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Department of Medical Microbiology, Faculty of Health Sciences, Linköping University, SE-581 85 Linköping, Sweden
Correspondence: Vesa-Matti Loitto, Division of Medical Microbiology, Department of Health and Environment, Faculty of Health Sciences, Linköping University, SE-581 85 Linköping, Sweden. E-mail: veslo{at}ihm.liu.se
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: neutrophil • fMLF receptor • cell motility • F-actin • video microscopy
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The receptors for N-formyl peptide chemoattractants are structurally specific, seven-transmembrane-domain-containing surface molecules that mediate signal transduction through heterotrimeric G-proteins [4 5 6 ]. Once the receptor is occupied, neutrophils undergo an as-yet-incompletely defined sequence of intracellular signaling events and morphological alterations [reviewed in ref. 2 3, and 7–10]. One of these effects, cell activation, leads to cytoskeletal rearrangements, making N-formyl peptides potent inducers of neutrophil filamentous-actin (F-actin) formation [5 , 11 ]. The present concept of chemoattractant-receptor dynamics includes internalization of the ligand-receptor complex, cycling of receptors between the backs and fronts of cells, and shedding of receptors from the posterior area of each cell. All of these events have been shown to depend on interactions with or modification of the actin cytoskeleton, which thus is a key modulator of both cell polarity and cell motility.
To migrate, neutrophils might acquire a polarized morphology that helps them to transform intracellularly generated forces into net cell body translocation [12 ]. In a chemotactic gradient, moving neutrophils regularly assume such a polarized morphology, with an anterior lamellipodium extended in the direction of movement, an elongated cell body parallel to the axis of lamellar protrusion, and a posterior knob-like uropodium [8 ]. Motile neutrophils tend to preserve the same leading edge and often seem to prefer to turn toward an advancing new chemotactic gradient rather than form a new front [13 , 14 ].
It has been suggested that membrane components, and thus functional signals, segregate into distinct domains whenever cell shape and cytoskeletal architecture become asymmetric [10 , 15 , 16 ]. Neutrophils are capable of sensing differences in chemoattractant concentrations over the length of the cell of as little as 1%, i.e., >10–20 µm [13 ]. How cells are able to compare and amplify such minute differences in concentrations of extracellular attractants has, however, not been fully clarified. Also, it is not known how the receptor-ligand interaction is translated into directional movement [17 ].
Two principal models have been proposed to explain the mechanisms of directional sensing. The first mechanism suggests a temporal regulation by which a cell senses the concentration of a chemoattractant, moves toward it, and compares the encountered level with the earlier one [18 ]. This model assumes that there are pilot sensors, in the form of numerous filopodia, that extend in all directions from the cell. Indeed, it has been shown that inhibition of the Rho family guanosine triphosphatase Cdc42, which is required for filopodium formation, prevents recognition of chemoattractant gradients in macrophages [19 ]. The second proposed mechanism assumes that the cell uses spatial sensing to compare the concentrations of a chemoattractant at two or more locations on its surface and then directs its movement toward increasing levels of chemotactic substance [18 ]. The latter model requires that cells be able to intrinsically detect differences in receptor occupancy between their two ends, even when there is no temporal change in receptor occupancy [20 ]. Thus, if more receptors are occupied on the leading front than on the trailing uropod [21 ], this must somehow be sensed by or must indirectly influence the motility machinery.
Directional sensing of chemoattractants has been explained as both an intracellular and a membrane receptor asymmetry-associated mechanism. The intracellular mechanisms include clustering of specific protein complexes, e.g., Akt/PKB [22 ], cofilin [23 ], coronin [24 ], and cyclase-associated proteins [25 ], to the inner face of the plasma membrane. These clusters may spatially regulate the activity of the signal transduction cascade [3 ]. In polarized neutrophils, asymmetric distributions of membrane receptors, such as those for the Fc portion of immunoglobulin G (IgG) [26 ], concanavalin A [27 ], succinyl-conA [15 ], and N-formyl peptides [28 , 29 ], have been observed. These asymmetries have been suggested to be necessary for cells to respond to a shallow gradient of chemoattractant.
Endogenous chemoattractant receptors tagged with green fluorescent protein (GFP) and expressed in PLB-985 cells [30 ] and in the amoeba Dictyostelium discoideum [20 , 31 ] display a more-or-less uniform distribution on the plasma membrane. It has been suggested that cells such as D. discoideum, which need to respond very quickly to changes in chemoattractant, are helped by a uniform distribution of receptors on the cell surface [17 ].
