for.

1. Action: Prepare slide as described above and allow to air dry.

    Rationale: Use of heat may damage the cells.

2. Action: Place the slide on a draining rack over a sink or over a staining bath.

    Rationale: This allows the stain to drain away without causing a mess.

3. Action: Flood the slide with Sudan 3 stain and leave for 3 minutes.

    Rationale: Sudan 3 binds with the fat in the sample.

4. Action: Wash gently with tap water.

    Rationale: This stops the action of the stain and removes any surplus.

5. Action: Place a coverslip over the smear.

    Rationale: This prevents the objective lens from being contaminated.

6. Action: Examine under the microscope.

    Rationale: Fat globules will stain orange.

1. Action: Label the Petri dish on the underside with a marker pen using the client’s name and the animal’s name or the case number.

    Rationale: All agar plates look the same and it is important not to mix up the samples.

2. Action: Unwrap a single-use disposable loop and dip it into the sample.

    Rationale: These are ready sterilized. The use of platinum loops is discouraged as sterilizing, them by passing them through a Bunsen burner flame may cause aerosol spread of bacteria.

3. Action: Pick up the half of the Petri dish containing the agar gel and turn it over so that the surface of the gel is uppermost.

    Rationale: Petri dishes have a base containing the set agar gel and a lid that looks very similar.

4. Action: Smear the material on the loop over a small area on the left of the agar (Fig. 5.12).

    Rationale: This ‘well’ is the start of your inoculation area.

5. Action: Replace the lid of the Petri dish.

    Rationale: It is not a good idea to leave the dish uncovered for too long as organisms in the air may contaminate it.

6. Action: Unwrap another loop.

    Rationale: To make sure that the only bacteria on the plate are from the original sample.

7. Action: Pick up the Petri dish as before and using the loop, make 3–4 streaks, all in the same direction from the edge of the well. Take care not to tear the agar (Fig. 5.12).

    Rationale: This technique helps to spread the contents of the sample evenly over the plate.

8. Action: Continue to spread the sample over the agar as shown in Figure 5.12.

    Rationale: The aim is to spread the sample ever more thinly so that single colonies will grow in the area of the final stroke.

9. Action: Place the lid back on the dish and put it in the incubator upside down (i.e. with the agar side on top).

    Rationale: If the lid is uppermost then water droplets, formed by condensation during incubation, will drip onto the agar causing the bacterial colonies to spread.

10. Action: Do not stack the agar plates more than two or three high in the incubator.

    Rationale: Overcrowding will prevent free circulation of air within the incubator.

11. Action: Incubate at 37°C for 18–24 hours.

    Rationale: Most pathogenic bacteria grow at normal body temperature (i.e. 37°C).

12. Action: Remove from the incubator and examine for bacterial colonies.

    Rationale: Colonies of bacteria appear as raised ‘bumps’ along the streak lines. Take note of their colour and any change in the agar around them as this may help in their identification.

13. Action: If there is no visible growth, incubate the plate for another 18–24 hours.

    Rationale: Some bacteria take longer to grow than others.

1. Action: Prepare a bacterial culture from the patient’s sample as described previously.

    Rationale: This test cannot be done straight from the sample.

2. Action: Take a single-use disposable loop from its packaging.

    Rationale: These loops are designed to be used once and are presterilized.

3. Action: Holding the Petri dish as described above, use the loop to pick up an isolated colony and transfer it to a buffered peptone broth.

    Rationale: If there appears to be more than one type of colony growing on the plate you will have to test each one separately, but start with the predominant type.

4. Action: Incubate the peptone broth at 37°C for 4 hours.

    Rationale: Buffered peptone broth is available commercially and enhances bacterial growth, particularly of more fastidious organisms such as Pasteurella and Corynebacterium.

5. Action: Immerse a sterile swab in the broth and use it to completely cover a Mueller Hinton blood agar plate.

    Rationale: Using a swab ensures good coverage of the agar. Mueller Hinton blood agar contains sheep’s blood and is available commercially. It is the standard agar for sensitivity tests.

