3: Detection of Parasites in the Blood

Detection of Parasites in the Blood

Various pathogenic and nonpathogenic protozoa and nematodes may be detected in blood samples from domestic animals. Most of these parasites are ingested by arthropod vectors during feeding and are present in the blood of their vertebrate hosts as a normal part of their life cycles.


Although the focus of this book is the morphologic diagnosis of parasitism, it is important to recognize that immunologic tests are widely used in conjunction with or in place of microscopic examination of blood smears for some blood‐borne parasites, and the use of these tests can be expected to increase in the future. Immunologic and molecular tests offer increased sensitivity compared with morphologic techniques in many cases and are especially valuable in some chronic hemoprotozoan infections and in canine and feline heartworm infection. In both cases, many infections are undetectable by routine microscopic tests. The commercial tests used most widely are the indirect fluorescent antibody (IFA) test, the enzyme‐linked immunosorbent assay (ELISA), and polymerase chain reaction (PCR). For additional discussion of the basis and use of immunodiagnostic and molecular diagnostic procedures in veterinary parasitology, see Chapter 4.


Most hemoprotozoan parasites are intracellular in erythrocytes or white blood cells and may cause anemia. A routine thin blood smear is therefore useful both for assessing erythrocyte abnormalities and for detecting the presence of parasites. Parasites are most likely to be detected in blood smears during acute infection. Once infections become chronic, immunologic or molecular diagnostic techniques are usually more sensitive as parasitemias can drop below detectable limits by light microscopy.

Schematic illustration of the technique for making a blood smear. (A) Bring a spreader slide back at an angle until it touches the drop of blood; wait until the drop flows laterally. (B) Draw the spreader slide away from the drop, maintaining an angle. The blood will spread into a smooth, thin film.

Fig. 3.1 Technique for making a blood smear. (A) Bring a spreader slide back at an angle until it touches the drop of blood; wait until the drop flows laterally. (B) Draw the spreader slide away from the drop, maintaining an angle. The blood will spread into a smooth, thin film.

For microscopic examination of blood smears for hemoprotozoa, Giemsa stain is most effective, but Wright’s stain can also be used in most cases. Commercial stain kits used in many veterinary practices (an example is Dip Quick Stain, Jorgensen Laboratories, Loveland, CO, www.jorvet.com) will also stain hemoprotozoa when used as directed, but the stain will be of poorer quality. The following procedure can be used for Giemsa stain.

To prepare a thin blood smear, place a drop of blood on one end of a microscope slide and draw the blood into a thin film as shown in Figure 3.1.

Giemsa Stain

  1. Air‐dry the blood film, protecting it from flies and other insects if it is not to be stained immediately.
  2. Fix in absolute methanol for 5 minutes and air‐dry.
  3. Dilute stock Giemsa stain 1 : 20 with distilled water and flood the film (or place slide in staining jar). Fresh stain should be prepared at least every 2 days.
  4. Stain for 30 minutes.
  5. Wash stain away gently with tap water.
  6. Air‐dry; parasite cytoplasm will stain blue, and nuclei will stain magenta.

Table 3.1. Average diameters of erythrocytes

Source: Measurements from Weiss and Wardrop (2010).

Animal Erythrocyte diameter (μm)
Horse 5.5
Cattle 5.8
Sheep 4.5
Goat 3.2
Dog 7.0
Cat 5.8
Chicken 7.0 × 12.0

The stained blood film can be scanned using the 40× objective of the microscope with use of the oil immersion lens for greater detail when suspected parasites are found.

The dimensions of blood parasites are best determined by means of an ocular micrometer (see Chapter 1). A micrometer is highly recommended for accurate measurement of parasites in blood and fecal samples. If a micrometer is not available, the size of the parasite on a blood film may be approximated by comparison with the dimensions of host erythrocytes (Table 3.1).


