WBC White Blood Cell. This is the number of leukocytes measured directly, multiplied by the calibration constant, and expressed as n x 103 cells/µL. Beckman Coulter. Perform calibration: -At the start of each stand, before you begin analyzing samples. -After you replace any component dealing with dilution. Learn more about Beckman Coulter. We enable science by offering product choice, services, process excellence and our people make it happen.

Contents • • • • • Principle of CASY technology [ ] Cell viability can be assessed based on the integrity of: the living cells have intact plasma membranes whereas membranes of dead cells are broken. When a cell is exposed to a low voltage field, the electric current cannot go through the intact membrane, which is an, if it is viable.

Otherwise, as the cellular membrane is broken, electric field can go through the injured cell as there are pores on their membrane. For a normal cell, its size cannot be smaller than its nuclear size, which is the criterion to distinguish between living cells and dead cells. As a result, when cells in an or a particular, they are aligned one by one to a precision measuring pore and exposed to the electric field, each of their information can be captured and the culture condition, including its concentration, viability and volume, can be analyzed. For example, when the living cells get greater volume and pass though the current flow, a greater pulse, in amp-1, can be generated and then amplified. As the cell size is related to the cell volume, a cell size profile in cell population can be produced in terms of pulse height. Since the cells are scanned at such high frequency, a precise result and a high resolution can be produced.

These results from each cell are cumulated and assigned in a calibrated multi-channel analyser with over 500,000 channels. So, for the CASY technology, as the cell, it can present data of each cell as a cell size distribution graph, which has 2 variables, the change in cell volume and that in cell viability. The materials passing though the apparatus can be gated.

For the newly invented equipments, they have an automatically lower threshold at 7 um, which can exclude small particles and cell debris in the cell culture. At the same time, there will be an upper threshold to prevent from cell aggregation for counting. However, some of users may set upper threshold to unlimited for cell size. Since the cell size of each cell type is varied, before doing gating, it should ensure the correct cell size is included during the cell size related experiment. Advantages [ ] Since the cell viability is determined by electric current exclusion, viability dyes such as and are not needed. Hence, cell viability determination need no longer be a terminal experiment.

This advantage permits subsequent tests using the cells such as viability after a further time interval. The given result will be very accurate because not only are all steps performed robotically, but are high throughput (e.g. A million events/second). CASY technology it is fast but also reliable and reproducible because of features such as the multi-channel analyzer for detecting and analyzing the pulse height generation. In fact, a channel means the pulse counted in a particular energy.

In the past, single channel analyzers were used the instruments. They can count the pulse in a narrow range only. So, they can analyze the cells for one or few times at the set frequency only. Once the electric current is changed during the cell transfer, it could not be detected. This can use not only a much time for analysis but also inaccurate cell counting result. However, for the multi- channel analyzer, it can scan the entire energy range and the pulses in each channel.

As there are more than 500, 000 channels for a cell counting, once a cell is pass through the measuring pore, there would be a lot of channel monitoring for 1 cell. As a result, the speed of CASY technology to obtain the information of cells can be very high. Applications [ ] One of CASY technology applications is electronic cell counter for determining cell number and their viability in a sample.

The equipment is shown as Fig. 3 and the Fig. 4 states the result, including the total cell count, as well as the percentage of debris, living cells and dead cells, displayed on the screen of the cell counter. (2005) has compared the CASY technology to two standard methods for cell viability measurement, including the neutral red uptake and.

They found that the most sensitive IC50 values, which were the closest to those in the literature, were performed by this electronic cell counter. Some toxicants in these experiments by using chemical methods would affect the mechanisms of the assays. So, the results would become invalid. However, for the electronic cell counter, it can not only monitor all the cells changes, even the cell necrosis, by various toxicants types and concentration, but also a complex mixture of toxicants in the cell culture. It would be seen that the progress changes of dying cells can be detected as well. On the other hand, all the results from electric cell counter could be transferred to the computers with common spreadsheet programs. No other specific software would be set up to every computer to obtain the result.

Differences between CASY and Coulter cell counting [ ] A is one of the other devices used for cell counting. Like CASY technology, this also uses electric current for cell counting. However, the difference between them is that there is an aperture called “sensing zone”, with a known volume of electrolyte in a coulter counter. When suspended cells pass through it, they would displace the equivalent volume of electrolyte in the sensing zone and cause a short term change of electric current across the aperture.

Since the circuit is to detect the change of current across it, any particles that can displace the electrolyte will be counted. It would be seen that the measurement of cells would be from a volume to another volume in the same sample. In contrast, the CASY technology incorporates no electrolytic reservoir in the aperture and the cells in the electrolyte can pass through the measuring pore. It would not be necessary to detect the cells from a batch to another batch but measure them continuously and smoothly. References [ ].