Here we report that N-formyl peptide receptors labeled with fluorescently tagged N-formyl peptide receptor ligands [1 ], i.e., antagonistic fluoresceinated tert-butyl-oxycarbonyl-Phe-(D)-Leu-Phe-(D)-Leu-Phe-OH (Boc-FLFLF) [32 ], agonistic formyl-Nle-Leu-Phe-Nle-Tyr-Lys (fnLLFnLYK) [33 ], and monoclonal antibodies, were distributed homogeneously at the plasma membrane of unstimulated neutrophils. We suggest that after cell activation occurs, the first binding site, or possibly the site where most receptors are engaged, determines the initial direction of migration. Chemoattractant receptors are then actively accumulated in this cell region by recruitment of receptors from other membrane domains and by up-regulation from intracellular stores. On morphologically polarized neutrophils, N-formyl peptide receptors distribute nonuniformly and concentrate in motile regions at the cell periphery. Thus, our results can be explained by a spatial mechanism for directional sensing. The results also suggest that asymmetric receptor activation is essential for initiation of motility and that a chemoattractant gradient further promotes sustained migration in a certain direction. In fixed neutrophils, N-formyl peptide receptors labeled with monoclonal antibodies colocalize with F-actin-rich domains at the leading edge of polarized cells, whereas receptors labeled with fluorescent peptides lag slightly behind in these locations. We suggest that the observed difference between fnLLFnLYK-peptide- and antibody-labeled receptors reflects different populations of N-formyl peptide receptors, e.g., in distinct stages of receptor activation. Other investigators have previously proposed the basic ideas presented here, but this is the first time such receptor pools have been visualized.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Isolation of neutrophils
Human neutrophils were isolated from venous blood, obtained from
healthy adult volunteers, essentially as described by Böyum
[34
]. Briefly, heparinized whole blood was allowed to
equilibrate to room temperature, separated on PolymorphprepTM dextran
gradients (Nycomed Pharma AS, Oslo, Norway) in 15-mL test tubes, and
centrifuged at 450 g for 35 min at room temperature. The
band containing granulocytes was transferred to a 50-mL test tube and
washed in phosphate-buffered saline (PBS; pH 7.3). Remaining
erythrocytes were lysed by a 30-s hypotonic treatment with deionized
water. The granulocytes were washed twice with calcium-free
Krebs-Ringer phosphate buffer supplemented with 10 mM glucose and 1.5
mM Mg2+ (KRG; pH 7.3), suspended in KRG (107
cells/mL), and then stored on melting ice until used. The resulting
cell suspension contained at least 95% neutrophils, with the remaining
cells being predominantly eosinophils.
Treatment with fnLLFnLYK and Boc-FLFLF
Isolated neutrophils (5 x 104 cells) were
allowed to sediment and adhere to HSA-coated or uncoated glass
coverslip-bottomed wells for 10 min. Fluorescent N-formyl
peptide receptor agonist, i.e., fnLLFnLYK (10 nM), or antagonist,
i.e., Boc-FLFLF (50 nM), was then added to the experimental well.
Images were captured 1–2 min after addition of the fluorescent ligand.
To reduce receptor-ligand internalization, the experiments with
fnLLFnLYK were performed at temperatures below 14°C
[35
, 36
]. Experiments with Boc-FLFLF were
done at room temperature because the probe has been shown to
internalize very little upon binding to the receptor
[1
]. The buffer used in all experiments was KRG
supplemented with 1 mM Ca2+ [calcium-containing medium
(CCM)].
fnLLFnLYK is a more potent activator of the cell than is fMLF [1 ]. However, to obtain sufficient labeling for visualization, fnLLFnLYK was used at slightly higher concentrations than those required for cell migration. We used a concentration of 10 nM, which initiates a neutrophil respiratory burst at 37°C [1 ]. According to Davis et al. [15 ], the sequence of events in cell morphology alterations should be the same for concentrations between 10 and 100 nM fMLF, independent of whether the stimulus is added as a gradient or as a homogeneous mixture.
To examine the extent of internalization, binding of fnLLFnLYK to live cells was studied using a confocal laser-scanning microscope. Adherent neutrophils were exposed to 50 nM fluoresceinated peptide and then monitored from initial binding to spreading and motility. The confocal laser-scanning microscopy (CLSM) experiments were performed at 16 and 37°C.
Labeling of the cell membrane
Neutrophils were first isolated as described above.
