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*
Pulmonary and Critical Care Medicine Division,
Hematology/Oncology Division, Department of Internal Medicine, and
Department of Pediatrics and Communicable Diseases, University of Michigan, Ann Arbor, MI 48109
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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50%
occupied with uPA, a receptor occupancy rate that does not change
appreciably even as in vitro activation up-regulates uPA and uPAR
expression (23, 24). PMNs reportedly express a lower
proportion (10–15%) of occupied receptors, again with little change
after in vitro stimulation (25). One of the signature
changes in leukocyte uPAR is that it aggregates at the cell-substratum
interface during adhesion and at the leading edge of polarized,
migrating cells (7, 26, 27, 28). We recently demonstrated in
promonocyte-like U937 cells and human monocytes that uPAR
aggregation, achieved by Ab-mediated cross-linking, stimulates prompt
increases in intracellular levels of d-myo-inositol
(1, 4, 5) trisphosphate, followed by increases in the
intracellular Ca2+ concentration
([Ca2+]i) that were
blocked by 1) depleting intracellular Ca2+ stores
with thapsigargin, 2) inhibiting phospholipase C with U73122, and 3)
inhibiting tyrosine kinase with herbimycin A, but were less affected by
reducing extracellular Ca2+ (27).
From these findings, it was concluded that uPAR aggregation triggers
phosphoinositide hydrolysis and mobilization of intracellular
Ca2+, with secondary influxes of extracellular
Ca2+. Recognizing that human PMN express uPAR
both on the plasma membrane and in a readily mobilized pool in
intracellular granules, we sought in this study to determine whether
uPAR aggregation can also initiate activation signaling in human PMN
and regulate the proinflammatory phenotype and function of these
cells. |
Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Purified murine IgG Fc fragments and F(ab')2 of goat Ab reactive against murine IgG F(ab')2 were obtained from Jackson ImmunoResearch Laboratories (West Grove, PA). Anti-uPAR mAb (IgG2a; clone 3B10) was purified from mouse ascites by protein A-Sepharose and quantitated by protein content (29). The 3B10 anti-uPAR mAb recognizes an epitope near the uPA binding site and thus preferentially binds to unoccupied uPAR, as demonstrated by a partial (70%) reduction in 3B10 binding after saturating uPAR with exogenous uPA (30). High m.w. uPA (HMW-uPA; 90,000 IU/mg protein) was obtained from American Diagnostica (Greenwich, CT). FITC-labeled anti-CD11b mAb, anti-L-selectin mAb, and IgG1 control mAb were obtained from BioSource (Camarillo, CA).
Purification and stimulation of human PMNs
Peripheral blood was obtained from healthy volunteers according
to the provisions of the University of Michigan institutional review
board for human subject research. PMNs were isolated to >95% purity
by sedimentation in 6% dextran in
Ca2+/Mg2+-free PBS,
followed by hypotonic lysis of RBCs with H2O for
30 s and centrifugation through Ficoll-Paque (Amersham Pharmacia
Biotech, Piscataway NJ). To cross-link uPAR, cells were suspended in
experimental buffer (145 mM NaCl/5 mM KCl/1 mM
MgCl2/10 mM glucose/1 mM
CaCl2/1% (w/v) BSA/10 mM HEPES, pH 7.4). Cells
were first incubated with murine IgG Fc fragments (150 µg/ml) at
4°C for 15 min to block binding of the primary Abs to Fc receptors.
Cells were then incubated with the anti-uPAR mAb or a control IgG2a
mAb (100 µg/ml) at 4°C for 30 min, and washed in experimental
buffer at 4°C. To initiate receptor cross-linking,
F(ab')2 of goat anti-mouse
F(ab')2 Ab (100 µg/ml) were added after warming
the cells to 37°C. Previous studies have shown by immunofluorescence
microscopy that virtually identical protocols for Ab-mediated uPAR
cross-linking in human neutrophils produce readily evident receptor
capping in approximately one-half of labeled cells (10, 22). To selectively aggregate uPA/uPAR complexes, cells were
blocked with Fc fragments as described above and then treated with an
anti-uPA mAb (100 µg/ml; 394OA, American Diagnostica) or an IgG1
control, followed by the F(ab')2 of goat
anti-mouse F(ab')2 Ab (100 µg/ml) as
described above. In some instances, available uPAR were preloaded with
an excess of HMW-uPA (1 µg/ml, 20 nM) for 15 min at 4°C in
experimental buffer, washed, and then subjected to cross-linking with
the anti-uPA mAb. To confirm that HMW-uPA added in this way binds
predominately to uPAR, PMNs were pretreated with a polyclonal rabbit
anti-human uPAR Ab or control rabbit IgG, followed by FITC-labeled
uPA (1 µg/ml for 15 min, 4°C; American Diagnostica). Flow cytometry
confirmed that the anti-uPAR Ab inhibited binding of the FITC-uPA
by 93 ± 3.7% relative to the control Ab (mean ± SEM;
n = 3).