Abstract The enumeration of absolute levels of cells and their subsets in clinical samples is of primary importance in human immunodeficiency virus (HIV)+ individuals (CD4+ T- lymphocyte enumeration), in patients who are candidates for autotransplantation (CD34+ hematopoietic progenitor cells), and in evaluating leukoreduced blood products (residual white blood cells). These measurements share a number of technical options, namely, single- or multiple-color cell staining and logical gating strategies. These can be accomplished using single- or dual-platform counting technologies employing cytometric methods. Dual-platform counting technologies couple the percentage of positive cell subsets obtained by cytometry and the absolute cell count obtained by automated hematology analyzers to derive the absolute value of such subsets. Despite having many conceptual and technical limitations, this approach is traditionally considered as the reference method for absolute cell count enumeration. As a result, the development of single-platform technologies has recently attracted attention with several different technical approaches now being readily available.

These single-platform approaches have less sources of variability. A number of reports clearly demonstrate that they provide better coefficients of variation (CVs) in multicenter studies and a lower chance to generate aberrant results. These methods are therefore candidates for the new gold standard for absolute cell assessments.

The currently available technical options are discussed in this review together with the results of some cross-comparative studies. Each analytical system has its own specific requirements as far as the dispensing precision steps are concerned. The importance of precision reverse pipetting is emphasized. Issues still under development include the establishment of the critical error ranges, which are different in each test setting, and the applicability of simplified low-cost techniques to be used in countries with limited resources. Cytometry (Comm. Cytometry) 42:327–346, 2000.

© 2000 Wiley-Liss, Inc. • • Counting the absolute numbers of CD4+ T cells, CD34+ hematopoietic precursor cells (HPC), and residual white blood cells (rWBC) in whole blood by flow cytometry (FCM) represents different aspects of the same technical problem: each setting requires single- or multiple-color surface or nuclear fluorescence analysis linked to an absolute cell counting technology as well as to a logical gating strategy to isolate the relevant cells.

However, as the percentages and absolute cell levels in these clinical settings are widely different, diverging technological requirements are set in terms of sensitivity, precision, and counting accuracy for the tests performed. In this review, we critically assess the cytofluorometric and noncytofluorometric methods for absolute cell and cell subset counting in order to investigate how they can fulfill their clinical requirements. Most of the techniques we summarize here are commercially available as cell counting systems and/or kit packages.

Clinical Utility of Absolute CD4+ T-Cell Counting Selective CD4 lymphopenia had been shown to be the hallmark of acquired immune deficiency syndrome (AIDS), even before the human immunodeficiency virus (HIV, LAV, HTLV-III) had been discovered (). Ever since, absolute counting of CD4+ T lymphocytes remains the major laboratory tool for staging HIV-infected patients (). It provides information about how far the patients have progressed along the path of HIV disease, this parameter being complementary to HIV plasma viral load (). It is now universally accepted that the most suitable CD4 value for the laboratory diagnosis occurs when CD4+ T-cell counts fall below 200/μL blood or 14% of total lymphocytes (). CD4 counts are therefore important clinical tools to assess the likelihood of opportunistic complications, for the appropriate timing of preventive medications, and to document the effects of intensive antiretroviral (,,, ), and possibly cytokine, therapy. The absolute T CD4 level continues to be the best validated predictor of the likelihood of an opportunistic infection, whereas such risk has not yet been adequately related to plasma viral load ().

Moreover, the best predictor of opportunistic infections is the most recently confirmed CD4 cell count, despite the occurrence of substantially lower prior CD4 cell counts. The risk varies continuously over the spectrum of absolute CD4 cell counts (). Additional surrogate indicators of disease progression include the assessment of CD38 expression, as a measure of activation status of CD8+ T cells, of CD45RA/RO, and of CD28 on T cells (). In diseases other than HIV infection, the percent and absolute CD3+CD4+ T-cell count are also of clinical relevance.

These areas include solid-organ transplantation in patients in whom polyclonal antisera to human lymphocytes and monoclonal antibodies to CD3 and T-cell receptor (TCR) are used posttransplant to avert organ rejection (). The T-cell depletion in blood is observed by FCM and the dosage of antiserum or CD3 antibody may accordingly be adjusted. In addition, the emergence of T cells (CD2+,CD4+ and CD2+,CD8+ lymphoid cells) that down-regulated their CD3-TCR marker is monitored, pointing to one of the immunosuppressive mechanisms that contributes to the action of anti-CD3 antibody therapy (). Using this FCM approach, the relative immunosuppressive potency of the different antilymphocyte preparations and the duration of their long-term effects can also be studied (, ). Sustained CD4+ T depletion has been shown to be associated with the emergence of opportunistic infections and neoplasms including skin cancer (). During the periods of postchemotherapy and the recovery phase following bone marrow (stem cell) transplants (BMT), faster thymus-dependent T-cell regeneration is seen in juveniles. In adults, CD4+ T-cell regeneration can remain slow due to impaired thymic function ().

One important aspect is that following BMT and the reinfusion of CD34+ stem cells (HPC), the speed of CD4+ T-cell recovery may vary according to the technique used and the purity of the preparation (). This is because CD45R0+ CD4+ T cells of “memory” type in adults appear to regenerate from peripheral T cells and not from CD34+ HPC (). A prolonged reduction of CD4 counts is also a known problem following therapy with purine nucleosides, e.g., 2-chlorodeoxy-adenosine (cladribine) for hairy cell leukemia (, ). In other diseases, infections are also frequently associated with a reduced CD4+ T-cell level, both as a predisposing factor and as a consequence of an infected state (, ).