DiIC16(3) (a 1-mg/mL solution in DMSO) was added to the
cell suspension (in KRG) to a final concentration of 5.5 µg/mL. After
gentle mixing, the cell suspension was incubated for 20 min at 37°C.
The cells were washed three times with KRG and kept, protected from
light, on melting ice until used. Samples (5 x 104
cells) were withdrawn from the cell suspension and transferred to
experimental wells in the bottoms of which had been placed coverslips.
The neutrophils were allowed to adhere to the glass surface for 5–10
min at room temperature in CCM. Images were captured at room
temperature, both directly after adherence and after 15 min of
incubation.
The neutrophil membrane also was visualized using DiD (2.5 mg/mL, in DMSO). Labeling and subsequent washing procedures were the same as those used for DiIC16(3) except that the final concentration of DiD during labeling was 2.5 µg/mL. Samples were withdrawn, and the cells were allowed to adhere to HSA-coated coverslips at temperatures <14°C. Double labeling of membrane and N-formyl peptide receptors was attained by adding fnLLFnLYK (25 or 50 nM solution) to the experimental well containing adherent DiD-labeled neutrophils.
F-actin and fnLLFnLYK
Isolated neutrophils (5 x 104 cells/coverslip)
in CCM were allowed to adhere to both HSA-coated and uncoated glass
coverslips for 5 min at room temperature and then for 10 min at 37°C.
To induce a polarized cell morphology, fMLF (10 nM) was added. After an
additional incubation for 2–3 min, the medium was replaced with fresh
4% PFA in CCM (pH 7.3). Unstimulated cells were treated similarly,
although without exposure to fMLF. The neutrophils were fixed for 20
min at 37°C and washed three times with PBS (pH 7.3) supplemented
with 1% BSA (PBS-BSA). The cells were then incubated with
fnLLFnLYK (50 nM) for 60 min at 37°C. After being washed, the
cells were permeabilized with freshly prepared 0.1% Triton X-100 for 3
min at room temperature. The F-actin network was labeled by incubation
for 20 min with AlexaTM 594 phalloidin in PBS-BSA, as stipulated by the
manufacturer. The coverslips were mounted in ProLongTM mounting medium.
Monoclonal fMLF receptor antibodies and F-actin
Cell adhesion, preincubation with and without 10 nM fMLF, and
fixation were performed as described above. After undergoing fixation,
the cells were rinsed with PBS-BSA and incubated for 60 min at 37°C
with 10% goat serum in PBS-BSA. The cells were then incubated for 60
min at 37°C with 10-µg/mL of monoclonal mouse anti-human antibody
directed against the fMLF receptor. After being thoroughly rinsed with
PBS-BSA, the cells were labeled with secondary FITC- or AlexaTM
594-conjugated goat anti-mouse antibody for 30 min at 37°C. After
incubation with the secondary antibody, the cells were washed with
PBS-BSA. The cells were then permeabilized, labeled with AlexaTM 594-
or Bodipy Fl-conjugated phalloidin, and mounted, as described above.
Monoclonal fMLF receptor antibodies and fnLLFnLYK
Cells also were double labeled with the monoclonal fMLF receptor
antibody and the fluoresceinated N-formyl peptide
fnLLFnLYK. Cell adhesion, preincubation with and without 10 nM
fMLF, and PFA fixation were performed as described above. The fMLF
receptors were labeled with antibodies as described above and then
incubated with 50 nM fnLLFnLYK for 60 min at 37°C. After being
washed, the preparations were mounted in ProLongTM.
Controls for the fMLF receptor antibody were incubated with a monoclonal mouse antibody to human epithelial membrane antigen rather than the primary antibody. In some experiments, the 10% goat serum solution was replaced with a 100-µg/mL solution of purified goat IgG in PBS-BSA. We also examined the specificity of the antibody by mixing the antibody with increasing concentrations of fMLF prior to labeling.
Fluorescence microscopy
A Zeiss (Oberkochen, Germany) Axiovert 135M microscope, equipped
with a 100-W type HBO mercury arc lamp and a Zeiss 100x oil immersion
neofluar objective (1.3 numerical aperture) for epifluorescence
procedures, was used to study the fluorescent samples. Images of the
fnLLFnLYK-, Boc-FLFLF-, and FITC-conjugated antibody distributions
were obtained by using the FITC channel with a 470-nm ± 20-nm
band-pass filter for excitation and a 540-nm ± 25-nm band-pass
filter for emission. The DiIC16(3)-, DiD-, and AlexaTM 594
phalloidin-labeled cells were excited with a 546-nm ± 12-nm
band-pass filter, and the emission was selected with a 590-nm long-pass
filter, i.e., a Texas Red channel. All filters were obtained from
Zeiss. When possible, a 95% neutral-density filter was inserted
in front of the excitation beam to reduce photobleaching and to
minimize heating when working with the cooled stage drive.
