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T-Cell Motility: does motility require direction specific actin polymerization?

T-Cell Motility: does motility require direction specific actin polymerization?


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T-cells have been shown to migrate inside concentration gradients - both in the direction of the source or away. Even under shallow gradients, t-cells move. I argue that, to be able to move in a directed manner, a cell must be able to have a differential response of actin polymerization and philipodia. But I find it hard to believe that it is a differential response unless the sensor resolution not only communicates centrally, but is also extremely sensitive. How does this work?


It's understood at least that leukocyte motility is dependent on cytoskeleton remodeling and actin dynamics. In the blood, as the T cells tumble past the activated endothelium of an affected site, they're grabbed by selectins and integrins expressed on the endothelial surface. These are often illustrated as the adhesion and rolling phases of T cell migration.

Chemokines, which normally form a gradient for T cell chemotaxis, induce changes in the cellular morphology that promote migration into the surround tissues. The cells polarize into a force-generating lamellopodium with a leading edge in the front, and a uropod in the back. It appears as though actin polymerizes in the lamellopodium and juts the cell out toward the direction of the signal, and the uropod contracts in a myosin-dependent manner, releasing molecules bound there, thereby allowing the cell to tug in the direction of the lamellopodia (1, 2).

The mechanism has in fact been characterized by leading-edge redistribution of chemokine receptors such as CCR5 and CXCR4, which may have to do with actin/myosin dynamics and lipid rafts. This makes the receptor distribution asymmetric as well, and so a weaker response to the chemoattractant in the back end of the cell. The best way I could describe it is using the formation of the cSMAC in T cell receptor biology (Figure 1) (3). The SMAC is basically a synapse formed during T cell activation where receptors are concentrated to a particular area where the T cell is meeting the APC.

Figure 1. TCR-mediated cytoskeletal remodeling

The way the authors describe it is the downstream signals from the TCR localize actin pathway molecules to the site of signaling, and causes cytoskeletal remodeling at that particular site.

Studies in neutrophils have shown that polarization leads to a frontness and backness to the cell, regulated by different proteins and inhibit the formation of one another. This is a good paper to read through on that matter, though it's a tougher read in my opinion: https://www.researchgate.net/publication/8957894_Cytoskeletal_remodeling_in_leukocyte_function


Confinement-optimized three-dimensional T cell amoeboid motility is modulated via myosin IIA–regulated adhesions

During trafficking through tissues, T cells fine-tune their motility to balance the extent and duration of cell-surface contacts versus the need to traverse an entire organ. Here we show that in vivo, myosin IIA–deficient T cells had a triad of defects, including overadherence to high-endothelial venules, less interstitial migration and inefficient completion of recirculation through lymph nodes. Spatiotemporal analysis of three-dimensional motility in microchannels showed that the degree of confinement and myosin IIA function, rather than integrin adhesion (as proposed by the haptokinetic model), optimized motility rate. This motility occurred via a myosin IIA–dependent rapid 'walking' mode with multiple small and simultaneous adhesions to the substrate, which prevented spurious and prolonged adhesions. Adhesion discrimination provided by myosin IIA is thus necessary for the optimization of motility through complex tissues.


REVIEW article

Alison Gaylo † , Dillon C. Schrock † , Ninoshka R. J. Fernandes and Deborah J. Fowell*
  • Department of Microbiology and Immunology, David H. Smith Center for Vaccine Biology and Immunology, Aab Institute of Biomedical Sciences, University of Rochester, Rochester, NY, USA

Effector T cells exit the inflamed vasculature into an environment shaped by tissue-specific structural configurations and inflammation-imposed extrinsic modifications. Once within interstitial spaces of non-lymphoid tissues, T cells migrate in an apparent random, non-directional, fashion. Efficient T cell scanning of the tissue environment is essential for successful location of infected target cells or encounter with antigen-presenting cells that activate the T cell’s antimicrobial effector functions. The mechanisms of interstitial T cell motility and the environmental cues that may promote or hinder efficient tissue scanning are poorly understood. The extracellular matrix (ECM) appears to play an important scaffolding role in guidance of T cell migration and likely provides a platform for the display of chemotactic factors that may help to direct the positioning of T cells. Here, we discuss how intravital imaging has provided insight into the motility patterns and cellular machinery that facilitates T cell interstitial migration and the critical environmental factors that may optimize the efficiency of effector T cell scanning of the inflamed tissue. Specifically, we highlight the local micro-positioning cues T cells encounter as they migrate within inflamed tissues, from surrounding ECM and signaling molecules, as well as a requirement for appropriate long-range macro-positioning within distinct tissue compartments or at discrete foci of infection or tissue damage. The central nervous system (CNS) responds to injury and infection by extensively remodeling the ECM and with the de novo generation of a fibroblastic reticular network that likely influences T cell motility. We examine how inflammation-induced changes to the CNS landscape may regulate T cell tissue exploration and modulate function.


MATERIALS AND METHODS

Parasite Culture

T. gondii tachyzoites of the RH strain were maintained by serial 2-d passage in HFF cell monolayers as previously described (Morisaki et al., 1995). RH-strain parasites expressing β-galactosidase (2F) were used in biochemical assays (Dobrowolski et al., 1997). RH-strain parasites that expressed a fusion of the last 82 amino acid residues of TgMyoA with green fluorescent protein (GFP TgM-Atail) were provided by Dr. Dominique Soldati (Hettman et al., 2000). Parasites were isolated soon after host cell lysis, passed through a 3-μm filter, and washed once with Hanks' balanced salt solution (Life Technologies, Gaithersburg, MD) containing 0.001 M EGTA, 0.01 M HEPES (HHE).

YFP-ACT1–Strain Construction

A plasmid expressing yellow fluorescent protein (YFP)-actin as a fusion protein was designed to have a 10-alanine linker between YFP and actin (Doyle and Botstei, 1996). The ACT1 gene was amplified from a T. gondii cDNA plasmid by use of the PCR primers 5′-ACCAATGCATGCTGCCGCAGCGGCGGCAGCCGCTGCAGCA-ATGGCGGATGAAGAAGTGCAAGC-3′ (ACT1F primer, containing coding sequence for 10 alanines [italicized]) and 5′-CATGCTTAATTAATTAGAAGCACTTGCGGTGGACG-3′ (ACT1R primer) and digested with NsiI and PacI. YFP was amplified by PCR using the primers 5′-ACGTCAGATCTAAAATGGTGAGCAAGGGC- GAGG AGC-3′ (YFPF primer) and 5′-CAGTATGCATCTTGTACAGCTCGTCCATGCCG-3′ (YFPR primer) and digested with BglII and NsiI. This construct, referred to as pTUB/YFP-ACT1/SAGCAT, contained the YFP-ACT1 fusion flanked by 5′-α-tubulin (Nagel and Boothroyd, 1988) and 3′-DHFR (Roos, 1993) regulatory sequences, followed by a chloramphenicol resistance cassette including the CAT gene flanked bySAG1 regulatory sequences (Striepen et al., 1998). Freshly isolated parasites (n = 10 7 ) were transfected with 75 μg plasmid DNA and inoculated into host cells as previously described (Roos et al., 1994). To produce stable transgenics, chloramphenicol was added 24 h later to a final concentration of 6 μg/ml, and drug-resistant clones were isolated by limiting dilution after several rounds of selection. Transformants were further subcloned in the presence of drug clone YA2 was selected for further use. Rabbit polyclonal anti-actin sera (Dobrowolski et al., 1997) and mouse monoclonal antibody (mAb) 3E6 against GFP (Clontech, Cambridge, UK) were used to detect the YFP-actin protein in Western blots. Mouse mAb Tg17–43 against GRA1 (Cesbron-Delauw et al., 1989) was used as a loading control.

