MONOCYTE AND NEUTROPHIL ISOLATION

12/23/97

This procedure describes the method used for isolation of monocytes and neutrophils used in both the chemotaxis assay and microbicidal assay described below.

Principle

Heparinized whole blood is layered on top of a density gradient material ( Ficoll / Hypaque ) and subjected to a centrifugal force. Since erythrocytes (RBC) and PMN are more dense than the gradient material, they penetrate the gradient material and sediment to the bottom of the centrifuge tube. The lighter mononuclear cells (lymphocytes and monocytes) sediment to the plasma-density gradient interface. At equilibrium, the mononuclear cells are carefully aspirated and saved. The remaining plasma and gradient material are aspirated down to the RBC interface. The mononuclear cells are layered onto gelatin-coated flasks, allowing the monocytes to adhere. The nonadherent mononuclear cells (lymphocytes) are carefully decanted. The adherent monocytes are dislodged from the gelatin-coated surfaces with vigorous pipeting . The RBC cushion (which also contains PMN ) is resuspended to the initial whole blood volume. Dextran 500 is added to a final concentration of 1% and mixed. The RBC are allowed to sediment to the bottom of the centrifuge tube. The PMN remain in suspension. The PMN -rich supernatant is carefully aspirated.

Specimen

Ideally, 15 to 20 ml of heparinized whole blood (40 ml of monocyte chemotaxis assay) should be collected from both the patient and a normal control subject. Depending on the patient's absolute monocyte and/or neutrophil count, a sufficient number of cells may be obtained from a smaller volume.

Reagents and Supplies

Hanks balanced salt solution ( HBSS )

1% Gelatin: dissolve 2 g of endotoxin -free gelatin in 100 ml of phosphate-buffered saline (PBS)

Dextran T-500 ( Pharmacia )

6% Dextran: dissolve 6 g of Dextran T-500 in 100 ml of 0.85% of Nacl

0.05% Gentian violet in 2% acetic acid

Ficoll - Paque ( Pharmacia )

Procedure

1.	Dilute heparinized whole blood 1:2 with HBSS .

2.	Carefully layer a maximum of 30 ml of diluted whole blood over 10 ml of Ficoll -
Paque .

3.	Centrifuge for 30 min at 400 x g at room temperature.

4.	Carefully aspirate the mononuclear cells located at the plasma-Ficoll- Paque interface. Wash the mononuclear cells once in 10 ml of HBSS and resuspend in 3 ml of HBSS . The monuclear cells are now ready for adherence to gelatin-coated flasks as describe in step 14 below.

5.	Carefully aspirate the remaining plasma and ficoll- Paque down to the RBC peller (this fraction also contains PMN ).

6.	Resuspend the RBC- PMN pellet in HBSS to twice the original whole blood volume.

7.	Add 6% dextran to a final concentration of 1% and allow the RBC to settle for 1 h at room temperature.

8.	Carefully aspirate the PMN -rich supernatant and dilute 1:2 with HBSS .

9.	Centrifuge at 600 x g for 10 min.

10.	Vortex the PMN cell pellet. If there is significant RBC contamination, briefly lyse the RBC with 2 ml of ammonium chloride lysing buffer and dilute with 20 ml of HBSS .

11.	Centrifuge at 600 x for 10 min.

12.	Resuspend the PMN cell pellet in 3 ml of HBSS .

13.	To 0.190 ml of 0.05% gentian violet add 0.010 ml of PMN cell suspension (1:20 dil.) and determine cell density with a hemacytometer . At this point, the PMN are ready to be evaluated for their chemotactic and/or microbicidal activity as described below.

14.	Mononuclear cells prepared as described above for step 6 are adjusted to 107 cells per ml. Add 3 ml to 75 cm2 tissue culture flasks which have been precoated with 1% endotoxin -free gelatin and stored at 4°C until used.

15.	Incubate flasks for 1 h at 37°C to allow adherence of monocytes.

16.	Wash flask five times with 10 ml of warm HBSS to remove nonadherent cells.

17.	Remove adherent monocytes by vigorous pipeting with 10 ml of cold HBSS .

18.	Centrifuge cells at 100 x g for 10 min and resuspend in 1 ml of cold HBSS .

19.	To 0.190 ml of 0.05% gentian violet add 0.010 ml of monocyte cell suspension (1:20 dilution) and determine cell density with a hemacytometer . At this point the isolated monocytes are prepared to be studied for their chemotactic and/or microbicidal activity as described below.

