| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
University of British Columbia Pulmonary Research Laboratory, St. Paul's Hospital, Vancouver, British Columbia, Canada V6Z 1Y6
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
ACKNOWLEDGEMENTS
FOOTNOTES
REFERENCES
ABSTRACT 
Kitagawa, Yuko, Stephan F. Van Eeden, Darlene M. Redenbach,
Maleki Daya, Blair A. M. Walker, Maria E. Klut, Barry R. Wiggs, and
James C. Hogg. Effect of mechanical deformation on structure and
function of polymorphonuclear leukocytes. J. Appl.
Physiol. 82(5): 1397-1405, 1997.
The present
studies were designed to test the hypothesis that mechanical
deformation of polymorphonuclear leukocytes (PMN) leads to functional
changes that might influence their transit in the pulmonary
capillaries. Human leukocytes were passed through 5- or 3-µm-pore
polycarbonate filters under controlled conditions. Morphometric
analysis showed that the majority of PMN were deformed and that this
deformation persisted longer after filtration through 3-µm filters
than through 5-µm filters (P < 0.05) but did not result in the cytoskeletal polarization
characteristic of migrating cells. Flow cytometric studies of the
filtered PMN showed that there was a transient increase in the
cytosolic free Ca2+ concentration
after both 3- and 5-µm filtration (P < 0.01) with an increase in F-actin content after 3-µm filtration
(P < 0.05). Although
L-selectin expression on PMN was
not changed by either 5- or 3-µm filtration, CD18 and CD11b were
increased by 3-µm filtration (P < 0.05). Priming of the PMN with
N-formyl-methionyl-leucyl-phenylalanine (0.5 nM) before filtration resulted in an increase of CD11b by both 5 (P < 0.05)- and 3-µm
(P < 0.01) filtration. Neither 5- nor 3-µm filtration induced hydrogen peroxide production. We conclude that mechanical deformation of PMN, similar to what occurs in the
pulmonary microvessels, induces both structural and functional changes
in the cells, which might influence their passage through the pulmonary
capillary bed.
neutrophils; deformability; F-actin; adhesion molecules
INTRODUCTION THE PULMONARY MICROVESSELS restrict the passage of
polymorphonuclear leukocytes (PMN) because of the discrepancy between
the size of the PMN and the size of the lung capillary segments (5, 14,
15). Although erythrocytes [red blood cells (RBC)] have similar
maximum dimensions to PMN, their greater deformability allows them to
negotiate these restrictions more quickly (14). This results in
pulmonary capillary transit times that are 60-100 times longer for
PMN than for RBC, and this concentrates PMN with respect to RBC in
pulmonary capillaries (14, 15).
Stimuli such as peptide chemoattractants, endotoxin, and smoking are
known to decrease PMN deformability and further increase the
concentration of PMN in pulmonary capillaries (6, 18, 27, 30). Several
in vitro filtration studies have measured the biophysical properties of
PMN, particularly their size and deformability, and have shown that
these factors determine the magnitude of PMN sequestration in the
pulmonary microvessels (6, 18, 27, 30). The importance of the
discrepancy between PMN and pulmonary capillary dimensions in causing
PMN sequestration in the lung has also been demonstrated during forced
expiratory maneuvers in which raised intra-alveolar pressure compresses
alveolar microvessels and delays PMN (21). Studies in humans undergoing cardiac catheterization show that alveolar compression results in an
immediate arteriovenous difference of PMN across the lung that reverses
when alveolar pressure is returned to normal (21). Mechanical
deformation of PMN has been demonstrated in pulmonary microvessels (5),
and Wiggs et al. (29) have shown that PMN of ~7 µm in diameter
negotiate the capillary bed in the same way as 4-µm nondeformable
beads. These findings indicate that the PMN deform in the pulmonary
capillary bed, but whether this deformation is an active or passive
process remains to be determined.
Our working hypothesis is that the mechanical deformation of PMN in the
pulmonary capillaries may induce functional changes that could
influence their passage through these restrictions. To test this
hypothesis, we used an in vitro filtration system with polycarbonate
membranes to simulate the in vivo mechanical deformation of PMN. This
system has been used by others to study the deformability of PMN (6, 8,
10, 11, 18, 23, 27, 30), and we have modified it to study the effect of
PMN deformation on their structure and function. We measured the
deformation that occurs as the cells pass through the polycarbonate
filters and assessed cell polarity for evidence of active locomotion. Flow cytometry was used to determine whether this mechanical
deformation produced changes in the conversion of globular actin
(G-actin) to filamentous actin (F-actin), cytoplasmic free
Ca2+ concentration
([Ca2+]i),
the expression of cell adhesion molecules, and degranulation with
oxygen radical production.