Southern Blotting

Genomic DNAs from HFF cells or T. gondii(wild-type or YFP-ACT1 transgenics) were digested with restriction endonucleases at 37°C overnight, electrophoresed in 0.8% agarose gels, and transferred to nylon membranes by alkaline capillary transfer (Maniatis et al., 1982). Blots were probed overnight with a 32 P-labeled probe encompassing a partial actin sequence of 241 nucleotides generated from the plasmid pTUB/YFP-ACT1/SAGCAT by PCR using the primers 5′-GACGACATGGAGAAAATCTGGCATCACACC-3′ (probeF primer)and 5′-CATGCTTAATTAATTAGAAGCACTTGCGGTGGACG-3′ (probeR primer). Blots were hybridized at 42°C overnight in 4× SSPE, 0.8% SDS, 40% formamide, 4× Denhardt's solution, and 100 μg/ml calf thymus DNA and washed at a final stringency of 68°C in 0.1× SSPE, 0.1% SDS. Filters were exposed to Kodak XAR film at −70°C for autoradiography.

Immunofluorescence Labeling

Parasites were resuspended in HHE and treated with 2 μM JAS (Molecular Probes, Eugene, OR) that had been dissolved in dimethylsulfoxide (DMSO) or 1% DMSO alone as a control for 10 min at room temperature, then allowed to adhere to poly- l -lysine–coated coverslips for 10 min in a hydrated chamber. Coverslips were rinsed, fixed for 5 min with 100% methanol at −20°C, and blocked with 20% FBS (Hyclone, Logan, UT) in PBS for 30 min. After labeling with rabbit polyclonal anti-ACT1 sera (Dobrowolski et al., 1997) at 1:250 in PBS, rabbit anti-α-tubulin sera (Morrissette and Sibley, 2002b) at 1:1000, mouse monoclonal anti-SAG1 antibody (DG52) (Morisaki et al., 1995) at 1:250, and/or mouse monoclonal anti-IMC1 (inner membrane complex) (45.15) antibody at 1:1000 in PBS (Mann and Beckers, 2001), parasites were labeled with goat anti-mouse or anti-rabbit Alexa 488 and/or 594 (1:500 in PBS, Molecular Probes). Samples were mounted in Vectashield + DAPI (Life Technologies) and examined with a Zeiss Axioskop or a Zeiss 510 confocal microscope (Zeiss, Oberkochen, Germany). Wide-field fluorescence images were collected with a Zeiss Axiocam and Zeiss Axiovision software version 2.0.5, then processed and merged with Adobe Photoshop (Adobe Systems, Mountain View, CA). Confocal micrographs were taken with a 63× oil plan-apochromat objective lens (numerical aperture, 1.4 Zeiss) and He-Ne and Kr-Ar lasers. Images were imported into Zeiss Imagebrowser software and then processed in Adobe Photoshop.

Gliding Motility Assay

Coverslips were coated in 50% FBS in PBS for 1 h at 37°C and rinsed in PBS. Freshly harvested tachyzoites were resuspended in HHE, treated with varying concentrations of JAS or 1% DMSO for 10 min, added to precoated coverslips, and incubated at 37°C for 15 min. Slides were fixed in −20°C methanol and blocked with 20% FBS in PBS. The presence of the surface membrane protein SAG1 in trails was detected with mAb DG52 (1:250) conjugated directly to Oregon green (Molecular Probes). Coverslips were rinsed, mounted in Vectashield (Vector Laboratories, Burlingame, CA) plus DAPI, and examined with wide-field fluorescence microscopy. Average trail length in parasite body lengths (7 μm) was determined from five randomly selected 63× fields that contained ∼50 parasites per field in each of three separate experiments (mean ± SEM).

Invasion Assay

HFF cells were plated on coverslips 24 h before experiments. Freshly harvested tachyzoites were resuspended in invasion media (DMEM, Life Technologies), 3% FBS, 10 mM HEPES) and used to challenge HFF monolayers for 15 min at 37°C. Monolayers were rinsed in HHE, fixed in 3% formaldehyde in PBS for 15 min at room temperature, blocked with 20% FBS in PBS, and stained with mAb DG52 (1:250) conjugated to Texas Red (Molecular Probes). Samples were then rinsed, permeabilized with 0.1% saponin for 15 min at room temperature, and stained with DG52 conjugated to Oregon green (1:250). Coverslips were rinsed, mounted in Vectashield, and examined with wide-field fluorescence microscopy. The percentage of intracellular parasites (green-stained but not red-stained) was determined from five randomly selected 63× fields that contained ∼50 parasites per field in each of three separate experiments (mean ± SEM).

Quantification of Filamentous Actin by Western Blotting

Freshly harvested 2F-strain parasites (RH transfected with β-galactosidase) were placed in actin stabilization buffer (60 mM PIPES, 25 mM HEPES, 10 mM EDTA, 2 mM MgCl2, 125 mM KCl) and treated with varying concentrations of JAS or 1% DMSO for 10 min at 37°C. Glycerol was added to a final concentration of 10%, 100× protease inhibitor cocktail (1 mg/ml E64, 10 mg/ml phenylmethylsulfonyl fluoride, 10 mg/ml TLCK (Nα-p-Tosyl-L-lysine-chloromethyl ketone hydrochloride), and 1 mg/ml leupeptin) was added to a final concentration of 1×, and Triton X-100 was added to a final concentration of 1%. Samples were incubated at 0°C for 1 h, then spun at 16,000 × g in an Eppendorf centrifuge 1415C at 4°C for 30 min to pellet actin filaments. The supernatant was mixed with an equal volume of acetone (Fisher Scientific, Houston, TX) and pelleted at 16,000 ×g at 4°C for 20 min. These samples and parasite lysates (Dobrowolski and Sibley, 1996) were resuspended in protein gel sample buffer. Supernatants were loaded onto protein gels at 25% the concentration of the pellet samples. Actin was detected with a rabbit anti-ACT1 polyclonal antibody at 1:10,000 (Dobrowolski et al., 1997). As a control for lysis, samples were blotted with a mouse monoclonal anti-β-galactosidase antibody (40–1a) at 1:30 (Dobrowolski et al., 1997). Signals were enhanced with a horseradish peroxidase–conjugated secondary antibody to rabbit or mouse (1:10,000) and detected with the ECL+Plus Western blotting detection system (Amersham Biosciences, Piscataway, NJ). Western blots were quantified with a Molecular Dynamics (Sunnyvale, CA) Storm 860 phosphoimager and Bio-Rad (Hercules, CA) Molecular Analyst software.