MICROBICIDAL ASSAY

Principle

The microbicidal assay has long been considered the best functional assay for the evaluation of phagocyte function (12). It requires an active respiratory burst and functional phagocytic mechanism. The test is based on the addition of opsonized bacteria to purified monocytes or neutrophils at known bacteria-phagocyte ratios, followed by incubation at 37° C with sampling of reactions at a 30-min intervals up to 2 h. The samples are then plated on blood agar plates followed by overnight incubation at 37° C to allow viable bacteria to grow into visible colonies. The colonies are counted for each time point. The number of colonies for each time are then plotted on semi-log graph paper. Alternatively, the percentage of bacteria killed for each time point is determined from reaction mixtures containing only opsonized bacteria. In either case, control PMN -bacteria reactions should yield greater than 90% bacterial killing at 2 h of incubation.

Specimen

Heparinized whole blood is obtained from the patient and a normal adult control. The volume of whole blood needed is as described above for the isolation of monocytes and neutrophils. The specimens should be maintained at room temperature and processed for evaluation within 6 h of collection.

Reagents and Supplies

Mueller-Hinton broth

Staphylococcus aureus ( ATCC 27217)

0.1% Gelatin in distilled water

HBSS

Control opsonin -pooled sera obtained from at least five normal adult controls, which

has been aliquoted and stored at -70° C. The complement activity should be determined prior to use by routine 50% hemolytic complement assay.

0.05% Gentian violet in 2% acetic acid

Blood agar plates

Procedure

1.	On the day before the scheduled assay, thaw one vial of stock S. aureus and inoculate 10 ml of Mueller-Hinton broth with 5 l of bacteria. Incubate overnight at 37° C.

2.	Aliquot 2.5 ml of 0.1% gelatin into sterile polystyrene tubes (12 by 75 mm). These tubes may be prepared in advance and stored at 4° C. A total of 15 tubes is needed for one assay.

3.	Isolate monocytes and/or neutrophils as described above.

4.	Remove S. aureus from incubator and centrifuge at 900 x g for 15 min.

5.	Resuspend the cell pellet in 10 ml of 0.1% gelatin and centrifuge at 900 x g for 15 min.

6.	Resuspend the cell pellet in 1 ml of 0.1% gelatin.

7.	Prepare 1/30, 1/60, 1/120, and 1/240 dilutions of S. aureus in 0.1% gelatin.

8.	Zero or blank a spectrophotometer set at 620 mm with 0.1% gelatin to determine the optical density of each dilution. The optical density of the 1/120 dilution should be close to 0.1. (Use cuvettes with 1 cm path length.) Determine the dilution that will give 0.1 optical density, which equals 108 bacteria per ml and will yield - 250 bacteria at final plating from baseline PMN -bacteria reactions and reactions containing only opsonized bacteria. The assay has been formulated to provide a phagocyte/bacteria ratio of 1/1. The diluted bacteria are maintained at 4° C until added to reaction tubes.

9.	Thaw an aliquot of control opsonin and maintain at 4°C until added to reaction rubes.

10.	Remove 30 blood agar plates from 4° C storage. Label plates AO , A30 , A60 , A90 , and Al20 in duplicate for reactions containing opsonized bacteria only. Label plates BO, B30 , B60 , B90 and B120 in duplicate for reactions containing control PMN and opsonized bacteria. Label plates CO, C30 , C60 , C90 , and C120 in duplicate for reactions containing patient's PMN and opsonized bacteria.

11.	Adjust the PMN cell density to 2.94 x 106/ml with HBSS .

12.	Label and prepare PMN -bacteria- opsonin reaction tubes (12 by 75 mm) as follows:

(A)	0.85 ml of HBSS plus 0.150 ml of opsonin

(B)	0.85 ml of control PMN plus 0.150 ml of opsonin

(C)	0.85 ml of patient PMN plus 0.150 ml of opsonin

13.	Add 0.025 ml of S. auereus (0.1 optical density at 620 nm ) to each reaction tube and vortex.

14.	Immediately remove 10 l from each reaction tube and add to the 12-by-75-mm tubes containing 2.5 ml of 0.1% gelatin. These tubes should be previously labeled as described above in step 10 for the blood agar plates.