Many species of parasitic worms enter the bloodstream of the host to reach certain organs, where they develop to maturity. These parasites usually stay in the blood only minutes or hours; thus, they are seldom seen in blood samples taken for diagnostic purposes. There are some filarial nematodes, however, whose larvae (e.g., microfilariae) are normally found in the peripheral blood. The microfilarial stage of these parasite species remains in the circulation until ingestion by the bloodsucking intermediate host. Microfilaria testing is often performed for detection of canine heartworm infection. The following discussion of techniques for microfilaria detection is directed specifically to Dirofilaria immitis testing. However, all species of microfilariae in the blood could be detected by the same microscopic techniques.

Although the techniques for microscopic detection of heartworm microfilariae are presented below, the ELISA antigen test for diagnosis of canine heartworm infection is a commonly used screening test. Antigen tests are significantly more sensitive than microfilaria tests because many heartworm infections are amicrofilaremic (occult infections). The absence of microfilariae may be due to low or single‐sex worm burdens or immune clearance of microfilariae. Moreover, some heartworm preventives are microfilaricidal and may render infected dogs amicrofilaremic after one or several months of administration.

The American Heartworm Society and the Companion Animal Parasite Council currently recommend annual testing using an antigen test and a microfilaria test to identify D. immitis infection in dogs. Dogs testing positive on an antigen test should always be tested for microfilariae to determine if microfilaricidal treatment is necessary. Antigen tests currently available in the United States are available in ELISA and immunochromatographic formats. Differences in sensitivity among these tests have been found experimentally and are particularly evident when only a few adult worms are present. False‐negative antigen results can occur, especially when immune complexes have formed, precluding detection. Pre‐treatment of the serum or plasma to disrupt immune complexes using heat or chemicals prior to performing the antigen test releases the antigen and allows detection. Because specialized equipment is required, sending a sample to a diagnostic laboratory is recommended when blocked antigen is suspected. Available tests are considered highly specific although false‐positive results have been reported both before and after treatment to reveal blocked antigen.

Diagnosis of heartworm infection in cats is more difficult than in dogs. Several of the antigen tests can be used in cats, but false‐negative results are common because of immune complex formation as well as the low worm burdens usually found in cats. Similarly, cats are rarely microfilaremic and may develop disease before the adult stage, detectable by antigen testing, is present. To improve the sensitivity of heartworm detection in cats, heartworm antibody tests have been developed. These tests can detect infection earlier than antigen tests but may only indicate exposure to the parasite rather than active infection. Care should be taken in interpreting a feline antibody test, and the results of that test alone should not be used to establish a diagnosis of heartworm infection. In a cat showing clinical signs consistent with heartworm infection, both antigen and antibody tests should be performed as part of the diagnostic workup.

For current recommendations of the American Heartworm Society and the Companion Animal Parasite Council relating to diagnosis and treatment of heartworm infection in dogs and cats, consult the websites of the two organizations: www.heartwormsociety.org and www.capcvet.org.

Tests for Canine Heartworm Microfilariae in Blood Samples

The following techniques can be used to detect microfilariae in blood samples. The canine heartworm, Dirofilaria immitis, is found throughout the world. In North America, dogs may also be infected with Acanthocheilonema (= Dipetalonema) reconditum or, rarely, with Dirofilaria striata, a parasite of wild felids in North and South America. In parts of Europe, Asia, and Africa, Dirofilaria repens and Acanthocheilonema dracunculoides parasitize dogs. When a microfilaria test is used for heartworm diagnosis, the microfilariae of other species must be differentiated from those of D. immitis.

Staining characteristics can be used in discriminating among species, but are not usually performed in veterinary practices. Measurement of total length, width, and the shape of the head can also aid microfilaria identification (Table 3.2). Sizes should be determined with an eyepiece micrometer (see Chapter 1 for micrometer calibration procedure). The standard measurements of microfilariae in Table 3.2 were determined with formalin‐fixed specimens; use of other fixatives or lysing solutions may alter the size of the organisms. Similarly, storage of microfilariae in blood samples for more than 3 days may cause D. immitis microfilariae to shrink in length to the size of A. reconditum.

