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*
Clinical Research Division, Fred Hutchinson Cancer Research Center, Seattle, WA 98109; and Departments of
Medicine and
Obstetrics and Gynecology, University of Washington School of Medicine, Seattle, WA 98145
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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,
IL-4, and IL-5 revealed a Th1-type pattern in cervical T cells, with
significantly higher percentages of IL-2- and IFN--producing T cells
in the mucosa than in blood (mean 67% vs 29%). Coexpression of
surface CCR5 with intracellular IL-2 and IFN- was observed only
among T cells in the mucosa, but not among those in circulation. Thus,
we postulate that T cell homing to the genital mucosa leads to
differentiation into the combined CCR5+ Th1 phenotype.
Moreover, the predominance of CCR5+ Th1-type T cells in
normal cervical mucosa provides targets accessible for the efficient
transmission of macrophage-tropic HIV-1 variants in women following
sexual exposure. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Both in mice and humans, Th cells have been subdivided into Th1
cells secreting IL-2 and IFN-; Th2 cells secreting IL-4, IL-5, and
IL-10, and Th0 cells secreting various cytokine combinations
of the two subtypes (12, 13, 14). Like
CCR5+ T cells, Th1 cells are believed to
preferentially home to inflammatory sites, with the exception of
parasite-induced and allergic infiltrates where Th2 cells predominate
(15, 16). The assumption that CCR5+
and Th1 T cells share homing properties and represent an identical cell
type is supported by the recent finding that Th1 clones or T cells
polarized in vivo toward the Th1 phenotype characteristically express
high levels of CCR5 (17, 18). Concomitant expression of
CCR5 and secretion of Th1 cytokines by T cells may occur during
circulation or after homing to tissue. However, the sequence of events
leading to the differentiation of the CCR5+ Th1 T
cell phenotype remains unclear. Thus, one of the goals of this study
was to understand this process by comparing the phenotypes of blood and
genital mucosal T cells.
Regulation of the CCR5+ Th1 phenotype in the genital tract has particular relevance to acquisition of HIV-1 infection by sexual contact. CCR5 functions as the major coreceptor for HIV-1 strains transmitted sexually (19, 20, 21, 22). Moreover, HIV-1 preferentially infects CD4+ T cells with the Th1 phenotype (23, 24). Thus, infiltration into the genital mucosa of CD4+ Th1 cells which bear the HIV-1 coreceptor as a result of local environmental stimuli as well as sexually transmitted infections provides susceptible target cells for HIV-1 infection. To better understand host factors associated with HIV-1 transmission, here we present an extensive phenotypic analysis of T cells isolated from normal human cervicovaginal mucosa. By correlating CCR5 expression and cytokine production on a single cell level and comparing those findings with T cells from peripheral blood, we provide evidence that the link between CCR5 expression and Th1 phenotype is only established when T cells home to the genital mucosa and does not exist in peripheral blood.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Blood and tissue blocks containing portions of ectocervix or vagina were obtained from women undergoing hysterectomy or vaginal repair operation at the University of Washington Medical Center and affiliated hospitals. Subjects were either at low risk for HIV-1 infection or tested HIV-1-negative and had no cervicovaginal inflammation when examined preoperatively. The University of Washington Human Subjects Committee approved the study, and volunteers provided written consent before the procedure.
