Measuring Neutrophil Phagocytosis

Transcription

1 JOURNAL OF CLINICAL MICROBIOLOGY, Aug. 1992, p /92/ $02.00/0 Copyright 1992, American Society for Microbiology Vol. 30, No. 8 Rapid Whole-Blood Microassay Using Flow Cytometry for Measuring Neutrophil Phagocytosis CATHY WHITE OWEN,12* J. WESLEY ALEXANDER,1'2 R. MICHAEL SRAMKOSKI,2 AND GEORGE F. BABCOCK1' 2 Department of Surgery, University of Cincinnati College ofmedicine, 231 Bethesda Avenue, Cincinnat4 Ohio 45267, 1* and Cincinnati Unit, Shriners Burns Institute, Cincinnati Ohio Received 18 February 1992/Accepted 19 May 1992 A simple flow cytometric method (FCM) for measuring phagocytosis of Staphylococcus aureus by human neutrophils (polymorphonuclear leukocytes [PMNs]) is described. This assay utilizes 100 I of EDTAanticoagulated whole blood and a simplified method of fluorescently labeling bacteria. A commercially available whole-blood lysing reagent allows for the removal of erythrocytes and the exclusion of external free or adherent bacteria. Phagocytized bacteria are unaffected by this reagent, so PMNs containing internalized bacteria can be easily identified by FCM. Advantages of this method include the following: (i) small sample size, (ii) no requirement for PMN separation, (iii) rapid reliable method of labeling the bacteria, (iv) ability to distinguish between adherent bacteria and those which are actually internalized, (v) avoidance of vital dyes as quenching agents, and (vi) ability to fix cells and store for future FCM analysis. The clinical importance of neutrophil (polymorphonuclear leukocyte [PMN]) function, particularly with regard to phagocytosis of microorganisms, has been well established (1, 8, 10). The use of flow cytometry (FCM) with fluorescently labeled particles (bacteria, yeast cells, or latex beads) has facilitated the investigation of this important PMN function (3, 7). Early FCM methods provided no reliable distinction between internalized particles and those which may be simply adherent to the cell surface (3, 12). More recently, quenching of external fluorescence has been accomplished with the use of vital dyes such as ethidium bromide (EtBr) (9), trypan blue (2), or crystal violet (8, 13). The use of these dyes, however, is quite cumbersome, creates technical problems in FCM methods, and precludes the use of fixatives which would otherwise allow delayed FCM analysis. It is well known that ph changes may significantly affect the fluorescence of fluorescein compounds, most notably fluorescein isothiocyanate (FITC), which is a common agent used to label particles for phagocytic assays (4, 5, 6). In this article, we describe a new FCM method which utilizes FITC-labeled Staphylococcus aureus bacteria and a 100-pl whole-blood sample requiring no leukocyte (WBC) separation. After incubation of blood with bacteria, phagocytosis is halted and erythrocytes (RBCs) are lysed with a formic-acidbased commercially available reagent (Immuno-Lyse; Coulter Corp., Hialeah, Fla.) which concurrently quenches the fluorescence of external nonphagocytized bacteria. The use of vital dyes is avoided, delayed analysis is possible, and only small blood volumes are required. MATERIALS AND METHODS Labeling of bacteria. S. aureus organisms were cultured for 16 h at 37 C in Trypticase soy broth (Baltimore Biological Labs, Baltimore, Md.) with FITC (Research Organics, Inc., Cleveland, Ohio) at a concentration of 30 to 50 jig/ml. Bacteria were then washed twice in Dulbecco's phosphatebuffered saline (DPBS) (GIBCO Laboratories, Grand Island, * Corresponding author N.Y.) and heat inactivated at 60 C for 30 min. They were again washed and resuspended in DPBS to a final concentration of 109 bacteria per ml. The organisms were examined by fluorescence microscopy for uniformity of FITC staining. Confirmation was provided by FCM. The bacterial suspension was then aliquoted and frozen at -70 C in a lightprotected