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- LITERATURE CITED
Single-platform flow cytometric absolute cell counting protocols provide increased robustness for CD34+ cell enumeration by limiting potential sources of imprecision. However, samples with any cellular fragmentation or debris, such as cord blood samples, provide challenges for these assays. We describe a simple, robust absolute CD34+ cell counting protocol, suitable for cord blood, using TRUCOUNT™ absolute count tubes (BD Biosciences, San Jose, CA) and a modified ISHAGE (International Society for Hematotherapy and Graft Engineering) gating strategy. An advantage of TRUCOUNT tubes is that each tube is supplied with a known number of lyophilized fluorescent beads. The method includes no-wash fixative-free ammonium chloride red blood cell lysis and the viability dye, 7-amino actinomycin D, to exclude dead cells. The threshold was set on CD45 expression in the FL1 channel and an exclusion gate in the forward scatter channel reduced debris. No manual adjustment of the gating regions was required, even for samples in less than optimal condition. Comparison of the TRUCOUNT-ISHAGE protocol with the original dual-platform ISHAGE assay (n = 30) and the single-platform ISHAGE protocol using Flow-Count™ Fluorospheres (Beckman Coulter, Fullerton, CA; n = 22) showed high correlation (R2 = 0.949 and 0.989, respectively) and no significant difference or bias for samples ranging from 22 to 600 CD34+ cells per microliter. Results are presented that demonstrate the detrimental effect of a fixative-containing lysis reagent when used in a lyse-and-wash procedure. The TRUCOUNT-ISHAGE protocol combines the attributes of TRUCOUNT tubes and the ISHAGE gating strategy to provide a single-platform protocol capable of achieving readily standardization of CD34+ cell enumeration. Cytometry (Comm. Clin. Cytometry) 46:254–261, 2001. © 2001 Wiley-Liss, Inc.
Human umbilical cord blood is a rich source of hematopoietic stem cells (HSC) that is being used increasingly as an alternative to bone marrow for HSC transplantation (1–3). The cellular content, in particular the number of HSC, is a principal factor in assessing the adequacy of cord blood units for clinical transplantation. HSC are contained within the CD34+ cell population, which represents typically less than 1% of the total leukocytes in cord blood (4). This places CD34 analysis in the category of “rare-event” analysis and has presented a challenge to the development of standardized, robust methods for the quantitation of HSC. Cord blood poses additional challenges. These include the presence of variable numbers of nucleated red blood cells (RBC; which can lead to overestimation of the leukocyte count) and the age of the samples, which is often greater than 24 h old. Increased apoptosis, cell death, and debris can potentially confound the analysis.
CD34+ cell enumeration has relied predominantly on flow cytometric procedures (5–7). Guidelines published by the International Society for Hematotherapy and Graft Engineering (ISHAGE; 8) were a significant attempt towards standardization of the CD34 assay. These guidelines were based on a dual-platform method whereby the percentage of CD34+ cells was determined by flow cytometry and the leukocyte count by an automated hematology analyzer. A lyse-and-wash method for sample preparation, a gating strategy based on a forward scatter threshold, CD45 staining to define the total leukocyte population, and light scatter characteristics were recommended. Nevertheless, the use of two instruments that were used to determine the absolute CD34+ cell number, the potential loss of cells during the lysis and wash steps, and the influence of the choice of lysis reagent were sources of variability and obstacles to standardization of the assay.
The development of single-platform methods enable the absolute CD34+ cell count to be determined by a single instrument. Most of these new methods rely on flow cytometry. An exception is a microvolume fluorometry method that utilizes a novel instrument, the IMAGN® 2000 with the STELLer™ CD34 assay kit (BD Biosciences, San Jose, CA). This instrument is fully automated. However, the STELLer CD34 assay has not been optimized for cord blood and samples in less than pristine conditions, such as cord blood samples greater than 24 h old, are problematic (Sparrow, unpublished observations). Furthermore, it cannot distinguish live from dead cells.
