Laboratory techniques

Chapter 5


Laboratory techniques




Chapter Contents


Laboratory Equipment



Practical Techniques



Haematology



Cytology



Bacteriology



Urinalysis



Parasitology



Ringworm



Faecal analysis



Laboratory tests are used as an aid to diagnosis following taking a history and making a clinical examination. This evidence leads to a differential diagnosis and laboratory tests will then help to eliminate some possibilities and confirm others.


Some results will come back as normal; these are used to rule out some of the possible diagnoses and you can then pay more attention to the abnormal findings. Never discount an abnormal or unexpected result. It is easy to have a fixed idea of the diagnosis in your mind and to fit all the evidence to support it; however, if you think a result contradicts your diagnosis, repeat the test – it may have been an error due to inaccuracy, but it may be that the result is indicating something other than your original theory.


Many laboratory tests are carried out within the practice, although some will be sent away to a commercial laboratory; the proportion varies between practices, but most practices have a small ‘in-house’ lab. Some larger practices have designated lab technicians who do all the work, but it is important for the veterinary surgeon to know how to carry out the basic techniques, as there will be times, such as weekends or at night, when the support staff are not available.


As with all aspects of veterinary practice, the laboratory is subject to health and safety legislation. It is not the brief of this chapter to go into any detail except to say that precautions must be taken to ensure the safety of yourself and everyone around you. Make sure that you take steps to prevent yourself becoming infected by the samples; for example, wear protective clothing and eye protection, always wash your hands, avoid mouth pipetting and do not eat in the lab. Make sure that the lab itself is kept clean and tidy and that used equipment is cleaned immediately or disposed of in the correct manner (e.g. sharps bins or clinical waste bags).


Laboratory tests must be carried out correctly and accurately. Remember that the quality of your result reflects the quality of your sample – correct methods of collection, preservation and storage play a significant part. There are some rules that should be observed for all tests:




Laboratory equipment


A routine practice laboratory will have the following pieces of equipment.



The microscope


The type of microscope found in a practice laboratory is the light microscope, which will have an inbuilt light source, although older ones may use a mirror and an external light source. Microscopes are expensive precision instruments and should be handled with care. They may be monocular or binocular, but both types are equally effective.


General rules for the care of microscopes are shown in Box 5.1.



Box 5.1   General rules for the care of light microscopes






Procedure: Care and use of the microscope (Fig. 5.1)




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.



Procedure: Use of the oil immersion lens

When used correctly, this technique increases the magnification to the maximum achievable with a light microscope. This is required for the examination of bacteria and blood smears.



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.



Procedure: Use of the Vernier scales

On most microscopes these scales are located on both sides of the stage, arranged at right angles to each other – known as the vertical and horizontal axes. Their function is to allow location of a particular point on the slide so that you can find it again (Fig. 5.3).




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.



The centrifuge


Centrifuges are an essential piece of laboratory equipment and, although they may vary, they all work on the principle of centrifugal force. Samples enclosed in special containers are spun around at speed, which results in the heavier particles settling at the bottom while the lighter ones go to the top. This would occur naturally in response to gravity if the container was left alone for some time, but the centrifuge accelerates the process.


The use of a centrifuge is required in many tests including:



There are two main types:



1. Angle head – the specimen tubes are held in a fixed position, usually at 25–50° to the vertical. Higher rotational speeds can be achieved because of the aerodynamic shape of the rotor; however, because of the angulation of the tubes, the sediment settles at this angle making it difficult to remove.


2. Swing-out head – the specimen tubes are placed in buckets that swing out from the vertical to the horizontal as the speed of rotation increases. As the machine stops the buckets return to the vertical position. The surface of the sediment is level, which means that the supernatant can be more easily removed with a pipette.


    A microhaematocrit centrifuge, which is required to spin down blood samples prior to doing a packed cell volume estimation (PCV), consists of a metal plate on which fixed horizontal grooves carry the microhaematocrit capillary tubes.




Procedure: Care and use of the centrifuge



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.



Electronic analysers


Many veterinary practices now use various types of electronic analyser to assist in the diagnosis of their clinical cases. These include those designed to analyse biochemistry, haematology, electrolytes and hormones. The variety of design makes it impossible to accurately describe their use within this book; however, you should take note of the following:




Practical techniques



Haematology


This is the study of the cellular elements of the blood and its associated clotting factors. Many haematology tests are nowadays done using a haematology analyser which automatically determines red and white cell counts, differential white cell counts, packed cell volume, haemoglobin levels and platelet counts; however, it is important to be able to know how to perform some of the more basic tests.