environment. Blood samples. Blood was obtained from 20 normal healthy adult volunteers, collected in sodium EDTA, and stored at room temperature for no longer than 6 h prior to use. More prolonged storage resulted in suboptimal PMN isolation and altered phagocytosis (data not shown). Total WBC counts were performed on a Neubauer hemacytometer by using 3% acetic acid lysis of RBCs. If the WBC count exceeded 107 cells per ml, the whole blood was diluted with DPBS to bring the WBC count within the range of 3.0 x 106 to 9.9 x 106 cells per ml. When isolated PMNs were used for the assay, as a comparison control for the whole-blood assay, they were isolated in the following manner. Five milliliters of whole blood, collected in sodium EDTA, was layered over a Ficoll gradient consisting of 4.4 ml of Ficoll- Hypaque (Pharmacia, Uppsala, Sweden) and 0.6 ml of 85% Hypaque (57% meglamine ditrizoate-28% sodium ditrizoate in H20). The gradient was centrifuged at 500 x g for 30 min at room temperature. This process resulted in two clearly defined bands of WBCs, the upper band consisting of mononuclear cells (including monocytes and lymphocytes) and the lower band consisting of PMNs and some minor RBC contamination. The majority of the RBCs went through the gradient and pelleted at the bottom. Contaminating RBCs were removed by H20 lysis. This separation technique yielded a >98% pure population of granulocytes with >98% viability by trypan blue exclusion. The average PMN yield was approximately 65%. PMNs were collected and washed twice in DPBS and resuspended to a concentration of 5 x 106 cells per ml. Phagocytosis assay. In 20 separate experiments, 100,ul of whole blood, diluted whole blood, or isolated PMNs was dispensed into polystyrene tubes for each time point, namely, at time zero and at 2, 5, 10, and 15 min. Whole blood or diluted blood samples were washed twice in DPBS to

2 2072 WHITE-OWEN ET AL. remove EDTA and plasma. Opsonization was accomplished by resuspending cells in 100,ul of pooled normal human type AB serum in each tube. After a 3- to 5-s sonication, 10,ul of stock FITC-labeled bacterial solution (109 bacteria per ml) was added to each tube (10:1, bacteria/cells). Cytochalasin D (Sigma Chemical Co., St. Louis, Mo.) (final concentration, 5,uM) was added to the time zero tube immediately. The remainder of the tubes were incubated for the appropriate time periods in an agitating 37 C water bath. Phagocytosis was stopped in each tube by the addition of cytochalasin D at a final concentration of 5 pm. Tubes were placed immediately on ice, and 3 ml of ice-cold DPBS was added to each. For each blood sample, one tube containing cells only (no bacteria) was carried through the procedure as a control to evaluate the background fluorescence of the PMNs alone. Whole-blood lysis. All tubes were washed twice in cold DPBS. RBCs were lysed with Immuno-Lyse (Coulter) by following the prescribed protocol. Briefly, the lysis reagent was diluted 1:25 with DPBS. One milliliter of this solution was then added to each tube. All tubes were briefly vortexed immediately and at 1 min after addition of the reagent. Tubes were maintained on ice, and at 2 min, the lysis was halted by the addition of 250,ul of Coulter fixing reagent. After sufficient washing with DPBS to remove all free hemoglobin, the cells in each tube were either fixed with 0.5 ml of ice-cold 1% paraformaldehyde solution containing 1% sodium cacodylate and 0.8% NaCl or prepared for immediate FCM analysis by the addition of 0.5 ml of ice-cold DPBS. Paraformaldehyde-fixed samples were stored in a light-protected environment at 4 C for future FCM analysis. Those experiments involving separated PMNs also included treatment with Immuno-Lyse so that the effects of the process on adherent external bacteria and on isolated PMNs could be evaluated. Again, cells were either fixed with paraformaldehyde and stored or placed in DPBS for