Fascin Is Involved in the Antigen Presentation Activity of Mature Dendritic Cells¹

Dendritic cells (DC)³ are now recognized as the most potent professional APC involved in initiating primary immune responses (1). Numerous studies have confirmed that mature interdigitating dendritic cells (IDC) are the primary, if not the only, APC involved in the activation of naive Th cells (2, 3). The importance of DC maturation for efficient T cell activation has been extensively discussed in several previous reports (1, 4, 5). Immature DC that reside in the peripheral tissues (areas of high Ag encounter) are equipped to capture and process Ag (6, 7). In response to activation signals, such as inflammatory cytokines, these immature DC undergo maturation, where they develop into potent T cell stimulators by up-regulating MHC class II as well as costimulatory and adhesion molecules. At the same time these DC migrate into secondary lymphoid organs to stimulate Ag-specific T cells (8, 9, 10, 11). In the secondary lymphoid organs, DC express high levels of the C-C chemokine, DC-CK1, which has been shown to preferentially attract naive (CD45RA⁺) T cells (12). In addition, DC maturation is associated with morphological changes, resulting in the expression of numerous delicate dendrites (13, 14). This dendrite formation has been associated, in other cells, with the expression of fascin.

Fascin is a 55-kDa, actin-bundling protein that regulates the rearrangement of cytoskeletal elements, and the interaction between the cytoskeleton and the cell membrane in response to extra- or intracellular signals (15). There is substantial evidence to link fascin expression with dendrite formation. High levels of fascin expression were found to be critical for dendrite development in neurons (16, 17). Recently, fascin expression was found to participate in the development of dendrites in mouse epidermal Langerhans cells (LC) (18). Moreover, when fascin was transfected into a pig epithelial cell line, the cells became dendritic and their motility was markedly increased (19).

Of interest is the fact that DC are the only leukocytes that have been demonstrated to express fascin (20). In the lymphoid organs only IDC, which are involved in APC activity, show high levels of fascin expression. This suggests a direct link between fascin expression and the ability of DC to function as potent APC.

In this study, we evaluated fascin expression and function during DC maturation. The data presented here demonstrate that fascin expression is tightly regulated during maturation of DC. They also demonstrate that fascin expression is critical for the Ag presentation activity of mature DC.

Adult BALB/c and C57BL/6 mice were purchased from Charles River Breeding Laboratories (St. Constant, Quebec, Canada) and housed in the Carleton Animal Care Facility (Sir Charles Tupper Medical Building, Dalhousie University, Halifax, Canada). All animals were housed in compliance with the guidelines established by the Canadian Council on Animal Care and were given standard rodent chow and water ad libitum. The medium used for bone marrow-derived DC (BM-DC) culture was RPMI 1640 (Sigma-Aldrich, Oakville, Ontario, Canada) supplemented with 5% heat-inactivated (30 min, 65°C) FCS (Life Technologies, Grand Island NY), 100 U/ml penicillin, 100 μg/ml streptomycin, and 5 mM 2-ME (British Drug House, Toronto, Ontario, Canada). In this report, we referred to this medium as complete RPMI (cRPMI).

A panel of mAbs was used in the immunocytochemistry and flow cytometry studies. All Abs were titered, and the optimal dilution was used. The anti-fascin mAb (mouse IgG1) was a gift of Dr. Erik Langhoff, Pennsylvania State University (Hershey, PA). Anti-Ia (mouse IgG1), anti-Thy1.2 (CD90; mouse IgG2b), anti-CD11c (N418; hamster IgG), purified mouse IgG1, PE-conjugated rat IgG-2a, FITC-labeled streptavidin, FITC- and PE-labeled rabbit anti-rat IgG2a, and PE-conjugated goat anti-hamster (IgG) Abs were all purchased from Cedarlane Laboratories (Hornby, Ontario, Canada). PE-conjugated anti-I-A^d/I-E^d mAb (rat IgG2a) was purchased from PharMingen (San Diego, CA). Alexa 488 (green) goat anti-mouse IgG conjugate was purchased from Molecular Probes (Eugene, OR). The hybridoma, GL1, producing the anti-B7-2 (rat IgG2a) mAb was purchased from the American Type Culture Collection (ATCC; no. HB-253; Manassas, VA).