METHODS Cell Preparation
In Vitro Filtration of PMN
LRP or purified PMN were filtered in vitro according to the filtration method described by Lennie et al. (20). Briefly, a 20-ml polypropylene syringe (Sherwood Medical, St. Louis, MO) was filled with sample solution (LRP or purified PMN) and filtered through polycarbonate filters (Poretics, Livermore, CA) with defined pore size [pore diameter 5 µm, length 10 µm, and pore density 4 × 105/cm2 or pore diameter 3 µm, length 9 µm, and pore density 2 × 106/cm2 (manufacturer's data)] by using a syringe-infusion pump (pump 22, Harvard Apparatus, Millis, MA), which provides a constant flow rate of solution across the filter. Hydrostatic pressure was continuously monitored upstream from the filter by using a pressure transducer (Validyne Engineering, Northridge, CA) connected to a recording system. The pressure-sensing system was calibrated by using a water manometer under conditions of no flow before each filtration. Cell viability after filtration was 98% as tested by trypan blue exclusion.Morphometric Analysis of PMN
LRP (4 × 105 cells/ml) prepared by the methods described above was filtered through polycarbonate filters with pore sizes of 5 and 3 µm at two different flow rates (3.0 or 1.0 ml/min). Samples were fixed in paraformaldehyde (final 1.6%) before filtration, immediately after (time 0), and 10, 30, 60, and 180 s after filtration. Cells allowed to settle were stained with Giemsa in suspension. The degree of deformation was quantified by dividing the longest by the shortest diameter (perpendicular at midpoint of longest diameter) to obtain a long-to-short ratio (L/S) in 20 randomly selected PMN per time point (n = 4). These measurements were made on a standardized image-analysis system (Bioquant System IV, R&M Biometrics, Nashville, TN).Analysis of PMN Polarity
The distribution of microtubules and F-actin was determined in PMN after either 3- or 5-µm filtration at 3.0 ml/min at time = 0, 10, 30, 60, or 180 s (n = 3). The PMN were double stained with anti-tubulin (5A6 mouse monoclonal antibody raised against
-tubulin, gift from Dr. David Brown, final
dilution 1:4,000) (16) to localize microtubules or fluorescein
isothiocyanate (FITC)-conjugated phalloidin (1:200 dilution; Molecular
Probes, Eugene, OR) to localize F-actin. Experiments were carried out with materials in suspension at room temperature, except
if otherwise noted. Cells were permeabilized with a mixture of
methanol-acetone (1:1) at 
20°C for 10 min, followed by
sequential incubation in 2 µM
L--lysophosphatidylcholine
palmitoyl (Sigma Chemical) and 0.2% Triton X-100 in PBS for 15 min at
37°C. Non-specific binding was blocked with 5% normal goat serum
in 0.1% bovine serum albumin (BSA)/PBS for 30 min at 37°C. Samples
were incubated in 200 µl of anti-tubulin antibody 5A6 diluted in 1%
normal goat serum in 0.1% BSA/PBS for 60 min at 37°C, followed by
tetramethylrhoadamine isothiocyanate (TRITC)-conjugated goat anti-mouse
immunoglobulin (Ig) G secondary antibody for 60 min at 37°C (final
dilution 1:200). For F-actin, cells were incubated with
FITC-conjugated phalloidin for 20 min (final dilution 1:200) at
37°C. Cell suspensions were rinsed three times in 500 µl of
0.1% BSA/PBS between all steps. Cells were visualized on a
Zeiss Axiophot microscope equipped for epifluorescence with filters
for FITC (excitation 450-490; barrier 510; emission
515-556 nm) and TRITC (excitation 546; barrier 580; emission
590 nm).
Cytoskeletal polarity was evaluated without knowledge of the sample source. Twenty cells were scored from each time point. Cells were classified visually as either "deformed" or "round." The microtubule organizing center (MTOC) was scored as either "polarized," when microtubules converged at any pole of the nucleus of a deformed PMN, or "nonpolar" when they did not. Cells were scored for F-actin polarity as either "nonpolarized" (symmetrical distribution) or polarized (asymmetrical distribution) to identify F-actin-rich lamellapodia or filipodia as an indication of active cell migration.
Flow Cytometry
F-actin content of PMN. F-actin content of PMN was determined immediately after filtration by using a modification of a previously described method (17). Leukocytes filtered through polycarbonate filters (5- or 3-µm pore) at a flow rate of 3.0 ml/min were fixed immediately for 30 min at room temperature with 3% paraformaldehyde in PBS (pH 7.3) and washed for 5 min with PBS. The cells were simultaneously permeabilized and stained, in the dark at 37°C for 30 min, with a fresh mixture of 2 µM L--lysophosphatidylcholine
palmitoyl (Sigma Chemical) and 0.1 µg/ml
N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-phallacidin (NBD-phallacidin; Molecular Probes). After staining, samples were washed and resuspended in PBS. F-actin content was measured with a flow
cytometer (model Profile Epics II, Coulter Electronics, Hialeah, FL).
Analysis gates for PMN were established by using distinctive forward
and side scatter profiles and were expressed as mean fluorescent
intensity (log) of 3,000-6,000 cells. As a positive control, PMN
were stimulated with
N-formyl-methionyl-leucyl-phenylalanine (FMLP; 100 nM, Sigma Chemical) for 10 s, fixed with 3%
paraformaldehyde, and analyzed as above.