A reduced CD4+ T-cell level has been reported to be of prognostic value in cytomegalovirus (CMV) infection in immunocompromised patients (). Conversely, in pediatric respiratory syncytial virus, the infectious state seems to be associated with an increase in the CD4+ T-cell count (). Honeywell Dial Set Chronotherm Manual Arts there. T-cell count derangements have also been described in association with protein-caloric malnutrition, anorexia nervosa, lactation and iron deficiency (, ), smoking habit (), autoimmune thyroiditis (), bronchial asthma (), multiple sclerosis (), polymyalgia rheumatica (), and among synovial fluid cells taken from patients with rheumatoid arthritis ().

Finally, in some cancers, a reduced CD4+ T-cell count is often observed as an associated, nonspecific finding (). Clinical Utility of Absolute CD34+ Progenitor Cell (HPC) Counting In normal blood, CD34+ HPC circulate in very low numbers, i.e., 1,000/μL and CD34+ cells are at least 40–50/μL, or at least 10–20/μL according to certain protocols (, ). The CD34+ HPC yield of a leukapheresis procedure can be predicted rather accurately from the peripheral blood starting absolute CD34+ cell level (). Also, the comparison between the expected and obtained CD34+ HPC yield is a useful consistency control of the entire procedure. The final CD34+ HPC content of the leukapheresis bag is then counted as the percent and absolute CD34+ count per milliliter as well as the total CD34+ cell number per bag. The exact determination of CD34+ cells is important because the autotransplantation engraftment rate is proportional to the number of viable and functional CD34+ per kilogram patient's body weight (, ). Similar CD34+ HPC counts are also useful for cord blood transplants () and during allogeneic mobilized peripheral blood transplants () and BMT ().

The biological and technical aspects concerning the percent and absolute CD34+ cell assessment have been extensively reviewed by another EWGCCA task force () and validated procedure and guidelines are now available (, ). Robust and reliable percent and absolute CD34+ HPC counting methods and guidelines have been developed that can be applied to the widest range of specimens (peripheral blood from mobilized cancer patients and normal subjects, native and manipulated apheresis products, bone marrow, and cord blood; 47,51,53,59). An experimental stain-no-lyse procedure has been recently described (). Clinical Utility of rWBC Counting in Blood Products Leukocyte depletion of red cell, platelet, and fresh plasma preparations for transfusion to critical patients has been demonstrated to be a mandatory requirement to prevent a number of untoward events, such as febrile reactions, microorganism transfer (CMV, Epstein-Barr virus [EBV], Creutzfeldt-Jacob disease variants), alloimmunization, platelet refractoriness, and immunosuppression (, ).

Leukoreduction is performed by separation and/or filtering techniques that may take place both in blood banks and at the patient's bedside. Typically, a leukoreduced 250–300 mL blood product bag must contain. • • Lymphocyte selection by CD45 gating in lyse-no-wash (ammonium chloride) single-platform multicolor IF and TruCount beads. Left: Clearcut lymphocyte (L), monocyte (M), and polymorphonuclear (PMN) cell discrimination. Right: Presence of lymphocyte aggregates (arrow A) and immature erythroid cells or basophils (arrow B) that render the lymphocyte identification problematic. Stain-no-lyse procedures have been implemented so far on the FACSCount (Becton-Dickinson Immunocytometry Systems [BDIS], Braintree, MA) for CD4 counting (), and as an experimental analysis protocol for CD34+ cells ().

Precise Identification of Reference Populations From the definitions above, it may appear that dual-platform methods are difficult to perform due to their reliance on reference populations. Nevertheless, the identification of lymphocytes as reference standards has been greatly facilitated by the introduction of CD45 gating for the following reasons. When lymphocytes are identified by scatter (or morphological) gating using FSC versus orthogonal SSC, nonlymphoid cells (i.e., basophils, nucleated unlysed erythrocytes, monocytes) may contaminate the gates, whereas large lymphocytes and lymphoblasts may be inadvertently excluded. In 1990, Loken et al. () described an IF gating method based on CD45-fluorescein isothiocyante (FITC)/CD14-phycoerythrin (PE; 79). Using this method, also referred to as fluorescence backgating, the lymphocyte gate purity and the lymphoid cell recovery are easily calculated on the basis of the differential CD45 antigen density expression. CD45 is a pan-leukocyte marker expressed on polymorphonuclear cells (CD45+), monocytes (CD45++), and lymphocytes (CD45+++) at different intensities, respectively.

Monocytes are identified by their selective expression of the CD14 antigen. Once CD45 gating is employed, scatter gates can also be redefined as follows (): the highest possible fraction of lymphocytes (>95% CD45+++, CD14-) is included in the light scatter analysis gate and the purity of these CD45+++,CD14- lymphocytes in the light scatter analysis gate should be >90%. The percentage value of any lymphocyte subset obtained by a light scatter gate must then be corrected for this purity value. The purity and recovery rates are inversely related variables. When a gating policy is defined, a compromise must be sought between these two conflicting requirements.