Electron Microscopy

Parasites were resuspended in mammalian Ringer's solution (in mM: 155 NaCl, 3 KCl, 2 CaCl2, 1 MgCl2, 3 NaH2PO4, 10 HEPES, 10 glucose) at ∼10 8 parasites per milliliter and pretreated with 2 μM JAS, 1% DMSO, or 1 μM CytD (Molecular Probes) for 10 min at room temperature before being allowed to glide on poly- l -lysine–coated coverslips for 10 min at 37°C. Coverslips were sonicated in KHMgE (in mM: 70 KCl, 30 HEPES, 5 MgCl2, 3 EGTA at a pH of 7.2–7.4) plus 2 μM JAS, 1% DMSO, or 1 μM CytD with a microprobe mounted on a Mega-Mix Sonicator (GCA/Precision Scientific, Winchester, VA) for three pulses of ∼5 s each at maximum intensity. Samples were then fixed in KHMgE plus 2% glutaraldehyde, rapidly frozen in liquid nitrogen, freeze-dried, and platinum-coated as described previously (Håkanssonet al., 1999). Replicas were examined and photographed with a JEOL CX100 transmission electron microscope (Jeol, Peabody, MA).

Videomicroscopy

Freshly isolated parasites were resuspended in Ringer's solution containing 1% FBS, treated with varying concentrations of JAS, 1 μM latrunculin B (LatB) (Molecular Probes), or 1% DMSO, and allowed to glide on glass-bottom microwells (MatTek Corp, Ashland, MA) precoated with 50% FBS in PBS. Videomicroscopy was conducted using a Zeiss Axiovert equipped with phase contrast and epifluorescence microscopy and a temperature-controlled stage (Medical Systems Corp., Greenvale, NY) to maintain 37°C incubation as described previously (Håkansson et al., 1999). The optical path in this microscope is corrected to simulate an upright orientation, and thus objects appear in their true orientation with respect to thex and y planes. Motility was observed within minutes of addition of drug-treated parasites to the heated chamber and was recorded over a period of up to 30 min.

Images used for quantification were collected in real time under low-light illumination using an intensified CCD C2400 camera (Hamamatsu Photonics KK, Bridgewater, NJ) at 63× magnification. The video signal was digitally processed using a video-frame capture board (Perceptics Corp, Knoxville, TN) controlled by Biological Detection Systems image analysis software (BDS Image V 1.4.1, Oncor Imaging, Gaithersburg, MD) running on a Macintosh Quadra 900 computer (Apple Computers, Cupertino, CA). The analog output was recorded to S-VHS tape using a JVC model SR-S360U videocassette recorder. For analysis, parasite tracks were traced from the video monitor onto transparency sheets. Run lengths and start and stop frames for each movement were used to calculate velocities. The length of T. gondii (7 μm) provided a magnification standard. Five examples from three separate experiments were analyzed to obtain mean ± SEM values.

Time-lapse images used to construct Figure 4 and the supplemental videos were collected under low-light illumination with a Hamamatsu ORCA ER camera. Videos were recorded digitally at approximately eight frames per second with Openlab version 3.0.5 (Improvision, Lexington, MA), cropped, imported into Adobe Premiere 6.0, and saved as Quicktime movies.


Discussion

This study was undertaken to elucidate the observation whereby delocalization of β-actin mRNA can affect cell motility and polarity in fibroblasts. Delocalization of β-actin mRNA with antisense oligonucleotides has been observed to reduce the motility of CEFs (2) and smooth muscle cells (15).

Our results show that antisense but not control (sense) oligos caused a delocalization not only of β-actin mRNA, but also of β-actin protein and barbed ends from the leading edge of fibroblasts and resulted in a random distribution of all three. This was reversible upon removal of the antisense oligonucleotides. By investigating sites of actin filament nucleation, we showed that they were delocalized as a result of disrupting the targeting of β-actin mRNA. This result reveals a possible mechanism for establishing cell polarity: β-actin protein, and/or proteins with related zip codes, define the location of nucleation of actin polymerization and consequently, cell polarity and directional motility.

The molecular mechanism by which polarity of cell crawling is affected by β-actin mRNA localization could depend on several interdependent events: (i) Localized synthesis of β-actin from localized mRNA drives protrusion of the lamellipod. (ii) β-actin isoform specific protein interactions are responsible for the protrusion. (iii) Localization to the leading edge of the mRNAs for other proteins in addition to β-actin, e.g., the nucleating complex containing Arp 3 mRNA. A discussion of the evidence for each of these is detailed below.

Localized Synthesis of β-Actin from Localized mRNA.

Based on the estimated 2,500 β-actin mRNA molecules per cell, at the established translational rate of 1.5 actins per sec per mRNA molecule, the cell would synthesize 3,900 actin molecules per sec or 2.34 × 10 5 per min (2). In moving cells, the polymerization zone uses a minimum of 3.6 × 10 6 actin molecules per min (9). Therefore, it is unlikely that a 6.5% contribution of newly synthesized β-actin will significantly contribute to the rate of actin polymerization at the leading edge. However, if all of the β-actin is synthesized in a restricted volume, a consequence of localizing the β-actin mRNA to the leading edge, the local rate of synthesis of β-actin might significantly impact the actin polymerization events in this restricted volume and, therefore, establish a preferred location for actin polymerization.

The above model would be further supported if newly synthesized actin would have a faster rate of polymerization, or a higher affinity for a nucleation complex than “older” actin, which may be posttranslationally modified. For instance, interaction of a chaperone with the β-actin nascent chain (16) could promote assembly of a nucleation complex near the site of synthesis.

Β-Actin Isoform-Specific Protein Interactions.

The β isoform of actin may be preferentially stored as the monomer used for polymerization at the leading edge. In this hypothesis, local accumulation of a nonfilamentous form of actin that could be released suddenly upon stimulation of motility would determine the location of actin polymerization. Potential storage particles containing nonfilamentous actin have been identified by comparing the localization patterns of vitamin D-binding protein, which binds to G-actin with 5 nM kd, and phalloidin, which binds to actin (17). These stores of nonfilamentous actin are found at the leading edge and are located adjacent to sites of actin polymerization and in the region of the cell where the β-actin mRNA is also present. Possibly these sites could result from islands of β-actin synthesis.