[Ca2+]i
in PMN.
Changes in
[Ca2+]i
of PMN were determined by using flow cytometry with a
fluorescent Ca2+ indicator, fluo
3-aminomethyl ester (fluo 3-AM; Molecular Probes), at excitation and
emission wavelengths of 488 and 525 nm, respectively (24). Purified PMN
were suspended in HBSS (4 × 105 cells/ml) and incubated with
2.2 µM fluo 3-AM for 30 min in the dark at room temperature. Cells
were washed twice and resuspended in HBSS. Before the filtration or
stimulation, baseline
[Ca2+]i
was measured in triplicate. Changes in fluo 3 fluorescence, representing
[Ca2+]i,
were measured for up to 3 min after filtration through 5- or
3-µm-pore filters (flow rate 3.0 ml/min) or after addition of
stimuli. Ionomycin (10 µM, Calbiochem, La Jolla, CA) and 100 nM FMLP
stimulation were used as positive controls.
Expression of L-selectin, CD11b,
and CD18 on PMN.
After filtration of leukocytes (4 × 105 cells/ml) through 5- or
3-µm-pore filters by using a flow rate of either 1.0 or 3.0 ml/min,
the expression of L-selectin and
CD11b/CD18 was measured by indirect immunofluorescence. One hundred
microliters of prefiltered or filtered LRP were incubated with 200 µl
of PBS (pH 7.3) containing the primary antibodies against human
L-selectin (1 µg/ml, Leu-8, Becton-Dickinson, San Jose, CA), CD11b
(1.6 µg/ml, DAKO-CD11b, Dakopatts, Glostrup, Denmark), or CD18 (6.9 µg/ml, DAKO-CD18, Dakopatts) for 10 min in the dark at room
temperature. LRP incubated with mouse
IgG1 (2 µg/ml, Sigma Chemical)
served as negative control. Cells were washed with PBS and incubated
with FITC-conjugated anti-mouse IgG (5 µg/ml, Sigma Chemical) as a
secondary antibody for 10 min at room temperature in the dark. The
remaining RBC in the samples were lysed (Immunolyse and Coulter Clone,
Coulter Electronics), and the leukocytes were fixed with 1%
paraformaldehyde and stored at 4°C. The immunofluorescence was
measured by using flow cytometry as described above. Cells incubated
with 100 nM FMLP or 1 µM phorbol 12-myristate 13-acetate (PMA; Sigma
Chemical) served as positive controls. To test the effect of mechanical deformation of primed PMN on the expression of adhesion molecules, PMN
exposed to priming doses of FMLP (0.5 nM) were filtered through the
polycarbonate filters (5- or 3-µm pore) by using a 3.0 ml/min flow
rate. The expression of
L-selectin and CD11b was
determined by immunofluorescence as described above. The changes in
fluorescence after filtration on primed PMN were compared with those of
untreated PMN.
Hydrogen peroxide
(H2O2)
production by PMN.
The respiratory burst activity of PMN after mechanical deformation was
determined by measuring PMN
H2O2
production according to the method previously described by Bass et al.
(4). Briefly, PMN are loaded with dichlorofluorescin-diacetate
(DCFH-DA), which diffuses into the cells and is hydrolyzed
intracellularly to 2
,7-dichlorofluorescin (DCFH), a
nonfluorescent compound.
H2O2
produced during the respiratory burst oxidizes the DCFH to highly
fluorescent 2,7-dichlorofluorescein (DCF), which can be
measured by flow cytometry. LRP was incubated with PMN buffer
consisting of 5 µM DCFH-DA (prediluted in 95% ethanol; final
concentration of ethanol was 0.1%) and 0.1% gelatin and horizontal
agitation in a shaking water bath at 37°C and was kept at this
temperature for the whole experiment. Baseline DCF fluorescence was
measured as well as fluorescence of PMN after filtration through 5- or
3-µm-pore filters for up to 40 min. As positive controls, 100 nM FMLP
and 1 µM PMA (Sigma Chemical) were used.
Statistical Analysis
Changes in deformation and polarity between PMN filtered through 3- and 5-µm filters and over time were analyzed by using
2. Recovery of cell shape to
prefilter shape was examined by comparison of deformation at each time
point with prefilter values.
Differences between prefiltered or prestimulated control values for each condition were treated as individual parametric values, and groups of these values were compared with postfiltered or poststimulated values by using paired t-tests. Changes over time were analyzed by using analysis of variance for repeated measures. Corrections for multiple tests and comparisons were performed by using the sequential rejective Bonferroni procedure (18). Corrected P < 0.05 were considered significant. All values are expressed as the means ± SE.