The flow cytometric-based single–platform methods center on the incorporation of a known number of fluorescent beads into the sample. Using a lyse-no-wash procedure for sample preparation enables the ratio of CD34+ cells to beads to be determined and the absolute CD34+ cell count to be calculated. The ProCOUNT™ assay (BD Biosciences) was the first commercial kit based on this approach. This three-color assay utilizes TRUCOUNT™ absolute count tubes (BD Biosciences), which contain a predispensed lyophilized pellet of a known number of 4.2-μm fluorescent beads. Due to the small beads, a forward scatter threshold cannot be used, necessitating the threshold to be set on a fluorescence channel. For the ProCOUNT assay, a proprietary nucleic acid dye is used to discriminate nucleated cells from erythrocytes, platelets, and debris and to set the threshold in the FL1 channel. CD34+ and CD45+ events are detected in the FL2 and FL3 channels, respectively. Acquisition and analysis are performed according to the gating strategy recommended by the manufacturer or by using proprietary software (BD Biosciences). Most evaluations of the ProCOUNT assay have been favorable (9, 10). However, significant discrepancies have been observed in some instances (11, 12). Furthermore, sample variability may necessitate manual adjustment of the gating regions, thus lessening the robustness of the assay. The assay has not been optimized for cord blood or bone marrow samples and, for single-laser flow cytometers, cannot be used to assess viability or analyze subpopulations.
Hübl et al. (13) described a modification of the ProCOUNT assay that is applicable to cord blood. In this modified protocol, the nucleic acid dye was replaced with a viability dye, YO-PRO-1 (FL1 channel), and the threshold was set on CD45 staining (FL3 channel). Nevertheless, manual adjustment of the regions was still required for each sample, potentially reducing the robustness of the assay.
Fornas et al. (14) described a no-lysis, no-wash whole blood method using TRUCOUNT tubes. The method relies on a vital nucleic acid dye, SYTO-13, to discriminate erythrocytes and debris from viable nucleated cells and to set the threshold in the FL1 channel. This protocol eliminates the lysis step. However, efflux of the SYTO-13 dye, particularly in P-glycoprotein–expressing cells, necessitated the addition of cyclosporin A to the assay mixture.
Using an alternative source of fluorescent beads, Keeney et al. (15) refined the original ISHAGE method by incorporating Flow-Count™ Fluorospheres (Beckman Coulter, Fullerton, CA), along with a no-wash, fixative-free lysis method for sample preparation and inclusion of a viability dye, 7-amino actinomycin D (7-AAD). This method formed the basis of the Stem-Kit™ CD34 HPC enumeration kit (Beckman Coulter). A significant advantage of this approach is the robustness and sensitivity of the ISHAGE gating strategy. However, Flow-Count Fluorospheres are supplied as a bead suspension that necessitates careful handling to ensure accuracy. Recent data from a multicenter trial conducted by the CD34 Task Force of the European Working Group of Clinical Cell Analysis demonstrated good interlaboratory agreement using the Flow-Count procedure (16). This trial was designed specifically to ensure a high level of standardization across the participating laboratories. Stabilized blood samples configured to represent mobilized peripheral blood stem cell specimens were used with a standardized protocol and targeted training. A protocol using TRUCOUNT tubes was also assessed and yielded comparable results with good interlaboratory agreement.
We describe an absolute cell counting procedure for CD34 analysis of cord blood using TRUCOUNT tubes and a modification of the ISHAGE gating strategy. The method includes fixative-free ammonium chloride RBC lysis and elimination of nonviable CD34+ cells using 7-AAD. The threshold is set on CD45 expression in the FL1 channel and debris is removed from the CD34 gating region using an exclusion gate in the forward scatter channel. This TRUCOUNT-based method compared favorably with the original dual-platform ISHAGE method and single-platform method using Flow-Count Fluorospheres. In addition, we show that the commercial lyse-and-fix reagent FACSLyse (BD Biosciences) has a detrimental effect on CD34+ cell enumeration when using a lyse-and-wash procedure. This new protocol combines the advantages of TRUCOUNT tubes and the ISHAGE gating strategy in a simple, robust assay.
- Top of page
- MATERIALS AND METHODS
- LITERATURE CITED
The increasing clinical utilization of HSC transplantation, the establishment of cord blood banks, and the inclusion of HSC processing facilities within the control of national regulatory authorities have hastened the need for accurate and robust assays for CD34+ cell enumeration. Cord blood has posed additional challenges, particularly due to the presence of varying numbers of nucleated RBC and the extended age of the samples at the time of CD34+ cell enumeration. Single-platform absolute CD34+ cell-counting protocols offer the greatest potential for the development of robust assays capable of analytical accuracy and standardization. In this study, we described a modification of the single-platform ISHAGE protocol that incorporates an alternative source of fluorescent beads, presented in the format of TRUCOUNT tubes, which is suitable for analysis of cord blood samples.
Initial experiments were performed to ascertain the influence of the RBC lysis reagent. These experiments utilized the original dual-platform ISHAGE lyse-and-wash protocol. Our results demonstrated that the fixative-containing FACSlyse reagent gave significantly lower estimates of the CD34+ cell number compared with the fixative-free bicarbonate-buffered ammonium chloride RBC lysis solution. These results were consistent with those reported by others (19–24). Loss of CD34+ cells following treatment with fixative-containing solutions has been suggested to be a consequence of increased stickiness of fixed cells (24). Fixed cells may be lost from the cell suspension either by increased adhesion to the surface of the assay tubes or greater loss during washing. Gratama et al. (24) recommended that the wash steps be omitted to minimize cell loss associated with fixative-containing RBC lysis reagents.