6. Action: Pick up the antibiotic assay ring with forceps or tweezers and place it in the centre of the agar (Fig. 5.13).

    Rationale: The ring will have several antibiotic impregnated discs attached to it. Each disc is impregnated with a specific antibiotic (e.g. erythromycin, methicillin, tetracycline, etc.) and is printed with an identifying code. Some tests use individual antibiotic discs that are dropped onto the agar. Avoid handling the ring or discs with your fingers as this may contaminate them.

7. Action: Place the lid back on the dish and put it in the incubator upside down (i.e. with the agar side on top).

    Rationale: If the lid is uppermost then water droplets, formed by condensation during incubation, will drip onto the agar causing the bacterial colonies to spread.

8. Action: Do not stack the agar plates more than two or three high in the incubator.

    Rationale: Overcrowding will prevent free circulation of air within the incubator.

9. Action: Incubate at 37°C for 18–24 hours.

    Rationale: Most pathogenic bacteria grow at normal body temperature (i.e. 37°C).

10. Action: Remove from the incubator and examine for bacterial growth.

    Rationale: If the bacteria are sensitive to a particular antibiotic, growth will be suppressed, which appears as a clear zone around the relevant disc (Fig. 5.13).

11. Action: Record your results.

    Rationale: This information will allow you to prescribe the antibiotic to which the causal organism is most sensitive, thus promoting a rapid recovery.

1. Action: Prepare a glass microscope slide by cleaning it with alcohol and allowing it to dry.

    Rationale: This removes any residues or grease that may be sticking to the slide.

2. Action: Unwrap a single-use disposable loop and dip it into the saline.

    Rationale: These are ready sterilized. The use of platinum loops is discouraged as sterilizing them by passing them through a Bunsen burner flame may cause aerosol spread of bacteria.

3. Action: Use the loop to place 2–3 drops of sterile normal saline near to the centre of the slide.

    Rationale: This will dilute the bacterial colony and make it easier to spread.

4. Action: Unwrap another loop.

    Rationale: To ensure that it is sterile.

5. Action: Using the loop, select an isolated bacterial colony and carefully remove it from the agar plate.

    Rationale: If there appears to be more than one type of colony growing on the plate you will have to make a smear for each type.

6. Action: Place the colony on the slide close to the saline and mix the two together using the loop.

    Rationale: Mixing with saline results in a single layer of cells, allowing easier identification.

7. Action: Spread the mixture out over the slide to cover 1–2 cm.

    Rationale: A thin layer of cells enables you to see the shape of the cells.

8. Action: Dry the bacterial smear by passing the slide with the smear side uppermost through a Bunsen burner flame two or three times.

    Rationale: This fixes the slide. If the slide is stained while it is still wet then some of the bacteria will float off. Be careful not to overheat the slide.

9. Action: Stain the smear using Gram’s stain or methylene blue.

    Rationale: These stains show up the shape of the bacteria.

1. Action: Place the prepared slide with the smear facing uppermost on a staining rack over a sink or staining bath.

    Rationale: Staining is a messy process and the surplus stain must be able to drain away. You can also stain using upright Coplin jars.

2. Action: Using a pipette, flood the slide with crystal violet for 30 seconds.

    Rationale: At this point, both types of bacteria absorb the stain and become purple.

3. Action: Wash the slide with tap water.

    Rationale: To remove the crystal violet stain.

4. Action: Flood the slide with iodine for 60 seconds.

    Rationale: This acts as a mordant, fixing the stain. Gram’s iodine is stronger than Lugol’s iodine and is the stain of choice.

5. Action: Flood the slide with acetone for 2–3 seconds.

    Rationale: This decolourizes the smear and is very rapid. Gram negative bacteria lose their colour.

6. Action: Wash the slide with water.

    Rationale: To remove the acetone.