Immediately after sampling, the tissue was placed in a sterile specimen
container holding 50 ml of culture medium (RPMI 1640 supplemented with
100 U/ml penicillin, 100 µg/ml streptomycin, 2.5 µg/ml amphotericin
B, 2 mM L-glutamine (BioWhittaker, Walkersville, MD), and
10% FBS (Gemini, Calibasas, CA)) and transported on ice to the
laboratory. Processing began within 2 h of collection. After
washing the sample five times in cold PBS, a maximum of 2 x 2 cm
(cervix) or 4 x 4 cm (vagina) of mucosa per donor were dissected
from the underlying tissue. The mucosa was then placed in culture
medium containing 1µM DTT (Sigma, St. Louis, MO) for 10 min at 37°C
to dissolve mucus and cleave off loose epithelial cells. The mucosa was
placed in sterile water for 30 s to lyse contaminating blood cells
remaining on the surface of the tissue, extensively washed in cold PBS,
and cut into pieces <1 mm with a sterile razor. Tissue pieces were
transferred to T75 flasks (Costar, Cambridge, MA) and washed five times
to remove loose epithelial cells and remaining contaminating blood
cells by adding cold PBS. After the last wash, medium was added very
gently, the flasks were transferred to an incubator, carefully avoiding
agitation to prevent liberation of epithelial cells, and the tissue was
incubated at 37°C and 5% CO2. After 36 h,
Collagenase D (2 mg/ml, Boehringer Mannheim, Indianapolis, IN) was
added for 10 min at 37°C to reduce trapping of emigrated cells within
collagenous debris. The flasks were slowly tilted, the tissue pieces
were allowed to settle for 
10 min, and the supernatant containing
the emigrated cells, both intraepithelial and lamina propria, was
carefully harvested. The cell suspension was then underlayered with
Ficoll-Hypaque (Pharmacia, Uppsala, Sweden) and centrifuged for 20 min
at 895 x g. Cells collected at the interface were
washed three times and counted by trypan blue (Sigma) exclusion under
light microscopy. Lymphocyte yields were 0.3–3 x
106 for cervical epithelium and 1–10 x
106 for vaginal epithelium with >95% viability
and <5% contamination with epithelial cells. PBMC from the same
donors were isolated by Ficoll-Hypaque density centrifugation, and
washed, and counted as above.
Flow cytometry analysis of phenotypic markers
To define phenotypic properties of the mucosal lymphocytes,
cells were reacted in V-bottom 96-well plates (Costar) with
combinations of the following mAbs: anti-CD3 PE-Cy5, anti-CD4
FITC, anti-CD8 PE-Cy5, and anti-CD19 PE (all from Sigma);
anti-CD4 PE, anti-CCR5 FITC, anti-CXCR3 PE, and
anti-CCR5 PE (all from PharMingen, San Diego, CA); anti-CD49d
and anti-CD62L PE (both from Coulter, Hialeah, FL); anti-CD25
PE (Biosource, Camarillo, CA); anti-CD28 PE, anti-CD38 PE,
anti-CD44 FITC, anti-CD69 PE, anti-TCR
ß FITC, and
anti-TCR
PE (all from Becton Dickinson, San Jose, CA);
anti-CD45RA PE, anti-CD45RO FITC, anti-CD45RO PE, and
anti-CD103 FITC (all from Dako, Glostrup, Denmark); and
anti-CCR3 (clone 7B11, obtained through the AIDS Research and
Reference Reagent Program from LeukoSite, Cambridge, MA). For
conjugated Abs, a minimum of 4 x 104 cells
were incubated for 20 min at 4°C in 100 µl of PBS supplemented with
1% BSA (Sigma) and 0.1% sodium azide (Sigma) (FACS buffer) containing
10% normal mouse serum (Sigma) and a combination of three Abs
conjugated to three different fluorochromes (each 5–10 µg/ml) or
matching isotype controls (Becton Dickinson or Coulter). Cells were
washed twice with FACS buffer to remove unbound Abs and fixed in 0.5%
para-formaldehyde (Baker, Phillipsburg, NJ). For the two unconjugated
Abs, cells were first reacted for 20 min at 4°C in FACS buffer
containing 10 µg/ml of either anti-CD49d, anti-CCR3, or
matching isotype control and washed twice. Cells were then resuspended
in 100 µl of FACS buffer containing 10% normal goat serum and 5
µg/ml PE-labeled goat anti-mouse IgG (Fc)
F(ab')2 (Coulter), incubated for another 20 min
at 4°C, and washed twice. Finally, cells were resuspended in FACS
buffer containing 10% normal mouse serum and a combination of two Abs
or isotype controls (5–10 µg/ml) conjugated to FITC and PE-Cy5,
washed twice, and fixed for analysis. Cells were analyzed on a Calibur
flow cytometer (Becton Dickinson) for three-color fluorescence. Scatter
gates were chosen to acquire only small cells with low granularity,
which were >95% lymphocytes, and to exclude larger and more granular
cells, which were predominantly nonlymphoid mononuclear cells and
epithelial cells. Isotype controls were used to define background
staining and quadrant markers. Cells stained with only one specific Ab
were used to compensate for overlapping signals between different
fluorescence channels.
