ISSN 2398-2985      

Cytology: fluid sampling

Jreptile

Overview

  • As pleural, pericardial and peritoneal cavities normally contain only a small amount of fluid, little or no fluid can be obtained by aspiration unless effusion is present.
  • Effusion is the presence of excess fluid in a body cavity:
    • Peritoneal fluid.
    • Pleural fluid.
    • Pericardial fluid.
  • Investigation of effusions commonly includes fluid analysis (RBC, nucleated cell counts, and refractometer total protein) and cytology.
  • Prior to the preparation of fluid samples for cytologic examination, macroscopic observations of fluid samples should be made and recorded. These include the source, color, appearance (eg bloody, mucoid, serosanguinous, watery, milky, or cloudy), and refractometer-determined total solute (protein) concentration of the fluid, as well as the presence of any odor, clots, or tissue fragments.
  • This information will facilitate classification of effusions such as transudates, modified transudates, or exudates.

Sampling

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Tests

Methodologies

Sample collection

  • Fluids that have a plasma-like consistency should be handled in the same manner as preparing a blood film.
  • Direct smears should be made from aspirated fluids with good cellularity, ie 10 000 cell/mL or greater, and can be prepared using the conventional wedge method or the cover-glass method commonly used for making blood films.
  • Mucoid samples or fluids containing clots or solid tissue fragments should be prepared using the squash preparation method. When preparing mucoid samples, it may be necessary to pull the slides apart using a wavy motion while separating them in order to break apart the tenacious fluid. Raised tissue fragments and clots should be removed from the surface of the slide to avoid problems with overstaining and placement of a cover slip.
  • Fluid samples that are composed predominantly of blood should be handled as if preparing a blood film. Preparing a buffy coat smear will also improve the diagnostic yield of such samples:
    • To prepare a buffy coat smear, first transfer part of the sample from the EDTA tube to a microhematocrit tube.
    • The microhematocrit tube is centrifuged in the same manner as preparation of a microhematocrit in order to concentrate the buffy coat.
    • After centrifugation, the tube is broken at the cell–plasma interface to obtain the concentrated cells located in the buffy coat area.

Serum-coated slides

  • If the protein content of the fluid is low, the cells may be easily washed off the slides during the staining process. Use of serum-coated slides for the preparation of such samples will facilitate the fixing of the cells to the slide. 
  • Serum-coated slides can be made in advance by applying several drops of serum across the surface of glass microscope slides to create a film and allowing the slides to dry. After drying, the slides may be stored in the freezer to prevent bacterial growth. Slides are thawed and allowed to reach room temperature before using them to prepare cytologic specimens to prevent condensation which will cause cell lysis.

Cell concentration

  • Poorly-cellular fluids and wash samples require concentration methods in order to increase the sample cellularity, which will in turn increase the number of cells available for microscopic examination. Several methods of cell concentration may be used to improve the cellularity of the sample and include margination of the cells, preparation of a sediment smear, cytocentrifugation, and sedimentation.
  • The first method of cell concentration would be to marginate the cells while making the smear, using the conventional two-slide wedge technique:
    • Samples are prepared similarly to the same two-slide wedge technique previously described for preparing blood films except that the spreader slide is advanced more slowly and is abruptly lifted from the surface of the slide near the end of the smear.
    • It is important that the entire fluid sample applied to the slide should remain on the slide; the fluid
      should not be allowed to go off the end of the slide during sample preparation.
    • If sample is lost from the end of the slide, there is a potential for loss of diagnostic cellu-
      lar material, such as clumps of neoplastic cells and large infectious agents.
    • Marginated cells will be concentrated at the feathered end of the smear. The thinner area of the stained smear can be used to estimate cell numbers while the thicker part near the end of the slide is used to evaluate cell types.
  • The second method of concentrating cells is by preparation of a sediment smear following centrifugation:
    • The fluid sample is transferred to a plastic sample tube or test tube and centrifuged at a relative centrifugal force, 600×G (gravity) for 10 min. If possible, use disposable centrifuge tubes with plastic screw tops to prevent aerosolization of the sample during centrifugation. Glass tubes should not be used for the sample preparation because cells often adhere to glass surfaces and will be lost for sampling.
    • During centrifugation, the cellular content of the fluid is concentrated at the bottom of the tube and forms a button or pellet. After centrifugation, the clear supernatant is discarded, and the sediment is either aspirated from the tube using a disposable pipette or syringe or collected on a wire loop or cotton-tipped swab that has been premoistened by the supernatant fluid.
    • When a pipette or syringe is used to obtain the concentrated sample, a few drops are placed on the surface of the slide and another slide is used to make the smear. If a cotton swab is used, the swab is gently rolled in one direction across the slide.
  • A third technique for cell concentration of fluid samples involves the use of cytocentrifuge equipment:
    • This method involves the use of centrifugation to directly deposit the concentrated cells onto the glass slide while absorbing the noncellular fluid onto filter paper. 
    • The concentrated cells are located in a small circular area in the center of the slide.
  • A fourth cell-concentrating method is based on sedimentation:
    • Filter paper, eg Whatman no. 2 filter paper, is cut in the shape of the slide, and a standard 2-mm paper punch is used to create a hole in the center of the filter paper. 
    • The filter paper is then placed on the slide with the hole centered on the slide.
    • A sedimentation device is made using a base to support the microscope slide and a clamping mechanism to firmly support a fluid column made from the barrel of an 1-ml syringe (which has a 2 mm diameter equal to that of the hole in the filter paper) with the tip removed.
    • The base of the syringe barrel is positioned with the opening superimposed over the hole in the filter paper and secured to the slide with a tight seal.
    • A small amount of fluid, eg 0.5 mL or less, is placed in the syringe column and the apparatus is allowed to stand undisturbed. As the fluid falls in the column by gravity, it is absorbed by the filter paper. The cells adhere to the surface of the exposed glass slide.
    • After the fluid has drained, the apparatus is disassembled and the cellular sample remaining in the 2-mm circle created by the hole in the filter paper is allowed to air-dry before staining.
    • This method is somewhat complicated and time-consuming but has an advantage that cell morphology is better preserved compared to that of cells concentrated using centrifugation.