15.	Immediately place reaction tubes A, B, and C in a 37° C incubator with rotation for 30 min.

16.	Vortex AO , BO, and CO gel tubes, transfer 25 l in duplicate to appropriately labeled blood agar plates, and immediately spread the inoculum across the entire plate to obtain discrete colonies after overnight incubation.

17.	Remove the reaction tubes from the rotator after the 30-min incubation and repeat steps 13 through 16 for each 30-min interval up to 2 h of total incubation time.

18.	Incubate all blood agar plates at 3°C overnight.

19.	Next day, count the number of colonies on each plate, obtain the mean of each duplicate, and either plot the absolute number of bacteria for each time point on semi-log paper or plot the percent bacteria killed for each time point for BO through B120 and CO through C120 on linear graph paper, using AO through A120 as 0% kill values.

Interpretation

The reaction tube (B) containing control PMN along with opsonin and bacteria should demonstrate greater than 90% killing at 2 h.

CHEMOTAXIS ASSAY

Principle

Directed migration of both monocytes and neutrophils in response to specific chemotactic stimuli is the first crucial step in the sequence of events leading to acute and chronic inflammatory responses (14). Cell surface receptors for various stimuli function in concert with adhesion molecules on the surface of the cell along with intracellular recycling of F-actin polymerization. These events result in the directed migration of phagocytose to local sites of inflammation. The assay described below measures the radial migration of monocytes or neutrophils under agarose in response to a chemotactic gradient created by the diffusion of a chemoattractant through the agarose. After allowing for directed migration toward the chemoattractant in one direction and random migration in the opposite direction of applied chemoattractant, the monocytes or neutrophils are fixed and stained with Giemsa stain. After drying, the distance of directed migration and random migration of fixed and stained phagocytes is determined by microprojection (40%) net directed migration is determined by the subtraction of random migration distance (in centimeters) from directed migration toward the chemoattractant. Patient values are then compared with control values obtained with control phagocytes evaluated in parallel with patient phagocytes.

Specimen

Heparinized whole blood is collected from the patient and a control subject as described above for the isolation of monocytes and neutrophils. The specimens should be maintained at room temperature and processed for evaluation within 6 h of collection.

Reagents and Supplies

Agarose, type HSA (Hyman Serum Albumin)

10 X Eagle minimum essential medium with Earle's salts and L-glutamine (MEM)

Fetal calf serum

Leveling table with level

Template and well-cutting tool. The template is constructed from Plexiglas to

accommodate a 60-mm-by-15-mm petri dish. Replicates of three wells (3 mm inside diameter) each are oriented in a straight line (5 mm, center to center, at 60° intervals).

Dimethyl sulfoxide (DMSO)

N-Formyl-Met-Leu-Phe (FMLP): 10-2 M solution in DMSO, aliquoted in 0.025 ml

amounts and stored at -70° C

37% Formaldehyde

Giemsa stain

Projecting microscope

Procedure

1.	Isolate monocytes and/or neutrophils from patient and control heparinized whole blood as described above and adjust to 25 x 106/ml. Store cells at room temperature until added to agarose plates.

2.	Formulate 1.2% agarose plates, prepared in 60-mm petri dishes, by dissolving 0.24 g of agarose in 10 ml of dH2O (2.4%); maintain at 56° C. Prepare 10 ml of 2 x MEM containing 20% fetal calf serum, equilibrate at 56° C, and add it to the 10 ml of 2.4% agarose in dH2O. Pour 6.0 ml of the resulting 1.2% agarose-1 x MEM into each of three 60-mm petri dishes which have been placed an a leveling table. Allow the agarose to solidify (15 min) and place at 4° C for 30 min before cutting wells with the template.

3.	Place the template on each petri dish and employing the cutting tool, carefully cut 3-mm wells while avoiding the etching of the plastic surface. Carefully remove the agarose plugs by using a Pasteur pipette connected to a vacuum source. Store the plates at 4° C until ready for use.

4.	Remove a 0.025 ml aliquot of 10-2 MFMLP from - 70° C storage, thaw, and prepare 10-7 M and 5 X 10-8 M solutions in PBS.