RESULTS
Pressures and Shear Stress During PMN Filtration
The pressures 1 min after starting filtration of the LRP at a concentration of 4 × 105 cells/ml were 0.6 ± 0.2 (1.0 ml/min) and 3.9 ± 0.7 cmH2O (3.0 ml/min) through 5-µm pores and 5.6 ± 1.3 (1.0 ml/min) and 22.5 ± 3.7 cmH2O (3.0 ml/min) through 3-µm pores. We calculated the shear stress on leukocytes at the wall by using the method of Downey and Worthen (10). In this technique the wall shear stress (
) is related
to the diameter (d) and length
(l) of the tube as well as the
pressure drop across the tube (
P) by the relationship
= [Pd/(4l)].
The calculated shear stress in the 5-µm pores with a length of 10 µm was 487.5 dyn/cm2 at high
flow rates and 75 dyn/cm2 for the
low flow rates.
Morphometric Analysis of PMN
Figure 1 shows the representative features of PMN deformed by filtration through 5- or 3-µm-pore polycarbonate filters. The filtered PMN have elongated shapes, and their nuclei were often located off center after 3-µm filtration. Exposure to the same pressures without filtration had no effect on cell shape. Figure 2 shows the degree of PMN deformation measured by L/S. This ratio was greater for PMN filtered through 3- than through 5-µm polycarbonate filters at each time point (P < 0.01). PMN also remained deformed for a longer period when filtered through 3- than through 5-µm filters (Fig. 2).
Analysis of PMN Polarity
Table 1 shows the fractions of deformed PMN, MTOC polarization, and F-actin redistribution. Samples at different time points after 3- and 5-µm filtration at 3.0 ml/min were pooled (n = 3). These data show that 78 and 87% of cells were deformed immediately after 3- and 5-µm filtration, respectively, and recovery of spherical shape was significantly delayed in 3-µm compared with 5-µm filtration at each time point. There was an increased brightness in F-actin staining only at t = 0 s in the 3-µm filtered cells. Although many cells were greatly deformed, there was no immunocytochemical evidence of microtubular or microfilament reorganization or polarization characteristic of migrating cells.
|
|||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Flow Cytometry
F-actin content of PMN. Filtration of PMN through 3-µm-pore filters increased their F-actin content by 94 ± 44% from prefiltered values (P < 0.05, n = 4) immediately after filtration (t = 0 s), which was similar to that seen with FMLP stimulation (83 ± 17%, P < 0.01, n = 4, Fig. 3). Filtration through 5-µm filters did not change F-actin content of PMN at t = 0 s. The exposure of PMN to the pressure required to force them through the filters had no effect on their F-actin content.
Free [Ca2+]i in PMN. Filtration of PMN through both 5- and 3-µm pores caused an increase in cytosolic free [Ca2+]i (Fig. 4A). The degree of this increase was greater in 3-µm filtration than in 5-µm filtration (P < 0.05, n = 4) but was smaller than the value obtained with FMLP or ionomycin stimulation (P < 0.01, n = 4). Unlike the immediate increase of [Ca2+]i in PMN stimulated by FMLP or ionomycin, the peak increase of [Ca2+]i in filtered PMN was observed 30 s after filtration (Fig. 4B). The increase in [Ca2+]i was sustained for up to 3 min after stimulation by ionomycin, in contrast to a rapid decline to baseline after 3 min in the [Ca2+]i signal after FMLP stimulation or filtration of PMN.
To determine whether [Ca2+]i increase by filtration induces cross-desensitization to activation by FMLP, PMN were sequentially activated by either filtration or 100 nM FMLP, followed by 100 nM FMLP at 3 min. Three minutes after stimulation, fluo 3 fluorescence was back to baseline, and this time point was selected for the second stimulation. Figure 5A shows that sequential stimulation of PMN after initial FMLP stimulation did not result in a second increase in [Ca2+]i. In contrast, FMLP stimulation after 3-µm filtration induced a significant second [Ca2+]i signal (Fig. 5B).
Expression of L-selectin, CD11b, and CD18 on PMN. Changes in the surface expression of the cell adhesion molecules L-selectin and CD11b/CD18 on PMN after deformation by filtration through 5- or 3-µm-pore filters are shown in Fig. 6. PMN filtration through 5- or 3-µm-pore filters did not change the expression of L-selectin, in contrast to stimulation with either 100 nM FMLP or 1 µM PMA (Fig. 6A). CD11b and CD18 were significantly upregulated by 3-µm filtration (Fig. 6, B and C) by using both 1.0 and 3.0 ml/min flow rate (n = 4). Five-micrometer filtration did not change the expression level of CD11b/CD18 on PMN (Fig. 6, B and C). The exposure to the pressure required to pass the PMN through the filters did not affect expression of L-selectin or CD11b/CD18 (data not shown).
, Flow rate (ml/min); , pore size. Significantly
different vs. control: * P < 0.05; ** P < 0.01.
Neither a suboptimal (priming) dose of 0.5 nM FMLP nor 5-µm filtration alone induced upregulation of CD11b on PMN, as shown in Fig. 7. However, PMN primed with 0.5 nM FMLP showed a significant increase in their expression of CD11b when passed through 5-µm filters (P < 0.05, n = 4). Moreover, the upregulation of CD11b was enhanced when primed PMN were filtered through 3-µm filters (P < 0.01, n = 4). The filtration of primed PMN had no effect on the expression level of L-selectin (data not shown).