Epidermal skin sheets were prepared essentially as previously described by Baker et. al (21). Briefly, a mouse ear was dissected and rubbed with the back of a pair of forceps to separate the two halves. The separated skin was then cut into 3- to 4-mm² squares and incubated in 3.8% NH₄SCN (Sigma-Aldrich Canada) for 30 min at 37°C. The skin was then washed in PBS, and the epidermis was separated from the dermis under a dissecting microscope. The epidermis was then fixed in cold (−20°C) acetone (BDH) for 5 min and stored in PBS at 4°C until stained. All staining of the epidermal skin sheet was performed in 1.5-ml microfuge vials (O.H. Johns, Mississauga, Ontario, Canada). The sheets were then mounted on slides, and positive cells were counted under ×20 magnification power. The data is presented as the number of positive cells per mm² of the skin sheet.

DC were prepared from BALB/c bone marrow essentially as previously described (22, 23). Briefly, marrow was flushed out of mouse femurs and tibias using sterile cRPMI in a laminar flow hood. The marrow suspension was then passed through nylon mesh to remove bone marrow particles. The cells were washed, and RBC were lysed with lysing buffer, containing 155 mM NH₄Cl in 10 μM Tris-HCl buffer (Sigma-Aldrich Canada) for 2 min at room temperature. FcR-positive cells were depleted by incubation (at 37°C for 1 h) on a Petri dish coated with IgG. After washing, lymphocytes and Ia-positive cells were removed by incubation with a mixture of mAbs for 60 min at 4°C followed by Low-Tox rabbit complement (Cedarlane Laboratories) for 60 min at 37°C. The mAbs used for depletion were GK1.5 (anti-CD4), HO2.2 (anti-CD8), B21-2 (anti-Ia), and RA3-3A1/6.1 (anti-B220/CD45R) (TIB 207, 150, 229, and 146, respectively; ATCC). Cells (5 × 10⁶) were cultured in 50-ml flasks (Nunc, Naperville, IL) in 5 ml cRPMI supplemented with either 50 U/ml recombinant mouse GM-CSF (rmGM-CSF; Cedarlane Laboratories) ± 200 U/ml rmTNF-α (Life Technologies), or 50 U/ml rmGM-CSF + 0.5 ng/ml TGF-β (CalBiochem-NovaBiochem, La Jolla, CA). The cultures were fed every 3 days by aspirating 85% of the medium and adding back fresh medium with growth factors.

Immunostaining was performed on lymph node histological sections, epidermal skin sheets, or BM-DC cytospins. For cytospin preparation, 7 × 10⁴ cells were cytocentrifuged onto poly-l-lysine (Sigma)-coated slides. Cells were then fixed in cold (−20°C) acetone for 2 min and stored at −20°C until use. For fascin staining, where mouse anti-fascin mAb was used, slides were fixed in 10% acetate-buffered formalin and incubated in a reagent (Signet Kit; IDlabs, Ontario, Canada) designed to block nonspecific binding of mouse Ab to mouse tissue. For all other staining, slides were fixed in cold (−20°C) acetone (10 min), 4% paraformaldehyde (2 min), dextran-Tris buffer (15 min), and glycine-lysine buffer (15 min). These slides were then incubated (30 min) in 3% H₂O₂ to block endogenous peroxidase followed by 1 h in horse serum to block nonspecific binding. Titered primary Ab or isotype control (mouse IgG1) was then added to the slides for incubation overnight at room temperature followed by the appropriate biotinylated secondary Ab for 1 h at room temperature. The Ab was localized using streptavidin-HRP (Signet Kit; IDlabs) for 1 h at room temperature and 3-amino-9-ethyl-carbozole (AEC; Sigma) as a chromogen.

Cultured DC were harvested on different days and suspended in 100 μl of PBS supplemented with 1% BSA. The cell suspension was then incubated with the appropriate primary Ab. All Abs were incubated for 30 min at 4°C. Permeabilization was required to stain for fascin, which is a cytoplasmic protein. Cells were permeabilized by incubation with 100% methanol (BDH) for 30 min at room temperature. For fascin-MHC class II double staining, cells were stained with anti-fascin mAb followed by FITC-conjugated goat anti-mouse Ab. PE-conjugated anti-MHC class II mAb was then added as a second Ab. For fascin-B7-2 double staining, cells were stained with anti-fascin mAb followed by FITC-conjugated goat anti-mouse Ab. Anti-B7-2 was then added followed by a PE-conjugated rabbit anti-rat Ab. Cells were washed twice with 1% BSA-PBS after each step and fixed with 1% paraformaldehyde-PBS. Fluorescence was analyzed on a total of 10,000 cells per sample using a flow cytometer (FACScan; Becton Dickinson, Mountain View, CA).