Quantification of surface molecule expression by flow cytometry
Cells were incubated for 40 min at 4°C in FACS buffer containing 10% normal mouse serum and a saturating amount of anti-CCR5 PE (PharMingen) or matching isotype control in combination with anti-CD3 PE-Cy5, washed twice to remove unbound Abs, and fixed in 0.5% para-formaldehyde. Receptor density per cell, designated as molecules of equivalent PE (MEPE),3 was calculated by comparing staining of samples to a standard curve. Individual standard curves were established for each experiment by analysis of a mixture of six calibrated bead populations containing six different MEPE levels/bead (RCP-30-5, Spherotech, Libertyville, IL), plotting the fluorescence intensity of each bead population against its designated MEPE value, and interpolating between points by linear regression. Differences between MEPE values of circulating and mucosal T cells were tested for significance using the student t test for paired samples.
Intracellular cytokine staining
Mucosal cells and PBMC were stimulated immediately after
isolation with 50 ng/ml PMA and 1 µM ionomycin (both from Sigma) in
the presence of 2 µM monensin (GolgiStop; PharMingen) in culture
medium for 4 h at 37°C and 5% CO2. After
stimulation, cells were stained for surface markers using anti-CD3
PE-Cy5, anti-CD4 PE-Cy5 (PharMingen), anti-CCR5 PE, and isotype
controls as described above, then fixed and permeabilized
(Cytofix/Cytoperm Kit, PharMingen). Cells were then reacted in 100 µl
of FACS buffer for 30 min at 4°C with various combinations of
anti-IL-2 FITC (2 µg/ml), anti-IFN- FITC (2 µg/ml),
anti-IFN- PE (1.5 µg/ml), anti-IL-4 PE (1.3 µg/ml),
anti-IL-5 PE (1.5 µg/ml), and matching isotype controls
(PharMingen). Fluorescence was analyzed on a Calibur flow cytometer as
described above. Cells treated with monensin, but not PMA and
ionomycin, were always incubated in parallel as controls and never
stained positive for IL-2 and only occasionally for IL-4 within the
CD3- but not CD3+
compartment. As previously reported (25), frequencies of
IL-4-producing T cells were generally lower than IL-2-producing T
cells. However, the utility of the anti-IL-4 Ab used in this
analysis was demonstrated by positive staining in the majority of
bronchoalveolar mast cells isolated from an allergic control
individual.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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+ß+, CD45RO+
T cells
Following isolation and staining for flow cytometry, two distinct
cell populations from the cervicovaginal mucosa were observed in the
forward and side scatter plots (data not shown). The larger cells with
higher granularity mainly consisted of nonlymphoid mononuclear cells
and contaminating epithelial cells (data not shown), whereas the
smaller cells with lower granularity represented >90%
CD3+ T lymphocytes (Fig. 1
A). Nearly all T cells
expressed TCRß (data not shown) rather than the TCR (Fig. 1A; Table I).
CD8+ T cells were generally more common than
CD4+ T cells, with a mean CD4/CD8 ratio of 0.83
in 15 patients (Fig. 1B; Table I). A small fraction of T
cells coexpressed both the CD4 and the CD8 receptor (Fig. 1B). The remaining small cells with lower granularity were
CD16+ NK cells, ranging from 0 to 19.8% (median
0.6%, n = 12), and CD19+ B
lymphocytes, ranging from 0.1 to 2.5% (median 0.6%, n
= 12) (Fig. 1C; Table I). Cells isolated from the vaginal
and the ectocervical mucosa were not phenotypically different (data not
shown).
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D; Table I).
CD4+ T cells were more clearly dominated by the
CD45RO+ memory subtype than
CD8+ T cells (Fig. 1, E and
F; Table I). Most cervicovaginal T cells express early activation markers (CD69+) and mucosal homing receptors
Although the majority of mucosal T cells stained positive for CD69
(mean 94%) and HLA-DR (mean 68%), only a fraction expressed the IL-2
receptor (CD25) and CD38, indicating early but to a lesser extent, late
T cell activation (Fig. 2,
A–D; Table I). Of note, only 12% of
CD4+ T cells from peripheral blood expressed
HLA-DR (data not shown). The costimulatory molecule CD28 was expressed
on almost all mucosal CD4+ cells, but only on
about two-thirds of CD8+ T cells (Table I).