To further magnify images, an intermediate magnification factor of 1.6 in the microscope was utilized occasionally. Images were captured using a water-cooled TE4 Astromed 4200 slow-scan charge-coupled device camera (LSR Ltd., Cambridge, UK) controlled by PixCelTM software (LSR Ltd.). The figures presented here were obtained using 20–400-ms exposures.
Both instant images and time-lapse sequences, with 20–30 s between successive images, were recorded. All images were stored on an IBM-compatible personal computer. The extent of bleedover from the Texas Red channel to the FITC channel, and vice versa, was examined for all fluorophores used. Assembly of color-merged images of double-labeled specimens was accomplished using PhotoShop 5.5 software (Adobe Systems Inc., Tokyo, Japan) on a Power Macintosh computer.
CLSM
To characterize the binding and possible internalization of
fnLLFnLYK, CLSM was performed at 16 and 37°C. This was done on a
Sarastro 2000 instrument (Molecular Dynamics, Sunnyvale, CA) mounted on
a Nikon Optiphot microscope equipped with a 60x oil-immersion
objective (1.4 NA). An Indigo R4000 workstation (Silicon Graphics,
Mountain View, CA) with ImageSpace software (Molecular Dynamics) was
attached to handle the CLSM. CLSM was optimized for the green region of
the spectrum by using an illumination setup comprising a 488-nm
excitation argon laser wavelength, a 510-nm DRLP beam splitter, and a
510-nm EFLP long-pass emission filter. Cell fluorescence was scanned
within 0.5 s of initial contact with the fluorescent ligand
(spherical, nonactive cells), which was added under the microscope.
This resulted in more-or-less immediate activation of spreading and
motility. The delay between images was around 10 s.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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F-actin and fnLLFnLYK
N-formyl peptide receptors have been shown to associate
with actin-rich plasma membrane microdomains [37
]. We
therefore double labeled cells with fnLLFnLYK and
fluorophore-conjugated phallotoxin. fnLLFnLYK-labeled receptors
distributed uniformly over the surface of unstimulated cells
(Fig. 4
inserts), while morphologically polarized cells exhibited a
nonuniform distribution over the plasma membrane, with accumulations
extending from the posterior region toward the lamellipodium of the
cell (Fig. 4) . The F-actin network exhibited a distinct localization to
the cell edges, especially to the leading front of polarized
neutrophils. When the images were merged, no major colocalization of
the fnLLFnLYK-labeled receptors and F-actin was observed (Fig. 4)
.
As was also observed in the experiments using the membrane probe DiD,
the fnLLFnLYK-labeled receptors seemed to lag slightly behind
the front of the lamellipodia of the cells. Although capping of the
receptor-ligand complex at the uropodium was pronounced in living cells
studied over longer time periods (Fig. 2c
and 2d)
, no significantly
higher fluorescence intensities were observed at the rear of the cell
in fixed preparations.
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Monoclonal fMLF receptor antibodies and fnLLFnLYK
Double labeling of chemoattractant receptors with monoclonal
antibodies and fluoresceinated peptides supports the observations
described in the previous section. Receptors labeled with antibodies
were predominantly found in motile regions and at the cell periphery,
whereas fnLLFnLYK-labeled receptors lagged behind. This was
especially clear in radially spreading cells, in which the
antibody-labeled receptors formed an outer ring around the receptors
labeled with fnLLFnLYK (Fig. 7
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Servant et al. [30 ] recently observed a uniform distribution of GFP-tagged chemoattractant receptors over the neutrophil membrane. By contrast, our results indicate that, after cell activation, receptors redistribute to specific locations and domains of the plasma membrane. Furthermore, by two distinct labeling methods, N-formyl peptide receptors were observed to exist in two populations that separated into distinct domains of the membrane. This is in accordance with observations made by Walter and Marasco [39 ], who observed that receptors displaying high- and low-affinity characteristics distributed asymmetrically over the cell surface. Because of ligand binding, differences in anchorage to cytoplasmic components, or effects on various membrane constituents in close proximity to the receptors, the two populations could reflect differing states of activation. The division into two distinct populations would probably not be distinguished using GFP tagging.