5.	Remove agarose plates from 4° C storage and draw a line across the bottom of each plate, dividing the plates in half with three replicates of three wells on each half plate. Label one side of the plate "patient" and the other side "control." Two plates will be needed, one labeled "10-7 MFMLP," the other labeled "5 x 10-8 MFMLP."

6.	Vortex patient and control phagocytes and pipette 0.010 ml into the center wells of the appropriate section of both plates.

7.	Pipette 0.010 ml of PBS into the inner wells of both plate.

8.	Pipette 0.010 ml of 10-7 FMLP or 5 x 10-8 MFMLP into the outerwells of the appropriate section of each plate.

9.	Cover the plates and incubate for 2 h at 37° C under 5% C02.

10.	Remove plates from the incubator and flood the plate with enough absolute methanol to cover the agarose. Allowed the plates to stand for 30 min at room temperature or overnight at 4° C.

11.	Using a fume hood, remove the methanol from each plate and add enough 37% formaldehyde to cover the agarose. Allow the plates to stand for 30 min at room temperature.

12.	Remove the formaldehyde and carefully dislodge the agarose with an applicator stick or spatula.

13.	Allow the plates to air dry, and stain the adherent phagocytes by the addition of 1.5 ml of Giemsa stain. Incubate for 15 min at room temperature, then rinse each plate with dH20 and allow to air dry.

14.	Using a projection microscope (40x), record the distance in centimeters of phagocyte migration from the edge of each well to the leading edge of phagocyte migration toward wells containing chemoattractant (directed migration) and toward wells containing PBS (random migration).

15.	Calculate the average net directed migration for each dilution of FMLP as follows:

(directed migration-random migration)

number of wells

= average net directed migration

Interpretation

The patient results of each assay are interpreted relative to the control results. Net directed migration of control phagocytes should range from 1.5 cm to 4.5 cm, with random migration ranging from 0.5 cm to 1.5 cm. Patient values are interpreted as either comparable to or deviating from control values.

NBT SLIDE ASSAY

Principle

An important microbicidal mechanisms of normal phagocytes is their ability to gencnerace toxic oxygen radicals upon phagocytosis,leading directly or indirectly to bacterial cell death. One of the oxygen radicals, superoxide (02-), is easily detected in stimulated phagocytes by the reduction of the compound NBT to an insoluble form (formazan) (1). Formazan is observed by light microscopy as blue granules in the cytoplasm of stimulated phagocytes. The amount of superoxide produced by stimulated phagocytes (2). Phagocytes deficient in key enzymes of the hexose monophosphate shunt are impaired in their ability to generate superoxide, which is reflected in decreased reduction of NBT.

Specimen

Whole blood (1 ml), collected from both patient and control subject, is placed immediately on ice to delay clotting. The specimens should be processed within 1 h of collection.

Reagents and Supplies

Normal human serum (type AB)

NBT (grade III) (Sigma Chemical Co., St. Louis, Mo.)

Phorbol 12- myristate , 13-acetate ( PMA ): 1 mg/ml in DMSO, stored at -70° C

(Sigma Chemical Co., St. Louis, Mo.)

Absolute methanol

Safranin 0 1 g/100 ml of dH2O in 40 ml of glycerin, stored at room temperature

Permount

Procedure

1.	Dissolve 10 mg of NBT in 5.4 ml of PBS and add 3 ml of normal human serum. Filter the solution through a 4-5 m -pore-size filter. Remove 1 ml of solution, to which PMA will be added just before use.

2.	In a humid chamber, place acid-cleaned coverslips (in duplicate) onto slides labeled as (a) control PMN resting, (b) control PMN stimulated, (c) patient PMN resting, and (d) patient PMN stimulated.

3.	Place 2 to 3 drops of patient or control whole blood onto appropriate coverslips and incubate at 37°C under 5% CO2 for 30 to 45 min.

4.	Wash the clot off each coverslip with a gentle stream of PBS and place back on appropriately labeled slide.

5.	Prepare a 1/10 dilution of PMA (stock 1 mg/ml in DMSO) in PBS. Add 0.010 ml of diluted PMA to 1 ml of NBT -serum solution prepared as described in step 1 above.