immediate FCM analysis. FCM. FCM was performed with a Coulter Epics 753 FCM (Coulter Cytometry) by using a 488-nm line of an argon ion laser. Green fluorescence was collected by using a nm band pass filter and linear amplification. Data were collected and analyzed by using the Coulter Easy-2 software. Non-PMN debris were excluded from analysis by using forward and 90 -angle light scattering. Adherence versus phagocytosis. The effect of the wholeblood lysis technique on external free bacteria was tested by processing free FITC-labeled bacteria alone with the Immuno-Lyse reagents as described above. The effect of Immuno- Lyse on external membrane-bound bacteria was evaluated by arresting phagocytosis with cytochalasin D before incubation of cells with the fluorescent S. aureus organisms. Briefly, whole blood, diluted whole blood, or isolated PMNs were pretreated with cytochalasin D (final concentration, 5,M) to inhibit phagocytosis. Bacteria were then added to each tube, and the assay was carried out according to the protocol. In this system, phagocytosis, not bacterial adherence, is inhibited. FITC fluorescence of PMNs was evaluated in tubes which did not undergo lysis (purified PMNs only) and in tubes which were lysed (purified PMNs and whole blood). These results were compared with the results obtained by using the technique of Fattorossi et al. (9), in which EtBr is used to quench FITC fluorescence of free or membranebound microorganisms. Briefly, adherent or free FITClabeled organisms were stained with EtBr (final concentration, 50,ug/ml) for 5 min at 4 C after the phagocytosis assay had been performed. By the process of resonance energy CO) 0 Fluorescence Intensity FIG. 1. (A) FITC fluorescence intensity (FI) of heat-killed, sonicated bacteria grown in Trypticase soy broth in the presence of 30 pg of FITC per ml; (B) Fl of unsonicated FITC-labeled bacteria; (C) Fl of FITC-labeled bacteria after treatment with Immuno-Lyse. transfer, FITC fluorescence was quenched. Since it is well known that the EtBr does not penetrate intact cell membranes, internal ingested bacteria cannot be affected by EtBr, and, thus, the bacteria retain their green fluorescence. Therefore, green fluorescence of any PMN after EtBr quenching can only be explained by the presence of internal ingested bacteria. RESULTS J. CLIN. MICROBIOL. FITC labeling of S. aureus. In the initial experiment, S. aureus was grown in broth containing 50,g of FITC per ml. The bacteria were brightly and homogeneously stained. They retained their fluorescence through the heat killing process and through several freeze-thaw episodes as well (data not shown). In later experiments, similarly good results were obtained when the bacteria were grown in as little as 30,ug of FITC per ml of broth (Fig. 1A). Cultures of S. aureus

3 VOL. 30, 1992 were labeled with FITC on six separate occasions. In all experiments, the bacteria were adequately stained. However, the intensity of staining did vary among cultures. Therefore, it is recommended that with each new culture, FITC fluorescence should be checked on the FCM and the instrument should be recalibrated accordingly. Fluorescence was stable for at least 3 months when suspensions were stored at -70 C in a light-protected environment. After the bacterial suspension had been thawed, a brief (3- to 5-s) period of ultrasonification was required before the introduction of the bacteria into the phagocytosis system to break up clumps of bacteria which, on FCM, may be perceived as poor or heterogeneous staining of bacteria (Fig. 1B). This method of fluorescently labeling bacteria is also quite satisfactory for the gram-negative bacteria Escherichia coli (data not shown). Effect of Immuno-Lyse on free bacteria. When free FITClabeled bacteria were subjected to the lysis process, it was discovered that nearly all FITC fluorescence was quenched. Figure 1C demonstrates FITC fluorescence of