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stromal (Fig. 4) or subepithelial space distribution (Fig. 5). The
inflammatory cells in this dense area were counted to a total of
500 cells or until all inflammatory cells (EO, PMN, lymphocyte,
plasma cells) in the polyp sample were counted. Data were nor-
malized as a ratio to the total cell count. To determine the
cellularity of the polyp sample, five consecutive high power
fields (HPFs) (1,000
3
) were used to count the total number of
EO, PMN, lymphocyte, and plasma cells in one HPF. The aver-
age number of the total cell count from the five HPFs of each
patient was defined as the cellularity of the nasal polyp.
Ten HPFs (1,000
3
) of the epithelium were analyzed to
characterize and count the goblet cells present. The surface epi-
thelial morphology was categorized as either pseudostratified
ciliated columnar or transitional epithelium. The area of high-
est concentration of mast cells was identified. Ten consecutive
HPFs (400
3
) were used to count for mast cells. The average
number of mast cells for each patient was obtained.
Data analysis was performed using analysis of variance
(ANOVA) on SPSS statistical software version 17.0 (IBM SPSS,
Armonk, NY) with appropriate Tukey post hoc analyses.
P
val-
ues
<
.05 were considered to be statistically significant.
Flow Cytometry
The same polyp specimen used in the histologic examina-
tion was used for flow cytometry. Fresh tissue specimens were
placed in Royal Park Memorial Institute (RPMI) 1640 1
3
me-
dium (Cellgro, Manassas, VA) and processed within 1 hour of
extraction. Under a category 2 sterile hood, tissue samples were
disaggregated to allow separation of cells from the tissue. Cell
suspensions were prepared from the resulting eluent, using a
70
l
m BD Falcon cell strainer (BD Biosciences, Franklin Lakes,
NJ).
Pelleted cells (400
3
g
, 4 C, 5 minutes) were stimulated in
nonpolarizing stimulation media to facilitate production of in-
tracellular cytokines. Results were achieved for leukocytes by
reconstituting cells in 1 mL RPMI 1640 supplemented with 10%
fetal bovine serum (Lonza Group, Ltd., Basle, Switzerland), 1%
penicillin–streptomycin (Invitrogen, Carlsbad, CA), 1 ng/mL
phorbol 12-myristate 13-acetate (PMA) (Sigma-Aldrich, St.
Louis, MO), 0.6
l
L BD Golgi stop protein transport inhibitor
(BD Biosciences), and 500 ng/mL ionomycin (Sigma-Aldrich)
and cultured for 5 hours at 37 C according to established BD
Biosciences protocols.
A total of 100
l
L aliquots of cells were distributed to poly-
styrene fluorescent activated cell sorter (FACS) tubes at room
temperature, in the dark, for 30 minutes. Temperature single-
cell suspensions were stained for CD3, CD4, CD8, CD19, CD45,
and CD56 using antihuman fluorochrome conjugated antibodies
and incubated at 4 C for 30 minutes according to established
BD Biosciences protocols. Following the extracellular stain, 250
l
L of BD Cytofix/Cytoperm fixation and permeabilization solu-
tion (BD Biosciences) was added to each sample for 20 minutes
at 4 C in the dark.
Intracellular cytokines were stained for IL4, IL5, IL13,
IL17, and interferon (IFN)-
c
using specific antibodies or appro-
priate isotype controls for 30 minutes at 4 C in 1
3
BD perm/
wash buffer (BD Biosciences). Cells were washed twice with
perm/wash buffer according to the manufacturer’s instructions
(BD Biosciences). Cells were fixed with a preparation of 2%
paraformaldehyde (PFA v/v in phosphate-buffered saline) and
stored at 4 C covered in the dark.
Samples were run through a Cytek 8DXP upgraded (Cytek
Development, Fremont, CA) FACSCalibur (BD Biosciences).
Flow cytometer and fluorescence data were acquired using
FlowJo software version 4.6 (TreeStar, Ashland, OR). Gates
were created based on isotypes and fluorescence-minus-1 con-
trols. For the intracellular cytokines, positive signal for CD45
was used to gate on polyp leukocytes. T helper cells were sub-
gated from the leukocyte population using CD4 antibody. The
resulting CD4
1
leukocytes were then analyzed for IL4, IL5,
IL13, IL17, and IFN-
c
–producing cells. Intracellular cytokines
were also analyzed for CD45
1
CD4
2
s cells.
Baseline significance for this study was set at
a
5
.05. All
groups were compared using ANOVA on SPSS statistical soft-
ware (IBM SPSS) with appropriate Tukey post hoc analyses. In
some analyses, Student t tests with Bonferroni-Holme correc-
tions were conducted.
RESULTS
Phenotype
Eighty-four patients were included in the study,
with ages ranging from 7 to 83 years of age (median age,
46 years) (Table II). CF was the youngest subclass and
statistically lower than each asthmatic sinusitis group
(
P
<
.01). There were 48 females in the study, with signifi-
cant higher females in the asthmatic sinusitis (AScA
Fig. 5. Microscopic hematoxylin and eosin slide (2
3
) of a nasal
polyp demonstrating superficial subepithelial distribution of cells
within the rectangle.
TABLE II.
The Demographic Information for the CRS Subclasses.
CRS Subclass (n)
Mean Age (yr)
Female (n)
AERD (9)
46
5
AFS (11)
34
4
CF (7)
16
5
AScA (13)
48
11
ASsA (5)
35
4
NAScA (14)
54
7
NASsA (12)
59
3
Control (13)
50
9
Total (84)
46
48
AERD
5
aspirin exacerbated respiratory disease also known as aspirin
triad; AFS
5
allergic fungal sinusitis; AScA
5
asthmatic sinusitis with allergy;
ASsA
5
asthmatic sinusitis without allergy; CF
5
cystic fibrosis; CRS
5
chronic
rhinosinusitis;
NAScA
5
nonasthmatic sinusitis with allergy;
NAS-
sA
5
nonasthmatic sinusitis without allergy.
Han: Subclassification of Chronic Sinusitis
Laryngoscope
123: March
2013
50