7. Action: Flood the slide with carbol fuchsin for 30 seconds.

    Rationale: This counter-stains the bacteria and Gram negative bacteria become pink.

8. Action: Wash the slide with tap water.

    Rationale: To remove the stain.

9. Action: Wipe the back of the slide.

    Rationale: Do not wipe the smear off the front of the slide.

10. Action: Pass the slide rapidly over the flame of the Bunsen burner.

    Rationale: To dry the slide. Do not overheat the slide as it will shatter.

11. Action: Place the slide under the microscope and examine under high power.

    Rationale: Bacteria range in size from 0.5 µm to 5.0 µm so use ×100 oil immersion lens.

1. Action: Place the prepared slide with the smear facing uppermost on a staining rack over a sink or staining bath.

    Rationale: Staining is a messy process and the surplus stain must be able to drain away. You can also stain using upright Coplin jars.

2. Action: Using a pipette flood the slide with methylene blue stain and leave for 3 minutes.

    Rationale: This stains the bacterial cells.

3. Action: Wash the slide with tap water from the water bottle.

    Rationale: This removes the excess stain from the slide.

4. Action: Wipe the back of the slide.

    Rationale: Do not wipe off the smear from the front of the slide.

5. Action: Pass the slide rapidly over the flame of the Bunsen burner.

    Rationale: To dry the slide. Do not overheat the slide as it will shatter.

6. Action: Place the slide under the microscope and examine under high power.

    Rationale: Bacteria range in size from 0.5 µm to 5.0 µm so use ×100 oil immersion lens.

1. Action: Select the correct type of dipstick test.

    Rationale: Reagent pads vary in number and type according to manufacturer.

2. Action: Check the expiry date.

    Rationale: Out of date sticks may give unreliable results.

3. Action: Remove the lid of the container, remove one stick and replace the lid.

    Rationale: Avoid contaminating the rest of the sticks with urine.

4. Action: Dip the stick in the fresh urine sample until all the pads are wet.

    Rationale: Never perform this test on stale urine as the results may be affected.

5. Action: Remove the stick from the sample and gently tap it on the side of the sample pot.

    Rationale: To remove any surplus urine.

6. Action: Using a timer or the second hand on your watch, keep to the time specified on the side of the dipstick bottle to take your reading.

    Rationale: Each test pad requires a specific time in contact with the urine before it reacts appropriately.

7. Action: Hold the dipstick bottle in one hand and the dipstick in the other and compare the colour of each pad with the correct one on the side of the container.

    Rationale: Each reagent pad will change colour. The range of colour changes is illustrated on the side of the bottle along with the appropriate result.

8. Action: Check that you read the results from the correct end of the stick.

    Rationale: If you read from the wrong end you will get results that are incorrect for each test pad.

9. Action: Write down your results.

    Rationale: Do not rely on your memory – the results may need to be kept for some time before you use them to assist your diagnosis.

10. Action: Dispose of the dipstick in the clinical waste.

    Rationale: There may be pathogens on the stick.

1. Action: To calibrate the refractometer, wipe the glass prism (Fig. 5.14) with a piece of lint free tissue. Using a pipette place approximately 2 drops of distilled water onto the prism and close the lid.

    Rationale: The refractometer must be calibrated every time you use it to ensure that it measures the specific gravity accurately.

2. Action: Hold the refractometer up to the light and look through the eyepiece. Read the scale visible through the eyepiece.

    Rationale: The reading is taken at the point where there is a boundary between light and dark. As the liquid is pure water it should read 1.000.

3. Action: If the reading is not 1.000 then adjust the refractometer using the screw on the top of the instrument and the screwdriver provided. Adjust until the reading is 1.000.

    Rationale: Some instruments have different ways of making the adjustment – read the instruction leaflet.

4. Action: Wipe the water off the glass with lint free tissue.

    Rationale: Lint free tissue should not leave any fibres on the glass.

