References

Baermann G Eine einfache Methode zur Affindung von Ankylostomum-(Nematoden)-Larven in Erdproben. 1917; 57:131-37

Bowman DD Georgis' parasitology for veterinarians, 9th ed. St. Louis MO: Saunders; 2009

Bowman DD, Hendrix CM, Lindsay DS Feline clinical parasitology.Ames, Iowa: Wiley-Blackwell; 2002

Webster JM, Eriksson KB, McNamara DG An Anecdotal History of Nematology.: Pensoft Publishers; 2008

Zajac AM, Conboy GA Veterinary clinical parasitology, 8th ed. Ames, Iowa: Wiley-Blackwell; 2012

ABC series on diagnostic parasitology part 3: the Baermann technique

02 December 2017
Practical
5 mins read
02 December 2017
Volume 8 · Issue 10

Abstract

Diagnosis of parasitic larval forms is typically associated with complex, costly methods. As such, simple, low-cost diagnostic tools for larvae identification are of the utmost importance in diagnostic parasitology in the veterinary clinic. Herewith we describe the Baermann technique, one of the most used larvae identification methods.

The Baermann technique is a simple, low-cost diagnostic tool for diagnosis of parasitic larval forms, that parasitologists, health and veterinary personnel rely on. Historically, this technique is derived from attempts to identify nematodes from the soil, and relies on the fact that these parasites are aquatic animals and that almost all are mobile (Webster et al, 2008). In 1917, while working in Java, the Dutch physician Baermann devised a simple method of isolating nematodes, including infective hookworm larvae, from soil. It became known as the Baermann funnel technique, or Baermann technique (Baermann, 1917). Dr Baermann discovered, by putting soil in a muslin bag and into a funnel filled with water for several hours, that nematodes tended to migrate downwards out of the soil and through the muslin, and then could be seen in the water at the stemend of the funnel. Despite the fact that this was ground-breaking, Baermann´s discovery presented a drawback, with the water resulting from this method being often murky (as a result of the leaching of pigments and particles of small diameter through the muslin) (Webster et al, 2008). This has led to several modifications in the methodology.

Following its application to human and veterinary parasitology, it has been shown that the Baermann technique has advantages over other tests used for the same purpose (Bowman et al, 2002). In particular, since larvae may be present in low numbers or intermittently, the Baermann technique allows for the use of a larger amount of stools to be used compared with a faecal flotation test (Bowman, 2009).

Despite the fact that the Baermann technique is possibly the easiest morphology-based test to perform and evaluate in diagnostic parasitology, it still seems to be rarely used in veterinary practice. It is worth pointing out some of the difficulties that exist for the execution of the technique, such as the size of the equipment, space required in the lab and the large volumes of faeces required. Additionally, there is the need for experience when identifying specific parasitic larvae and therefore for increased specificity of the test. Nonetheless, it presents great value and potential for veterinary use in the extraction of live larval stages of nematode parasites from faeces (Zajac and Conboy, 2012).

The Baermann technique is based on the active migration or movement of larvae. In brief, faeces are suspended in water and the larvae move into the water. Since they are unable to swim, they sink to the bottom and can be collected for identification.

In the Baermann test it is essential to use a fresh faecal sample, collected immediately after passage from the animal, since the contact with the ground can contaminate faeces with freeliving nematodes that will also be recovered in the test and that can be hard to distinguish from parasite larvae. Also it is not recommended to refrigerate samples for a lengthy period (days), since larvae may die in that time, and the test requires the presence of live larvae, able to react to a stimulus such as the warm water used in this test (Bowman, 2009).

Herewith we describe the Baermann technique:

The Baermann technique

Equipment needed:

  • Funnel — size according to need
  • Funnel stand
  • Rubber or plastic tubing
  • Clamp or spring clip
  • Cheesecloth or dental napkin or gauze
  • Thin stick or metal rod
  • Strainer
  • Microscope
  • Test tube
  • Pasteur pipette
  • Small petri dishes
  • Scissors
  • Disposable paper towels
  • Spoon or spatula
  • Rubber band or length of string
  • Jug or flask
  • Microscope slides and coverslips
  • Iodine (Figure 1).
    • Figure 1. A, Funnel; B, Funnel stand; C, Rubber or plastic tubing; D, Clamp or spring clip; E, Gauze; F, Thin stick; G, Strainer; H, Scissors; I, Pasteur pipette; J, Spoon or spatula; K, Small petri dishes; L, Test tube; M, Rubber band; N, Jug or flask; O, Microscope slide and coverslip; P, Iodine solution.