6.	Add 3 drops of the NBT -serum solution to patient and control coverslips labeled "resting." Add 3 drops of the NBT -serum- PMA solution to patient and control coverslips labeled "stimulated."

7.	Close the humid chamber and incubate at 37°C under 5% CO2 for 20 min.

8.	Rinse each coverslip with a gentle stream of PBS and air dry.

9.	Fix PMN by the addition of absolute methanol for 60 s, rinse with distilled water, and air dry.

10.	Counterstain the fixed PMN by the addition of safranin O solution for 5 min.

11.	Rinse each coverslip with distilled water and air dry.

12.	Using Permount , mount each coverslip on appropriately labeled slides and allow to dry.

13.	Evaluate each coverslip by light microscopy at 100 X. Positive cells will have dark blue deposits of formazan distributed throughout the cytoplasm, with the counterstained segmented nuclei appearing red. Negative cells will have clear to pale blue cytoplasm. Count between 100 and 200 cells on each coverslip and record the percentage of positive cells.

Interpretation

With PMA stimulation, control PMN should produce 99 to 100% positive cells while control resting PMN should be between 5 and 40% positive cells.

DCF ASSAY

Principle

Hydrogen peroxide is an important microbicidal by product of the oxidative burst of stimulated phagocytes. This compound is derived from the dismutation of superoxide produced by the hexose monophosphate shunt. Hydrogen peroxide can be measured by the oxidation of the nonfluorescent compound 2',7' - dichlorofluorescin ( DCFH ) to the flourescent compound 2',7' - dichlorofluorescin ( DCF ). The nonpolar and nonfluorescent form of DCFH is 2',7' - dichlorofluorescin diacetate ( DCFH -DA). DCFH -DA readily diffuses across the cytoplasmic membrane of phagocytes and is trapped inside the cell by cleavage of the acetyl groups by cytoplasmic enzymes to the nonfluorescent , ionically charged DCFH . Stimulation of the oxidative burst of DCFH -labeled phagocytes can be readily detected by flow cytometry (3). The advantage of this assay is the ability to evaluate phagocytes in whole blood samples (8), thus avoiding the stimulatory effects often encountered in purification procedures. Further, extremely small volumes of whole blood (0.2 ml) are required for evaluation, which makes the assay well suited for infants and pediatric patients.

Specimen

EDTA - anticoagulated whole blood (0.5 ml) is obtained from the patient and a normal control subject. Blood samples maintained at room temperature may be evaluated up to 24 h after being drawn, which allows for transport of specimens from distant sites.

Reagents and Supplies

RBC lysing buffer. -8.26 g of NH4Cl , 1.0 g of KHCO3 , 0.037 g of (tetrasodium)

EDTA in 1 liter of distilled water, stored at 37° C under 5% CO2

DCFH -DA (Molecular Probes, Inc., Eugene, Oreg .): 10 mM stock prepared in absolute

ethanol and stored at -20° C

Catalase from bovine liver (13,000 U/mg of protein)

(Sigma Chemical Co. St. Louis, Mo.)

PMA (Sigma Chemical Co. St. Louis,Mo .): 1 mg/ml in DMSO, stored at -20° C

Procedure

1.	Prepare catalase solution by dissolving 30 mg of catalase in 10 ml of PBS at room temperature, followed by filtration through a 0.45 m -pore-size filter.

2.	Prepare 10 M DCFH -DA from 10 nM stock (10 l , to ml in PBS).

3.	Label 12-by-75-mm tubes as (a) patient resting, (b) patient stimulated, (c) control resting, and (d) control stimulated, and add 100 l of whole blood to appropriate tubes.

4.	Add 2 ml of RBC lysing buffer to each tube, vortex, and allow to stand for approximately 2 min.

5.	Pellet white blood cells (WBC) by centrifugation at 1,000 x g for 1 min.

6.	Aspirate supernatant, vortex cell pellet, and add 2 ml of PBS.

7.	Pellet WBC by centrifugation at 1,000 x g for 1 min and resuspend WBC pellet in 0.1 ml of PBS.

8.	Add 0.1 ml of 10 M DCFH -A to each tube, vortex, and incubate for 15 min at 37°C.

9.	Add 0.7 ml of catalase solution to each tube and vortex.

10.	Prepare working PMA solution (1 g /ml) in PBS from stock PMA (1 mg/ml in DMSO). Add 0.1 ml to tubes labeled "patient stimulated" and "control stimulated" and add 0.1 ml of PBS to tubes labeled "patient resting" and "control resting." Incubate tubes for 15 min at 37° C.