T cells were enriched from C57BL/6 spleen by filtration through a warm (37°C for 1 h) nylon wool column as previously described (24). Briefly, spleens were homogenized to achieve a single cell suspension. After RBC were lysed (as above), the cells washed with RPMI 1640, loaded into a nylon wool column, and incubated at 37°C for 1 h. Selection of nonadherent cells was followed by anti-B220 treatment followed by incubation with rabbit Low-Tox complements (as above). T cell purity was routinely between 80 and 86%, as assessed by flow cytometry using Thy1.2 staining.

BM-DC were harvested on day 9 and treated with 25 μg/ml mitomycin C (Sigma) for 30 min at 37°C. Treated DC were then applied in graded doses to 2 × 10⁵ T cell-enriched allogeneic spleen cells for 4 days. Cultures were maintained in U-shaped 96-well plates (Nunc), in 200 μl RPMI 1640 supplemented with 10% FCS, 50 μM 2-ME, 100 U/ml penicillin, and 100 μg/ml streptomycin. The cells were pulsed with 1 μCi/ml of [³H]thymidine (ICN Pharmaceuticals, Costa Mesa, CA) in the last 18 h of incubation. T cell proliferation was assessed by harvesting the cells on filtermats using a cell harvester (Skatron, Sterling, VA) and measuring the thymidine uptake in a liquid scintillation counter (Beckman Coulter, Fullerton, CA).

A fascin antisense oligonucleotide (oligo) and a matched sense control oligo of the same length (17 bp) were purchased from University Core DNA Services, University of Calgary (Calgary, Alberta, Canada). The sequence of the antisense oligo was provided by Dr. J. Bryan of Baylor College of Medicine (Houston, TX; J. Bryan, unpublished observation; Refs. 16, 17). To stabilize the antisense and the control sense oligo, three base pairs at the 3′ and the 5′ end were phosphorothioated. The sequence of the control oligo was CCGGCACCATGACCGCC and the antisense oligo was GGCGGTCATGGTGCCGG. Bone marrow precursors were cultivated in cRPMI supplemented with GM-CSF alone, GM-CSF plus control oligo (2 μM), or GM-CSF plus antisense oligo (2 μM). The cells were fed with new media supplemented with GM-CSF ± oligo every 3 days. On day 9, DC were harvested and an MLR assay was performed to investigate the effect of inhibiting fascin expression on DC allostimulatory activity.

Five areas on a fascin-immunostained cytospin were randomly selected. Microscopic images on these areas were captured using a JVC model TK-1070U video camera (JVC, Scarborough, Ontario, Canada) and a Nuvista⁺ frame grabber board (Truevision, Indianapolis, IN) connected to a Macintosh computer. The captured images were analyzed using the software package NIH Image (website: rsb.info.nih.gov/NIH-IMAGE/) on a Power Macintosh 6100/60 computer. The result is reported as percentage of fascin staining in the total DC cell area.

LC in the skin are the classical immature DC (25, 26). In contrast, lymph nodes contain mostly mature DC that have migrated from different interstitial sites in response to danger stimuli (1, 2). We chose to examine fascin expression in lymph node and skin as a first step to evaluate the correlation between fascin expression and DC maturation. Fascin binding was detected by immunohistochemistry using a biotinylated anti-mouse Ab as a secondary Ab and streptavidin-HRP for visualization. The results (Fig. 1,A) reveal strong fascin staining of DC in the T cell-dependent areas of the lymph node (IDC). However, in the germinal center (B cell) areas of the lymph node, the follicular DC were essentially fascin negative. This restriction of fascin expression to IDC is similar to what has been reported previously in human lymph nodes (27). Fig. 1 B shows the staining of the epidermal skin sheet for fascin. Very few fascin-positive cells could be seen. Image analysis revealed that only 10 ± 17 per mm² (n = 7) were fascin positive in this skin sheet. However, when these sheets were stained for MHC class II (image not shown), significant numbers of positive cells were observed (688 ± 158 cells per mm²).

These observations suggest that the expression of fascin in DC may be regulated during maturation. Furthermore, they showed that the IDC-restricted expression of this protein in the mouse lymph node is similar to that in humans. This data is consistent with a role for fascin in DC Ag presentation.