Measurement of [Ca2+]i
Cells were loaded (5 x 106/ml) with the Ca2+-sensitive fluorescent dye fluo-3/AM (2 µM; Molecular Probes, Eugene OR) at 30°C for 30 min in 145 mM NaCl/5 mM KCl/1 mM MgCl2/10 mM glucose/4 mM probenecid/10 mM HEPES, pH 7.4. After pretreatment with Abs as indicated, 2.5 x 106 cells were suspended in 1 ml of incubation buffer and prewarmed to 37°C. Fluorescence intensity was then measured with a SLM8000 spectrofluorometer equipped with SLM Spectrum Processor version 3.5 software (SLM Instruments, Urbana, IL), using a 1-cm light path cuvette with continuous stirring at an excitation wavelength of 505 nm and an emission wavelength of 530 nm. Fluorescence measurements were acquired at 2-s intervals for 300 s and converted to nanomolar concentrations of [Ca2+]i by the calibration method of Grynkiewycz et al. (31), using a Kd for fluo-3 of 864 nM (32).
Immunofluorescence flow cytometry
For immunolabeling, cells were resuspended in labeling buffer
(PBS with 0.1% human 
-globulin and 0.1% glucose, pH 7.4) and
incubated with the specified primary mAb for 30 min, 4°C, followed by
PE-conjugated goat anti-mouse Ab (30 min, 4°C). For negative
controls, cells were stained with secondary mAb alone and with an
irrelevant isotype-matched primary Ab. In some instances
FITC-conjugated primary Abs were used along with an FITC-labeled
isotype-matched Ab as a control. Fluorescence intensity was assessed as
a measure of relative Ag expression using an EPICS Elite ESP flow
cytometer (Coulter, Miami, FL; University of Michigan Flow Cytometry
Core Facility). Gating was determined by forward and 90° light
scatter characteristics. Mean fluorescence intensities (linear scale)
were determined from 
10,000 cells, and specific fluorescence
intensities were calculated by subtracting the corresponding value of
the nonspecific control. To maintain consistent results between
experiments, the flow cytometer was adjusted to provide constant
fluorescence intensities for Coulter Standard Brite beads.
Neutrophil degranulation
ß-Glucuronidase (a marker of azurophilic granules) was measured by incubating conditioned medium 1:4 (v/v) with 10% phenolpthalene ß-monoglucuronate in 70 mM sodium acetate buffer, pH 4.5, for 18 h at 37°C. The reaction was then stopped with 0.4 M glycine buffer, pH 10.5, and read in a microplate spectrophotometer at 540 nm. The reaction product was standardized against known quantities of phenolpthalene, and release was expressed as a percentage of the total cellular content in excess of unstimulated cells cultured in parallel (33). Lactoferrin (a marker for specific granules) was assayed by ELISA, using a rabbit anti-human lactoferrin capture IgG and a peroxidase-conjugated rabbit anti-human lactoferrin detection IgG (ICN/Cappel, Costa Mesa, CA), developed with a standard phenylene diamine substrate and standardized to purified human lactoferrin (34, 35). Data are expressed as nanograms per milliliter released in excess of unstimulated control cells.
Superoxide release
PMNs were incubated in 145 mM NaCl/5 mM KCl/1 mM MgCl2/10 mM glucose/1 mM CaCl2/1% (w/v) BSA/10 mM HEPES, pH 7.4, with Fe3+ cytochrome c (675 µg/ml; Sigma, St. Louis, MO) with or without superoxide dismutase (250 µg/ml; Sigma) for 45 min at 37°C. The reaction was terminated by rapid chilling to 4°C and removing the cells by centrifugation. Conditioned medium was then transferred to microtiter wells and read at 550 nm in a plate spectrophotometer. Superoxide production was determined from a standard curve relating absorbances to known quantities of sodium dithionite-reduced Fe3+ cytochrome c, and data are expressed as nanomoles of superoxide released per 45 min/106 cells (36).
