Blood Specimens – Specimen Processing

Preparing Blood Smears

If you are using venous blood, blood smears should be prepared as soon as possible after collection (delay can result in changes in parasite morphology and staining characteristics).

Thick smears

Thick smears consist of a thick layer of dehemoglobinized (lysed) red blood cells (RBCs). The blood elements (including parasites, if any) are more concentrated (app. 30×) than in an equal area of a thin smear. Thus, thick smears allow a more efficient detection of parasites (increased sensitivity). However, they do not permit an optimal review of parasite morphology. For example, they are often not adequate for species identification of malaria parasites: if the thick smear is positive for malaria parasites, the thin smear should be used for species identification.

Prepare at least 2 smears per patient!

1. Place a small drop of blood in the center of the pre-cleaned, labeled slide.
2. Using the corner of another slide or an applicator stick, spread the drop in a circular pattern until it is the size of a dime (1.5 cm2).
3. A thick smear of proper density is one which, if placed (wet) over newsprint, allows you to barely read the words.
4. Lay the slides flat and allow the smears to dry thoroughly (protect from dust and insects!). Insufficiently dried smears (and/or smears that are too thick) can detach from the slides during staining. The risk is increased in smears made with anticoagulated blood. At room temperature, drying can take several hours; 30 minutes is the minimum; in the latter case, handle the smear very delicately during staining. You can accelerate the drying by using a fan or hair dryer (use cool setting). Protect thick smears from hot environments to prevent heat-fixing the smear.
5. Do not fix thick smears with methanol or heat. If there will be a delay in staining smears, dip the thick smear briefly in water to hemolyse the RBCs.

Scratch Method for Thick smears

The scratch method is an alternate method for making thick films that allows for improved adherence and faster turnaround times. The process is similar to making a normal thick film, but instead of using a stick to spread the blood, the edge of a glass microscope slide is used, while applying firm pressure to create small scratches in the underlying slide.  The scratches allow for improved adherence of the blood film to the slide without affecting the smear morphology. The smear can then be stained as soon as it is dry, generally within 20-30 minutes of smear preparation.

Reference: Norgan AP, Arguello HE, Sloan LM, Fernholz EC, Pritt BS. Malaria Journal 2013; 12: 231.

Thin smears

Thin smears consist of blood spread in a layer such that the thickness decreases progressively toward the feathered edge. In the feathered edge, the cells should be in a monolayer, not touching one another.

Prepare at least 2 smears per patient!

1. Place a small drop of blood on the pre-cleaned, labeled slide, near its frosted end.
2. Bring another slide at a 30-45° angle up to the drop, allowing the drop to spread along the contact line of the 2 slides.
3. Quickly push the upper (spreader) slide toward the unfrosted end of the lower slide.
4. Make sure that the smears have a good feathered edge. This is achieved by using the correct amount of blood and spreading technique.
5. Allow the thin smears to dry. (They dry much faster than the thick smears, and are less subject to detachment because they will be fixed.)
6. Fix the smears by dipping them in absolute methanol.

Note: Under field conditions, where slides are scarce, national malaria programs (and CDC staff) prepare both a thick and a thin smear on the same slide. This works adequately if one makes sure that of the two smears, only the thin smear is fixed.

Special Procedures for Detecting Microfilariae

Blood microfilariae:

1. Capillary (fingerstick) blood
   Since microfilariae concentrate in the peripheral capillaries, thick and thin smears prepared from fingerstick blood are recommended.
2. Anticoagulated (EDTA) venous blood (1 ml) should be concentrated by one of the following methods:
   1. Centrifugation (Knott’s technique)
      1. Prepare 2% formaldehyde (2 ml of 37% formaldehyde + 98 ml H2O).
      2. Mix 9 ml of this 2% formaldehyde with 1 ml of patient’s venous blood. Centrifuge at 500 × g for 10 minutes; discard supernatant. Sediment is composed of WBCs and microfilariae (if present).
      3. Examine as temporary wet mounts.
      4. Prepare thick and thin smears; allow to dry; dip in absolute methanol before Giemsa staining to enhance staining of microfilariae.
   2. Filtration
      1. Place Millipore® or Nucleopore® membrane filter (5 µm pore) in filter holder with syringe attachment.
      2. Mix 1 ml of venous blood (in EDTA) with 10 ml of 10% Teepol® 610 (Shell Co.); allow to stand for several minutes to allow lysis; transfer to a 10 ml Luer-Loc® syringe; attach the filter apparatus.
      3. Force the solution through the 5 µm pore filter, followed by several syringes of water to wash out the remaining blood, then 1 or 2 syringes full of air to clear excess fluid.
      4. Prepare a temporary wet mount by removing the filter and placing it on a glass slide, adding a drop of stain or dye and a coverslip.
      5. For permanent preparations, pass 2 to 3 ml of methanol through the filter while it is still in the holder; remove filter and dry it on a glass slide; then stain it with Giemsa stain, horizontally (so that the filter does not wash off the slide); coverslip filter before examining.

For additional information on making blood smears, call the Division of Parasitic Diseases at (404) 718-4110.

Reference:

Eberhard ML, Lammie PJ. Laboratory diagnosis of filariasis. Clin Lab Med 1991;11:4.