    Step-by-step guide

  • Take a funnel and fit a short piece of tubing to the stem.
  • Close the tubing with a clamp or spring clip.
  • Support the funnel on a single stand (Figure 2).
  • Place a double layer of cheesecloth or dental napkin on a disposable paper towel or equivalent on the bench.
  • Using a spoon or spatula weigh or measure approximately 10 g of faecal material.
  • Place the faecal material in the centre of the cheesecloth.
  • Using a rubber band or length of string close the cheesecloth pouch (Figure 3).
  • Push the stick or short metal rod under the rubber band or string so that the pouch can be suspended (Figure 4).
  • Place the pouch containing the faecal material in the funnel.
  • Either trim off the excess cheesecloth, or let it hang over the rim of the funnel.
  • Fill the funnel with lukewarm water, ensuring that the faecal material is covered (Figure 5).
  • Leave the apparatus to stand for 24 hours (Figure 6).
  • Draw off a few millilitres of fluid from the stem of the funnel into a test tube (Figure 7) and leave to sediment for at least 30 minutes (if a centrifuge is available, the fluid can be drawn into a centrifuge tube and spun at 1000 rpm for 2 minutes).
  • Check sedimented sample in a petri dish for the presence of larvae. This may be all that is required to diagnose the presence of nematode parasites, but often more detailed examination is required since other parasitic or free-living nematode life-cycle stages may be present if the faecal sample was not fresh when processed or if it was collected from the ground.
  • Use a Pasteur pipette to transfer a small droplet of the sedimented fluid from the petri dish to a microscope slide (Figure 8).
  • Add a drop of iodine to fix the larvae and gently place a coverslip over the drop (Figure 9).
  • Any free-living nematodes will stain dark brown, uniformly, in a few seconds, while the larvae of parasitic species will stain in a few minutes, as the larval sheath protects the body.
  • Examine under compound microscope at 10 x 10 magnification (Figure 10).

    • Note: Free-living nematodes stain deeply brown in iodine and can be distinguished from parasitic species by the presence of a double bulbed (rhabditiform) oesophagus.

    Figure 2. Support the funnel and the tubbing on a single stand, and close the tubing with a clamp or spring clip.
    Figure 3. Using a spoon (or spatula) weigh or measure approximately 10 g of faecal material (A, B) and place it in the centre of the gauze and using a rubber band close the gauze pouch (C, D).
    Figure 4. Push the stick under the rubber band (A) so that the pouch can be suspended (B).
    Figure 5. Fill the funnel with lukewarm water, ensuring that the faecal material is covered.
    Figure 6. Leave the apparatus to stand for 24 hours.
    Figure 7. Draw off a few millilitres of fluid from the stem of the funnel into a test tube.
    Figure 8. With a Pasteur pipette transfer a small droplet of the sedimented fluid from the petri dish to a microscope slide.
    Figure 9. A drop of iodine is added to the sediment (A) and a coverslip is gently placed over the drop (B).
    Figure 10. The slide is examine under compound microscope at 10 x 10 magnification.

    Further considerations

    As with other diagnostic procedures in parasitology, it is important to highlight that stools are biological materials that carry risk of human infection, namely for the person carrying out the technique. As such and for the safety of the user, faecal material should always be presumed to be infectious and protective outerwear and disposable gloves should be used at all times. General health and safety regulations for biological material manipulations should always be followed.

    Conclusion

    The Baermann technique is not recommended as a primary diagnostic technique for evaluation of parasites in faeces, but is very useful to detect nematode larvae (L3). It is easily executed, however it requires experience to identify parasitic species present.

    KEY POINTS

  • The Baermann technique is a low-cost diagnostic tool.
  • The Baermann technique is based on the active migration of larvae.
  • Fresh faecal samples are essential for a reliable result.
  • Free living nematodes must be readly distinguishable from parasitic ones.
    • Baermann technique
    parasites
    diagnostic