5. Action: Using a clean pipette, place 2 drops of the urine sample on to the glass and close the lid.

    Rationale: Using the same pipette may dilute the urine sample. The instrument is now set up to read the specific gravity of urine.

6. Action: Hold the refractometer up to the light, look through the eyepiece and read the scale. Record your result.

    Rationale: Normal specific gravity for dog, cat and rabbit urine is shown in Table 5.5.

7. Action: Rinse off the urine with distilled water and wipe with lint free tissue and replace the instrument in its case.

    Rationale: Do not leave urine to dry on the glass. Refractometers are expensive precision instruments and should be cleaned and stored correctly.

1. Action: Using a pipette, place 3–5 ml of fresh urine in a centrifuge tube.

    Rationale: A conical centrifuge tube helps to separate the sediment from the supernatant.

2. Action: Place the tube in the centrifuge and balance it with a similar tube on the other side.

    Rationale: The machine must be balanced to prevent vibration.

3. Action: Set the centrifuge to spin at 1000–2000 r.p.m. for 2–3 minutes.

    Rationale: If you spin for longer you may damage any cells in the urine.

4. Action: Use a pipette to remove and discard most of the supernatant. Be careful not to disturb the sediment.

    Rationale: Only the sediment is needed for the examination.

5. Action: Flick the test tube with your fingers.

    Rationale: This resuspends the sediment in any remaining liquid and makes a more even smear.

6. Action: Add 1–2 drops of Sedistain® to the sediment in the tube.

    Rationale: This is optional, but it helps to make the contents of the sediment more visible and easier to identify. The test can be performed unstained.

7. Action: Use a pipette to remove 1–2 drops of the sediment and place them onto the centre of a clean glass slide. Cover with a coverslip.

    Rationale: This makes a uniform layer for examination.

8. Action: Place the slide under the microscope and examine using ×10 and ×40 objective lenses.

    Rationale: Look for evidence of blood cells, epithelial cells, crystals (Fig. 5.15), casts, sperm, mucin threads and bacteria.

1. Action: Place some of the skin scraping on to the centre of a clean glass microscope slide.

    Rationale: Do not use too much as the debris may mask the parasites.

2. Action: Use a pipette to add 2–3 drops of 10% potassium hydroxide to the debris.

    Rationale: Be careful – this is caustic!

3. Action: Place a coverslip over the sample.

    Rationale: This prevents the microscope lens from becoming contaminated and makes a uniform layer to examine.

4. Action: Holding the slide with forceps, warm it gently over the Bunsen burner flame – do not let it boil!

    Rationale: Warming the debris breaks it down so that it becomes clear, making the parasites easier to see.

5. Action: Allow the slide to cool and place it under the microscope. Examine using the ×4 and ×10 objective lenses.

    Rationale: Larger parasites such as Sarcoptes may be seen with low magnification whereas smaller species such as Demodex require higher magnification (Fig. 5.16).

6. Action: It is a good idea to prepare several slides for examination.

    Rationale: Numbers of parasites are likely to be low and they may be missed if you examine only one smear.

1. Action: Wearing disposable gloves, collect a sample from the patient by plucking hair from the edge of the lesion.

    Rationale: Ringworm fungi cause circular lesions that spread outwards. The centre of the lesion is dead and scaly but the outer ring should contain live fungus.

2. Action: Place a pair of forceps in the flame of the Bunsen burner for a few seconds and then allow to cool to room temperature.

    Rationale: The forceps must be sterilized to reduce the risk of bacterial contamination.

3. Action: Remove the lid of a Petri dish containing Sabouraud’s agar or a commercial medium such as Dermatofyt®.

    Rationale: Avoid contaminating the agar with your fingers.

4. Action: Use the forceps to take 6–8 hairs from the sample and place them in the centre of the agar. Replace the lid.

    Rationale: Avoid contamination from micro-organisms, including fungal spores, floating in the atmosphere. Sabouraud’s medium is selective for ringworm fungi.