these bacteria after treatment with Immuno-Lyse in one experiment. There was an average reduction of FITC fluorescence of 78 ± 3% (standard error of the mean) in all experiments after lysis. By using the EtBr quenching technique, there was an average reduction of FITC fluorescence of 69 ± 6%. Thus, our method compares favorably with that of Fattorossi et al. (9). These results were reproducible in 10 separate experiments. Effect of Immuno-Lyse on isolated PMNs. Figure 2 illustrates the effect of Immuno-Lyse treatment on Ficoll gradient-isolated PMNs. This figure represents the results of one experiment which typifies the outcome obtained in 10 experiments. There were significant changes in cell size and granularity as assessed by light scatter histograms after Immuno-Lyse treatment of these cells (Fig. 2B) when compared with isolated PMNs which had not undergone lysis (Fig. 2A). A similar effect was not seen after whole-blood lysis (Fig. 2C). PMNs from whole-blood lysis fell into essentially the same gate as purified PMNs which have not undergone lysis treatment. This would suggest that the use of Immuno-Lyse on purified PMN populations is not advisable and that, indeed, a whole-blood assay is preferable. However, to further clarify this effect of Immuno-Lyse on isolated PMNs, a series of experiments was performed in which various volumes of lysis reagents were used. Figure 3 shows the results of these experiments. Figure 3A again demonstrates the dramatic effect of Immuno-Lyse on isolated PMNs when 1.0 ml of lysis reagent was used. Figures 3B and C show the effects of using 0.5 ml or 0.25 ml of lysing reagent, respectively. Clearly, the effect on cell size and granularity is almost completely obliterated with the use of less lysis reagent. The use of these lower volumes of lysis reagent, even as little as 0.25 ml (data not shown), had no effect on the ability to quench external bacterial FITC fluorescence. Therefore, if Ficoll gradient-isolated PMNs must be used, reduction of the volume of Immuno-Lyse is strongly advised. Effect of Immuno-Lyse on membrane-bound bacteria. Table 1 indicates the percentage of PMNs which have cellassociated FITC fluorescence at each time point (time zero and at 2, 5, and 10 min) in 10 phagocytosis experiments which involved either purified PMNs which were not treated with Immuno-Lyse (group A), purified PMNs which were treated with Immuno-Lyse (group B), and purified PMNs which were prevented from phagocytizing with cytochalasin D and were not treated with Immuno-Lyse (group C). Cell-associated FITC fluorescence in the group A experi- WHOLE-BLOOD MICROASSAY FOR PHAGOCYTOSIS 2073 C') -J~~~~~~~~~~~~~~~~~~~N Ca > *~~ FALS FIG. 2. (A) Light scatter histogram of purified PMNs before treatment with Immuno-Lyse. (Note the small agranular population of cells represents contaminating RBCs.) (B) Histogram of purified PMNs after Immuno-Lyse treatment. (Note the significant change in cell size [FALS] and granularity [9OLS].) (C) Histogram of whole blood after Immuno-Lyse treatment. (Note a slight change in cell size but granularity is preserved.) This sample also contains lymphocytes and monocytes. ments was the result of both phagocytized and adherent bacteria, since these cells did not undergo treatment with Immuno-Lyse. Fluorescence in the group B experiments was the result of internalized bacteria only, while fluorescence in group C, the arrested phagocytosis group, resulted from membrane-bound bacteria only. Theoretically, the sum of the results of groups B and C should approximate the percentage of fluorescent cells in group A for each time point. Again, Table 1 demonstrates this comparison. Indeed, the value for groups B+C appears to approximate that of group A fairly well. However, values for B+C do tend to be slightly lower than A values at each time point. Because of the previously mentioned effect of