An acceptable FSC/SSC gate on lymphocytes using CD45/CD14 backgating is virtually impossible in samples that contain immature myeloid cells. In samples with very low proportions of lymphocytes versus monocytes, basophils may also cause additional problems. Some analysis software systems automatically use CD45 gating for selecting relevant cells (e.g., Becton Dickinson MultiSET, Beckman-Coulter TetraONE System). However, the major limitation of the CD45/CD14 gating strategy used in the two-color IF system with replicate tubes is obvious: the analysis gate is set in Tube 1, which may not be an exact replica of the other tubes (Tube 2 onward) that contain the lymphocyte markers. The tube-to-tube variability is difficult to control; it has indeed been shown that after lysing the cells, the characteristics were no longer identical between the replicate tubes (). As a consequence, the two-color IF approach is no longer recommended by the Centers for Disease Control (14; but see other regulatory bodies, e.g., the NCCLS in ref.

Instead, recent modern recommendations (, ) emphasize the advantage of utilizing three to four-color IF techniques including CD45 in single tubes (see below). Thus, in three- and four-color immunophenotyping techniques, CD45 staining has recently been included as a primary (anchor) gating marker: lymphocytes are defined as CD45+++ with low SSC enabling a putative 100% lymphocyte gate purity (). The lymphocyte recovery can be calculated as CD3+ (T cells) + CD19+ (B cells) + CD16/56+ (NK cells) and then compared with the total CD45+++ /low SSC events. The most compelling reason for CD45/SSC gating is that one can provide percentage T-cell subset values from the same tube simultaneously. The preferred use of CD45/SSC gating for CD4+ T lymphocytes has been given priority in recent UK guidelines (). Finally, it is important to stress that the CD45 gating approach is not the only way to obtain a reliable absolute CD4+ count (see below; 81,90,116,117).

Logical gate strategies can also be used to avoid isotypic controls, both in lymphocyte and CD34+ HPC analysis (,, ). In CD34+ HPC dual-platform analysis, the reference cells are the total leukocytes and no morphologically or phenotypically defined cell subset can be identified as the reference population.

CD45 gating is also highly recommended when counting CD34+ HPC using the logical gate strategy developed by the International Society of Hematotherapy and Graft Engineering (ISHAGE; 119), and subsequently modified by others (,,, ). The ISHAGE protocol and its modifications may be the best current examples of routinely applicable logical gating. They represent the most widely accepted procedure for CD34+ HPC enumeration (,,,, ). The CD34+ cell events are filtered according to their CD45+ and SSC low expression. The latter gating is further refined by the verification of a homogeneous FSC/SSC pattern.

ISHAGE gating is particularly helpful in eliminating nonspecifically CD34-stained platelets, especially when very low percent and absolute CD34+ HPC are present (, ). The definition of the true reference leukocytes is, however, further complicated by the particular features of WBC occurring in the CD34+ HPC analysis setting. Peripheral blood may contain very low or very high total WBC, cord blood is particularly rich in cell debris, the mobilization protocols induce the release of a great deal of immature WBC and erythrocytes, and the leukapheresis procedure facilitates platelet clumping.

Therefore, additional means to better define reference WBC may be required. The use of nuclear dyes like LDS-751 or SYTO-type has been suggested (, ) or included in commercial kits like ProCount (Becton Dickinson Biosciences, San Jose, CA; 53,122). Moreover, apheresis samples may require dilution before staining, to obtain WBC within 10–20 × 10 9/L and ensure proper counting assay performance (). Bovine serum albumin (BSA)-supplemented phosphate-buffered saline (PBS) should be used as a diluent instead of plain PBS to avoid the occurrence of the vanishing bead phenomenon when using microbead-based counting techniques (). Specific Technical Issues of rWBCC The accurate FCM detection of very rare rWBC events in blood products introduces a number of challenging technical issues. RWBC enumeration has been traditionally performed with visual microscopy and large-volume Nageotte-type hemocytometers (, ). The visual counting method accounts for an intrinsic variance ranging from 25% to 91% (, ), it is quite operator dependent, time-consuming, and may not be sensitive enough in the lowest rWBC range, where some hemocytometers can give zero rWBC counts in a number of cases ().

An FCM counting technique using fluorescence signal detection by propidium iodide (PI) incorporation into the nucleus of permeabilized rWBC was introduced in this field by Dzik 10 years ago (, ). The subsequent technical development has been slow and difficult. Until recently, the FCM technical requirements for both rare event detection and absolute cell counting were not entirely established (). Several alternative FCM and hematological methods involving cell detection by phenotype and light scatter analysis have also been described (, ), but also judged as overall inefficient () or inaccurate with platelet concentrates ().

The filtration process was demonstrated to introduce cell activation and undesired changes in WBC light scatter properties (). More recently, significant improvements in FCM counting technology led to the development of more reliable rWBC detection and enumeration procedures.

Modern FCM techniques are able to acquire and store a large number of events at higher speed. The need for adequate fluidic circuit cleaning and increased sample flow rate as prerequisites for rare rWBC analysis were stressed (). Single-platform counting approaches with microbeads (), marker chicken RBC (), or volumetric technique () were also introduced. Although the WBC nuclear signal induced by PI is strong and clearcut, the need for a prolonged run time leads to the acquisition of many undesired background events. These events show a nonspecific diagonal fluorescence pattern that differs from that of rWBC and counting particles, as demonstrable in FL2 versus FL1 or FL3 displays.