β-actin is found at the leading edge of crawling cells. β-actin does not substitute for muscle actin in either the formation of stress fibers (17) or myofibrils in cardiomyocytes (18). In addition, it seems to interact more tightly with certain actin binding proteins that may function at the leading edge of crawling cells. Ezrin (19), profilin (20), thymosin β 4 (21), and L-plastin (22) bind more strongly to β-actin than α-actin. A capping protein, β-cap 73, may cap the barbed end in an isoform-specific manner (23). There is growing evidence that the Arp2/3 complex is required for nucleation of actin filaments at the leading edge (12, 24–27). If the Arp2/3 complex is the dominant nucleation activity at the leading edge, a possible preference for the β-actin isoform by the Arp2/3 complex would require local synthesis of β-actin to supply the preferred monomer for polymerization. Therefore, the localization of β-actin synthesis at the leading edge may be functionally important for polarity and motility.

Localization to the Leading Edge of Motility-Related mRNAs.

The localization of β-actin mRNA may be representative of the localization of a family of mRNAs with related 3′ UTR zip codes, many of which function synergistically at the leading edge. Proteins coded for by these mRNAs therefore might have related functions. We have analyzed the 3′ UTRs of mRNAs, which code for proteins believed to have actin binding functions at the leading edge, for the presence of the zip code consensus sequence. This sequence GACUX7–38ACACC is found in β-actin mRNAs known to target to the leading edge from all vertebrates. Besides β-actin mRNAs, mRNA for Arp3 and myosin IIB heavy chain contain the consensus sequence and are predicted to be recognized by the localization mechanism that targets β-actin mRNA to the leading edge. It is known that the ACACCC consensus sequence, when mutated in β-actin mRNA, results in a failure to localize the mRNA to the leading edge of cells (2, 7), even if the β-actin coding sequence remains intact and is used as the reporter mRNA. Preliminary results indicate that Arp3 mRNA, like β-actin mRNA, also localizes to the leading edge (G. Liu, W. Grant, D. Persky, V. L. Lathaur, R.H.S., and J.C., unpublished work). Serum-dependent localization of β-actin mRNA suggests that signaling mechanisms are involved in the localization of motility-related mRNAs, thereby coordinating their temporal and spatial distribution and expression (28). Furthermore, it is possible that localized synthesis of, for instance, Arp3 could determine the localization of Arp2/3 complex in the leading edge of the cells even if mRNAs coding other components of Arp2/3 complex were more diffusely distributed. Arp2/3 complex and β-actin, both localized in the leading edge, could determine the nucleation sites for actin polymerization. Newly formed actin filaments could interact with β-actin isoform-specific binding proteins, thereby stabilizing the cell polarity and consequent directional motility (29).

The leading edge of the cell is a complex composite of asymmetrically distributed proteins many of which function in concert to produce the motility response. It is likely that other proteins like β-actin also are synthesized asymmetrically and therefore would provide not only a differential concentration of these proteins but also an increased likelihood of interactions among relevant proteins in a cellular region where function depends on these interactions. We presume therefore that a panoply of mRNAs comprising a significant complexity of sequences is localized to the lamella to effect the complex events required by motility. It is our expectation that these sequences will contain a common motif and/or structure in the 3′ UTR characterizing them as mRNAs for motility-related proteins. It is likely that further investigations will reveal the consensus sequences (see below).

The localization of β-actin mRNA is not restricted to fibroblasts, but seems to be a feature of other localized cells. Neurons localize β-actin mRNA to the growth cone of developing neurites (30, 31). The presence of the mRNA results in the specific translation of β-actin protein in the growth cone. Like fibroblasts, the delocalization of the mRNA results in growth cone retraction and nondirectionality of growth cone guidance (37). In addition to the neuronal growth cones, embryonic neural crest cells might localize β-actin mRNA to the front of the cell, in the direction of their migration. Disruption of the Xenopus homolog of ZBP1 appears to inhibit their migration and result in severe embryological defects in forebrain development (J. Yisraeli, personal communication). Furthermore, if the zip code for β-actin mRNA is transferred to another protein, not normally at the leading edge, in this case vimentin, a distorted morphology results wherein the cell structure at the leading edge is branched and attenuated (32). These results argue that synthesis of the correct protein in the correct place (near the leading edge) is an important requirement for cell structure and polarity.

In addition to β-actin mRNA localization in fibroblasts (1), the field of RNA localization has been advanced by the discovery of a number of systems where mislocalization of the RNA can lead to a significantly altered phenotype or lethality (33–36). In many of these cases mRNA localization is required for normal development and differentiation because the localized mRNA codes for nuclear factors and the resultant cell divisions segregate the mRNAs for these morphogenic determinants. However, the nature of the localization we describe here is important for a different reason: it determines the spatial orientation, morphology, and behavior of these somatic cells. In this second aspect of RNA localization, the complex of proteins involved in cell migration, cellular reaction to the environment and development of cell polarity are organized within the cytoplasm by virtue of the spatial segregation of their cognate mRNAs, and are not in the short term related to transcription of genes. In this way, components of the mechanism controlling cell behavior and structure can rapidly reassemble within the cell. In this model, the proteins involved in forming these multipolypeptide complexes (the nucleation complex, for instance) would be compartmentalized in response to environmental cues and subsequent signal transduction events and then synthesized in proximity to each other where they would interact preferentially because of their higher local concentrations. Possibly these higher concentrations of proteins could autoregulate their own synthesis. In this way, we propose that the localization of β-actin mRNA represents one mechanism for the spatially compartmentalized assembly of cellular complexes.


The arresting action of microtubules in cell motility

Cell migration plays a crucial role in embryogenesis, wound healing, immunity and other essential functions. The movement of cells is a complicated and coordinated maneuver that involves extension of a microfilament-rich lamellipodium at the leading edge, polarization of the cell in the direction of movement, establishment of focal contacts at the leading edge as well as removal of contacts at the trailing edge and stress fiber-mediated retraction of the tail. This regulated remodeling of the actin cytoskeleton provides the driving force for cell motility.