Other investigators have cautioned the use of ammonium chloride RBC lysis reagents due to the observation that CD34+ cells undergo light scatter changes characteristic of apoptosis when exposed to such reagents (21). In the experiments described here, sample analysis was completed typically within 30 min following the addition of the ammonium chloride lysis solution. If the samples were not analyzed immediately, they were placed on ice following the 15-min lysis step. In some instances, samples were analyzed up to 60 min following the addition of lysis solution (results not shown). No change in the light scatter characteristics was observed with delayed sample analysis, seemingly a benefit of placing the tubes on ice prior to analysis (23). Based on these results, we adopted a fixative-free bicarbonate-buffered ammonium chloride solution as the preferred RBC lysis reagent.
In order to have in place a more robust CD34 assay for routine analysis of HSC products, in particular cord blood, we investigated the options of single-platform flow cytometry protocols. We preferred a method that utilized the ISHAGE gating strategy because of its robustness, sensitivity, and widespread use. Although the single-platform ISHAGE method using Flow-Count Fluorospheres is well established (15), there are disadvantages with the use of Flow-Count or with similar fluorescent bead suspensions. These include (1) ensuring a homogenous bead suspension prior to dispensing; (2) precise pipetting of the fluorospheres, as well as the sample; (3) presence of preservative in the bead suspension, which can have an adverse effect on cell viability (25), necessitating that the beads be added immediately prior to flow cytometric analysis; and (4) comparing the practicality of pack size versus cost-effectiveness (i.e., the current 200-test pack size of Flow-Count Fluorospheres combined with a 1-month expiry upon opening may not be cost-effective for many laboratories).
We were attracted to using TRUCOUNT tubes, as this approach eliminates a potential source of error associated with dispensing the beads. This product is ready to use, does not contain preservatives that interfere with cell viability, and each 25-test pack has a 2-month expiry upon opening. We modified the ISHAGE protocol by performing CD34 and CD45 labeling, RBC lysis, and 7-AAD staining in the TRUCOUNT tubes. Because the TRUCOUNT beads were small, modifications were required to the single-platform ISHAGE gating strategy compared with those described previously (15). Unlike Flow-Count Fluorospheres, a forward scatter threshold cannot be used for TRUCOUNT beads. Consequently, the threshold must be set on a fluorescence channel that defines the denominator cell population. We chose CD45 expression in the FL1 channel to set the threshold, allowing the FL3 channel to be used for the viability dye, 7-AAD. In order to reduce debris in the acquisition region, a polygonal exclusion gate was set in the forward scatter channel between the bead events and cells. The results obtained from cord blood samples, which had a wide range of CD34+ cell concentrations, showed good agreement compared with the original dual-platform ISHAGE protocol and the single-platform Flow-Count–ISHAGE protocol. We found this TRUCOUNT-ISHAGE method to be technically simple and time efficient. As has been noted by others (5, 6), particular care must be taken in dispensing the sample and RBC lysis reagent to ensure accuracy and reproducibility of the results. Reverse pipetting using calibrated pipettes is essential. Coefficients of variation of less than 10% were readily achievable. The TRUCOUNT-ISHAGE gating strategy described proved to be robust and efficient. Data acquisition was achieved within 5–6 min per sample and no manual adjustment of the gating regions was required, even for samples that were in less than pristine condition.
We also assessed the TRUCOUNT-ISHAGE protocol for the enumeration of CD34+ cells in mobilized peripheral blood and leukapheresis samples. The results obtained were comparable with those achieved using the dual-platform ISHAGE protocol (results not shown), indicating that this method is applicable for samples other than cord blood. We did not observe the “vanishing counting bead” phenomenon (26) in our small series of leukapheresis samples. However, the recommendation of ensuring sufficient protein in diluting media is notable.
In conclusion, we described a protocol that combines the attributes of TRUCOUNT tubes and the ISHAGE gating strategy to provide a robust, accurate single-platform absolute viable CD34+ cell-counting method. Although we established this assay for cord blood samples, this protocol would be equally applicable to other types of HSC samples as indicated by our preliminary results with mobilized peripheral blood and leukapheresis samples. This TRUCOUNT-ISHAGE protocol provides an alternative robust, time-efficient assay that is capable of readily achieving standardization across clinical laboratories.