To confirm our in vivo observation regarding the correlation of fascin expression with DC maturation and to investigate the functional role of fascin in DC, we generated DC in vitro. These BM-DC mature over 7–9 days in culture, thus allowing enough time to examine fascin expression during maturation (22). Bone marrow precursor cells were allowed to grow for 9 days in the presence of cRPMI supplemented with GM-CSF. This regime pushes DC precursors to mature into DC (22, 28, 29). On day 9, cells were examined for fascin expression by immunocytochemistry. We observed a 29-fold increase in the percentage of cells staining for fascin at day 9 as compared with day 0. The staining was evenly distributed throughout the cytoplasm (Fig. 2,A). Moreover, cells that stained positively for fascin were large and displayed long dendrites, features normally seen in mature DC. We also examined BM-DC for MHC class II expression, a molecule known to be up-regulated during DC maturation (8, 30). We observed that BM-DC stain strongly positive for MHC class II, reflecting their degree of maturity (Fig. 2 B). These data demonstrated that our population of mature BM-DC expresses both fascin and MHC class II.

We used double color staining to compare the changes that occur in fascin expression and MHC class II expression during DC maturation. Day 9 BM-DC were double stained for fascin and MHC class II and examined by flow cytometry. Nearly all (98.5%) of the fascin-positive cells were MHC class II positive (Fig. 3,A). Furthermore, when we gated on the fascin-positive cells in Fig. 3,A, we found that 67.5% (square in Fig. 3,A) of these cells express high levels of MHC class II, reflecting their maturity. A similar pattern of fascin and MHC class II double positivity was seen with day 3 and day 6 BM-DC (data not shown). In comparison to day 3, we observed 6.9- and 9.4-fold increases in fascin expression on day 6 and day 9 BM-DC, respectively. Over the 9 days of examination, there was a direct correlation between the level of fascin expression and MHC class II^high expression with a correlation coefficient yield of 0.98 (Fig. 3,B). MHC class II^high expression has been used previously as a marker of mature DC (8, 30). To confirm that the strong correlation between fascin and MHC class II expression was indeed correlated with DC maturation, we examined a second marker of DC maturity, B7-2, during BM-DC maturation. B7-2 is a costimulatory molecule critical for T cell activation and is known to be up-regulated during DC maturation (31). We used double color staining to evaluate the correlation between fascin expression and B7-2 expression in mature DC. Day 9 DM-DC were double stained for fascin and B7-2 and examined by flow cytometry. Similar to the MHC class II double staining nearly all (96.0%) of the fascin-positive cells were B7-2 positive (Fig. 4,A). B7-2 was also up-regulated on the BM-DC with time in culture in parallel with fascin expression (Fig. 4 B). The highly significant correlation between fascin expression and MHC class II and the up-regulation of B7-2 expression in parallel with fascin strongly links fascin expression with DC maturation.

To demonstrate that the increase in fascin expression was due to BM-DC maturation rather than simply an effect of time in culture, we suppressed or enhanced BM-DC maturation with additional growth factors and examined the expression of fascin and MHC class II. BM-DC precursors were grown in cRPMI supplemented with 1) GM-CSF alone; 2) GM-CSF plus TGF-β, which has been reported to suppress DC maturation from bone marrow progenitors (23); or 3) GM-CSF plus TNF-α, which has been shown to enhance DC maturation from bone marrow progenitors (23, 32). Cells were harvested on days 0, 3, 6, and 9 from each group, double stained for fascin and MHC class II, and analyzed by flow cytometry.

Regardless of the treatment group, there was an increase in fascin expression over time and, again, most of the fascin-positive cells (94 ± 6%) were MHC class II positive. However, the number of fascin-positive DC was significantly (p < 0.001) reduced at day 6 (by 80%) and at day 9 (68% reduction; p < 0.001) in the group treated with GM-CSF plus TGF-β compared with those treated with GM-CSF alone (Fig. 5). The cells in the group treated with TGF-β were small and had few mature DC-like cells when examined in culture by light microscopy. In contrast, the number of fascin-positive DC was significantly (p < 0.001) increased at day 6 (by 12%) and at day 9 (26% increase; p < 0.001) in the group treated with GM-CSF plus TNF-α when compared with those treated with GM-CSF alone (Fig. 5). The number of CD11c-positive cells in the TGF-β (32%), GM-CSF alone (33%), and TNF-α (34%) groups were not statistically different (ns; p > 0.05), indicating that the overall numbers of DC were similar. Light microscopy examination demonstrated more mature DC-like cells in the group treated with TNF-α. This clearly demonstrates that enhancement of BM-DC maturation with TNF-α increased fascin expression. In contrast, suppression of BM-DC maturation with TGF-β markedly reduced fascin expression.