Microtubule involvement in cell motility is less clearly documented. Studies have indicated that drugs able to inhibit or promote microtubule assembly can reduce the migration of most cell types, but several exceptions have been reported (summarized in reference 1 ) e.g., drug treatment was shown to increase rather than inhibit the motility of leukocytes. 2 Exceptions such as these have led to the perception that the role of microtubules in motility is cell type-specific. 3 Although such an explanation is possible, it is equally likely that some of the reported differences can be attributed to indiscriminate use of high drug concentrations that may cause toxicity unrelated to effects on microtubule assembly. At the other extreme, it has been reported that very low drug concentrations that do not disrupt microtubules can inhibit cell migration, and the authors of one study speculated that the drugs might be acting by suppressing microtubule dynamics. 4 Additional confusion has arisen regarding the mechanism by which microtubules might influence cell motility, with various investigators proposing microtubule-mediated effects on actin polymerization, vesicle transport to the growing lamellipodia and turnover of adhesion plaques. 3 , 5

Recent studies have begun to clarify these issues. Although it is currently thought that microtubule drug action is mediated almost entirely by suppression of microtubule dynamics, experiments have now demonstrated that these agents cause distinct concentration-dependent changes in microtubule behavior. Low drug concentrations with no effect on microtubule organization or polymer levels were able to suppress microtubule dynamics and cause a parallel inhibition of cell migration. 6 , 7 These low concentrations had no influence on cell division in either wild-type or mutant cell lines that are drug-dependent for proliferation. At the 10-fold higher concentrations needed to inhibit mitosis, the drugs acted by a newly discovered mechanism involving altered stability of microtubule attachment to centrosomes and spindle poles. The relationship between microtubule dynamics and cell migration was further supported by studies with 㬣, a tubulin isotype that is normally restricted to neurons but whose expression in non-neuronal tumors has been associated with resistance to therapy. 8 Transfection of 㬣 was found to have no direct effect on cell motility, but it counteracted the ability of microtubule drugs to inhibit cell migration by preventing them from suppressing microtubule dynamics. 9 Unpublished studies from our laboratory have further shown that expression of 㬦, a tubulin isotype normally restricted to mammalian platelets, potently suppresses microtubule dynamics and also inhibits cell migration. 10 It has thus become clear that microtubules must be dynamic in order for cells to move, and that the dynamics can be suppressed by drug concentrations that are much lower than those needed to block cell division or alter microtubule polymer levels. These observations suggest the possibility of using low, non-toxic concentrations of microtubule inhibitors to block cell migration-mediated processes, such as tumor metastasis and angiogenesis.

The mechanism by which microtubule dynamics affect cell migration is also starting to come into focus. When dynamics are suppressed, cells located at the edge of a scratch wound continue to extend lamellipodia and polarize in the direction of the wound, but they remain stretched for an extended period of time and appear to have difficulty retracting their tails. 7 This observation could indicate that drug treated cells are defective in removing adhesion sites from the tail, a process that has been reported to require microtubules. 3 However, this interpretation is insufficient to explain recent experiments indicating that micro-tubules are not needed for motility (our unpublished studies). For example, a plot of cell migration as a function of drug concentration produces a “U-shaped” curve. Low drug concentrations suppress microtubule dynamics and inhibit cell migration with no obvious change in microtubule organization or assembly. Migration remains inhibited as the drug concentration increases to the point that microtubules depolymerize, whereupon cell migration recovers. It should be noted that still higher drug concentrations again inhibit cell migration, most likely because of additional toxic drug effects. Based on these and other observations, we propose a model in which microtubules act to inhibit, rather than promote, cell migration ( Fig. 1 ). The concept is similar to the tensegrity model, in which microtubules act as struts to oppose contractile forces, and it is consistent with studies showing that microtubules inhibit muscle contraction. 11 , 12 The model is also consistent with the observation that stable microtubules are oriented toward the leading edge of migrating cells, whereas dynamic micro-tubules are oriented toward the tail. 13 As long as the microtubules are dynamic, they can remodel and allow the tail to retract, but when dynamics are suppressed, microtubules remain immobile and physically retard tail retraction. It should be noted that, while this model explains many of the contradictory reports regarding drug effects on cell migration, it does not necessarily rule out the other actions proposed for microtubules in migrating cells. However, the ability of cells to move efficiently in the absence of microtubules argues that any involvement of the microtubule cytoskeleton in actions such as adhesion site turnover or delivery of membrane vesicles to the leading edge is not rate limiting for cell motility.

The role of microtubule dynamics in cell migration. Cells normally move by extending lamellipodia and elongating in the direction of movement. The more stable, less dynamic microtubules (solid lines) are oriented toward the leading edge the less stable, more dynamic microtubules (dotted lines) are oriented toward the trailing edge. Forward progression occurs when the tail retracts, a process facilitated by the presence of dynamic microtubules but opposed by stable microtubules. A low drug concentration sufficient to suppress microtubule dynamics does not prevent the extension of lamellipodia or cell elongation, but the cells remain in the elongated state for a protracted amount of time and appear unable to retract the tail. The morphologies that are shown were traced from HeLa cells moving into a scratch wound, and the numbers indicate the time in minutes from an arbitrary start.


Notes

Abbreviations used: CCL and CCR, CC chemokine ligand and receptor, respectively CMFDA, 5-chloromethylfluorescein diacetate CXCL, CXC chemokine ligand DOCK2, dedicator of cytokinesis 2 FRC, fibroblastic reticular cell GEF, guanine nucleotide exchange factor HEV, high endothelial venule PBT cell, peripheral blood T cell PH, pleckstrin homology PI3K, phosphoinositide 3–kinase PIP3, phosphatidylinositol 3,4,5-trisphosphate plt, paucity of LN T cell PTX, pertussis toxin WMN, wortmannin.


Chemotaxis pathways that work in parallel with PtdIns(3,4,5)P3 signaling

The recent experimental observations described in the previous two sections support a model in which the PI3K-PTEN pathway is important for regulating the actin cytoskeleton in Dictyostelium chemotaxis and in mammalian cells, but it is not the only pathway regulating chemotaxis. In support of this, other parallel pathways are now thought to contribute to chemotactic movement and gradient sensing.

Phospholipase A2 and phospholipase C in Dictyostelium

The chemoattractant cAMP stimulates several second-messenger systems in Dictyostelium, including adenylyl cyclase, guanylyl cyclase, the uptake of Ca 2+ and its release from internal stores, phospholipase C (PLC), PI3K and phospholipase A2 (PLA2 encoded by plaA). Two groups have independently demonstrated that PLA2 regulates chemotaxis in parallel with PI3K (Fig. 2) (Chen et al., 2007 van Haastert et al., 2007). Further analysis of the two pathways revealed that inhibition of either pathway in shallow gradients inhibits chemotaxis, whereas, in steep gradients, both pathways must be inhibited to prevent proper chemotaxis.