11.	Evaluate samples for fluorescence by flow cytometry as follows:

( i )	Use a standard protocol for whole blood samples and one-parameter fluorescence histogram acquisition, accumulating 5,000 events on either gated neutrophils or monocytes or both. Gating on lymphocytes will also allow the monitoring of any possible extracellular cross-feeding of cells with hydrogen peroxide produced by stimulated phagocytes that were not degraded by catalase.

(ii)	Evaluate the control resting sample first and set the fluorescence cursor to less than 2% positive cells. This cursor setting should be used for the evaluation of all remaining samples. Also, the photomultiplier tube used for the collection of fluorescence signals should be adjusted so that signals obtained with control resting PMN appear as a Gaussian distribution of signals within the first decade (0.1-1.0) of a 3- or 4-log scale. This will usually place the fluorescence histogram of control-stimulated PMN within the third decade of the log scale, which is particularly important for the evaluation of heterozygous carriers of the X-linked form of CGD in which two distinct fluorescence distributions are observed, one appearing in the negative region and one in the positive region of the fluorescence histogram.

(iii)	Evaluate the control stimulated sample, which should be greater than 95% positive cells.

(iv)	Evaluate patient resting and stimulated samples are record the percentage of positive cells. The mean channel fluorescence of the fluorcsrence histogram should also be recorded for all stimulated samples.

Interpretation

PMA -stimulated PMN obtained from normal subjects yield at least 95% positive cells with high mean channel fluorescence values within the third decade of a four-decade scale, indicating high levels of hydrogen peroxide production.

IMMUNOPHENOTYPING :

Principle

Monocytes/ macrophages and neutrophils play important roles in nonspecific primary host defense mechanisms and in the regulation of specific cellular and humoral immune responses. These effector mechanisms are mediated through important ligand receptors on the cytoplasmic membrane of phagocytes. Receptors for the Fc region of human immunoglobulin G ( CD16 , FcR III; CD 32, FcR II; CD64 , FcR I), as well as receptor), play crucial roles in the binding of immune- complexed antigens to phagocyte cell surfaces, thus facilitating phagocytosis and clearance of invading pathogens. Furthermore, the expression of cell surface adhesion molecules, such as CD18, is essential for the directed migration of phagocytes to sites of inflammation. These same receptors found on monocytes/macrophages serve to expedite the internalization of antigens that contribute to antigen processing with subsequent presentation to immunoregulatory T cells. In addition to out knowledge of patients with deficiencies of cell surface receptors (CD11b, CD16, CD18) which predispose them to infections (6,10,13), we have learned much about modulation of these receptors in response to various cytokines(5). This knowledge has led to the development of cytokine therapies in patients with secondary or acquired immune deficiency that are designed to upregulate receptors on the surface of phagocytes, leading to enhanced phagocytic effector mechanisms (4,7,11). For example, gamma interferon therapy leads to the upregulation of FcR I receptors (CD64) on monocytes and is reflected in enhanced microbicidal activity as well as in enhanced lymphoproliferative responses to recall antigens.

These new therapies have dictated the development of clinical laboratory assays that are both practical and clinically useful in monitoring the response to cytokine therapy. Flow cytometric, multiparametric analysis of subpopulations of cells derived from whole blood specimens allows for the accurate quantitation of receptor expression on the surface of monocytes and neutrophils. This methodology not only detects the presence or absence of receptors, but also evaluates the regulation of expression of these receptors in response to various stimuli. The technique employs multiple monoclonal antibodies labeled with different fluorochromes that identify receptor expression of specific subpopulations of phagocytes derived from whole blood, this circumventing the need for time consuming cell purification.

Specimen

EDTA anticoagulated whole blood is obtained from the patient and a normal control subject. The volume of whole blood needed is dependent upon the number of different cell surface markers evaluated, with 0.1 ml of whole blood needed for each marker. Usually 0.5 to 1.0 ml of whole blood is adequate for most studies. The specimens are maintained at room temperature until processed.

Reagents and Supplies

Standard Brite fluorescence calibration spheres (Coulter Immunology, Hialcah, Fla.)