Statistical analysis
Individual comparisons of group means were performed with
two-tailed Student’s t tests, with p 
0.05 considered significant. Multiple comparisons were performed using
one-way ANOVA, with Dunnett’s post-test for multiple comparisons to a
single control or Bonferroni’s post-test for multiple selected
comparisons, as indicated. All analyses were performed with GraphPad
Prism version 3.00 for Windows (GraphPad Software, San Diego,
CA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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uPAR was demonstrable by flow cytometry on 34 ± 3.6% (mean ± SEM; n = 9) of PMNs immediately after purification, in keeping with prior observations that uPAR is expressed on PMNs at relatively low levels before stimulation with proinflammatory agents (25). There was significant interdonor variability in the quantity of uPAR expressed, with the sampled population expressing 2.3 ± 1.24 arbitrary units (mean ± SD; coefficient of variation, 54%; range, 0.82–4.3). Flow cytometry was also used to determine the occupancy state of uPAR by measuring uPA on the cell surface relative to the quantity of uPA bound after prior treatment with HMW-uPA (1 µg/ml for 15 min at 4°C). Immediately upon purifying the PMNs, 30.3 ± 4.3% (mean ± SEM; n = 9) of available uPA binding sites were occupied with uPA.
Effect of uPAR aggregation on intracellular Ca2+ concentrations
Ab-mediated cross-linking of uPAR was performed to determine
whether uPAR aggregation stimulates a change in
[Ca2+]i. Cross-linking
uPAR increased [Ca2+]i
from a baseline of 207 ± 9.6 nM to a peak of 501 ± 41.7 nM
(p < 0.0001). The difference between baseline
and peak [Ca2+]i (
[Ca2+]i) was 295 ±
37 nM (Fig. 1
). By contrast,
cross-linking an isotype-matched control mAb did not significantly
affect [Ca2+]i (baseline
and peak [Ca2+]i,
188 ± 9.8 and 188 ± 10.4 nM, respectively;
n = 9). Likewise, neither the primary mAb nor
cross-linking Ab alone affected
[Ca2+]i (not shown). The
uPAR has a single binding site for uPA, and uPA is monovalent toward
uPAR (6), so it is expected that uPA/uPAR complexes have
no direct mechanism for aggregation without a cross-linking agent, in
this case the secondary Ab. The increase in
[Ca2+]i in response to
uPAR aggregation began 38 ± 4.8 s after introducing the
cross-linking Ab, consistent with a response contingent upon
receptor redistribution. The duration of the response was 127 ±
6.2 s. By comparison, the
[Ca2+]i was 2- to 3-fold
greater in PMNs than previously observed for U937 cells and freshly
purified human monocytes, while all cell types yielded
[Ca2+]i responses of very
similar duration (27). To provide another frame of
reference for PMNs, fluo-3/AM-labeled PMNs were stimulated with FMLP
(5 x 10-7 M), producing virtually
immediate, large, and transient increases in
[Ca2+]i, (Fig. 2). This result is consistent with a
synchronous response to a simple ligand-receptor interaction,
whereas uPAR aggregation probably occurs asynchronously among the
cells, yielding a [Ca2+]i
transient of longer duration and a lower apparent peak. To further
compare the [Ca2+]i
responses, the areas under the respective curves were quantitated to
measure the increases in
[Ca2+]i integrated over
time. In this respect, the magnitude of the
[Ca2+]i response to uPAR
aggregation compared more favorably with FMLP, producing approximately
one-half the integrated Ca2+ signal (Fig. 2B).
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44 µM) to compare the effects on Ca2+
signaling of uPAR ligation vs uPAR aggregation. Similar to findings
reported previously (22), binding uPA to uPAR elicited
increases in [Ca2+]i
(Fig. 3). However, the response differed
from that elicited by uPAR aggregation in several important respects.
First, like FMLP, the onset of the response to HMW-uPA was virtually
immediate, consistent with a simple ligand-receptor interaction.
Secondly, the response was relatively brief. As shown by a
representative tracing in Fig. 3A, there was an initial
spike of 10 s or less, followed by a gradual return to baseline.