H2O2 production by PMN. The time-related changes in DCF fluorescence (H2O2 production) in PMN are shown in Fig. 8. Significant increases in DCF fluorescence were induced by 100 nM FMLP (5 min after stimulation, P < 0.01, n = 4) or 1 µM PMA (30 min after stimulation, P < 0.05, n = 4). PMN filtration did not induce an increase in DCF fluorescence after either 5- or 3-µm filtration.
,7-dichlorofluorescein (DCF) fluorescence representing
H2O2
were measured for up to 40 min. Cells treated with 100 nM FMLP, and 1 µM PMA served as positive controls. Time-related changes of mean
fluorescent intensity of DCF were compared with fluorescence of
unfiltered controls at each time point. Significantly different:
* P < 0.05;
** P < 0.01.
DISCUSSION
The cells circulating through the pulmonary microvessels must negotiate an average of 50-100 capillary segments between the arterial to the venous side of the pulmonary vascular bed (14, 15). The average diameter of each segment is 7.4 ± 2.3 µm in humans with a range between 2 and 15 µm (14). We have used filters with 3- and 5-µm pore sizes to simulate the smaller 20-30% of pulmonary capillary segments to determine the effect of deformation on the phenotypic and functional characteristics of PMN (5, 16). The experimental procedure chosen to examine the problem has been extensively used by other investigators (6, 8, 10, 11, 18, 23, 27, 30), and Selby et al. (27) have demonstrated, by using a similar system, a relationship between in vitro deformability of PMN and the in vivo sequestration of radiolabeled PMN in the lungs of human subjects. Downey and Worthen (10) have investigated the factors that determine the passage of PMN through polycarbonate filters and have calculated that the shear stress in their in vitro filtration system (80-800 dyn/s2) was remarkably similar to the estimated wall shear stress in vivo of between 100 and 500 dyn/s2 (10, 12). In our experiments we have used similar flow rates (1.0 and 3.0 ml/min) and calculated similar wall shear stress values. All filters were coated with albumin, and cells were prepared and filtered by using techniques to eliminate any significant exposure of cells to endotoxin that could result in cell activation. Hydrodynamic forces of flow, driving pressure, shear stress, and pore size, as well as cell size and deformability, have been shown to be important determinants in PMN passage through filters of a defined pore size (10-12). In this study we have determined the functional and structural changes that occur when PMN are exposed to hydrodynamic forces similar to those when they pass through smaller capillary segments in the lung.
The morphometrical analysis showed greater and more prolonged deformation after filtration through 3-µm compared with 5-µm-pore filters. The degree of PMN deformation estimated by L/S in the present study is within the range of deformation that was previously observed in human pulmonary capillaries and rabbit lungs (5). The delayed recovery of PMN to a rounded shape is also consistent with recent studies from our laboratory on isolated perfused rabbit lungs. In these studies PMN deformed in the capillaries maintained this elongated shape as they exited the pulmonary circulation, indicating that they retain this change in shape after their exit from the capillary bed (26). Varying the flow rates produced different driving pressures and shear stress on the PMN but did not affect the degree to which the cells deform or their recovery time to a spherical shape. Therefore, we conclude that the degree of deformation is mainly determined by pore size.
F-actin assembly is important for cytoskeletal reorganization and stabilization necessary to induce a migratory phenotype in PMN and acts as a critical factor in the mechanisms leading to protrusion of lamellapodia and provision of motor functions for chemotaxis (7). In response to stimulation by chemotactic factors, there is a rapid and transient increase in the assembly of F-actin from the G-actin pool (7). In several other cell systems, such as with endothelial cells in vitro (28) and in vivo (19), mechanical stress caused reorganization of actin filaments. Our data demonstrate an immediate short-lived increase in PMN F-actin content only after filtration through 3-µm-pore filters. This observation was supported by our immunofluoresence data showing brighter F-actin staining only after 3-µm filtration at t = 0 s (Table 1). No evidence of actin-rich lamellapodia or filipodia, which would indicate active migration, was observed. Similarly, the MTOC did not show evidence of organelle polarization, indicating active migration of PMN (Table 1). The prolonged deformation of PMN associated with the lack of MTOC and F-actin polarity is more consistent with deformation-induced stabilization of the actin network by cross-linking events rather than actin polymerization in preparation for migration.
An increase in free [Ca2+]i is an important second messenger in cell signal transduction pathways by activation of Ca2+-dependent proteases, protein kinases, and phosphatases. Recent studies have shown that mechanical stress induced a Ca2+ signal in several other cell systems (1). We observed a modest but significant elevation in [Ca2+]i in these studies, but the amplitude and time course of the Ca2+ response associated with mechanical deformation were markedly different from those observed after FMLP treatment. This suggests that a different mechanism may be responsible for Ca2+ mobilization after deformation, and this is supported by the observation that, in contrast to sequential stimulation by FMLP, the Ca2+ response associated with mechanical deformation of PMN did not abolish or impair a subsequent Ca2+ response by FMLP stimulation. PMN deformation may induce a Ca2+ signal through a nonreceptor mechanism similar to the Ca2+ signal in RBC. The Ca2+ signal resulting from PMN filtration is also different from the signaling pathway induced by PMA, which can induce oxidative burst activity of PMN without cytoplasmic Ca2+ flux. This increase of [Ca2+]i after PMN filtration may contribute to activation of Ca2+-dependent protein kinases, proteases, or phosphatases.