They can be excluded by an appropriate instrument setup and gating (). The evaluation of virtually leukocyte-free blood products represents a challenge for FCM operators. In conventional rare event analysis, it is generally accepted that at least 100 positive events must be acquired to obtain adequate data representation. Following this assumption, the rWBC enumeration in a virtually leukocyte-free product may require the analysis of a very large sample volume over an unacceptably long acquisition time. This is an example where the concept of mathematical limits cannot be applied to practical FCM.

The monitoring of the absence of a rare population may require a different algorithm compared with the conventional detection of a rare cell subset. When microsphere-based methods are used, the number of beads acquired is a good indicator of the approximate blood volume analyzed, which is optimally 10–20 μL. An appropriate acquisition gate counter can be set in most instruments to evaluate this parameter ().

When volumetric approaches are used, the sample preparation protocols must be adapted to acquire at least 10–20 μL of the original blood sample (). In all currently available preparation methods, a detergent permeabilizes rWBC and lyses RBC and a nucleic acid dye (DAPI, PI, TO-PRO-3) stains the cell nuclei (). Two commercially available kits now exist that use TruCount microbeads (LeucoCount, Becton Dickinson Biosciences; 133) or volumetric capillary cytometry (CEQer PRP and RBC assays, Becton Dickinson Biosciences; 134). An appropriate analysis protocol has also been developed for the volumetric FCM Dako Galaxy (). Another simple counting protocol has been developed for Partec PA and CCA particle analyzers using UV-lamp excitation and DAPI incorporation.

Great care must be paid in pipetting and dispensing very viscous samples such as packed RBC and platelet concentrates. Platelet bags are definitely more difficult to check than RBC concentrates, because cell clumping and a high nucleic acid background tend to generate more nonspecific staining. The addition of RNAse to the staining medium seems to improve the detectability of rWBC in platelet concentrates.

The actual biological and clinical meaning of minimal rWBC transfusions is still being debated, mostly because the detection and counting methods available so far are not entirely satisfactory. THE STATE OF THE ART: SINGLE PLATFORM FOR ABSOLUTE CELL COUNTING To date, many different methods have been developed for commercial use in order to count the absolute number of CD4+ T cells, CD34+ HPC, and rWBC (Table ). Most of these methods have already been evaluated in multicenter studies (,,,,, ). Some analysis systems were marketed in the past and are briefly described here for historic purpose. All cytometric methods can be used to count both CD4+ T cells and CD34+ HPC.

For CD4 counts, a number of noncytometric options have also been available but these are not suited for CD34 assays. Most of the methods also include cell fixation and virus-inactivating reagents, although this commendable feature (and its efficacy) is not always detailed in the sample preparation protocols.

Digital Electronics Malvino Leach Ebook Free Download. • • Cross-comparison of conventional dual-color dual-platform FCM count versus FACSCount (A), Ortho Cytoron Absolute (B), and Trax CD4 ELISA (C). Study on 325 HIV+ subjects.

The Bland-Altman diagrams reporting the agreement between each method are shown in the lower row. The limits of agreement are calculated according to the bias (mean difference) ± 2 SD of the differences. The worst agreement is between FCM and TraX ELISA. Two-hundred and ten normal and HIV+ subjects were studied for absolute CD4+ and CD8+ T cells by dual-platform FCM and the IMAGN 2000 4T8 kit at the Legnano Hospital, Italy ().

The X slope and the correlation coefficient of FCM versus IMAGN 2000 were 0.87 and r 2 = 0.916 for CD4 and 0.84 and r 2 = 0.948 for CD8. The Bland-Altman statistics for CD4 analysis gave -224 and +242, respectively, as the limits of agreement and a bias of +9 over a range of 5–2,250 CD4+ T cells per microliter. For CD8, the limits of agreement were somewhat wider: -236 and +358, respectively, and the bias was +61 over a range of 165–3,230 CD8+ T cells per microliter.

Multicenter Studies on CD34+ Cell Counting: The Emerging Consensus on Single-Platform Techniques The first standardized dual-platform counting method for CD34+ cells was developed in 1994 as the Milan Protocol (). Despite its limitations (), it is still used in many centers. In the majority of early multicenter trials on CD34+ HPC enumeration, an alarming interlaboratory variability was seen. Result variation was generated by the usage of local nonstandardized staining and gating procedures () and by sample instability (). Using nonstandardized staining and analysis methods, the interlaboratory CVs of dual-platform CD34+ cell values ranged from 50% to 284% (, ), whereas the use of a predefined analysis protocol reduced the variability to a more acceptable 14–82% (,,, 177). Using the simple Milan Protocol, the interlaboratory CV was limited to 20% in a large study in northern Europe ().