The overall duration of the response was 63.3 ± 4.1 s, which
was one-half the duration of the
[Ca2+]i response to uPAR
aggregation. The
[Ca2+]i integrated over
time indicates that the magnitude of the Ca2+
signal to uPAR cross-linking exceeded the response to HMW-uPA ligation
by >3-fold (Fig. 3B). Stimulation with 2000 IU/ml HMW-uPA,
although still far exceeding the 0.1–0.5 nM
Kd reported for uPAR (6),
did not have a significant effect on
[Ca2+]i (not shown).
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, there was a relatively
small increase from baseline to peak
[Ca2+]i, corresponding to
a [Ca2+]i of 147
± 59.6 nM. Cross-linking a control IgG1 mAb had no effect (
[Ca2+]i = 17.7 ±
9.8 nM). When all available uPAR were fully saturated with uPA,
cross-linking uPA then induced Ca2+ transients
that were generally comparable to those induced by uPAR cross-linking,
with larger [Ca2+]i
that were statistically indistinguishable from the responses to
cross-linking the anti-uPAR mAb. The
[Ca2+]i without uPA
loading was 33% of the maximal
[Ca2+]i (elicited by
cross-linking uPA-loaded uPAR), which is essentially equivalent to the
30% occupancy rate of uPAR, as determined by flow cytometry.
Therefore, magnitude of the
[Ca2+]i signal appears to
be related to the quantity of uPAR cross-linked regardless of the state
of occupancy with uPA.
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Many routes of proinflammatory signaling induce PMNs to alter the
expression of adhesion molecules so as to facilitate the transition
from nonadherent or loosely adherent, rolling adhesion to firm,
integrin-mediated adhesion. To determine whether signals initiated by
uPAR can promptly alter expression of adhesion molecules, PMNs were
subjected to uPAR cross-linking, after which the cells were
immunolabeled with FITC-conjugated anti-CD11b, FITC-conjugated
anti-L-selectin, or an isotype-matched (IgG1) control mAb.
Cross-linking uPAR increased surface levels of CD11b by 80%, with
the maximal response occurring within 5 min (Fig. 5). While cross-linking the control IgG2a
mAb induced a smaller increase in CD11b expression, the response to
uPAR cross-linking was significantly greater after both 5 and 10 min
(p < 0.02). Progressive loss of L-selectin was
seen over 40 min even after cross-linking the control IgG2a mAb,
possibly triggered by the repeated manipulations inherent in the
cross-linking procedure. Cross-linking uPAR caused greater losses of
L-selectin consistently throughout the time course, but the differences
from control were not statistically significant.
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To further examine the downstream consequences of uPAR
aggregation, PMNs were subjected to the receptor aggregation protocols
described above and incubated at 37°C for 10 min in the presence of
cytochalasin B (5 µg/ml) to enhance the release of intracellular
granules (33). Cells were preloaded with fluo-3/AM so the
increase in [Ca2+]i could
be monitored in parallel. As shown in Fig. 6, uPAR aggregation significantly
increased the release of ß-glucuronidase (a marker of azurophilic
granules) and lactoferrin (a marker of specific granules). In both
cases, degranulation was significantly greater than the response to
cross-linking a control IgG2a mAb. Interestingly, the magnitude of both
degranulation responses correlated very closely with the log of the
concomitant [Ca2+]i
(Fig. 6). Selectively aggregating uPA-uPAR complexes with the
anti-uPA mAb produces minimal levels of degranulation, again in
keeping with the smaller increases in
[Ca2+]i (Fig. 7). However, preloading uPAR with
exogenous HMW-uPA at 4°C before aggregating the uPA-uPAR complexes
augmented the degranulation responses, virtually duplicating the effect
of aggregating uPAR directly. Preloading with HMW-uPA followed by sham
cross-linking (no primary mAb) did not induce degranulation (not
shown). The effects of acutely exposing cells to high concentrations of
HMW-uPA was determined by adding 4000 IU/ml to PMNs at 37°C and
measuring degranulation over 10 min. HMW-uPA triggered a modest degree
of degranulation over unstimulated controls (3.7 ± 2.1%
ß-glucuronidase release (mean ± SEM; n = 5; not
significant) and 214.3 ± 67.9 ng/ml lactoferrin release
(mean ± SEM; n = 6; p < 0.03)).
These findings indicate that both the magnitude of the
[Ca2+]i signal and the
degranulation responses are dependent upon the extent to which uPAR is
aggregated, and the occupancy state of uPAR with uPA has little or no
impact on degranulation.