5. Action: Incubate at room temperature for 28 days.

    Rationale: Ringworm fungi grow very slowly, although you may see some evidence of growth from the 4th day.

6. Action: When fungal growth is visible, examine and identify.

    Rationale: Growth appears as a fluffy white colony on Sabouraud’s medium. On commercial media such as Dermatofyt®, an indicator within the agar changes from yellow to red. However, there are some contaminants that may also cause this colour change so it is important to identify the fungus under the microscope.

• Microsporum canis – flat with white silky centre and a bright yellow edge. Underside is yellow.

• Microsporum gypseum – flat brown with a powdery yellow fringe. Underside is yellow / brown.

• Trichophyton mentagrophytes – flat, granular and tan coloured, or heaped with a white cottony appearance. Underside is yellow / red.

1. Action: Culture a colony of fungus from the patient’s sample as described.

    Rationale: Infected hairs can be used to make a direct slide, but identification is more difficult and requires experience.

2. Action: Take a clean glass microscope slide and, using a pipette, place 2–3 drops of lactophenol cotton blue stain in the centre.

    Rationale: The stain helps to spread the fungal colonies over the slide. The phenol in the stain inactivates the fungal spores and the blue stain aids visualization of the spores and the mycelia. You can use Quink® ink instead of the stain.

3. Action: Using a single-use loop, pick up a small amount of fungal colony and mix it with the stain on the slide. Discard the loop in the clinical waste.

    Rationale: To prevent contamination by the fungus on the loop.

4. Action: Place a coverslip over the material on the slide.

    Rationale: This provides a layer of uniform thickness for examination.

5. Action: Place the slide under the microscope and examine under the ×10 and ×40 objective lenses.

    Rationale: Microscopic examination will confirm the identification of the species.

• M. canis – spores are thick walled, boat shaped and have knob-like structures at the end.

• T. mentagrophytes – spores are cigar shaped and thin walled and the hyphae are spiral.

1. Action: Place the animal on a stable examination table in a darkened room.

    Rationale: The darkened room will enhance the fluorescent effect.

2. Action: Put on some form of eye protection.

    Rationale: UV light can cause damage to the conjunctiva and the retina.

3. Action: Switch on the Wood’s lamp and allow it to warm up for 5–10 minutes.

    Rationale: To ensure that it is emitting the correct wavelength of light.

4. Action: Ask the owner or an assistant (who should be wearing eye protection) to restrain the animal appropriately.

    Rationale: This is not a painful procedure, but the animal may try to escape.

5. Action: Hold the lamp over the suspect area and then screen the rest of the body thoroughly looking for signs of apple-green fluorescence.

    Rationale: Other particles in the coat (e.g. skin flakes, dirt and detergent) may fluoresce and petroleum jelly will fluoresce blue-white.

6. Action: When the examination is complete turn off the lamp.

    Rationale: Prolonged exposure to UV light can burn the skin.

1. Action: Take a small amount of faeces and place it in the centre of a clean glass microscope slide.

    Rationale: If the smear is too thick then it is difficult to see the eggs. The ideal smear should appear almost clear to the naked eye.

2. Action: Place 2–3 drops of water on the slide and mix with the faeces using a glass rod or a mounted needle. Keep the material as close to the centre of the slide as possible.

    Rationale: The water helps to dilute the faeces and makes a thinner solution.

3. Action: Remove any large pieces of material from the mixture.

    Rationale: These are likely to be undigested fibrous material, which will interfere with the coverslip.

4. Action: Carefully place a coverslip over the area of the sample. Try to avoid trapping bubbles under the slip.

    Rationale: Bubbles may be mistaken for eggs. Use a mounted needle to lower the edge of the slip onto the faeces.

5. Action: Make sure that the coverslip lies as flat as possible.

    Rationale: This forms a uniform layer to examine under the microscope. Any large pieces of debris will prevent the slip lying flat.

6. Action: Place the slide under the microscope and examine systematically under ×10 and ×40 objective lenses.

    Rationale: Systematic examination of the slide will ensure that you examine the whole slide. Air bubbles appear as black circles.

