Body Fluid Analysis:

The visual appearance of the body fluid aids in determining whether a dilution should be made for cell counting and what dilution should be prepared. Body fluids that are clear do not require a dilution, and the fluid can be loaded directly onto a hemacytometer. Fluids that are visibly cloudy or bloody must be diluted to obtain accurate cell counts. Table 17.1 is provided as a guide to dilution selection based on visual appearance. When diluents that do not lyse RBCs are used, a higher dilution may be necessary. Body fluids that are visibly clear indicate a low cell count and are evaluated undiluted; sometimes cell counts in hazy or slightly cloudy fluids can also be performed on the undiluted fluid. To enhance visualization of WBCs in fluids other than synovial fluid, the fluid can be “exposed” to glacial acetic acid (Box 17.1). Blood-tinged or bloody fluids can be diluted using diluents that (1) lyse RBCs and (2) enhance visualization of nucleated cells.

Often isotonic, particle-free commercial diluents used by the laboratory’s hematology analyzer can be used to dilute body fluids (e.g., Cellpack, Sysmex Corporation, Kobe, Japan). Laboratory-prepared isotonic solutions such as normal saline (0.85%) can also be used for RBC counts, whereas dilute acetic acid solutions are often used for the nucleated cell and WBC counts. Acetic acid in the diluent performs two functions: It lyses any RBCs present and it enhances the nuclei of WBCs. Table 17.2 summarizes diluents commonly used for body fluids when WBC and RBC counts are performed. Appendix D provides details on the preparation of these diluents.

The total nucleated cell or WBC count is an important count that is requested on almost all body fluids. In contrast, little clinical value is derived from an RBC count other than to identify a traumatic puncture procedure, so it may not be performed, particularly when counts are done manually. Box 17.2 summarizes a protocol for a manual cell count using an “improved” Neubauer hemacytometer (Fig. 17.1).

Note that the dimensions are not the same in different types of counting chambers. For example, the depth in a Neubauer chamber is 0.1 mm whereas in a Fuch-Rosenthal chamber the depth is 0.2 mm. Using the correct dimensions is crucial because the resultant calculations (i.e., cell count) are dependent on them.

Before filling a hemacytometer chamber with undiluted fluid or preparing a dilution, the specimen must be adequately mixed to evenly disperse cells. Body fluid specimens can be mixed for 2 to 5 minutes on an automated rocking mixer or by inverting the tube 10 to 15 times by hand.1 Note that synovial fluid specimens should be mixed for 5 to 10 minutes. When using nondisposable hemacytometers, they should be cleaned before use by flooding with 70% alcohol and wiping dry with lens paper. This cleans the chambers and removes dust, which can interfere with placement of the coverslip.

The number of squares counted in each chamber of the hemacytometer depends on the total number of cells present. Accuracy in cell counts is directly related to cell numbers; the more cells counted, the better the accuracy. Therefore dilutions should be avoided if possible. When necessary, a minimum volume of 50 μL should be used to avoid pipetting errors associated with smaller volumes.² Dilutions should not be excessive and should result in an adequate number of cells for counting. Ideally, a minimum of 100 cells should be counted, but this is not feasible in many body fluids (e.g., CSF). To compensate for fluids with extremely low cell counts, additional squares of the hemacytometer grid can be counted. With fluids that have a high cell count, fewer squares can be counted. Note that regardless of the variation used, calculations must be properly adjusted for the volume of body fluid actually counted.

To ensure detection of potential errors during the preparation of body fluids for manual cell counts, dilutions should be performed and analyzed in duplicate. In other words, two separate dilutions of the body fluid are prepared and loaded into a hemacytometer—each dilution is loaded into one chamber. The cell counts obtained for each chamber are compared and must agree with the criteria established by the laboratory, usually 20% or less. If the counts between chambers are unacceptable (exceed 20%), the body fluid dilutions and counts must be repeated. Note that counting the same chamber twice or comparing two different chambers filled using the same dilution is not true replication and will not detect errors in pipetting, dilution, and mixing.