The expression of adhesion molecules acts as a component of a molecular
cascade in leukocyte-endothelial interaction and is regulated according
to the activation status of PMN (2). Chemotactic stimulation of PMN
results in shedding of
L-selectin and translocation of
CD11b/CD18 (MAC-1) from intracellular storage pools to the cell surface
(17). Therefore, the quantitative evaluation of these molecules is a
useful indicator for PMN activation status. PMN deformation causes a
differential change of cell adhesion molecules. The upregulation of
CD11b/CD18 induced by 3-µm filtration shows that PMN are activated by
mechanical deformation. These changes were flow rate independent and
underline the importance of pore size and the degree of deformation,
rather than shear stress, as a determinant of CD11b/CD18 upregulation.
Furthermore, the calculated shear stress in our system (80-800
dyn/cm2) is three orders of
magnitude larger than that required to inhibit adherence to flat
endothelial surfaces (10). Unlike the upregulation of CD18/CD11b, PMN
deformation did not affect the expression of L-selectin. This differential
effect of PMN deformation on cell adhesion regulation suggests that
there are different signaling pathways that modify these molecules
(22). Molad et al. (22) recently demonstrated that immune complexes
induced the same type of differential effect on adhesion molecule
expression of PMN (impaired
L-selectin shedding after
stimulation). They speculated that upregulation of
2-integrins with impaired
L-selectin shedding causes
tighter adhesion of PMN onto microvascular endothelium without
migration, which may cause vascular endothelial injury if associated
with production of active oxygen metabolites. Our data show that
mechanical deformation of PMN did not cause
H2O2 production even in 3-µm filtration with flow rates of 3.0 ml/min. This result suggests that the mechanical deformation induced by filtration through small pores was not able to induce the respiratory burst associated with damage to the pulmonary microvessels.
The physiological significance of the structural and functional changes observed in PMN after mechanical deformation remains to be established in vivo. The difference in CD11b after 5- and 3-µm filtration suggests there is a threshold for the degree of deformation that produces CD11b upregulation and that priming the PMN with low doses of FMLP lowers this threshold (Fig. 7). This suggests that primed PMN may be more sensitive to the activation by mechanical stress, and we speculate that mechanical stress is one of the factors that influences PMN function at inflammatory sites.
In conclusion, this study establishes that mechanical deformation of PMN causes functional changes that include reorganization and stabilization of the cytoskeleton that could contribute to their passage through small restrictions in the pulmonary capillary bed. It also results in signal transduction with an increase in free [Ca2+]i and upregulation of cell-surface expression of CD11b/CD18. It further shows that the effect of deformation can be enhanced by priming PMN. These changes may be of relatively little importance under normal physiological conditions but could enhance the response of PMN sequestrated in the pulmonary capillary beds compressed by positive end-expiratory pressure induced by either external sources in mechanically ventilated patients or internal sources in patients with chronic airway obstruction. If these modest effects were amplified by other inflammatory stimuli, they might enhance the recruitment of PMN and contribute to lung injury.
ACKNOWLEDGEMENTS
We especially thank Stuart Greene and Beth Whalen for technical support and Lorraine Verburgt for statistical advice and analysis.
FOOTNOTES
Address for reprint requests: S. F. Van Eeden, Univ. of British Columbia, Pulmonary Research Laboratory, St. Paul's Hospital, 1081 Burrard St., Vancouver, British Columbia, Canada V6Z 1Y6.
Received 30 September 1996; accepted in final form 22 January 1997.
REFERENCES
| 1. |
Adams, D. S.
Mechanism of cell shape: the cytomechanics of cellular response to chemical environment and mechanical loading.
J. Cell Biol.
117:
83-93,
1992
|
| 2. | Adams, D. H., and S. Show. Leukocyte-endothelial interactions and regulation of leukocyte migration. Lancet 343: 831-836, 1994 [Medline] . |
| 3. | Aitchson, W. A., and D. L. Brown. Duplication of the flagellar apparatus and cytoskeletal microtubule system in the algae Polytomella. Cell Mol. Cytoskeleton 6: 122-127, 1986. |
| 4. | Bass, D. A., J. W. Parce, L. R. Dechatelet, P. Szejida, M. C. Seeds, and M. Thomas. Flow cytometric studies of oxidative formation by neutrophils: A graded response to membrane stimulation. J. Immunol. 130: 1910-1917, 1983 [Abstract] . |
| 5. |
Doerschuk, C. M.,
N. Beyers,
H. O. Coxson,
B. Wiggs,
and
J. C. Hogg.
Comparison of neutrophil and capillary diameters and their relation to neutrophil sequestration in the lung.