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4 subunit which
together with the ß1 subunit forms VLA-4 (Fig. 2, E and F; Table I). About two-thirds of
CD8+ and one-third of CD4+
mucosal T cells also stained positive for CD103, the
E subunit of the
Eß7 integrin (Fig. 2G; Table I). In addition, CD44 expression was abundant
(Fig. 2H; Table I). These integrins are typically expressed
on memory T cells homing to sites of Ag-exposure (VLA-4, CD44) and more
specifically to mucosal tissues (CD44, CD103) (26, 27). In
contrast, L-selectin (CD62L), preferentially expressed by naive T cells
homing to peripheral lymphoid organs (28), was found on
only very few of the cervicovaginal T lymphocytes (Fig. 2I;
Table I). The mucosal T cells were clearly distinguishable from
peripheral blood T cells, which expressed high CD62L ligand and low or
absent CD103 (data not shown). The frequency of CCR5-expressing T cells but not the receptor density on individual cells is markedly increased in the genital mucosa as compared with peripheral blood
The percentages of mucosal T cells expressing CCR5 ranged from 78
to 97% (mean 87%) in the 16 tissue donors tested, and no difference
was apparent between CD4+ and
CD8+ cells in 4 donors tested (Fig. 3A; Table I). Both
HLA-DR+ and HLA-DR-
CD4+ mucosal T cells expressed CCR5 (Fig. 3A), and nearly all (mean, 87%) CCR5+
CD4+ mucosal T cells were
CD45RO+ memory cells (Fig. 3B). In
contrast, expression of CCR3 on cervicovaginal T cells was not
observed (Table I). The proportion of T cells expressing CCR5 was
markedly lower in peripheral blood (range 4.2–55%, mean 24%) than in
the mucosal compartment (range 81–97%, mean 88%) in 13 individuals
(p < 0.001) (Fig. 3C). We also
examined in two donors the expression of the chemokine receptor CXCR3,
a T cell marker associated with inflammatory reactions. In these
analyses, 20–25% of the mucosal CD4+ T cells
expressed CXCR3, and of these, 95–96% coexpressed CCR5 (data not
shown).
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Our findings indicate a preponderance of CCR5+ T
cells in the lower female genital tract, but it was unclear whether
these cells also have a higher density of CCR5 on their surface. To
address this, we estimated the CCR5 receptor density on mucosal and
peripheral blood T cells from seven donors by comparing their
fluorescence intensities with calibrated beads containing known
quantities of PE molecules (MEPE) (Fig. 3, D and
E). As shown in one representative donor, anti-CCR5 PE
binding curves and peaks for the six bead populations (with six
different levels of PE fluorescence) were overlaid, and the arrows
depict the level of anti-CCR5 PE binding for which MEPE units per T
cell were calculated (Fig. 3D). Anti-CCR5 binding curves
usually had a broader shape in blood than in mucosal T cells and the
MEPE units were calculated within the last distinct peak of the curve.
This strategy quantifies CCR5 expression on T cells with high receptor
density rather than calculating an average for all CCR5 positive T
cells. In comparison to circulating T cells, CCR5 surface receptor
density was not significantly up-regulated on mucosal T cells
(p = 0.07) (Fig. 3E). Mean MEPE
units were 4191 (range 3009–6468) for circulating T cells and 5837
(range 3511–9365) for cervical T cells in six of seven tissue donors
tested. However, one individual demonstrated a pronounced up-regulation
from 2291 MEPE units on circulating T cells to 12050 MEPE units on
cervical T cells.
The potential to produce IL-2 and IFN- is significantly
increased in cervical T cells as compared with peripheral blood
To determine whether T cells infiltrating the human genital mucosa
preferentially exhibit the Th1 or Th2 phenotype, we analyzed their
ability to synthesize IL-2 and IFN- or IL-5 and IL-4, respectively,
upon stimulation with ionomycin and PMA (see Materials and
Methods). One representative comparison is shown in Fig. 4. On the forward and side scatter plots,
a small percentage of mucosal T cells (Fig. 4B, right arrow)
were larger, more granular, and phenotypically activated than the
predominant population which we further analyzed here. Among six donors
tested, the mean percentage of CD4+ T cells
producing IL-2 was clearly higher in the cervical mucosa (mean 61%,
range 43–83%) than in peripheral blood (mean 35%, range 17–58%)
(Fig. 4, C and D). Of the larger, more granular
mucosal T cells, practically all produced IL-2 (data not shown).