Calculations

When using a hemacytometer, regardless of the number of squares counted or the dilution used, the number of cells per microliter (or cubic millimeter) of fluid is determined using the following formula.

# cells per μL(mm3)=# cells counted(A)×dilution factor(B)[Area counted(mm2)×depth(0.1 mm)](C) Equation 17.1

Equation 17.1

There are two approaches to using this formula, and both will produce the same result. One approach is to count both chambers of the hemacytometer, compare the counts to ensure that precision criteria are met, average the count (if acceptable), and then use the formula to calculate the number of cells per microliter. An alternate approach is to count each chamber, compare the counts to ensure that precision criteria are met, and if acceptable, determine the sum of both chambers and use the formula to calculate the number of cells per microliter; see the example provided in Box 17.3.

Depending on the approach used, (A) is either (1) the average number of cells counted in a chamber (Option 1, Box 17.3) or the sum of the number of cells counted in both chambers (Option 2, Box 17.3) of the hemacytometer. This value (A) is multiplied by the dilution factor (B), which accounts for any dilution made of the fluid. If the fluid is analyzed without diluting, this factor is 1. The cells were distributed in the actual volume of fluid evaluated (C). This volume (mm³ = μL) is determined by multiplying the area counted (mm²) by the depth between the coverslip and the chamber, which is standardized at 0.1 mm. Disposable hemacytometers have a fixed (immovable) coverslip, which maintains the depth of 0.1 mm. However, when using a nondisposable hemacytometer with a removable coverslip, if the chamber is overfilled or underfilled, the depth is not the assumed value of 0.1 mm, which will cause erroneous results. The concentration of cells per microliter can easily be converted to the cell number per liter of fluid as follows.

# cells/μL(mm3)×106 μL/L=# ×106 cells/L Equation 17.2

Equation 17.2

Following are some examples using a Neubauer hemacytometer to determine cell counts of body fluids using undiluted and diluted fluid. The calculations used follow Option 2 described in Box 17.3. Note that an additional dilution factor is required when a fluid is “pretreated” before dilution for cell counting, which can occur with synovial fluid or semen (see Example C).

Example A: Using Undiluted Body Fluid

A well-mixed undiluted CSF specimen is loaded on a hemacytometer, and the WBCs in five large squares (4 “W”+Center) are counted in each chamber (i.e., 5 mm²). The following results are obtained.

	Chamber 1	Chamber 2
Cells counted (A):	15	18
Area each chamber:	5 mm2	5 mm2
Total volume counted (C):	10 mm2 × 0.1 mm = 1.0 mm3 (μL)
Dilution factor (B):	1
Precision assessment: 20% of 18 = (3.6) 4 cells	Count difference = 18 – 15 = 3 cells
(A×B)C=cells/μL	(15+18)×1a(1.0)=WBCs/μL =33 WBCs/μL
Unit Conversion:	33 WBCs/μL×106 μL/L =33×106 WBCs/L
aPretreatment dilution factor.

Note that the counts from both chambers agree within precision criteria of less than or equal to 20%, which indicates that the fluid was well mixed and equivalently dispensed in both chambers. The difference in cell number between side 1 and side 2 (18 – 15) is 3 cells, which is less than 20% (3.6 cells = 18 × 0.20) and acceptable.

Example C: Sperm Count Using Diluted Semen

A semen specimen was pretreated 1:2 using a bromelain enzyme preparation to get it to liquefy. For the sperm count, the fluid is diluted 1:20, in duplicate. Both dilutions are loaded on a hemacytometer—one dilution on each side. The five “R” squares in the large central square (i.e., four small corner+center squares) of the hemacytometer are counted in each chamber (i.e., 0.2 mm²). The following results are obtained. Note that sperm counts are reported as the number of sperm per milliliter (mL), which requires the use of a different unit conversion factor.

	Chamber 1	Chamber 2
Cells counted (A):	37	42
Area each chamber:	0.2 mm2	0.2 mm2
Total volume counted (C):	0.4 mm2 × 0.1 mm = 0.04 mm3 (μL)
Dilution factor (B):	20
Precision assessment: 20% of 42=(8.4)8 cells	Count difference = 42 – 37 = 5 sperm
(A×B)C×Da=cells/μL	(37+42)(0.04)×20×2=sperm/μL =79,000 sperm/μL
Unit Conversion:	79,000 sperm/μL×103 μL/mL =79×106 sperm/mL

^aPretreatment dilution factor.