4 2074 WHITE-OWEN ET AL. J. CLIN. MICROBIOL. A.' TABLE 1. Time Percentage of PMNs with FITC fluorescence after phagocytosis % PMNs with FITC fluorescence (±SEM)a Group A Group B Group C Group B+C Time zero minT min T minT10 63±2 25±3 32±5 57±6 a Group A, purified PMNs which did not undergo lysis assay; group B, purified PMNs which did undergo lysis assay; group C, purified PMNs in arrested phagocytosis assay which were not treated with RBC lysis reagent. Groups B+C should approximate the percentage of FITC fluorescent cells in group A for each time point. SEM, standard error of the mean. 00 FALS FIG. 3. Effect of various volumes of Immuno-Lyse on the light scatter histogram of Ficoll gradient-isolated PMNs. (A) A 1.0-ml volume of Immuno-Lyse was used. (Note the dramatic change in cell size [FALS] and granularity [90LS].) (B) An 0.5-ml volume of Immuno-Lyse was used. (C) An 0.25-ml volume of Immuno-Lyse was used. (Note that granularity and cell size are preserved.) Immuno-Lyse on purified PMNs (see above), we hypothesize that both B and C values may be artificially lowered since some of the fluorescent cells may not fall within the PMN gates on FCM after lysis. To reiterate this point, Table 2 compares the percentages of FITC fluorescent cells observed at each time point for purified PMNs which were treated with Immuno-Lyse with those of whole blood treated with Immuno-Lyse. These data support the hypothesis that Immuno-Lyse has the same effect on membrane-bound FITC-labeled bacteria as it does on free fluorescent organisms. Namely, Immuno-Lyse treatment extinguishes FITC fluorescence of external adherent bacteria. When the Immuno-Lyse method was compared with the method of Fattorossi et al. (9), which uses the vital dye EtBr to quench FITC fluorescence of external membrane-bound bacteria, favorable results were again obtained (Table 3). In Table 3, the percentages of cells demonstrating cell-associated fluorescence in three groups are compared: group I, purified PMNs which were not treated with Immuno-Lyse after phagocytosis; group II, treated in the same manner as group I but quenched with EtBr prior to FCM analysis; and group III, PMNs from the whole-blood Immuno-Lyse assay. A similar, if not somewhat greater, degree of FITC quenching is seen with the Immuno-Lyse method, thus confirming that this method does allow differentiation of intracellular phagocytized organisms from external membrane-bound bacteria. Ability of assay to detect phagocytosis. Figure 4 typifies the histograms obtained from 10 experiments using normal PMNs in this assay. Figure 4A demonstrates the fluorescence intensity (FI) at time zero, while Fig. 4B, C, and D demonstrate the FI at 2, 5, and 10 min, respectively. By 10 min, maximal FI is obtained when normal PMNs are used. However, PMNs from a variety of clinical patient populations may vary in terms of the time when maximal FI is obtained (data not shown). Effect of paraformaldehyde fixation on use of quenching vital dyes. When EtBr was used to quench bacterial FITC fluorescence in unfixed PMNs, our results were consistent with those of Fattorossi et al. (9), i.e., there was adequate quenching observed. However, when previously paraformaldehyde-fixed cells were treated with EtBr, no such quenching occurred. In fact, there was a dramatic increase in what was read as FITC fluorescence in cells treated in this way. These results would indicate that the use of quenching vital dyes such as EtBr after fixation with paraformaldehyde is not possible, contrary to the findings of Fattorossi et al. Reliability of results after paraformaldehyde fixation in the whole-blood Immuno-Lyse method. Comparisons of results TABLE 2. Effect of Immuno-Lyse on purified PMNs versus whole blood % Fluorescent PMNs (±SEM)r Time Immuno-Lyse-treated Immuno-Lyse-treated purified PMNs whole blood Time zero 6 ± 2 3 ± 1 2min 6 ± 1 4 ± 2 5 min 8 ± 3 17 ± 3 10 min 25 ± 3 46 ± 6 a SEM, standard error of the mean.