Previous studies in Dictyostelium have suggested that products of PLA2 action, such as arachidonic acid, affect chemoattractant-induced Ca 2+ influx and can trigger Ca 2+ influx directly (Schaloske and Malchow, 1997). In mammalian cells, arachidonic acid is involved in the release of Ca 2+ from internal stores by regulating calcium channels (Osterhout and Shuttleworth, 2000 Shuttleworth and Thompson, 1999). Cells lacking PLA2 display a decrease in the levels of 3 H-arachidonic acid, or a closely related derivative, after stimulation (Chen et al., 2007). Although disrupting PLA2 has no effect on Ca 2+ uptake (Chen et al., 2007), simultaneous inhibition of Ca 2+ uptake and Ca 2+ release does make cells sensitive to PI3K inhibitors (van Haastert et al., 2007). These results suggest that the PLA2-dependent pathway involves a rise in intracellular Ca 2+ that might be regulated by arachidonic acid or derivatives via the release of Ca 2+ from internal stores. Cytosolic Ca 2+ might also have a regulatory effect on PLA2 activation (Chen et al., 2007 van Haastert et al., 2007).

Inhibition of both PLA2 and PLC almost completely inhibits the cAMP-mediated PtdIns(3,4,5)P3 response and causes drastic chemotactic defects (van Haastert et al., 2007), although inhibition of PLC alone does not affect chemotaxis (Drayer et al., 1994). This result implicates PLC in the regulation of the PI3K-mediated chemotaxis pathway – it probably acts by regulating phosphatidylinositol (4,5)-bisphosphate [PtdIns(4,5)P2] levels and PTEN (Fig. 2). In addition, PLC signaling might cross-talk with the PLA2 pathway at the level of intracellular Ca 2+ regulation. PLC controls intracellular Ca 2+ levels by generating inositol phosphates [inositol (1,4,5)-trisphosphate Ins(1,4,5)P3] and diacylglycerol (DAG), which activates Ca 2+ -sensitive enzymes such as protein kinase C (PKC) (Drin and Scarlata, 2007).

The exact role of PLA2 and its derivatives in chemotaxis and the degree to which the pathways have overlapping functions or influence different aspects of chemotaxis remain to be determined. The fact that cells are still able to sense gradients and move with a high directionality in the absence of PI3K or PLA2 confirms that the machinery responsible for directional sensing must act downstream of G-protein activation by chemoattractants but upstream of the PI3K-PTEN and PLA2 pathways. Components of this direction-sensing machinery could include Ras proteins, which help to control chemotaxis in Dictyostelium, in part via the regulation of PI3K and TORC2 (Funamoto et al., 2002 Lee et al., 2005 Lee et al., 1999 Sasaki et al., 2004). Interestingly, the expression of a dominant-negative RasG in Dictyostelium cells lacking the RasGEF Aimless (AleA) severely impairs directional sensing (Sasaki et al., 2004).

The regulation of PtdIns(4,5)P2 and PtdIns(3,4,5)P3 levels by PLC is also involved in the response to chemorepellents in Dictyostelium (Keizer-Gunnink et al., 2007). cAMP analogs, such as 8-para-chlorphenylthio-cAMP (8CPT-cAMP), can induce a repellent response in Dictyostelium by binding to the cAMP receptor cAR1 (Johnson et al., 1992). This induces a localized inhibition of PLC, which is normally activated by cAMP. Inhibition of PLC is proposed to cause the local accumulation of PtdIns(4,5)P2, PTEN binding and PtdIns(3,4,5)P3 degradation at the front of the cell. This leads to dominant PtdIns(3,4,5)P3 signaling at the rear of the cell, resulting in a movement away from the repellent source. PLC therefore can act as a polarity switch, controlling the response to signals in the environment.

PLA2 and PI3K/PTEN regulate chemotaxis in Dictyostelium. In Dictyostelium, chemotaxis is regulated by at least two intertwined and partly redundant pathways involving PI3K and PLA2. Both pathways are regulated by extracellular cAMP. The PI3K pathway is regulated, via PtdIns(4,5)P2 (PIP2)/PTEN, by PLC. The PLA2 pathway depends on cytosolic Ca 2+ , which is regulated by IP3 (thus partly by PLC), fatty acids and Ca 2+ uptake. In steep gradients, either pathway is dispensable in shallow gradients, both pathways are necessary to allow efficient chemotaxis (see text for details). Red arrows indicate enzymatic reactions. PL, phospholipids Lyso-PL, lyso-phospholipids Gαβγ, heterotrimeric G protein cAR1, cAMP receptor PIP3, PtdIns(3,4,5)P3.

PLA2 and PI3K/PTEN regulate chemotaxis in Dictyostelium. In Dictyostelium, chemotaxis is regulated by at least two intertwined and partly redundant pathways involving PI3K and PLA2. Both pathways are regulated by extracellular cAMP. The PI3K pathway is regulated, via PtdIns(4,5)P2 (PIP2)/PTEN, by PLC. The PLA2 pathway depends on cytosolic Ca 2+ , which is regulated by IP3 (thus partly by PLC), fatty acids and Ca 2+ uptake. In steep gradients, either pathway is dispensable in shallow gradients, both pathways are necessary to allow efficient chemotaxis (see text for details). Red arrows indicate enzymatic reactions. PL, phospholipids Lyso-PL, lyso-phospholipids Gαβγ, heterotrimeric G protein cAR1, cAMP receptor PIP3, PtdIns(3,4,5)P3.

PLC and PLD in mammalian chemotaxis

PtdIns(4,5)P2 has a pivotal role in both the PLC and PLD cellular signaling pathways. It serves as the major substrate for PLC proteins and simultaneously influences the subcellular localization and activity of PLD proteins. Studies on mouse neutrophils lacking PLC-β2 and PLC-β3 isoforms have indicated that the PLC pathways play an important role in chemoattractant-mediated production of superoxide, and in the regulation of protein kinases and chemokine-induced Ca 2+ signaling, but not in chemotaxis (Li et al., 2000 McNeill et al., 2007). Nevertheless, treatment of human neutrophils with PLC inhibitors blocks chemotactic responses to interleukin 8 (IL8) and leukotriene B4 (LTB4) (Hou et al., 2004). Thus, the function of PLC in neutrophil chemotaxis is not fully understood and might vary according to the chemoattractant. By contrast, PLC-β is clearly necessary for T-cell chemotaxis. It acts by transiently raising cytoplasmic Ca 2+ concentrations via inositol (1,4,5)-trisphosphate [Ins(1,4,5)P3, IP3] but does not influence PKC activity (Bach et al., 2007 Smit et al., 2003). The identities of the Ca 2+ -dependent downstream components (such as calmodulin kinases, myosin light-chain kinase or Rho kinase) that mediate the chemotactic response in T-cells remain elusive.

The PH domain of PLC is thought to target it to particular lipids and membrane surfaces (Rebecchi and Scarlata, 1998). There is evidence that, in differentiating promyelocytes, PLC-β2 interacts with actin via the PH domain of PLC-β2 (Brugnoli et al., 2007). This promotes the association with cytoskeleton-associated PtdIns(4,5)P2 in the plasma membrane, resulting in hydrolysis of PtdIns(4,5)P2 and consequent cytoskeletal rearrangements and motility.