N-formyl peptide receptors distribute nonuniformly
Distribution of chemoattractant receptors on neutrophil plasma
membranes has been intensively studied using different approaches and
visualization techniques [1
, 15
,
26
27
28
29
30
, 40
]. In our experiments we used
fluoresceinated N-formyl peptides—i.e., Boc-FLFLF (an
antagonist) and fnLLFnLYK (an agonist)—and a novel monoclonal
antibody directed against the human fMLF receptor, together with
sensitive fluorescence video microscopy. When adherent, live
neutrophils were exposed to the agonist form of fluoresceinated
N-formyl peptides, the ligand initially distributed
uniformly (i.e., randomly) on the cell membrane (Fig. 2A)
. However,
after cell activation, receptor enrichment at motile regions of the
cell was an early event during morphological polarization (Fig. 1
; Fig. 2B
and 2C
). Although a uniform distribution of receptors is consistent
with the hypothesis that motile cells are equally chemotactically
responsive at all points on their perimeter [3
], we
observed chemoattractant receptor accumulation in motile regions of the
cell upon stimulation.
Insertion of receptors at the motile region
Fluoresceinated N-formyl ligands bound preferentially
to receptors lagging slightly behind the outermost lamellipodia of the
cell. This observation was reached by double labeling cells with
membrane dyes, or fluorophore-conjugated phallotoxin (Fig. 4)
, in
combination with receptor antibodies or with fnLLFnLYK (Fig. 7)
.
Apparently, the extremely thin leading edge constitutes an effective
barrier for insertion of vesicles with "fresh" receptors into the
very front of the cell. The lamellipodium of a cell is in its simplest
model a dense F-actin meshwork between two lipid bilayers. Such a
compact organization of the extreme leading edge would thereby exclude
an active fusion of receptor-containing transport vesicles having a
minimum radius of perhaps 20–50 nm. Therefore, fusion possibly occurs
at locations slightly behind the leading edge. In this region, the
cytoplasmic motile machinery is reshaped and the actin nucleation
activity of a motile cell is especially pronounced [41
].
Indeed, studies of protein insertion at the electron-microscopic level
indicate that fusion actually occurs nearer the forward boundary of the
endoplasm than to the leading edge of the lamellipodium
[42
]. Furthermore, in the region of the leading edge
itself, there was no evidence of vesicle fusion [42
].
After being disengaged from the signaling domain, the dorsally located
receptor aggregates move rearward [43
], leading to an
accumulation at the uropod, where they eventually are endocytosed
[15
]. Although the life cycle of chemoattractant
receptors is largely unknown, the receptors could then follow a
polarized endocytic cycle [44
] similar to that described
for 
vß3-integrins [45
].
Differences between fluoresceinated fnLLFnLYK and antibodies
The apparent receptor distribution in fixed neutrophils differed
considerably depending on whether labeling was done with
fluoresceinated N-formyl peptides or antibodies. When
fnLLFnLYK was used to localize the N-formyl
peptide receptors, fluorescence was distributed primarily from the
lamellipodia toward the rear of the cell. At the leading edge of the
lamellipodia of adherent cells, the fluorescence decreased and did not
extend all the way out to the F-actin-rich regions. Cluster formation
was reduced compared with the level observed for labeling with
antibodies, whereas capping of the peptide-receptor complex at the
uropodium was pronounced in living cells studied over longer time
periods (Fig. 2c
and 2d)
. Davis and colleagues [15
]
proposed that pinocytic activity is restricted to the tail region of
the cell, implying that part of the fnLLFnLYK-derived fluorescence
at the uropodium in living neutrophils could be of intracellular
origin. However, when adherent neutrophils were studied at 16°C by
CLSM, the fluoresceinated peptide remained membrane associated
throughout the experiment (Fig. 2)
.
Antibody-labeled fMLF receptors, on the other hand, clearly colocalized with F-actin-rich areas at the cell edges (Fig. 6) . This observation agrees well with earlier studies showing that chemoattractant receptors segregate into actin-rich domains in the neutrophil [37 ]. These observations support the interaction of the fMLF receptor with detergent-insoluble F-actin networks described by Jesaitis and co-workers [5 ]. There is compelling biochemical evidence that, upon activation, receptors redistribute from membrane domains rich in G-proteins to areas depleted of G-proteins but rich in cytoskeletal elements, preferably F-actin [5 ].