5 VOL. 30, 1992 a, _ E z 0 0 D FITC A WHOLE-BLOOD MICROASSAY FOR PHAGOCYTOSIS s30 Soo B T so~~~~~ MP i 2075 U- C T5 D A ltc II E M w. z 0~ ama., 0. FIG. 4. Fluorescence Intensity awm Soo to sb Fluorescence Intensity FITC Fl of normal neutrophils in phagocytosis assay at time zero (A), 2 min (B), 5 min (C), and 10 min (D). obtained immediately after the assay in unfixed cells and assays which were fixed and delayed (up to 3 months) demonstrated virtually no variation in results. All delayed results fell well within ±2 standard deviations of initial mean values of percentage of cells with FITC fluorescence. DISCUSSION Traditionally, FITC labeling of bacteria has been accomplished by the addition of FITC to heat-killed organisms, followed by a relatively prolonged incubation to ensure even and adequate staining. We have been able to eliminate the need for this step by adding the FITC directly to the medium at the time of inoculation with bacteria. Good results were obtained when the bacteria were grown in the presence of as little as 30,ug of FITC per ml of medium. Fluorescence was stable for at least 3 months when suspensions were stored at -70 C in a light-protected environment. The use of Immuno- Lyse in a whole-blood phagocytosis assay solved many of the problems associated with other FCM phagocytosis methods. Most methods require the isolation of PMNs by using density gradient centrifugation, which has been shown by several authors to alter morphologic as well as functional TABLE 3. Percentage of FITC fluorescence cells after phagocytosis in purified PMNs not treated with Immuno-Lyse (group I), EtBr-quenched purified PMNs not treated with Immuno-Lyse (group II), and PMNs from whole-blood lysis assay (group III) Time % FlTC fluorescence cells (±SEM)a in: Group I Group II Group III Time zero 13 ± 3 14 ± 3 3 ± 1 2 min 17 ± 1 12 ± 1 4 ± 2 5min 28 ± 5 15 ± 1 17 ± 3 10 min 63 ± 2 52 ± 7 46 ± 3 a SEM, standard error of the mean. properties of PMNs and which allows for potential uncontrolled selection of subpopulations of PMNs (15-17). It is, therefore, desirable to avoid these separation techniques whenever possible. The method described in the present article excludes the need for PMN separation by Ficoll density gradients or dextran sedimentation and allows the use of very small volumes of sample blood, thus providing a feasible way to study small volumes of blood such as those obtained from pediatric populations. Additionally, the use of Immuno-Lyse allows for the discrimination between external and internal bacteria, thus providing an accurate assessment of true phagocytosis. The use of vital dyes as quenching agents requires an additional incubation period and may, in some circumstances, foul the tubing and flow cell of the instrument. This can result in unwanted quenching of subsequent sample runs, and extensive cleaning of the tubing and flow cell is required after such experiments. Cantinieaux et al. (8) have described the use of paraformaldehyde as a cell fixative to allow delayed FCM analysis, a technique which is quite useful in a busy laboratory or where a core FCM facility is utilized. However, our attempts to use EtBr to quench FITC fluorescence in cells fixed with paraformaldehyde were totally unsuccessful. The fact that paraformaldehyde fixation permeabilizes the PMN cell membrane, as evidenced by extensive intracellular staining with EtBr, prevents the use of this agent for fixation. Because an intact cell membrane is essential to the principle of EtBr quenching of external fluorescent bacteria, it cannot be used as a quenching agent in paraformaldehyde-fixed cells. Indeed, there was an increase in fluorescence when EtBr was added to paraformaldehyde-fixed cells. This was most likely due to excessive intracellular EtBr staining. Although the latter fact would not be a problem with other vital dyes such as crystal violet or trypan blue, the effect of paraformaldehyde on the integrity of the PMN cell membrane would obviate the use of these agents when paraformaldehyde fixation is planned. These problems are completely elimi-