PLD hydrolyzes the phosphodiester bond in phosphatidylcholine (PC), resulting in the production of choline and the second messenger phosphatidic acid (PA) (Oude Weernink et al., 2007). The function of PLD during neutrophil chemotaxis has not been fully determined. Azuma et al. (Azuma et al., 2007) propose that PLD is required for activation of p38MAPK upon stimulation with fMLP under conditions that stimulate superoxide production, but not chemotaxis. By contrast, Powner et al. (Powner et al., 2007) have recently demonstrated that PLD regulates integrins that support stable adhesion during neutrophil migration. In that study, PA produced by PLD activity stimulated the generation of PtdIns(4,5)P2 by stimulating phosphatidylinositol 4-phosphate 5-kinase activity in response to fMLP. PtdIns(4,5)P2 promoted the binding of talin to the surface-expressed β2-integrin CD18 and hence caused activation of the integrins CD11b/CD18, required for stable adhesion and migration. The study also pointed to the involvement of PLD in the distribution and function of actin stress fibers. In Dictyostelium, inhibition of PLD causes a dramatic decrease in PtdIns(4,5)P2 synthesis, resulting in severe defects in actin-based motility (Zouwail et al., 2005). In mast cells, human PLD binds to actin, which is important for the regulation of PLD1b activity (Farquhar et al., 2007). Evidence thus supports a link between phosphatidylcholine hydrolysis and remodeling of the actin cytoskeleton in a variety of processes and cell types.

In neutrophils, PI3Kγ becomes localized to the plasma membrane in response to stimulation by fMLP, increasing the formation of PtdIns(3,4,5)P3 and thereby the recruitment of other factors that activate PLD (e.g. Rho and Arf-GTPases, as well as PKC isoforms) (Chen and Exton, 2004 Henage et al., 2006). The activity of PLD can be inhibited by prostaglandin E2 (PGE2), which stimulates protein kinase A (PKA). This in turn inhibits the translocation of the PLD-activating factors, possibly by inhibiting PI3K (Burelout et al., 2007 Burelout et al., 2004). However, it is unclear how PI3K might be inhibited, because the PI3K regulatory and catalytic subunits are not targets for PKA phosphorylation in vitro and are unlikely targets in vivo. PI3K activity is regulated by the binding of the subunits p101/p110γ to the Gβγ subunits released upon receptor activation (Brock et al., 2003 Stephens et al., 1997), but Gβγ-subunit protein sequences do not contain consensus sites for PKA phosphorylation, and phosphorylation by PKA has not been demonstrated. The fMLP receptor is also not phosphorylated by PKA (Burelout et al., 2007). Therefore, the mechanism that underlies the inhibitory effect of PKA on PI3K requires further analysis.

PLC and cofilin in adenocarcinoma cells

In adenocarcinoma cells, epidermal growth factor (EGF) stimulation induces two peaks of actin polymerization, which is similar to the biphasic F-actin-polymerization response to cAMP that is seen in Dictyostelium (Chan et al., 2000 Chan et al., 1998 Chen et al., 2003). The second peak is dependent on PI3K activity, both in carcinoma cells and in Dictyostelium (Chen et al., 2003 Hill et al., 2000). In carcinoma cells, the first peak was recently demonstrated to depend on PLC-γ and cofilin (Mouneimne et al., 2004). These results, and those from other studies, strongly argue that PLC, together with cofilin, mediates gradient sensing in these cells (Ghosh et al., 2004 Mouneimne et al., 2006). By contrast, in Dictyostelium PLC regulates PIP2 levels and therefore cell motility, but not directional sensing. In Dictyostelium, cofilin is involved in actin remodeling and localizes to the leading edge during chemotaxis (Aizawa et al., 1995 Aizawa et al., 1997) but there is no evidence that it regulates gradient sensing and, in contrast to mammalian cofilin, Dictyostelium cofilin lacks the regulatory Ser at position 3. Interestingly, in cells lacking both PLA2 and PI3K activities (plaA – /pi3k1 – /2 – cells), the first peak of actin polymerization is significantly decreased, although both plaA – cells and pi3k1 – /2 – cells have nearly normal first peaks of actin polymerization, indicating that both pathways might be involved in the initial actin polymerization (Chen et al., 2007).

In carcinoma cells, cofilin is activated via PLC-γ, which acts by locally decreasing PtdIns(4,5)P2. Cofilin is essential for the localized formation of barbed ends, which act as sites for new local actin polymerization cofilin thus determines the direction of cell protrusion and movement (Condeelis, 2001 DesMarais et al., 2005 Ghosh et al., 2004) (Fig. 3). Cofilin activity seems to be mainly dependent on PLC-γ-mediated PtdIns(4,5)P2 hydrolysis and does not involve an IP3-mediated Ca 2+ release (Ma et al., 2000 Mouneimne et al., 2006 Yonezawa et al., 1991). In addition to activation by PLC-γ, mammalian cofilin is also phosphorylated at Ser3 by LIM kinase (LIMK), which inhibits the ability of cofilin to bind actin (Zebda et al., 2000), and is dephosphorylated by the phosphatase Slingshot (SSH) (Nishita et al., 2005 Niwa et al., 2002 Ohta et al., 2003). Phosphorylation of cofilin increases upon EGF stimulation (Mouneimne et al., 2004 Song et al., 2006) and is required for chemotactic sensing, although the exact function is not fully understood (Mouneimne et al., 2006). In Jurkat T-cells, cofilin is thought to be inactivated by phosphorylation by LIMK after stimulation with stromal cell-derived factor-1α (SDF-1α). This results in the formation of F-actin-rich lamellipodial protrusions (Nishita et al., 2005). Via the association with F-actin, the phosphatase Slingshot-1L becomes locally activated in the protrusions and dephosphorylates, and thereby re-activates, cofilin in the lamellipodium. This allows actin-filament turnover and ensures the dynamic nature of the lamellipodium (Nishita et al., 2005).

Recent studies of carcinoma cells indicate that the initial activation of cofilin does not involve dephosphorylation in response to chemoattractant stimulation (Mouneimne et al., 2004 Song et al., 2006). Cofilin is instead thought to be locally released and activated by hydrolysis of PtdIns(4,5)P2 by PLC-γ and, simultaneously, be globally inactivated via phosphorylation by LIMK (Hitchcock-DeGregori, 2006 Mouneimne et al., 2006). This leads to an asymmetric distribution of cofilin activity, setting the direction of lamellipodium formation and subsequent migration. This model is consistent with earlier findings that cofilin is recruited to the leading edge immediately before lamellipod extension and is followed by the Arp2/3 complex and the extension of the lamellipod (DesMarais et al., 2004). However, it is not fully understood whether or not cofilin is completely deactivated by LIMK and whether the phosphatase SSH or 14-3-3 proteins might be involved in this process (Soosairajah et al., 2005).