Wet Mount

The wet mount is the simplest and most rapid of the procedures for microfilariae detection. It is not a very sensitive technique but can be used in conjunction with an adult heartworm antigen test to determine if microfilariae are present or to evaluate the pattern of movement of microfilariae when attempting to differentiate between Dirofilaria and Acanthocheilonema

  1. Place one drop of anticoagulated venous blood onto a clean microscope slide and coverslip.

    Table 3.2. Characteristics of Dirofilaria spp. and other microfilariae found in canine blood based on formalin‐fixed specimens

    Dirofilaria immitis Dirofilaria repens Dirofilaria striata Acanthocheilonema reconditum Acanthocheilonema dracunculoides
    Length (μm) 295–325 268–360 360–385 250–288 189–230
    Width (μm) 5–7.5 5–8 5–6 4.5–5.5 5–6
    Head Tapered Blunt Tapered Blunt Blunt
    Tail Straight Variable— straight or hooked Curved Variable—hooked (30%) or curved Sharp and extended
    Body shape Straight
    S‐shaped Curved
    Motion (live) Stationary
    Stationary Progressive
    Relative number Few to many
    Few Few
    Location of adult Pulmonary arteries, right heart Subcutaneous intramuscular tissues Subcutaneous intramuscular tissues Subcutaneous tissues Peritoneum
    Geographic location Worldwide Europe, Africa, Asia North and South America Africa, Europe, North America Africa, Europe, India

  2. Examine the coverslip area under low magnification (10×) of the microscope. Look for undulating movements of larvae, which may retain motility for as long as 24 hours.

Hematocrit Test

This technique is only slightly more sensitive than the wet mount:

  1. Draw fresh whole blood into a microhematocrit tube.
  2. Spin for 3 minutes in a hematocrit tube centrifuge.
  3. Examine the plasma portion of the separated blood, while still in the tube, under low magnification (10×). Moving microfilariae will be present in the plasma above the buffy coat (Fig. 3.2).
Photo depicts the results of a hematocrit test using a blood sample containing D. immitis microfilariae. The microfilariae can be seen as a hazy layer to the right of the buffy coat layer (arrow). Closer microscopic examination would show individual moving microfilariae.

Fig. 3.2 Results of a hematocrit test using a blood sample containing D. immitis microfilariae. The microfilariae can be seen as a hazy layer to the right of the buffy coat layer (arrow). Closer microscopic examination would show individual moving microfilariae.

The wet mount and microhematocrit techniques may not detect infections with only small numbers of microfilariae. Therefore, if a microfilariae test is used as a screening procedure for heartworm infection, one of the following concentration techniques should be used.

Modified Knott’s Test

The modified Knott’s technique is the preferred concentration method for the detection and identification of microfilariae in blood:

  1. Draw a sample of blood into a syringe containing anticoagulant such as EDTA or heparin.
  2. Mix 1 mL of the blood with 9 mL of a 2% formalin solution. If not well mixed, the red cells will not be thoroughly lysed by the hypotonic formalin solution, making the test much more difficult to read. Microfilariae, but not red cells, will be fixed by 2% formalin. If 10% formalin is used (the concentration used for fixation of tissues), red cells will also be fixed and not lysed.
  3. Centrifuge the mixture at 1200 rpm for 5 minutes (or as for fecal flotation procedures) and discard the supernatant.
  4. Add one drop of 0.1% methylene blue to the sediment, mix well, and transfer the entire stained sediment to a microscope slide using a Pasteur pipette.
  5. Examine using the 10× microscope objective. Microfilariae will be fixed in an extended position with nuclei stained blue.

An alternative procedure using the same amount of blood is the filter test, which traps microfilariae on a filter that is examined with the microscope. This technique can be performed more quickly than the modified Knott’s test, but microfilariae are not easily measured for identification. Materials for performing the filter test were sold as a kit (Difil‐Test®), which is no longer available. Components of the test can be purchased individually if desired.