Similarly, of the CD4+ T cells derived from the
genital mucosa in three of these donors examined, an average of 61%
(range 55–70%) produced IFN-, in contrast to 12% (range 6–20%)
from peripheral blood (Fig. 4E). In addition, a mean of 43%
(range 32–49%) of CD4+ T cells in the genital
mucosa simultaneously produced both, and a mean of 87% (range
77–93%) at least one of the Th1 cytokines (Fig. 4E, and
data not shown). By contrast, IL-4-producing CD4+
T cells from the six donors were infrequent in both mucosal (mean
7.6%, range 2.6–18%) and blood-derived CD4+ T
cells (mean 4%, range 1.1–6.6%) (Fig. 4F, and data not
shown). Of note, the majority of IL-4-producing mucosal
CD4+ T cells were coproducing IL-2 (mean 53%,
range 38–64%) and thus corresponded to the Th0 rather than the Th2
phenotype (data not shown). Production of IL-5 never exceeded 2% in
both the mucosal and blood derived CD4+ T cells
of three donors tested.
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Approximately two-thirds of T cells isolated from the genital
mucosa either produce IL-2 or IFN- or express CCR5, suggesting
relatively tight correlation of both traits. However, it is not clear
whether differentiation to the combined IL-2+,
CCR5+ or IFN-+,
CCR5+ phenotype occurs during circulation or when
homing to the genital tract. To address this, we first investigated
coexpression of CCR5 and IL-2 (six donors) or IFN- (three donors) in
peripheral blood T cells (Fig. 4C). Interestingly, although
appreciable numbers of circulating CD4+ T cells
produced IL-2 (mean 35%, range 17–58%) coexpression of intracellular
IL-2 and CCR5 was infrequently observed (Fig. 4C). On
average, only 3% (range 1–6%) of IL-2-producing
CD4+ T cells simultaneously expressed CCR5.
Similarly, in three donors, only 17% (range 3.5–29%) of the
IFN--producing peripheral blood cells coexpressed CCR5. By contrast,
examining the mucosal T cells from the same donors, the majority of
IL-2- and IFN--producing mucosal CD4+ T cells
also expressed CCR5 (mean 88% (range 80–94%) and 82% (range
79–85%), respectively) (Fig. 4D). Approximately 100% of
the larger and more granular lymphocytes coexpressed intracellular IL-2
and CCR5. Thus, production of Th1-type cytokines and CCR5 expression
are usually mutually exclusive in circulating T cells but tightly
coupled in mucosal T cells. This finding indicates that differentiation
to the complete CCR5+ Th1 phenotype does not
occur while in circulation but during or after homing of T cells to the
genital mucosa.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Eß7 and
4ß1 that are commonly
involved in T cell recruitment to other extralymphoid mucosal sites. In
addition, chemokines and their receptors as well as other intercellular
adhesion molecules may be involved in T cell trafficking to the genital
mucosa. Of note, our findings indicate that T cells expressing the chemokine receptor CCR5 are abundant in the human female genital mucosa, which stands in marked contrast to the low numbers of CCR5-expressing T cells in the circulation. One possible explanation for these findings is that CCR5 is up-regulated during the 36-h interval used to acquire emigrated T cells from the cervicovaginal tissue in vitro. We believe this is unlikely for several reasons. The time period employed was short, our medium lacks stimulatory agents such as IL-2, PHA, or anti-CD3, the application of similar conditions to isolate peripheral blood T cells actually resulted in down-regulation of CCR5, and only prolonged activation of T cells over many days led to CCR5 up-regulation in vitro in a recent report (10). Thus, our data suggest that memory T cells expressing CCR5 are either selectively recruited to the genital mucosa or they up-regulate CCR5 during extravasation or once localized within the mucosa. That the peak density of CCR5 receptors on individual cells was not significantly different between circulating and mucosal T cells in six of seven individuals argues for a mechanism that selectively recruits CCR5+ T cells, and favorably those with higher receptor densities, to the genital mucosa. In addition, interactions between locally secreted chemokines and the CCR5 receptor may retain T cells in the mucosa during antigenic challenge.