Even using a carefully standardized procedure, however, 46% of variation resulted from nontechnical factors (i.e., sample type and quality), 10% from various documented technical flaws, and 44% from unexplained factors (). The lysing, fixing, and washing steps have been shown to be of major influence in CD34+ cell counting. Such cells are particularly vulnerable to fixative-containing lysing agents, which can cause a remarkable drop of CD34 molecule expression and absolute cell level, especially when lyse-and-wash procedures are used (,,,, ). Using lyse-no-wash techniques, consistently higher absolute CD34+ values were observed (, ). This suggests that the interlaboratory CD34+ cell counting variation might be further reduced if a standardized lyse-no-wash technique is coupled with single-platform technology. The single-platform counting techniques were introduced rather recently in the CD34+ PBSC analysis. Commercially available analysis kits and systems rapidly took place.

The use of the BDIS ProCount kit was shown to reduce remarkably both the intralaboratory and interlaboratory assay variability (,, ). The same was demonstrated for the Beckman-Coulter StemKit assay in a single-center study (). Less impressive results were obtained with the STELLer and IMAGN 2000 counting system both in single and multicenter studies, especially in the very low CD34+ level range (, ). Cross-comparison studies have shown an overall good agreement between different single-platform analysis methods (,,,, ), provided samples do not contain significant proportions of dead cells ().

Moreover, the positive effect on interlaboratory CV of specific technical training, along with the use stabilized samples and standardized analysis protocols, has been demonstrated. In a multicenter study, 24 expert laboratories performed repeated single-platform analysis of stabilized blood using a modified ISHAGE protocol and FlowCount beads. The interlaboratory CVs were 23.3%, 18.7%, and 10.8%, respectively, in three consecutive trials, indicating a clear learning effect on measurement quality (). Around 5% CV in intralaboratory reproducibility studies and 10% CV in interlaboratory trials on absolute CD34+ cell counting is now a target at hand.

This is a prerequisite for robust measurements in multicenter clinical trials involving CD34+ PBSC transplantation. In conclusion, single-platform enumeration of CD34+ cells can be now recommended as the new technical golden standard due to its definitely lower intralaboratory and interlaboratory variability (). The Issue of Clinically Significant Errors In monitoring the CD4 or CD34 absolute counts, we must take into account a poorly defined issue, namely, the magnitude of the measurement variability that can influence clinical decisions. A preliminary attempt to define the critical error boundaries for absolute T CD4+ cell count has been performed at the S. Martino Hospital/University of Genova by Kunkl et al.

The data came from comparative analyses of 24 HIV+ and normal blood samples by 18 laboratories participating in a regional quality control program. For each dual-platform sample, true CD4+ cell value was defined by the consensus mean and the 99.9% CI of the mean was calculated after the elimination of outliers and data without internal consistency (a total of some 300 valid data). Regression of the upper and lower confidence limits of the 24 samples was used to extrapolate confidence ranges for any theoretical CD4+ value. CIs of true CD4+ values have been taken as a measure of the error due to the variation of CD4+ counts. As an example, with this method, the 99.9% CI in the case of measured CD4+ of 100 and 300 CD4+ T cells per microliter were 80 and 120/μL and 254 and 346/μL, respectively.

This matter must also be viewed in the light of physiological, diurnal, and circannual variability of CD4+ T-cell counts (). CONCLUDING REMARKS Reliability of FCM in Clinical Service Until recently, the commercially available conjugated antibodies were limited to FITC/PE/third-color combinations. These reagents could be employed on the majority of FCM. In recent years, however, the development of various four-color staining and analysis stategies has reduced the wide reagent interchangeability that existed in the past. This technical advance may partly impair the full cross-comparison of methods.

Pipetting precision is an overall major issue in every absolute CD4+ T-cell, CD34+ cell, and rWBC counting procedure. High precision pipetting requires appropriate calibrated dispensers, manual or electronic reverse pipetting, periodic maintenance, volume calibration of dispensing devices, and the training of personnel ().

When looking at external quality assurance programs on CD4+ and CD34+ cell count with reverse pipetting, the interlaboratory CV could be greatly reduced with the use of stabilized blood samples, unified staining and analysis protocols (, ), and the targeted training of the involved personnel (,, ). Even people from very experienced laboratories may experience difficulty when approaching single-platform absolute counting procedures for the first time.

The single-platform techniques can be now considered as the new golden standard for CD4+ and CD34+ cell counting. This is because many well-controlled studies clearly indicate the definitely lower intrainstitutional and interinstitutional variability of these methods. Each center must perform adequate training on single-platform procedures, which may require more technical skill than the ordinary dual-platform techniques. The CD4+ T-cell counting methods so far credited with the lowest interlaboratory CV are the single-platform Ortho Cytoron Absolute and the Becton Dickinson FACSCount with values both less than 4% CV, a considerable technical achievement in FCM (, ). To date, several alternative FCM techniques exist for absolute CD4+ T-cell absolute counting. Conceptually, every methodological aspect can be criticized (see above), but most of them can be credited as valid for routine clinical use.

It is interesting to note that the major single- and dual-platform FCM techniques are based on entirely different concepts and biological premises but perform equally well. After adequate technical cross-comparison studies are completed, clearcut data on the superiority of FCM counting of rWBC over conventional techniques will also be available. A Dutch consortium between blood banks is currently working intensively on that matter and they are going to publish a paper shortly. Future Developments Stain-no-lyse procedures for absolute cell subset counting can be considered as a remarkable step forward because the flaws introduced by lysing agents have been clearly demonstrated. The first applicable no-lyse technology was developed for CD4 counting on the FACSCount instrument (). Strangely, no further developments of such a promising and effective technique have occurred since that time.