Filter Test

  1. Mix 1 mL of blood with 9 mL lysing solution (2% formalin) in a syringe.
  2. Attach the syringe to a filter holder containing a transparent 25 mm filter with a 5‐μm pore size and empty the syringe.
  3. Refill syringe with water and pass it through the filter to wash away remaining small debris.
  4. Refill syringe with air, reattach to the filter apparatus, and express.
  5. Unscrew the filter assembly, remove the filter with forceps, and place the filter on a microscope slide.
  6. Add one drop of 0.1% methylene blue, coverslip, and examine at 10×.


PARASITE: Hepatozoon spp. (Fig. 3.3)

Taxonomy: Protozoa (hemogregarine).

Geographic Distribution: Hepatozoon canis occurs worldwide, while the distribution of Hepatozoon americanum appears to be limited to the southeastern United States.

Location in Host: Gamonts are found in polymorphonuclear leukocytes (H. americanum, H. canis) and meronts in skeletal muscle (H. americanum) or various organs (H. canis) of dogs, cats, and various wild carnivores.

Life Cycle: Ticks acquire infection during feeding. Dogs become infected by ingesting infected ticks. Hepatozoon americanum is transmitted by Amblyomma maculatum (the Gulf Coast tick), and the vector of H. canis is Rhipicephalus sanguineus (the brown dog tick).

Laboratory Diagnosis: Sausage‐shaped Hepatozoon gamonts can be detected in polymorphonuclear leukocytes in Wright‐ or Geimsa‐stained blood smears. Although this method of diagnosis readily reveals H. canis, H. americanum is rarely found on blood smears, and a molecular diagnostic test may be necessary. Morphologic diagnosis of this species generally occurs by the detection of meronts in skeletal muscle biopsies or on histopathology after necropsy.

Size:Gamonts 8–12 × 3–6 μm

Clinical Importance: Hepatozoon americanum can cause severe disease, with fever, depression, joint pain, myositis, periosteal bone proliferation, and chronic wasting. Hepatozoon canis infections are usually subclinical.

Photo depicts Hepatozoon gamont in a polymorphonucleocyte. The parasite is sausage-shaped with a centrally compact nucleus that stains only faintly in this specimen (arrow). Hepatozoon americanum is rarely present in blood films, and muscle biopsies are more commonly used for diagnosis.

Fig. 3.3 Hepatozoon gamont in a polymorphonucleocyte. The parasite is sausage‐shaped with a centrally compact nucleus that stains only faintly in this specimen (arrow). Hepatozoon americanum is rarely detected in blood films, and muscle biopsies are a more sensitive means of diagnosis.

PARASITE: Large (e.g., Babesia canis) and small (e.g., B. gibsoni) Babesia spp (Figs. 3.4 and 3.5) Babesia spp.

Taxonomy: Protozoa (piroplasm). Babesia spp. are divided into large (>4 μm) and small (<3 μm). Large species include B. canis vogeli, B. canis rossi, B. canis canis, Babesia sp. (Coco), and an unnamed British isolate. Small Babesia include B. gibsoni, B. conradae, and B. vulpes.

Geographic Distribution: Babesia mostly occurs in the tropical and subtropical regions of the world. B. canis vogeli is found worldwide, B. canis rossi in Africa, B. canis canis in Europe, and Babesia sp. (Coco) has been reported sporadically in immunocompromised dogs in various U.S. states. Babesia gibsoni is widely distributed throughout most of the world, B. conradae in dogs from California and Oklahoma, and B. vulpes infects a variety of wild canids (primarily foxes) and occasionally domestic dogs in Europe, North America, and western Asia.

Location in Host: Canine red blood cells. Babesia spp. have been described in cats but are not widely distributed and do not appear to be present in North America.

Life Cycle: Ticks are definitive hosts for Babesia spp. In North America, dogs acquire B. canis vogeli from the brown dog tick, Rhipicephalus sanguineus. Other tick vectors include Dermacentor reticulatus in Europe and Haemaphysalis leachi in Africa. A definitive tick vector for B. gibsoni has not been demonstrated in North America and transmission is thought to occur primarily or only through the transfer of blood contaminated with piroplasms. Dog fighting increases the risk of infection with B. gibsoni.

Laboratory Diagnosis:

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