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Freshly purified PMNs were treated to cross-link uPAR directly
(3B10 anti-uPAR mAb, binding preferentially to unoccupied uPAR) or
only uPA-uPAR complexes (using the anti-uPA mAb). Cells were then
incubated 45 min to measure superoxide release. Experiments were also
performed where cross-linking was initiated 30 min before adding FMLP,
after which superoxide release was measured over 45 min. Neither
saturating available uPAR with HMW-uPA (1 µg/ml) nor uPAR aggregation
with the 3B10 mAb had any direct effect on superoxide production (Fig. 8). By contrast, selectively
cross-linking uPA-uPAR complexes significantly increased superoxide
release, approximately equivalent to a half-maximal response to FMLP.
Moreover, the amount of uPA bound to uPAR, although occupying only 30%
of the available binding sites, was sufficient to support this response
fully, because superoxide release was not increased further by
preloading uPAR with HMW-uPA before cross-linking. Aggregating uPA-uPAR
complexes also increased the subsequent superoxide response to FMLP
over a range of concentrations from
10-9–10-6 M (Fig. 9). At the higher FMLP concentrations,
the additive effect was approximately twice the increment in superoxide
release induced by uPA/uPAR cross-linking alone. Again, saturating uPAR
with uPA before cross-linking with the anti-uPA mAb did not magnify
this effect. Directly cross-linking uPAR with the 3B10 mAb enhanced
FMLP-induced superoxide release as well, but the effect was
comparatively small (Fig. 9). These findings indicate that aggregating
uPA-uPAR complexes is capable of enhancing superoxide release both
alone and as a costimulus with FMLP, while aggregating the receptor
without its ligand cannot fully duplicate these responses.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The present study demonstrates that uPAR, although expressed at
relatively low levels in resting human PMNs, is capable, when
aggregated, of initiating intracellular signaling, manifested by
increased [Ca2+]i. The
response to uPAR aggregation was substantial, approximately one-half of
the response elicited by FMLP (Figs. 1
and 2), as reflected by the
integrated Ca2+ signal (area under the
[Ca2+]i vs time curve),
which was used to contend with the distinctly different
Ca2+ waveforms generated by the two stimuli.
Analyzing the results in this way, it is also apparent that uPAR
aggregation induces a much larger Ca2+ response
than supersaturating concentrations of exogenous HMW-uPA. The magnitude
of the increase in
[Ca2+]i also exceeds the
responses to uPAR aggregation seen in U937 cells and monocytes
(27). Certainly, the consequences of
Ca2+ signaling cannot be judged only by the
magnitude of the change in
[Ca2+]i. Further studies
of individual cells using quantitative fluorescence microscopy will be
necessary to determine not only the size but also the subcellular
distribution and configuration (transient vs sustained or oscillatory)
of the Ca2+ responses, all factors that impact
downstream activation signaling (47).
The immediate effects of aggregating uPAR include significant
up-regulation of CR3 expression (Fig. 5), suggesting that uPAR
aggregation may be a factor in the progression of PMNs toward firm,
CR3-dependent adhesion. This transition is not uniquely elicited by
uPAR cross-linking, as aggregating L-selectin and CR3 produce similar
results (3, 4, 5). Nevertheless, CR3 and L-selectin
colocalize during L-selectin aggregation, just as uPAR can
colocalize with CR3, so it possible that these activation schemes
are all converging on shared signaling pathways rather than
representing redundant, but independent, mechanisms of PMN activation
(3, 14).
As shown in Fig. 6, uPAR aggregation elicits PMN degranulation, as
represented by enhanced release of ß-glucuronidase and lactoferrin
(48). The magnitude of the degranulation response was
closely related to the magnitude of the increase in
[Ca2+]i, and neither was
significantly affected by the state of receptor occupancy, i.e., the
presence or the absence of associated uPA. Although few if any agonists
can match the massive degranulation induced by FMLP, the magnitude of
the degranulation response to uPAR aggregation was comparable to that
described for PMN extravasating into a wound in vivo or to the
responses in vitro to other proinflammatory agonists, such as
leukotriene B4 and IL-8 (49, 50, 51). Because uPAR aggregation
occurs as leukocytes adhere and migrate (7, 14), it is
reasonable to speculate that coordinating with granule release could
enhance vascular permeability and matrix degradation, and thereby
facilitate PMN extravasation. The role of intracellular
Ca2+ mobilization in triggering PMN degranulation
remains controversial, but it appears that increased
[Ca2+]i is variably
associated with degranulation depending on the agonist
(52). The close correlation between the two events in this
study certainly implies that in the case of uPAR aggregation, they are
proximate downstream events of a common pathway.