Quite recently, an experimental stain-no-lyse procedure for absolute CD34+ cell counting using SYTO-13 nuclear staining was described (). Besides the established absolute cell counting techniques, other experimental issues are on the verge of a full clinical application. These include the absolute counting of blood CD34+ cells in myelodysplastic syndromes () and of total cells in bone marrow aspirates (), which may have relevance in BMT outcome and possibly also in leukemia diagnosis and staging. Interesting applications are emerging in quantitative and qualitative sperm analysis () and in the detection and counting of rare antigen-specific CD8+ cell subsets by HLA-tetramer technology (, ). Single-tube, multiple-color systems are brilliantly simple and precise, but expensive.

Counting microbeads are delicate, high-technology products, which introduce a remarkable extra cost to single-platform procedures. CD4+ T cell per microliter count, along with the minimum internal consistency control, can be obtained now with the use of four antibodies in a single tube using four-color bead-based procedures provided by Becton Dickinson and Beckman-Coulter, which represent state-of-the-art technology. However, an issue of great concern is the cost of each methodology (,,, ).

Cost is a major limiting factor for each laboratory testing, including CD4+ T-cell and CD34+ cell count. It is still questionable whether the use of alternative non-FCM technology may determine a real cost-saving; the answer is likely no. Why is FCM, the golden standard technology for absolute cell counting, still such a high-cost procedure?

What can be done to enable countries with limited health care resources to satisfy the increasing demand of locally affordable testing to cope with the ever increasing AIDS epidemics? We spent the last decade learning how to identify and count cells properly. This process led to increasing precision, but also to increasing complexity and costs. Now that all the assay variables have been identified in detail, some effort to simplify the technical burden and save resources seems warranted. It is indeed a necessary process that took place similarly in many technical fields. Reliable FCM CD4+ cell counts may be generated with newly designed low-cost minimalist procedures that are demanded by countries with limited resources (Fig.

The widespread clinical applicability of greatly simplified and cheaper assay systems (including newly designed instruments) seems, however, in contrast with the current recommendations of influential institutions like the CDC. • • Bland-Altman comparison of single-platform absolute CD4 counting employing full technology (Ortho Cytoron plus Trio Immunocount) versus reevaluation of single-color primary CD4+ cell gating on the same FCS files (minimal technology). The mean difference is -4, the CIs are -6 and +15, the lower and upper limits of agreement are indicated, along with linear regression. An isolated outlier (-650) had T-cell lymphoma. Data modified from Janossy et al. The procedure cost and assay complexity must also be weighed in comparison to another crucial issue, namely, the critical error we can afford with CD4+ T-cell and CD34+ cell count assays, which is much tighter in the latter case.

In other words, if the clinical decision-making threshold is narrow, every technical effort should be made to ensure a very precise and accurate absolute cell level measurement. This is typically the case of the delicate CD34+ PBSC transplantation. Looking back at the past few years, it is interesting to note that a number of consensus threshold values for biological testings have been established without any concern about the assay sensitivity, accuracy, and intrinsic variability (i.e., 10. In case sample dilution is required, always use PBS supplemented with 1% BSA to ensure an adequate protein content. • 3At least 1,000 bead events must be included in the sample list mode file (best 2–3,000). The relevant cell events must not exceed more than four to five times the bead events to ensure proper statistic robustness.

• 4When the total analyzed sample amount is the measurement endpoint (e.g., in leukoreduced blood products analysis), the number of acquired microbeads is a rough indicator of how much sample has been processed. • 5Ensure proper instrument setup and sample injection pressure to acquire at rates not higher than 3,000–4,000 events per second. This helps to minimize event loss for coincidence and to reduce bead peak CV. • 6Ensure that highly intense microbead fluorescence signals are as much as possible on scale. This helps to avoid prolonged photomultiplier tube (PMT) blinding and event loss due to PMT oversaturation (increased PMT dead time). • 7Ensure that sheath fluid tank is over 50% full.

This seems to increase flow stability. • 8Avoid bubble formation in any instance. Bubbles capture beads by capillarity and remove them from the suspension.

• 9Vortexing can be applied with care, provided proteins are present in the resuspension medium. Vortexing start must be gradual, and the mechanical mixing must not exceed 5 s at maximum 50% of tube height. Mixing samples by inversion after tube capping is also applicable.

• 10During very lengthy sample runs (i.e., more than 8–10 min), beads tend either to sediment or to float, thus causing a change in the proportion of cells and beads simultaneously aspirated. An additional mixing is therefore warranted if the run lasts more than 8–10 min.

Additional Usage Tips for Beckman-Coulter FlowCount Beads The following guidelines may be in some contrast with official manufacturer's instructions. • 1Always keep the bead bottle in upright position and tightly stopped. • 2FlowCount beads completely float at the suspension medium surface if the bottle is kept unperturbed for more than 12 h.