In support of recent reports that chemokine receptors are
differentially expressed in Th1 and Th2 cells, our findings in the
human genital tract link CCR5 expression with IL-2 and IFN- rather
than IL-4 and IL-5 production. The frequency of IL-2- and
IFN--producing cells paralleled CCR5 expansion and was significantly
higher in cervical than in circulating T cells. CCR5 expression in
genital tract T cells was therefore coupled with a Th1 (and
infrequently Th0) rather than the Th2 phenotype. This was further
substantiated by the demonstration that individual mucosal
CCR5+ T cells consistently displayed concomitant
production of IL-2 and IFN-, and that expression of CCR3, associated
with the Th2 phenotype (16, 18), was absent. By contrast,
CCR5-expressing and Th1 cytokine-producing T cells constitute two
separate populations in peripheral blood of the same individuals. This
result suggests that differentiation to the Th1 phenotype is coupled
with CCR5 up-regulation during or after homing of T cells to the
genital mucosa.
If CCR5 is involved in leukocyte recruitment to inflammatory sites, as
has been suggested previously (10, 11, 17, 18), then the
CCR5+ IL-2- rather than
CCR5- IL-2+ T cells are
more likely to be recruited to the genital mucosa. Local events such as
secretion of cytokines and chemokines may subsequently drive
differentiation into the combined CCR5+ Th1
phenotype. CCR5 may play a role in this process, which is supported by
findings in a murine model system that macrophage inflammatory
protein-1 (MIP-1), a natural ligand of CCR5, drives
differentiation from Th0 to Th1 cells (30). Once the link
between CCR5 expression and Th1 phenotype is established, it may be
maintained by autocrine loops whereupon continued expression of CCR5
depends on the presence of IL-2 (31), and synthesis of
MIP-1, MIP-1ß, and RANTES retains CCR5+
cells in the mucosa (32).
The high frequency of CCR5+ T cells in the female genital mucosa may explain the relatively high efficiency of infection of these cells with macrophage-tropic HIV-1 isolates, particularly when conjugated to mucosal dendritic cells. Thus, these T cells may serve as early targets of infection soon after HIV-1 exposure and contribute to the selective expansion of macrophage- and CCR5-tropic HIV-1 strains. Th1-dominated immune responses to coinfections or tissue damage may further enhance the infection locally, which is supported by studies demonstrating increased risk of HIV-1 transmission among women with genital ulcer disease and other sexually transmitted infections (33, 34).
To extend these findings, additional studies now are needed to identify
the chemokine and cytokine milieu that regulate chemotaxis and
differentiation of these CCR5-expressing, IL-2- and IFN--producing T
cells, as well as the induction of Ag-specific immune repertoire in the
genital mucosa. Our work establishes the feasibility to address these
issues, and clearly new insights may have utility in understanding
acquired immunity to HIV-1 and other sexually transmitted diseases.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. M. Juliana McElrath, Program in Infectious Diseases, Fred Hutchinson Cancer Research Center, 1100 Fairview Avenue North, D3-100, Seattle, WA 98109-1024. E-mail address:
3 Abbreviation used in this paper: MEPE, molecules of equivalent PE.
Received for publication September 11, 1998. Accepted for publication June 7, 1999.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, MIP-1ß receptor as a fusion cofactor for macrophage-tropic HIV-1. Science 272:1955.[Abstract]
and MCP-1 differentially regulate acute and relapsing autoimmune encephalomyelitis as well as Th1/Th2 lymphocyte differentiation. J. Leukocyte Biol. 62:681.[Abstract]
, MIP-1ß, and RANTES is associated with a type 1 immune response. J. Immunol. 157:3598.[Abstract]
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R. Krzysiek, E. A. Lefevre, J. Bernard, A. Foussat, P. Galanaud, F. Louache, and Y. Richard Regulation of CCR6 chemokine receptor expression and responsiveness to macrophage inflammatory protein-3alpha /CCL20 in human B cells Blood, October 1, 2000; 96(7): 2338 - 2345. [Abstract] [Full Text] [PDF]
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). D, To determine
relative CCR5 receptor density, cells were simultaneously stained with
anti-CD3 PE-Cy5 and saturating amounts of anti-CCR5 PE, and a
mixture of calibrated PE bead populations (MEPE) were run in parallel.
Histograms show the fluorescence intensity for binding of anti-CCR5
PE to CD3-gated T cells in one representative individual. Black arrows
indicate the fluorescence intensities for which MEPE values were
calculated (see text). Histograms depicting the bead mixture are
overlayed as black lines and were used to establish a standard curve
for calculation of MEPE values. E. MEPE values for
CD3-gated T cells in seven individuals tested. The lines connect values
from the two sites, peripheral blood and genital mucosa, in each
donor.