1. Action: Weigh 3 g of faeces into a glass beaker.

    Rationale: The faeces should be fresh and moist.

2. Action: Measure 45 ml of saturated sugar solution into a measuring cylinder, pour into the beaker and mix with a spatula.

    Rationale: Saturated sugar solution helps the eggs to float out of the faeces. The addition of several glass beads may help to break up the faecal matter – cat faeces are particularly hard to break up.

3. Action: Pour the solution through a sieve into a second beaker. Discard the debris in the sieve.

    Rationale: This removes any large particles but allows the eggs to go through.

4. Action: Mix the solution gently and allow to stand at room temperature for 5–10 minutes.

    Rationale: This enables the worm eggs to float to the top of the saturated sugar solution.

5. Action: If using a single chamber McMaster slide, prepare it by placing the coverslip grid side down on the slide.

    Rationale: Some chambers have a single chamber and use a coverslip grid whereas others have two chambers and an integral grid (Fig. 5.19).

6. Action: Using a pipette, withdraw approximately 2 ml of the solution and fill the counting chamber.

    Rationale: Make sure that you have enough fluid to completely fill the chamber. If there is insufficient fluid then bubbles may form over the grid.

7. Action: Apply the coverslip.

    Rationale: To avoid the inclusion of bubbles, which may distort the image, make sure that the coverslip makes contact with the chamber.

8. Leave the counting chamber on the bench for 5–10 minutes.

    Rationale: This allows any worm eggs or coccidial oocysts to float to the top, making them visible.

9. Action: Place the chamber under the microscope and examine using the ×10 objective. Focus on the grid and count all the eggs seen over the grid area.

    Rationale: Count eggs that lie over the lines and between the lines. If there is more than one type of egg then repeat the count for each type separately (Fig. 5.18). Air bubbles will appear as black circles.

10. Action: Calculate the number of eggs as follows:

    Rationale: This gives you the number of eggs per gram of faeces.

1. Action: Attach a short length of tubing to a funnel and clamp the funnel into a stand.

    Rationale: This should be as stable as possible so that it does not tip over when liquid is poured through.

2. Action: Place a clip across the end of the tubing so that it forms a watertight seal.

    Rationale: To prevent the faecal solution dripping out.

3. Action: Fill the funnel with lukewarm water to within about 1 cm of the rim.

    Rationale: This will soften the faeces and allow the larvae to run down to the bottom.

4. Action: Take a small piece of muslin and thread a strong wire through each side.

    Rationale: The loops will suspend the muslin across the top of the water.

5. Action: Place the muslin across the top of the funnel so that the wires lie across the top to suspend it.

    Rationale: The muslin forms a ‘lid’ that is held in place by the wires.

6. Action: Place about 10 g of faeces onto the muslin, which will now dip down into the water.

    Rationale: Make sure that the faeces sample is completely covered by the water.

7. Action: Leave the apparatus overnight.

    Rationale: This gives enough time for the larvae to migrate to the bottom of the funnel and enter the tubing.

8. Action: In the morning draw off a centrifuge tube of liquid by releasing the clip on the tubing.

    Rationale: If larvae are present they should accumulate in the tubing.

9. Action: Allow the tube to stand for 4 hours, or centrifuge at 1500 r.p.m. for 1 minute.

    Rationale: As the larvae are heavier than water, gravity or centrifugal force will pull them down to the bottom of the tube.

10. Action: Using a pipette, remove the supernatant fluid from the tube.

    Rationale: This should not contain anything significant.

11. Action: Place the sediment on a clean glass microscope slide, add a drop of Lugol’s iodine and cover with a coverslip.

    Rationale: Lugol’s iodine will kill the larvae and stop them moving. It also stains them brown, which increases visibility.

12. Action: Place the slide under the microscope and examine under ×10 and ×40 objective lenses.

    Rationale: Look for the presence of nematode larvae.