• 3The first bottle opening must be preceded by a very thorough mixing and the aluminum foil seal must be completely removed after the first mixing. • 4Mark the date of the first opening and use the bottle preferably within 1 month. • 5The bottle must be vortexed once a day, in the morning, then gently mixed by inversion just before any usage during the same day. • 6The same pipetting precautions described above must be applied to the drawing and dispensing FlowCount beads. • 7FlowCount beads must be dispensed using exactly the same pipette and tip type used for the primary sample. • 8After adding FlowCount to lysed sample, mix by gentle vortexing (as described above) or by inversion after tube capping.

• 9Analyze immediately after mixing or keep the samples in melting ice in the dark until analysis. • 10Instrument setup can include FSC as the primary threshold and trigger, provided low-volume debris and electrical noise are appropriately gated out during acquisition. • 11Take into account only bead singlets. Include time as an additional parameter to better define bead singlets and to monitor possible fluidic perturbations in the acquisition process. Try to keep bead singlet CV for FSC and fluorescence within 3%. • 12FlowCount beads generate end-scale fluorescence signals when used on Becton Dickinson instruments set up for routine IF analysis.

Ensure that the bead events are fully on scale in at least one fluorescence channel. • 13FlowCount beads are not excited by He-Ne red lasers in instruments equipped with dual-laser configuration. Additional Usage Tips for Becton Dickinson TruCount Beads The following guidelines may be in some contrast with official manufacturer's instructions. • 1Always keep the TruCount tube bag carefully airtight (best with adhesive tape) and use an opened bag within 1 month. • 2Discard tubes where bead bolus is fragmented or if bead dust is evident.

• 3Discard the entire sample if the bead bolus does not dissolve quickly and completely. • 4Ensure maximum pipetting precision when using TruCount beads. All the measurement precisions rely on this step and it is not compensated by the dual-step pipetting (i.e., sample and bead pipetting) as with FlowCount. • 5TruCount beads must be used with primary threshold and trigger on a fluorescence channel, because they are too small to be acquired when FSC is used as a threshold parameter.

• 6A fluorescence marker must be used to define at least the relevant cell population to be analyzed. • 7TruCount beads are some 10 times less brilliant than FlowCount beads. This implies that end-scale bead events are usually not generated, but that some interference between cell and bead signals can occur. • 8All bead events, including doublets and multiple aggregates, must be taken into account for absolute count calculation. This is best accomplished with a dual-fluorescence display and a upperright corner window including end-scale events. • 9TruCount beads tend to sediment with time.

• 10TruCount beads are excited also by the He-Ne red laser. Additional Tips for Volumetric Counting Systems • 1Maximum dispensing precision and accuracy must be applied in any step. However, the primary sample dispensing still has the major influence on overall accuracy. • 2If a limited amount of final sample is aspirated by volumetric instruments, verify that the relative proportion between primary sample and lysing is set to ensure the acquisition of the minimum positive cell events or the minimum primary sample volume. • 3Avoid inadvertent final sample dilution by spillage of sheath fluid drops from the sample injection port.

Ancillary Article Information. • 1 Fauci AS, Macher AM, Longo DL, Lane HC, Rook AH, Masur H, Gelmann EP. NIH conference. Acquired immunodeficiency syndrome: epidemiologic, clinical, immunologic, and therapeutic considerations. Ann Intern Med 1984; 100: 92– 106. • • • • • 2 Valentine ME, Jackson CR, Vavro C, Wilfert CM, McClernon D, St Clair M, Katz SL, McKinney RE Jr.

Evaluation of surrogate markers and clinical outcomes in two-year follow-up of 86 HIV-infected pediatric patients. Pediatr Infect Dis J 1998; 17: 18– 23. • • • • • 3 Guarner J, Montoya P, del Rio C, Hernandez-Tepichin G. CD4+ T-lymphocyte variations in patients with advanced human immunodeficiency virus infection and counts below 100 cells per microliter. Cytometry 1997; 30: 178– 180. • • • • • 4 Spino C, Kahn JO, Dolin R, Phair JP.

Predictors of survival in HIV-infected persons with 50 or fewer CD4 cells/mm 3. J Acquir Immune Defic Syndr Hum Retrovirol 1997; 15: 346– 355. • • • • • 5 Vanhems P, Allard R, Toma E, Cyr L, Beaulieu R. Prognostic value of the CD4+ T cell count for HIV-1 infected patients with advanced immunosuppression. Int J STD AIDS 1996; 7: 495– 501. • • • • • 6 Rabeneck L, Hartigan PM, Huang IW, Souchek J, Wray NP.

Predicting outcomes in HIV-infected veterans: I. Progression to AIDS.

Survival after AIDS. J Clin Epidemiol 1997; 50: 1231– 1248. • • • • • 7 Keet IP, Janssen M, Veugelers PJ, Miedema F, Klein MR, Goudsmit J, Coutinho RA, de Wolf F. Longitudinal analysis of CD4 T cell counts, T cell reactivity, and human immunodeficiency virus type 1 RNA levels in persons remaining AIDS-free despite CD4 cell counts 5 years. J Infect Dis 1997; 176: 665– 671. • • • • • 8 Sabin CA, Mocroft A, Bofill M, Janossy G, Johnson M, Lee CA, Phillips AN. Survival after a very low (.