One of the more remarkable findings of this study was the enhanced
superoxide release triggered specifically by aggregating uPA-uPAR
complexes (Figs. 8 and 9). In many ways, this observation expands on
the prior observations of Cao et al. (22), who reported
that HMW-uPA primes PMNs for superoxide release by forming a ternary
signaling complex with uPAR and CR3. The present study shows that
aggregating uPA circumvents the need for very high, and arguably
supraphysiologic, concentrations of HMW-uPA needed to elicit this
response. In fact, the relatively small proportion (30%) of uPAR
occupied with uPA was fully capable of triggering a maximal response,
as loading available uPAR with exogenous HMW-uPA did not enhance
superoxide release (Figs. 8 and 9). Moreover, HMW-uPA, having no direct
effect of its own, only enhanced FMLP-induced superoxide release
(22), while aggregating uPA-uPAR complexes directly
increased superoxide release in addition to enhancing the effects of
FLMP. Cooperative signaling between uPA-uPAR aggregation and FMLP
produced somewhat more than additive superoxide responses, although the
increase was not large enough to firmly conclude that the two stimuli
are truly synergistic. It remains to be determined whether CR3 serves
as the signal transduction device for uPA-uPAR complexes in modulating
superoxide release. This certainly is a plausible mechanism, because
CR3 is involved in superoxide release (53). However, the
Ca2+ signal elicited by uPAR aggregation appears
to be independent of CR3 (27), suggesting that uPAR may
engage discrete signaling pathways depending upon the choice of
signaling partner with which it associates. CR4 (CD11c/CD18; p150/95)
is another candidate signaling partner for uPAR, because cytosolic
NAD(P)H autofluorescence oscillations vary in synchrony, but out of
phase, with proximity oscillations between aggregated uPAR and CR4 in
polarized PMNs (12). It is not known whether the
association with CR4 diminishes the intrinsic ability of uPA-uPAR
complexes to trigger superoxide release or displaces another partner
protein from associating with uPA-uPAR. Regardless of the mechanisms
involved, the present results indicate that aggregating uPA-uPAR
complexes may contribute to enhanced oxidant-mediated tissue injury
during PMN adhesion and migration. Moreover, it is clear that
aggregation of uPAR initiates two distinct pathways of activation
signaling, one being independent of receptor occupancy state and
resulting in [Ca2+]i
mobilization and degranulation, and the other critically dependent on
occupancy with uPA and resulting in enhanced superoxide release.
Prior work has emphasized the role that uPAR plays as an adapter protein that articulates its effects on cellular functions through its interactions with integrins. The present study has examined uPAR clustering as an isolated stimulus for proinflammatory signaling in nonadherent PMNs. However, one would expect that activation signaling initiated by uPAR aggregation in vivo would occur in adherent PMNs. It is not yet clear how uPAR clustering occurs in vivo, but it may occur actively as PMNs encounter a uPAR counterligand such as vitronectin (54), or it may occur passively as integrins or other proteins capable of binding uPAR are drawn into a clustered configuration. In either case, the signaling pathways engaged by uPAR clustering may be coupled to complimentary chemokine and integrin-mediated signaling events, and it will be necessary to investigate the role of uPAR-mediated activation signaling in this context.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Robert G. Sitrin, 6301 MSRB III, Box 0642, 1150 West Medical Center Drive, Ann Arbor, MI 48109-0642.
3 Abbreviations used in this paper: PMN, polymorphonuclear neutrophil; uPAR, urokinase plasminogen activator receptor; [Ca2+]i, intracellular Ca2+ concentration; uPA, urokinase plasminogen activator; [Ca2+]i, difference between baseline and peak [Ca2+]i; CR3, complement receptor 3; HMW-uPA, high m.w. urokinase plasminogen activator.