The differences between fnLLFnLYK- and antibody-labeled chemoattractant receptors suggest that neutrophils have two populations of N-formyl peptide receptors. The distinction of two populations may have several origins. First, it could reflect differing dependence on cell topography. Albeit an unlikely scenario, extensive folding of plasma membrane on the main cell body and posterior regions might sterically hinder antibodies from binding to the fMLF receptors. In contrast, the small hexapeptide easily enters the tight folds, resulting in labeling of the chemoattractant receptors. Alternatively, the receptor density at the leading edge may be so low that it can barely be visualized by direct labeling with the weakly fluorescent N-formyl peptide, whereas the antibody binding is amplified with the fluorophore-conjugated secondary antibody. Furthermore, it had been reported that there are at least two different populations of fMLF receptors, of high and low affinity, asymmetrically distributed over the cell surface [39 ]. The high-affinity-type receptors were found predominantly on the anterior half of the cell. The division into high- and low-affinity receptors is assumed to represent distinct conformational stages at which association of the receptors with intracellular complexes, such as with F-actin, or specific plasma membrane lipid rafts alters the ligand binding site or overall conformation of the receptor. Third, since 10 nM fMLF was used in the prestimulatory step of the experiment, a large fraction of the receptors may still be associated with the chemoattractant [6 ]. This could hinder in vivo binding of the fluoresceinated hexapeptide [compare ref 38 ]. Only at the insertion site of fresh receptors could the fluorescent ligand compete with the standard fMLF. Furthermore, it is likely that ligand-activated chemoattractant receptors have altered molecular conformations. If the fixation process preserves this receptor conformation, although the actual ligand is released, this would support the observed difference between the monoclonal antibodies and fluoresceinated peptides.
Directional sensing by neutrophils
It has been reported that neutrophil pseudopodia rarely form from
the tail regions. Even when a gradient is reversed, the cells prefer to
reorient themselves by extending pseudopodia from the front and making
a U-turn [14
, 46
]. This asymmetry in
responsiveness indicates that motile neutrophils have a nonuniform
distribution of chemoattractant receptors on their plasma membrane. The
proposal is consistent with our results. We observed a nonuniform
receptor distribution on morphologically polarized neutrophils when
using either fluoresceinated N-formyl peptides or antibodies
directed against the fMLF receptor. The chemoattractant receptors
distributed heterogeneously over the neutrophil plasma membrane and
were preferentially localized to motile regions of the cell.
Furthermore, the fluorescent ligands and the receptor antibodies seem
to distinguish two different populations with distinct distributions
(Fig. 7) , which helps the neutrophils control and maintain their
directional sensitivity to chemoattractant gradients during cell
migration.
Assuming that the cells sense a surface-bound gradient of chemoattractant, our results are in line with a spatial mechanism for directional sensing [18 ]; i.e., the cell senses and responds to chemoattractant concentrations at locations with the highest ligand density. The site with the largest fraction of occupied receptors will generate the strongest signal(s), which initiates, for instance, remodeling of the cytoskeleton and insertion of fresh receptors. Thus, the direction of subsequent chemotaxis depends on recruitment of receptors from other sites of the membrane and on up-regulation of receptors from intracellular stores. Assuming that the insertion of fresh receptors into the region of initial stimulation does not compensate for desensitization after the initial stimulation, the cells could spatially respond and turn either left or right, thus moving in a zigzag fashion along the gradient. Should there be no gradient, the cells would display stimulated random locomotion. Thus, spatial sensing and the rates of receptor desensitization and up-regulation determine the temporal behavior of the cells. If there is no substratum-associated ligand, but rather an instantaneous flux of ligand occurs, spatial inhomogeneity among randomly distributed receptors will probably translate into stimulated, random locomotion. So, to conclude, we think that temporal variation in the receptor distribution over the cell membrane allows for spatial sensing, which is reinforced when cells start to move along a chemoattractant gradient.
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ACKNOWLEDGEMENTS |
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This research was supported by the Foundation for Strategic Research (graduate student position for Vesa-Matti Loitto via Forum Scientum), the FöreningsSparbanken Linköping Graduate Student Program, the Swedish Research Council for Engineering Sciences, the Swedish Medical Research Council (project no. 6251), the King Gustaf V 80-Year Foundation, and the Professor Nanna Svartz Foundation.
Received August 3, 2000; revised December 4, 2000; accepted December 20, 2000.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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