Sample preparation

  • Once the sample has been collected and a smear has been made, the specimen must be properly fixed to the slide. The method of fixation will depend on the staining procedure used. Whenever possible, more than one smear should be made per sample to allow for specific staining procedures, eg fresh air-dried smears are adequate for Romanowsky stains, ie Giemsa and Wright’s stains, but proper fixation is required for other stains, as directed by specific staining procedures.
  • Staining procedures for the stains used are listed in the Cytochemical staining reactions: normal leukocytes/thrombocytes table.
  • When using stains, especially the quick stains, the solutions should be changed frequently. The frequency of refreshing staining solutions depends upon the number of slides being stained and the amount of con- tamination appearing in the solutions as a result of cells and debris falling off the slides. For this reason, two sets of staining stations are recommended: one for relatively 'clean' samples (such as blood films, wash samples, effusions, and organ biopsies), and a second set
    for 'dirty' specimens (such as abscesses, fecal samples and skin scrapings).

Evaluation

  • Scanning (40× or 100×) and low (200×) magnifications are used initially to obtain a general impression of the smear quality. At these magnifications, the examiner is able to estimate the smear cellularity, examine cellular aggregates, identify large infectious agents, eg yeast and fungal elements, and determine the best locations for examination at higher magnifications (monolayers of cells). High-dry (400× or 500×) and oil-immersion (500× and 1000×) magnifications are used to examine cell structure, bacteria, and other small structures, eg cellular inclusions.
  • Cellular distribution should also be noted when scanning the entire smear. The conventional blood film technique for making cytology slides often causes margination of bacteria, macrophages, and large neoplastic cells at the periphery of the smear. For this reason, peripheral margins of smears made from fluid samples should be examined closely.
  • Cellular interpretation is best made in monolayer areas of cells because the thickness of the smear will affect the appearance of the cells and the quality of the cellular sample. Areas containing a heavy accumulation of cells and background material will not allow the cells to expand on the slide, and the cells appear smaller and denser compared to the same cell type in a thinner area of the smear.
  • Optimum evaluation of cytologic specimens requires knowledge of the sample origin, method of collection, and normal cellular morphology.

Availability

  • Standard laboratory procedure.

Validity

Sensitivity

  • As with any cytologic specimens, absence of evidence does not provide conclusive support for absence of a particular condition.
  • Biopsy from abnormal tissues may be more sensitive in diagnosing pathology in many cases.

Specificity

  • Technical expertise in cytological assessment is important in diagnosis of specific pathology.

Technique (intrinsic) limitations

  • Cytologic evaluation may be compromised by excessive blood contamination.

Result Data

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Further Reading

Publications

Refereed papers

Other sources of information

  • Campbell T W (2015) Cytological Sampling Techniques and Evaluation. In: Exotic Animal Hematology and Cytology. 4th edn. Wiley-Blackwell. pp 348.
  • Campbell T W (2006) Clinical Pathology of Reptiles. In: Reptile Medicine and Surgery. 2nd edn. Ed: Mader D R. Saunders-Elsevier, USA. pp 453-470.

Reproduced with permission from Terry W Campbell: Exotic Animal Hematology and Cytology © 2015, published by John Wiley & Sons.
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