Received for publication March 13, 2000. Accepted for publication June 22, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Mß2' CD11b/CD18) during neutrophil polarization. J. Immunol. 156:297.[Abstract]
vß5 vitronectin receptor, promoting urokinase-dependent cell migration in breast cancer. Cancer Res. 59:5307., tumor necrosis factor-, and urokinase regulate the expression of urokinase receptors on human monocytes. J. Immunol. 141:4229.[Abstract]
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 O. D. Liang, K. Bdeir, R. L. Matz, T. Chavakis, and K. T. Preissner Intermolecular Contact Regions in Urokinase Plasminogen Activator Receptor J. Biochem., November 1, 2003; 134(5): 661 - 666. [Abstract] [Full Text] [PDF]
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 J. L. Coleman and J. L. Benach The Urokinase Receptor Can Be Induced by Borrelia burgdorferi through Receptors of the Innate Immune System Infect. Immun., October 1, 2003; 71(10): 5556 - 5564. [Abstract] [Full Text] [PDF]
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G. Zhang, H. Kim, X. Cai, J. M. Lopez-Guisa, P. Carmeliet, and A. A. Eddy Urokinase Receptor Modulates Cellular and Angiogenic Responses in Obstructive Nephropathy J. Am. Soc. Nephrol., May 1, 2003; 14(5): 1234 - 1253. [Abstract] [Full Text] [PDF]
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 Y. Xia, G. Borland, J. Huang, I. F. Mizukami, H. R. Petty, R. F. Todd III, and G. D. Ross Function of the Lectin Domain of Mac-1/Complement Receptor Type 3 (CD11b/CD18) in Regulating Neutrophil Adhesion J. Immunol., December 1, 2002; 169(11): 6417 - 6426. [Abstract] [Full Text] [PDF]
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 F. Winkler, S. Kastenbauer, U. Koedel, and H. W. Pfister Role of the urokinase plasminogen activator system in patients with bacterial meningitis Neurology, November 12, 2002; 59(9): 1350 - 1355. [Abstract] [Full Text] [PDF]
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 A. Berezov, J. Chen, Q. Liu, H.-T. Zhang, M. I. Greene, and R. Murali Disabling Receptor Ensembles with Rationally Designed Interface Peptidomimetics J. Biol. Chem., July 26, 2002; 277(31): 28330 - 28339. [Abstract] [Full Text] [PDF]
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V. Stepanova, U. Jerke, V. Sagach, C. Lindschau, R. Dietz, H. Haller, and I. Dumler Urokinase-dependent Human Vascular Smooth Muscle Cell Adhesion Requires Selective Vitronectin Phosphorylation by Ectoprotein Kinase CK2 J. Biol. Chem., March 15, 2002; 277(12): 10265 - 10272. [Abstract] [Full Text] [PDF]
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H. Yoshitake, Y. Takeda, T. Nitto, and F. Sendo Cross-linking of GPI-80, a possible regulatory molecule of cell adhesion, induces up-regulation of CD11b/CD18 expression on neutrophil surfaces and shedding of L-selectin J. Leukoc. Biol., February 1, 2002; 71(2): 205 - 211. [Abstract] [Full Text] [PDF]
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 M. Resnati, I. Pallavicini, J. M. Wang, J. Oppenheim, C. N. Serhan, M. Romano, and F. Blasi The fibrinolytic receptor for urokinase activates the G protein-coupled chemotactic receptor FPRL1/LXA4R PNAS, January 24, 2002; (2002) 22652999. [Abstract] [Full Text] [PDF]
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R. G. Sitrin, P. M. Pan, R. A. Blackwood, J. Huang, and H. R. Petty Cutting Edge: Evidence for a Signaling Partnership Between Urokinase Receptors (CD87) and L-Selectin (CD62L) in Human Polymorphonuclear Neutrophils J. Immunol., April 15, 2001; 166(8): 4822 - 4825. [Abstract] [Full Text] [PDF]
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O. D. Liang, T. Chavakis, S. M. Kanse, and K. T. Preissner Ligand Binding Regions in the Receptor for Urokinase-type Plasminogen Activator. STRUCTURAL REQUIREMENTS FOR A MULTIDOMAIN BINDING REGION AND RECEPTOR-RECEPTOR INTERACTION J. Biol. Chem., July 27, 2001; 276(31): 28946 - 28953. [Abstract] [Full Text] [PDF]
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M. Resnati, I. Pallavicini, J. M. Wang, J. Oppenheim, C. N. Serhan, M. Romano, and F. Blasi The fibrinolytic receptor for urokinase activates the G protein-coupled chemotactic receptor FPRL1/LXA4R PNAS, February 5, 2002; 99(3): 1359 - 1364. [Abstract] [Full Text] [PDF]
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