J. Appl. Physiol.
74:
3040-3045,
1993
|
| 6. | Dorst, E. M., C. Selby, S. Lannan, G. D. O. Lowe, and W. MacNee. Changes in neutrophil deformability following in vitro smoke exposure: mechanism and protection. Am. J. Respir. Cell Mol. Biol. 6: 287-295, 1992. |
| 7. | Downey, G. P. Mechanism of leukocyte motility and chemotaxis. Curr. Opin. Immunol. 6: 113-124, 1994 [Medline] . |
| 8. |
Downey, G. P.,
D. E. Doherty,
B. Schwab III,
E. L. Elson,
P. M. Henson,
and
S. Worthen.
Retention of leukocytes in capillaries: role of cell size and deformability.
J. Appl. Physiol.
69:
1767-1778,
1990
|
| 9. |
Downey, G. P.,
E. L. Elson,
B. Schwab III,
S. C. Erzurum,
S. K. Young,
and
G. S. Worthen.
Biophysical properties and microfilament assembly in neutrophils: modulation by cyclic AMP.
J. Cell Biol.
114:
1179-1190,
1991
|
| 10. |
Downey, G. P.,
and
G. S. Worthen.
Neutrophil retention in model capillaries: deformability, geometry, and hydrodynamic forces.
J. Appl. Physiol.
65:
1861-1871,
1988
|
| 11. | Eppihimer, M. J., and H. H. Lipowsky. The mean filtration pressure of leukocyte suspensions and its relation to the passage of leukocytes through Nuclepore filters and capillary networks. Microcirculation 1: 237-250, 1994 [Medline] . |
| 12. | Erzurm, S. C., G. P. Downey, D. E. Doherty, B. Scwab III, E. L. Elson, and G. S. Worthen. Mechanism of lipopolysaccharide-induced neutrophil retention. Relative contributions of adhesive and cellular mechanical properties. J. Immmunol. 149: 154-162, 1991. [Abstract] |
| 13. | Erzurm, S. C., M. L. Kus, C. Bohse, E. L. Elson, and G. S. Worthen. Mechanical properties of HL60 cells: role of stimulation and differentiation in retention in capillary-sized pores. Am. J. Respir. Cell. Mol. Biol. 5: 230-241, 1991. |
| 14. |
Hogg, J. C.,
H. O. Coxson,
M.-L. Brumwell,
N. Beyers,
C. M. Doerschuk,
W. MacNee,
and
B. Wiggs.
Erythrocyte and polymorphonuclear cell transit time and concentration in human pulmonary capillaries.
J. Appl. Physiol.
77:
1795-1800,
1994
|
| 15. |
Hogg, J. C.,
T. McLean,
B. A. Martin,
and
B. Wiggs.
Erythrocyte transit and neutrophil concentration in the dog lung.
J. Appl. Physiol.
65:
1217-1225,
1988
|
| 16. | Holland, B. S., and M. D. Copenhaven. An improved sequential rejective Bonferroni test procedure. Biometrics 43: 417-423, 1987. |
| 17. | Hughes, B. J., J. C. Hollers, E. C. Torabi, and C. W. Smith. Recruitment of CD11b/CD18 to the neutrophil surface and adherence-dependent cell locomotion. J. Clin. Invest. 90: 1687-1696, 1992 . |
| 18. | Inano, H., D. English, and C. M. Doerschuk. Effect of zymosan-activated plasma on the deformability of rabbit polymorphonuclear leukocytes. J. Appl. Physiol. 74: 1370-1376, 1992. |
| 19. |
Kim, D. W.,
A. I. Gotlieb,
and
B. L. Langille.
In vivo modulation of endothelial F-actin microfilaments by experimental alterations in shear stress.
Arteriosclerosis
9:
439-445,
1989
|
| 20. | Lennie, S. E., G. D. O. Lowe, and J. C. Barbenel. Filterability of white blood cell subpopulations separated by an improved method. Clin. Hemorheol. 7: 811-816, 1987. |
| 21. |
Marcos, J.,
R. O. Hooper,
D. K. Gray,
B. R. Wiggs,
and
J. C. Hogg.
Effect of raised alveolar pressure on leukocyte retention in the human lung.
J. Appl. Physiol.
69:
214-221,
1990.
|
| 22. |
Molad, Y.,
K. A. Haines,
D. C. Anderson,
J. P. Buyon,
and
B. N. Cronstein.
Immunocomplexes stimulate different signaling events to chemoattractants in the neutrophil and regulate L-selectin and 2-integrin expression differently.
Biochem. J.