Similar patterns of MHC class II expression were observed on BM-DC when treated with these growth combinations (data not shown). These results provide further evidence linking fascin expression with DC maturity.

Upon maturation, DC increase their potency at stimulating naive T cell proliferation in a MLR assay (1, 4). Because our data correlated fascin expression with DC maturation, we examined whether the increase in fascin expression would be associated with enhanced allostimulatory activity.

BM-DC were generated in the presence of different growth factor combinations that we have shown above to alter fascin expression and BM-DC maturation. At day 9, BM-DC were harvested from the different groups and added in graded doses to a fixed number of naive allogeneic T cells. T cell proliferation was then assessed in a 4-day MLR assay. In all groups, there was an increase in T cell proliferation when DC numbers increased (Fig. 6). DC that were treated with GM-CSF were potent APC. However, the allostimulatory activity of BM-DC treated with GM-CSF plus TGF-β, which express a lower level of fascin, was markedly reduced. This reduction become more evident at a higher DC/T cell ratio, with a 91% (p < 0.001) reduction at 1:25 ratio. In contrast, the allostimulatory activity of BM-DC treated with GM-CSF plus TNF-α, which express a higher level of fascin, was increased by 37% (p < 0.001) at the same DC/T cell ratio as compared with those treated with GM-CSF alone. In addition, when we plotted the correlation between fascin-positive cells and the levels of T cell alloactivation in MLR, we found a strong correlation between fascin levels and T cell alloactivation with a correlation coefficient yield of 0.97. These data demonstrate a strong correlation between the level of fascin expression and T cell allostimulation.

The correlation between fascin expression and allostimulatory activity does not prove a role for fascin in Ag presentation because DC maturation is also associated with the up-regulation of MHC class II and costimulatory molecules. To isolate the effects of fascin on BM-DC allostimulatory activity, we used antisense oligonucleotides to inhibit fascin expression during maturation. Bone marrow precursor cells were seeded in cRPMI supplemented with GM-CSF alone, GM-CSF plus antisense oligo, or GM-CSF plus control oligo for 9 days, and then evaluated by MLR. BM-DC viability in the no oligo (97%), control oligo (95%), and antisense oligo (95%) treatment groups was not significantly different (ns; p > 0.05). The change in DC morphology following antisense oligo or control oligo treatment was assessed using double staining with anti-fascin and anti-CD11c Abs. Consistent with previous studies that demonstrated the role of fascin expression in dendrite formation, antisense oligo-treated BM-DC became smaller and had fewer dendrites, whereas control-treated DC had a normal morphology (Fig. 7). In the control oligo-treated group, 37 ± 1% of the CD11c⁺ cells were dendritic as compared with 6 ± 0.1% in the antisense oligo-treated group (p < 0.001). Fascin expression in the antisense oligo-treated groups was suppressed by 70%, as judged by image analysis, when compared with control oligo-treated group (Fig. 8,A). However, MHC class II and B7-2 expression on BM-DC were not significantly different (ns; p > 0.05) between control oligo- and antisense oligo-treated groups (Fig. 8, B and C).

In all groups, there was an increase in T cell proliferation with an increase in the number of DC (Fig. 9). We observed no significant difference (ns; p > 0.05) in the allostimulatory activity between DC that had been treated with GM-CSF alone and those treated with GM-CSF plus control oligo. However, the allostimulatory activity of BM-DC that were treated with the GM-CSF plus antisense oligo was markedly suppressed (p < 0.001) compared with those treated with the GM-CSF plus control oligo or GM-CSF alone. Similar results were observed for the fascin antisense experiments when we used a population that contained 90% BM-DC generated by a different protocol (33). These data demonstrate for the first time a direct role for fascin in the interaction between DC and T cells.

In this study, we have shown that fascin expression is critical in the Ag presentation activity of mature DC. By phenotypical and functional examinations, we observed that fascin expression is strongly correlated with DC maturity. Furthermore, we found that fascin expression plays a significant role in the function of DC as APC. The mechanisms by which fascin influences Ag presentation by DC remain to be determined.

Although DC have been described as the most potent professional APC, their function in this respect is critically dependent on their degree of maturity (1, 4, 5). When DC-precursors leave the bone marrow they circulate in the blood and eventually reside in nonlymphoid tissues as immature DC. At these sites, especially at interfaces with the environment, immature DC are highly efficient in Ag uptake and processing but are poor in Ag presentation and T cell activation (7, 34). Upon maturation, DC migrate via the lymphatics to the secondary lymphoid organs and become extremely potent in Ag presentation, but poor in Ag uptake and processing (9, 10, 11). The increase in the Ag presentation ability of mature DC is associated with up-regulation of a variety of molecules, including MHC class II and B7-2, the major triggers for the initiation of T cell activation (35, 36, 37, 38, 39). DC express higher levels of MHC class II and costimulatory molecules than other professional APC. However, this alone does not account for their greater potency in activating Ag-dependent immune responses (8, 31).