The dilution factor (D) accounts for the dilution made when the fluid was pretreated to get it to liquefy using the bromelain enzyme solution. In this example, the semen counts from both chambers agree within 20%. The difference in sperm number between side 1 and side 2 (42 – 37) is 5 sperm. This value is less than 20% (8.4 sperm = 42 × 0.20), which indicates that the steps used to prepare both dilutions (mixing, pipetting, and diluting) were equivalent.

Numerous cytocentrifuges are commercially available and require the use of specially designed assemblies for each sample (Fig. 17.2). An assembly consists of a microscope slide, filter paper with a circular opening, and a chamber that holds the fluid specimen (Fig. 17.3).

When the body fluid is clear, usually 5 drops (≈0.25 mL) of body fluid is added directly to the chamber. However, depending on the nucleated cell count, as few as 2 drops or as many as 10 may be used. Table 17.3 provides a guideline based on the nucleated cell count for the volume of body fluid to use when preparing a slide by cytocentrifugation. When a dilution is needed, normal saline is most often used; however, some laboratories use a diluent that lyses RBCs for bloody samples (e.g., hypotonic saline). Appropriate dilutions will reduce distortions associated with the overcrowding of cells and ensure a monolayer of cells for viewing. Note that laboratories must establish their own dilution protocols because the appropriate dilution depends on the amount of sample used, as well as the duration and speed of cytocentrifugation.

Each assembly is placed onto the rotor of the cytocentrifuge, and when the instrument is activated, centrifugal force pulls the body fluid from the sample chamber to the microscope slide. The cells adhere to the glass slide while the liquid is absorbed by the surrounding absorbent filter paper. During centrifugation the cells concentrate as a monolayer in the open circular area of the filter to form a “cell button” on the microscope slide. The centrifugal force is low (e.g., 600–800 × g) to minimize cell distortion; the time is long enough to ensure adequate drying of the cell button. Microscope slides specifically designed for cytocentrifugation are available, and they have a white circular ring that surrounds the cell button area. This assists the microscopist when the body fluid sample contains few cells that are not visually apparent macroscopically on the slide. Another option is to use a wax pencil to mark the backside of the glass microscope slide, indicating the region of the cell button (Fig. 17.4).

During cytocentrifugation, some cells are lost to the filter paper, but this loss is not selective (i.e., all cells are equally affected); consequently, the remaining cell distribution in the cell button is accurate and representative. Several predictable cellular distortions that may be observed are listed in Box 17.4. Most are associated with high cell counts, the cytocentrifugation process (speed and time), or a time delay when an older specimen is used (i.e., not fresh).³ To reduce these artifacts, laboratories should determine the optimal speed and time of cytocentrifugation for their instrument, use fresh specimens, and prepare appropriate dilutions of fluids with high cell counts.

For specimens that have a low protein content (e.g., CSF), adding a drop of 22% albumin to the sample chamber before adding the body fluid enhances adherence of cells to the glass slide and reduces cell distortion (smudging) or disintegration.1,3

Slide preparations are stained using Wright or Wright-Giemsa stain performed manually or automatically using a slide stainer. The hand-drawn or premarked circle on the microscope slide indicates the location of the cell button (see Fig. 17.4).

Adjust the microscope to low-power (×100) magnification and thoroughly scan the entire cell button looking for cell clumps, which are characteristic of malignancies. Note that malignant cells can be present in low numbers, and even a single malignant cell is clinically significant. However, remember that not all cell clumps are composed of malignant cells.

Using oil immersion magnification (e.g., ×50 or ×1000), the nucleated cell or WBC differential is performed using any representative area of the cell button. A systematic approach to viewing should be used (similar to that used with blood smears) to prevent erroneous repeat counting of the same cells. Ideally, 100 to 300 cells should be evaluated.