299:
881-887,
1994
.
|
| 23. | Nash, G. B., J. G. Jones, J. Mikita, and J. A. Dormandy. Methods and theory for analysis of flow of white cell subpopulations. Br. J. Haematol. 70: 165-170, 1988 [Medline] . |
| 24. | Omann, M., and J. M. Harter. Pertussis toxin effects on chemoattractant-induced response heterogeneity in human PMNs utilizing fluo-3 and flow cytometry. Cytometry 12: 252-259, 1991. [Medline] |
| 25. | Rao, K. M., J. Padmanabhan, and H. J. Cohen. Cytochalasins induce actin polymerization in human leukocytes. Cell Motil. Cytoskeleton 21: 58-64, 1992 [Medline] . |
| 26. | Redenbach, D. M., D. English, L. Verburgt, and J. C. Hogg. White blood cells maintain shape changes imposed by restrictions in the pulmonary capillary bed (Abstract). Am. J. Respir. Crit. Care Med. 151: A453, 1995. |
| 27. |
Selby, C.,
E. Dorst,
P. K. Wraith,
and
W. MacNee.
In vivo neutrophil sequestration within lungs of humans is determined by in vitro "filterability."
J. Appl. Physiol.
71:
1996-2003,
1991
|
| 28. |
Shirnsky, V. P.,
A. S. Antonov,
K. G. Birukov,
A. V. Sobolevsky,
Y. A. Romanov,
N. V. Kabaeva,
G. N. Antonova,
and
V. N. Smirnov.
Mechano-chemical control of human endothelium orientation and size.
J. Cell Biol.
109:
331-339,
1989.
|
| 29. |
Wiggs, B. R.,
D. English,
W. N. Quinlin,
N. A. Doyle,
and
J. C. Hogg.
The distribution of capillary pathway size and neutrophil deformability to neutrophil transit through rabbit lungs.
J. Appl. Physiol.
77:
463-470,
1994
|
| 30. |
Worthen, G. S.,
B. Schwab III,
E. L. Elson,
and
G. P. Downey.
Mechanics of stimulated neutrophils: cell stiffening induces retention in capillaries.
Science
245:
183-184,
1989
|
This article has been cited by other articles:
|
|
 |
|
  X.-P. Gao, X. Zhu, J. Fu, Q. Liu, R. S. Frey, and A. B. Malik Blockade of Class IA Phosphoinositide 3-Kinase in Neutrophils Prevents NADPH Oxidase Activation- and Adhesion-dependent Inflammation J. Biol. Chem., March 2, 2007; 282(9): 6116 - 6125. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. Yap and R. D. Kamm Cytoskeletal remodeling and cellular activation during deformation of neutrophils into narrow channels J Appl Physiol, December 1, 2005; 99(6): 2323 - 2330. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. Yap and R. D. Kamm Mechanical deformation of neutrophils into narrow channels induces pseudopod projection and changes in biomechanical properties J Appl Physiol, May 1, 2005; 98(5): 1930 - 1939. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. J. Feldhaus, A. S. Weyrich, G. A. Zimmerman, and T. M. McIntyre Ceramide Generation in Situ Alters Leukocyte Cytoskeletal Organization and beta 2-Integrin Function and Causes Complete Degranulation J. Biol. Chem., February 1, 2002; 277(6): 4285 - 4293. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. T. Suratt, S. K. Young, J. Lieber, J. A. Nick, P. M. Henson, and G. S. Worthen Neutrophil maturation and activation determine anatomic site of clearance from circulation Am J Physiol Lung Cell Mol Physiol, October 1, 2001; 281(4): L913 - L921. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. J. Anderson, W. T. Roswit, M. J. Holtzman, J. C. Hogg, and S. F. Van Eeden Effect of mechanical deformation of neutrophils on their CD18/ICAM-1-dependent adhesion J Appl Physiol, September 1, 2001; 91(3): 1084 - 1090. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Sato, J. C. Hogg, D. English, and S. F. van Eeden Endothelin-1 Changes Polymorphonuclear Leukocytes' Deformability and CD11b Expression and Promotes Their Retention in the Lung Am. J. Respir. Cell Mol. Biol., September 1, 2000; 23(3): 404 - 410. [Abstract] [Full Text]
|
|
|
|
 |
|
 S. F. van Eeden, Y. Kitagawa, Y. Sato, and J. C. Hogg Polymorphonuclear Leukocytes Released From the Bone Marrow and Acute Lung Injury Chest, July 1, 1999; 116(2007): 43S - 46S. [Full Text]
|
|
|
|
|
|
S. F. van Eeden, J. Granton, J. M. Hards, B. Moore, and J. C. Hogg Expression of the cell adhesion molecules on leukocytes that demarginate during acute maximal exercise J Appl Physiol, March 1, 1999; 86(3): 970 - 976. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Sato, S. F. Van Eeden, D. English, and J. C. Hogg Pulmonary sequestration of polymorphonuclear leukocytes released from bone marrow in bacteremic infection Am J Physiol Lung Cell Mol Physiol, August 1, 1998; 275(2): L255 - L261. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. P. Downey Invited Editorial on "Effect of mechanical deformation on structure and function of polymorphonuclear leukocytes" J Appl Physiol, May 1, 1997; 82(5): 1395 - 1396. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