Blood collection


The sites and methods of restraint for the collection of blood from dogs, cats and rabbits have been described in Chapter 1, but once the sample has been collected it must be put into some form of container to preserve it and, if required, to prevent it from clotting. There are several forms of container:



1. Screw-topped tubes – these are usually made of plastic and have lids whose colour conforms to a standard colour code indicating the type of anticoagulant or preservative that they contain. This is shown in Table 5.1. Some brands may have push-on tops.



    These tubes are cheap and require only a small amount of blood to fill them. Remove the needle from the syringe and quickly transfer the blood into the tube. Immediately replace the cap and then mix the contents by repeated inversion unless you require a clotted sample. If you forcefully squirt the blood through a small gauge needle or shake the tube to mix it up you run the risk of breaking the blood cells resulting in a haemolysed sample.


2. Evacuated glass tubes (Vacutainers®) – these consist of an evacuated glass tube sealed with a colour-coded rubber bung. They require the use of a double-pointed needle, which fits into a special needle holder. One end of the needle is used to penetrate the vein while the other end penetrates the rubber bung. The vacuum within the tube draws blood out of the vein into the tube. These are expensive and are more widely used in large animal practice.


3. Plastic blood-collecting syringes (Monovette®) – these are syringes that are designed for aspirating blood and contain anticoagulant. Once blood has been collected the needle is replaced with a cap and the plunger is unscrewed creating a leak-proof container.



Plasma vs serum


Blood consists of the cellular fraction consisting of erythrocytes (red cells), leucocytes (white cells) and thrombocytes (platelets) and the fluid part – the plasma or serum.




Procedure: Packed cell volume

Packed cell volume (PCV) is used to measure the volume of the erythrocytes in whole blood when packed tightly together. It is used to assess the degree of anaemia and of dehydration in a patient.


Equipment list: Blood sample collected in an EDTA tube, plain capillary tubes, microhaematocrit centrifuge and reader, soft Plasticine® or Cristaseal®.




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 image 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:



NB PCV ranges for the dog, cat and rabbit are shown in Table 5.2, and normal PCV values are shown in Table 5.3.




Table 5.3


Red blood cell indices in the dog, cat and rabbit*



























Name / definition Measurement Normal values
Packed cell volume (PCV): percentage of packed red cells in a sample Centrifuge capillary tube containing blood Dog: 37–57%
Cat: 27–50%
Rabbit: 34–50%
Haematocrit (HCT): percentage of blood composed of red cells (often interchanged with PCV) HCT = MCV × total RBC (1012/l)
Less accurate than PCV		 —
Haemoglobin (Hb): amount of Hb within red cells – estimation of O2-carrying capacity Estimated using haematology analyser Dog: 12–18 g/dl
Cat: 8–15 g/dl
Rabbit: 10–17.5 g/dl
Mean corpuscular volume (MCV): measure of red cell size MCV (fl) = (PCV × 1000)/total RBC (1012/l)
Measured directly by analysers		 Dog: 70 fl
Cat: 45 fl
Rabbit: 69 fl
Mean corpuscular haemoglobin concentration (MCHC): average concentration per red blood cell MCHC (g/dl) = total haemoglobin (g/dl) /PCV 35 g/dl for all species

*Taken from a range of sources. fl = femtolitre: 1 fl = 10−15l.


Total white blood cell count (TWBC) and total red blood cell count (TRBC) are useful diagnostic parameters (Table 5.2), but are nowadays usually done by an electronic haematology analyser. They used to be done using a Neubauer haemocytometer, but this takes time and experience. Commercial labs can get the result back to the practice within 12 hours so manual analysis is rarely done.



Procedure: Preparation of a blood smear

Blood samples can be examined and preserved by smearing a drop of blood onto a glass slide. The skill of preparing a blood smear can be achieved by practice, and it is worth the effort as a smear can provide a great deal of diagnostic information. They may be used to evaluate the relative proportions of the cellular components of blood and even to check the cell counts done by a machine. They may also be used to indicate the presence of cellular abnormalities, provide a rough estimation of platelet numbers and identify the presence of blood parasites.


Blood smears may be fixed by air drying or, if examination is to be delayed, by immersing in 100% methyl alcohol for 1 minute. If the smear is to be sent to an external laboratory, it should be carefully packaged to prevent damage.


Equipment list: blood sample in an EDTA tube, glass microscope slides previously soaked in methanol and dried, glass cutter and marker pen.




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 image 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:


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Related posts:

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  2. Diagnostic imaging
  3. Medical diagnostic and treatment techniques
  4. Restraint, handling and administration of medication

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Jul 24, 2016 | Posted by in SMALL ANIMAL | Comments Off on Laboratory techniques

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