RBC lysing buffer. 8.26 g of NH4Cl + 1.0 g of KHCO3 + 0.037 g of tetrasodium EDTA

in 1 liter of distilled water, stored at 370C under 5% CO2

Fluorochrome-conjugated monoclonal antibodies: phycoerythrin (PE)- or fluorescence

isotiocyanate (FITC) conjugated monoclonal antibodies to CD14, CD11b, CD18, CD64, CD32, and CD16: appropriate fluorochrome conjugated isotype control monoclonal antibodies.

Paraformaldehyde: 1% solution prepared in PBS with pH adjusted to 7.3 (commercial

RBC lysing buffers which also contain fixative can also be used)

Flow cytometer

Procedure

1.	To appropriately labeled tubes (12 by 75 mm), add appropriate combinations of fluorochrome conjugated isotype control or antigen specific antibodies for both patient and control samples. If commercially prepared combinations are not used, titration of antibodies is necessary before use.

2.	Add 0.1 ml of whole blood from patient or control, vortex, and incubate for 30 min at 40C.

3.	Lyse RBC by the addition of 2 ml of lysing buffer.

4.	Pellet WBC and wash with 2 ml of PBS.

5.	Vortex the wet pellet and add 0.5 ml of 1% paraformaldehye. Store at 40C in the dark until evaluated for fluorescence by flow cytometry. Although fixed samples may be stored for up to 2 weeks without altering the light scattering characteristics of the cells, the samples should be evaluated as soon as possible after fixation. Alternatively unfixed samples may be evaluated immediately after staining.

6.	Before evaluating samples by flow cytometry, the instrument must be calibrated using a fluorescence standard for the measurement of fluorescence intensity. Using a standard two color protocol, evaluate Standard Brite spheres for fluorescence intensity. Adjust the laser power so that the mean channel fluorescence value observed for both red and green fluorescence is essential identical to values recorded from the previous instrument calibration. Minor adjustments in voltage setting of each photomultiplier tube used for the detection of either red or green fluorescence may also be required.

7.	Evaluate control samples first, observing the light scatter characteristics of the sample stained with PE-CD14 or FITC-CD11b.

8.	Construct a gate around monocytes and neutrophils and assign each gate to an individual two-parameter fluorescence histogram.

9.	Adjust the monocyte gate and red-green fluorescence cursors until greater than 95% of red-green signals appear in the red-green positive quadrant (quadrant 2). Adjust the neutrophil gate until greater than 95% of the red-green signals appear in the red negative-green positive quadrant (quadrant 4).

10.	Evaluate the e sample stained with the isotype control for the PE-CD14/FTTC-C11b sample. Greater than 95% of the red-green signals should appear within the red negative-green negative quadrant (quadrant 3) for both monocytes and neutrophils.

11.	Reevaluate the sample stained with PE-CD14/FITC-CD11b and accumulate 2,000 events with the monocyte gate. Record both percent positive cells and mean channel fluorescence values for both red and green fluorescence distributions for both monocytes and neutrophils.

12.	Evaluate remaining samples, recording both percent positive cells and mean channel fluorescence values for both red and green fluorescence from appropriate quadrants.

13.	Repeat steps 7 through 12 for patient samples.

Interpretation

Patients with intense deficiencies of CD11b and CD18 will show no expression of these markers on CD14-positive monocytes or neutrophils, while control monocytes and neutrophils will be greater than 95% positive for both markers. Modulation of CD11b, CD18, CD64, CD32, or CD16 must be carefully interpreted on the basis of establishment of a normal mean channel fluorescence range for each marker on either monocytes or neutrophils obtained from control subjects. Because these markers can be modulated by acute or chronic inflammation, interpretation of results in the setting of cytokine therapy must also take the clinical status of the patient into consideration . Mean channel values obtained for patient monocytes or neutrophils are compared with control ranges or with values obtained with patient samples collected prior to cytokine therapy (baseline value). Figure 2 illustratesthe flow cytometric evaluation of whole blood-derived monocytes for the expression of CD11b and CD18. In each instance, gating on monocytes (gate C) yielded greater than 95% CD14-positive cells which coexpressed both CD11b and CD18. These data illustrate the ease in which evaluations of monocytes from whole blood samples can be achieved.

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