It has recently been recognized that other features unique to DC enhance their ability to present Ag to T cells. All of these functions are dependent on the state of maturation of the DC. Mature DC retain MHC class II peptide complexes on their surface for prolonged periods in culture, whereas other APC have a turnover measured in hours (40, 41). Migration of DC from the periphery into the lymph node is closely regulated during DC maturation through changes in chemokine receptor expression (42, 43, 44). In the lymphoid organs only mature DC produce high levels of the DC-CK1 chemokine, which preferentially attracts naive T cells (12). These findings indicate that the changes that occur during DC maturation play a significant role in their ability to act as potent APC.

DC also undergo a variety of morphological changes during maturation, including the development of numerous long dendrites (13, 14). Dendrites are a feature common to both neurons and DC, and both cell types express a cytoskeletal protein known as fascin (16, 20). Indeed, several studies have clearly demonstrated that fascin is important for the development of dendrites (16, 18, 19). If dendrite formation is linked to APC activity then one would expect a correlation between fascin expression, dendrite formation, and APC activity. We first confirmed that the DC in the IDC compartment of the lymph node, where APC activity is highest, were fascin positive. This has been previously shown in human IDC (20), and we have here confirmed it in mice. We then confirmed that LC in the skin, which are the classic immature DC, do not stain for fascin.

The in vivo demonstration of fascin expression in mature DC, and not in immature DC, suggests a link between maturation and fascin expression. To confirm this association we examined DC maturation, in vitro, from bone marrow precursors. We found that fascin was expressed in mature DC as evaluated by morphology. Fascin expression was also correlated with the up-regulation of MHC class II and B7-2. These data establish an association between fascin expression and maturation and a tentative link with APC activity. To directly relate this to APC activity of mature DC we performed MLR to assess APC activity under different conditions of reduced fascin expression. We found a strong correlation between the level of fascin and the ability of DC to activate T cells in MLR. More specifically, fascin antisense oligonucleotides effectively reduced fascin expression by 70%, and the cells treated with these oligonucleotides showed reduced alloactivation. This reduced allostimulatory effect was not due to changes in MHC class II or B7-2 expression or reduced DC viability. Therefore, fascin is another DC protein regulated during maturation that is critical for APC activity. These data provide the first evidence suggesting that dendrite formation plays a functional role in the interaction between DC and T cells.

Although we have shown that fascin expression is directly involved in enhancing the ability of DC to activate T cells, the exact mechanism underlying this process is not fully understood. There are a number of explanations that might account for this observation. Fascin expression resulting in dendrite formation may increase the DC surface area, and this may favor interaction with a greater number of T cells. However, it may also be a much more active process. Fascin expression in epithelial cells results in active extension of lamellipodia (19). Likewise, fascin might permit coordinated extension of dendrites maximizing the surface contact area between DC and T cells (15).

Cell polarity may be another mechanism by which cytoskeletal proteins influence APC-T cell interactions. Recently, it has been demonstrated that rearrangements of the T cell actin cytoskeleton result in clustering of TCR molecules, thereby enhancing TCR cross-linking (45). These changes result in sustained T cell signaling, which is an important step in T cell activation (46). Through its actin-bundling function fascin may induce a similar rearrangement of MHC molecules on the DC contributing to the immunological synapse that develops between APC and T cells. Cell polarity may also result in the directional secretion of cytokines by DC as has been demonstrated for T cells (47, 48). Finally, fascin is important in the motility and migration of cells (15, 19). Migration of DC to lymph nodes during their maturation is critical for the generation of the immune response, and this may be influenced by fascin expression.

In summary, this study clearly demonstrates that fascin is expressed in DC upon maturation. More importantly, it indicates that fascin expression in mature DC is critical for their generation of dendrites and their ability to activate T cells. This observation is the first evidence linking dendrite formation with the ability of DC to activate T cells. Although the exact nature of this interaction remains to be elucidated, further studies of the mechanisms that control fascin expression in DC may improve our understanding of the interaction between DC and T cells.

We thank Patricia Colp for her excellent technical assistance, and Robert Douglas and Lynn Thomas for their assistance on the use of the image analysis program.