Whereas the inhibition of PLC-γ/cofilin leads to defects in gradient sensing, inhibition of PI3K or PTEN decreases motility and speed in carcinoma cells, as it does in Dictyostelium (Mouneimne et al., 2006 Mouneimne et al., 2004). Full lamellipod extension requires PI3K activity, because the second peak of actin polymerization is dependent on PI3K (Hill et al., 2000). PI3K has been postulated to signal to WAVE and the Arp2/3 complex, which is necessary for lamellipod protrusion (Bailly et al., 2001 Higgs and Pollard, 2001 Takenawa and Miki, 2001). The PLC-γ/cofilin and PI3K/Arp2/3 signaling pathways thus cooperate in chemotactic gradient sensing and efficient lamellipod generation in response to EGF stimulation. By hydrolyzing PtdIns(4,5)P2, PLC-γ activates cofilin, which promotes F-actin severing. This creates free barbed ends, defining the site for Arp2/3 activation. The Arp2/3 complex nucleates new filaments, which become elongated by Ena/VASP proteins, creating a branched actin network that allows stable lamellipod protrusion and migration.


Introduction

Bacterial motility is important for a wide variety of biological functions such as swarming, chemotaxis, biofilm formation, and virulence. For example, the Gram-negative bacterium Myxococcus xanthus exhibits a complex life cycle that includes swarming, predation, and fruiting body formation: motility is important for all of these functions. M. xanthus does not contain flagella, but is able to move across solid surfaces using two very different motility systems ( Hodgkin and Kaiser, 1979 ). The first motility system, called social (S-) motility, is similar to twitching motility in Pseudomonas aeruginosa and is powered by type-IV pili localized at the leading cell pole ( Wall and Kaiser, 1999 ). Cell movement occurs because the polar pili bind to polysaccharides on the substrate or on the surface of other cells: this triggers pilus retraction, which pulls the cells forward ( Wall and Kaiser, 1999 Sun et al, 2000 Li et al, 2003 ). The second motility system, called adventurous (A-) motility, is still not very well understood. Multiple genetic screens have led to the identification of over 40 genes required for A-motility, but their specific functions are mostly uncharacterized and a molecular motility engine has not been identified ( Rodriguez and Spormann, 1999 Youderian et al, 2003 Yu and Kaiser, 2007 ). During the last 40 years, many models have been proposed to explain the mechanism of A-motility, including surfactant effects, moving chains of adhesions, rotating membrane embedded rotors, and, more recently, slime extrusion through nozzles ( Wolgemuth et al, 2002 Mignot, 2007 ). However, clear-cut evidence for any of these models has been lacking.

Recently, several findings suggested that A-motility involves distributed motors and focal adhesion complexes. For example, observations of filamentous cells indicated that the A-motility gliding motors are not located at the lagging cell pole, but distributed along the cell bodies ( Sun et al, 1999 Sliusarenko et al, 2007 ). Furthermore, cytological studies showed that AglZ, an A-motility protein ( Yang et al, 2004 ), is localized in clusters that originate at the leading cell pole. As cells moved forward, the clusters were localized at regular intervals along the cell body, where they remained at fixed positions relative to the substratum ( Mignot et al, 2007 ). Based on this evidence, it was proposed that the AglZ clusters were associated with A-motility motors that power motility by coupling movement on a rigid cytoskeletal filament with adhesion complexes on the surface ( Wozniak et al, 2004 Mignot, 2007 Mignot et al, 2007 ). This proposed motility mechanism has similarities to eukaryotic focal adhesion complexes, where cell-surface ligands that provide anchor points with the extracellular matrix are connected to the actin–myosin network in the interior of the cell ( Wozniak et al, 2004 ).

Motility in M. xanthus exhibits an additional complexity in that cells periodically reverse. During reversals, which usually occur about every 7–14 min depending on the cultural conditions, the polarity of cells inverts. Thus, the leading cell pole becomes the lagging pole and the old lagging pole becomes the new leading pole ( Mauriello and Zusman, 2007 ). During cell reversals, the A- and S-engines reverse direction coordinately. For example, the S-motility protein FrzS and the A-motility protein AglZ are transferred together from the old to the new leading pole ( Ward et al, 2000 Yang et al, 2004 Mignot et al, 2005 , 2007 ). At the same time, proteins associated with the lagging cell pole, like the A-motility protein RomR ( Leonardy et al, 2007 ), track to the new lagging pole. The frequency of cell reversals is controlled by the Frz (frizzy) signal transduction system ( Blackhart and Zusman, 1985 ). It is hypothesized that periodic cell reversals allows cells to reorient themselves to achieve directed motility ( Zusman et al, 2007 ). Thus, most frz mutants rarely reverse and are defective in swarming and fruiting body formation in contrast, some frz mutants hyper-reverse and form very compact colonies as the cells show very little net surface translocation ( Bustamante et al, 2004 ).

In this paper, we identified the actin-like protein MreB and the Ras-like protein MglA as critical components in the localization of the A- and S-motility proteins, FrzS and AglZ. We also found that MreB acts upstream of MglA in the positioning of polar motility proteins and the focal adhesion complexes. Finally, our data suggest that assembly of the focal adhesion clusters is an essential requirement for cells to achieve A-motility.


Concluding Remarks

Two photon microscopy-based investigation of T cell� interactions has uncovered factors that, in addition to pMHC abundance and affinity, regulate the dynamic interplay between these two cell types. Although tightly regulated T cell attraction to activated DCs promote adaptive immune responses by allowing rare cells to meet, excessive interactions with low-affinity pMHC-presenting DCs deteriorate the overall quality of clonal expansion. Furthermore, intrinsic wiring of the migratory behavior of T cells facilitates their scanning function. Thus, programmed Myo1g-induced meandering behavior permits sufficiently long interactions between T cells and DCs. In addition, continuous F-actin treadmilling during both migration and IS formation endows T cells with the capacity to quickly switch between migratory versus stationary modes. Thus, upon cessation of TCR signaling, T cells resume their motility, presumably to avoid overstimulation and to prepare for egress. These observations raise new interesting questions. For example, is the duration of phase 2-like stable interactions regulated by external chemoattractant gradients or cell-intrinsic mechanisms? Furthermore, desensitization of chemokine receptors, such as CCR5, and its impact on T cell motility patterns in vivo has not been investigated in detail. Such studies are relevant since CCR5 ligands attract cognate and non-cognate naïve CD8 + T cells, and would rapidly limit access to DCs unless CCR5 desensitization is allowing cell turnover. The continued examination of mechanisms that control T cell motility in lymphoid tissue and their impact on adaptive immune responses will remain a productive field of research in years to come.


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