6 2076 WHITE-OWEN ET AL. nated by using this methodology. Paraformaldehyde fixation and delayed FCM are advantages of this methodology. Preliminary experiments have indicated an additional advantage of this technique. Since whole-blood lysis FCM techniques have been used for several years to identify cells which react with these fluorescently labeled monoclonal antibodies, cell surface staining with monoclonal antibodies labeled with fluorochromes other than FITC can be used in multicolor fluorescence procedures. By using a double- or triple-color immunofluorescence FCM method with phycoerythrin, Texas red, or allophycocyanin, we have been able to assess the phagocytosis of PMNs which have previously been stained with fluorescent monoclonal antibodies, thus evaluating the phagocytosis of subpopulations of PMNs without time-consuming panning or cell-sorting methods. One disadvantage of this technique, as with other methods which have used bacteria, is that the actual number of microorganisms ingested by each cell is difficult to assess. It should be noted that as bacteria are phagocytized, oxidative metabolic activity is initiated and phagolysosome acidification begins to occur. This will result in a decreased intensity of FITC fluorescence emitted by those bacteria contained within the phagolysosomes. In normal adults, it has been found that this effect becomes evident sometime between time zero and 15 min in this assay. A moderate degree of individual variability exists and may be more evident in ill-patient populations. Thus, it is advisable to use several time points when performing this assay to determine the point at which maximal intracellular fluorescence occurs. ACKNOWLEDGMENTS This work was supported in part by a grant from the Shriners of North America. We thank Lois Marchionne for her assistance in preparing this manuscript and Chris Haviland for technical assistance. REFERENCES 1. Axtell, R. A Evaluation of the patients with a possible phagocytic disorder. Hematol. Oncol. Clin. N. Am. 2: Bjerknes, R Flow cytometric assay for combined measurement of phagocytosis and intracellular killing of Candida albicans. J. Immunol. Methods 72: Bjerknes, R., and C. F. Bassoe Human leukocytes phagocytosis of zymosan particles measured by flow cytometry. Acta Pathol. Microbiol. Immunol. Scand. Sect. C 91: Bjerknes, R., C. F. Bassoe, H. Sjursen, 0. D. Laerum, and C. 0. J. CLIN. MICROBIOL. Solberg Flow cytometry for the study of phagocyte functions. Rev. Infect. Dis. 11: Bassoe, C. F., and R. Bjerknes Phagocytosis by human leukocytes, phagosomal ph and degradation of seven species of bacteria measured by flow cytometry. J. Med. Microbiol. 19: Bassoe, C. F., 0. D. Laerun, J. Glette, G. Hopen, B. Haneburg, and C. 0. Solberg Simultaneous measurement of phagocytosis and phagosomal ph by flow cytometry: role of polymorphonuclear neutrophilic leukocyte granules in phagosome acidification. Cytometry 4: Braunstein, J. D., A. Gorski, T. K. Sharpless, and M. R. Melamed Quantitation of granulocyte phagocytosis by flow cytometry. Fed. Proc. 35: Cantinieaux, B., C. Hariga, P. Courtoy, J. Hupin, and P. Fondu Staphylococcus aureus phagocytosis: a new cytofluorometric method using FITC and paraformaldehyde. J. Immunol. Methods 121: Fattorossi, A., R. Nisini, J. G. Pizzolo, and R. D'Amelio New, simple flow cytometry technique to discriminate between internalized and membrane-bound particles in phagocytosis. Cytometry 10: Galin, J Disorders of phagocytic function, p In J. Verhoef, P. K. Peterson, and P. G. Quie (ed.), Infections in the immunocompromised host: pathogenesis, prevention and therapy. Elsevier/North-Holland Publishing Co., Amsterdam. 11. Gelfand, J. A., A. S. Fauci, I. Green, and M. M. Frantz A simple method for the determination of complement receptor. J. Immunol. 116: Hausi, M., Y. Hirabayashi, and Y. Kobayashi Simultaneous measurement by flow cytometry of phagocytosis and hydrogen peroxide production of neutrophils in whole blood. J. Immunol. Methods 117: Hed, J The extinction of fluorescence by crystal violet and its use to differentiate between attached and ingested microorganisms in phagocytosis. FEMS Lett. 1: Hed, J., G. Haliden, S. G. 0. Johansson, and P. Larsson et al The use of fluorescence quenching in flow cytometry to measure the attachment and ingestion phases in phagocytosis in peripheral blood without prior cell separation. J. Immunol. Methods 101: Ogle, J. D., C. K. Ogle, J. G. Noel, P. Hurtubise, and J. W. Alexander Studies on the binding of C3b-coated microspheres to human neutrophils. J. Immunol. Methods 47: Seeds, M. C., J. W. Parce, P. Szejka, and D. A. Bass Independent stimulation of membrane potential changes and the oxidative metabolic burst in polymorphonuclear leukocytes. Blood 65: Verhoef, J., and F. A. Waldvogel Testing phagocytic cell function. Eur. J. Clin. Microbiol. 4: