Lymphocyte CD44 interactions with hyaluronan localized on the endothelium have been demonstrated to mediate rolling and regulate lymphocyte entry into sites of chronic inflammation. Because neutrophils also express CD44, we investigated the role of CD44 and hyaluronan in the multistep process of neutrophil recruitment. CD44−/− and wild-type control mice were intrascrotally injected with the neutrophil-activating chemokine, MIP-2, and leukocyte kinetics in the cremasteric microcirculation were investigated 4 h subsequently using intravital microscopy. Neither the rolling flux nor the rolling velocities were decreased in CD44−/− mice relative to wild-type mice. In vitro, neutrophils did not roll on the CD44 ligand hyaluronan, consistent with the in vivo data that CD44/hyaluronan did not mediate rolling. However, the number of adherent leukocytes in the venule was decreased by 65% in CD44−/− mice compared with wild-type mice. Leukocyte emigration was also greatly decreased in the CD44−/− mice. The same decrease in adhesion and emigration was observed in the wild-type mice given hyaluronidase. Histology revealed neutrophils as being the dominant infiltrating population. We generated chimeric mice that express CD44 either on their leukocytes or on their endothelium and found that CD44 on both the endothelium and neutrophils was important for optimal leukocyte recruitment into tissues. Of those neutrophils that emigrated in wild-type and CD44−/− mice, there was no impairment in migration through the interstitium. This study suggests that CD44 can mediate some neutrophil adhesion and emigration, but does not appear to affect subsequent migration within tissues.

Belonging to a highly heterogeneous family of hyaluronan-binding type I transmembrane glycoproteins, CD44 is present on a wide variety of cell types, including erythroid and myeloid cell lineages, fibroblasts, neurons, and endothelial cells (1). The family consists of some 20 different isoforms that are generated through differential splicing of the exons, with CD44H (hemopoietic), also known as CD44s (standard), being the most abundant form (2). Hyaluronan, a ubiquitously expressed polysaccharide, is the principal ligand for CD44 (2). Together, CD44 and hyaluronan may mediate a number of physiological and pathophysiological processes, including the inflammatory response (3, 4, 5, 6). The hallmark feature of acute inflammation is neutrophil infiltration into tissues; however, a role for neutrophil CD44 has not been elucidated.

Neutrophil recruitment into tissues particularly during acute inflammation has been well studied (7, 8). This paradigm involves a series of adhesion molecules that act in an interdependent fashion to allow fast flowing cells in the mainstream of blood to leave the circulation and enter the adjacent tissue. The selectins mediate the initial tethering and rolling event. The rolling process brings neutrophils into close proximity of chemokines on the surface of the endothelium, which activates the rolling leukocytes to firmly adhere via integrins and members of the Ig superfamily. Subsequently, the adherent neutrophils emigrate through the endothelial wall via PECAM-1, ICAM-1, and almost certainly other less well-established mechanisms (8, 9). CD44 has also been proposed to be an important adhesion molecule, but primarily in activated lymphocytes. Despite extensive expression on neutrophils, the role of CD44 on these cells is not well established.

The strategic position of the CD44 ligand, hyaluronan, on the endothelium (10) raises the possibility that CD44/hyaluronan could mediate neutrophil-endothelial cell interactions. Indeed, cross-linking of neutrophil CD44 induces cellular activation (11, 12), an important step in the recruitment of leukocytes in inflammation. Furthermore, on colorectal carcinoma cells, CD44 cross-linking has been shown to induce expression of the activation epitope of LFA-1 (CD11a/CD18), thus increasing adhesion to endothelium (13). CD44 has also been suggested to participate in the migration of cells through the interstitium (14, 15), with its relocation to the uropod during T cell chemotaxis (16). Recently, an in vitro study investigating the role of CD44 in neutrophil transmigration across epithelial monolayers showed that incubation of neutrophils with either an activating CD44 Ab or hyaluronic acid impaired transmigration possibly due to activation of adenylate cyclase, a negative regulator of neutrophil emigration (14). In support of this, another study demonstrated that CD44-deficient (CD44−/−) neutrophils crawl more quickly in hyaluronan-containing Matrigel (15). Thus, both these sets of results suggest a negative regulatory role for CD44 in emigration/motility. Moreover, in vivo, CD44−/− neutrophils accumulated in the lung to a much greater extent than wild-type (WT)3 neutrophils in response to Escherichia coli, but not Streptococcus pneumonia (15). Clearly, a comprehensive assessment of neutrophil CD44 is required to understand its role in neutrophil recruitment in vivo.

We systematically assessed the importance of CD44 in the recruitment of neutrophils, by using a neutrophil-specific chemokine, MIP-2, and intravital microscopy to visualize neutrophil rolling, adhesion, emigration, and migration through the interstitium in WT and CD44−/− mice. Because CD44 can be expressed on both neutrophils and the endothelium, we also made chimeric mice lacking CD44 on neutrophils or endothelium using bone marrow transplantation. Our data do not support the view that CD44 is important in neutrophil migration through the interstitium, but suggest that neutrophil CD44 is important for neutrophil migration across the endothelium. There was also decreased neutrophil adhesion, but this was dependent primarily upon endothelial CD44.

B6/129 mice deficient in CD44 (CD44−/−) were obtained from The Jackson Laboratory (Bar Harbor, ME). For controls, the WT B6/129 mouse strain was used from our own colony housed at the University of Calgary Animal Resource Center. All mice weighed between 20 and 35 g and were between 6 and 10 wk of age at the time of use. The animals were anesthetized with an i.p. injection of a mixture of 10 mg/kg xylazine (MTC Pharmaceuticals, Cambridge, Canada) and 200 mg/kg ketamine hydrochloride (Rogar/STB, Montreal, Canada). For all protocols, the right jugular vein was cannulated to administer additional anesthetic, if necessary. All experimental procedures were approved by the University of Calgary Animal Care Committee and conform to the guidelines established by the Canadian Council for Animal Care.

Briefly, bone marrow chimeras were generated following a standard protocol (17). B6/129 and CD44-deficient B6/129 mice were used as donors and/or recipients in bone marrow transplant experiments. Bone marrow was isolated from donor mice euthanized by spinal cord dislocation. Recipient mice were irradiated with 2 doses of 5 Gy (Gammacell 40 137Cs γ-irradiation source), with an interval of 3 h between the first and second irradiations. Cells (8 × 106) of the donor bone marrow were injected into the tail vein of recipient, irradiated mice. In the following 8 wk, the mice were kept in clean, germfree microisolator cages to allow full humoral reconstitution. Preliminary work confirmed that ∼99% of cells were from donor bone marrow, as assessed using Thy-1.1 and Thy-1.2 congenic C57B6 mice. C57B6 and 129Sv share major histocompatibility complexes; nevertheless, control B6/129 donor bone marrow was transplanted into B6/129 recipients to ensure that no untoward effects of transplant occurred. Inflammatory responses were identical with nontransplanted mice.

The mouse cremaster preparation was used to study the behavior of leukocytes in the microcirculation and adjacent connective tissue, as previously described (18). Briefly, an incision was made in the scrotal skin to expose the left cremaster muscle, which was then carefully removed from the associated fascia. A lengthwise incision was made on the ventral surface of the cremaster muscle using a cautery. The testicle and the epididymis were separated from the underlying muscle and were moved into the abdominal cavity. The muscle was then spread out over an optically clear viewing pedestal and was secured along the edges with 4-0 sutures. The exposed tissue was superfused with warm bicarbonate-buffered saline (pH 7.4). An intravital microscope (Axiolskip; Carl Zeiss, Don Mills, Canada) with ×25 objective lens (Weltzar L25/0.35; E. Leitz, Munich, Germany) and ×10 eyepiece was used to examine the cremasteric microcirculation. A video camera (5100 HS; Panasonic, Osaka, Japan) was used to project the images onto a monitor, and the images were recorded for playback analysis using a conventional videocassette recorder or time-lapse videocassette recorder.

Six single unbranched cremasteric venules ranging in diameter from 25 to 40 μm were selected per cremaster, and leukocyte kinetics were recorded for 2 min/vessel. The number of rolling, adherent, and emigrated leukocytes was determined off-line during video playback analysis. Rolling leukocytes were defined as those cells moving at a velocity of less than the erythrocytes in a given vessel. Leukocyte rolling velocity was determined by measuring the time required for a leukocyte to roll along a 100-μm length of venule. Rolling velocity was measured for 10 leukocytes entering the field of view over 2 min. The flux of rolling cells was measured as the number of rolling cells passing a given point in the venule per minute. A leukocyte was considered to be adherent if it remained stationary for at least 30 s, and the total adherent leukocytes were defined as the number of adherent cells within a 100-μm length of venule. The numbers of emigrated leukocytes were quantified as the average of the total cells counted in the field of view (a region of ∼200 × 300 μm), on either side of the vessel under study.

For each experiment, 1 μg of murine rMIP-2 in 0.2 ml of saline was administered locally by s.c. injection beneath the right scrotal skin using a 30G needle, 4 h before exteriorization. In some experiments, mice were i.p. administered with 20 U/g hyaluronidase (19). The left cremaster was then prepared for intravital microscopy. Leukocyte kinetics were investigated, as described above, and at the end of each experiment whole blood was drawn by cardiac puncture. Total leukocyte counts were performed using a Bright-line hemocytometer (Hausser Scientific, Horsham, PA).

An agarose gel containing MIP-2 (R&D Systems, Minneapolis, MN) was used to induce chemotaxis in the cremaster preparation. The agarose gel was prepared, as previously described in our laboratory (20), and carefully placed on the surface of the cremaster in a preselected avascular area, 350 μm (two television monitor screens wide) from a postcapillary venule. The gel was held in place using a coverslip, and the tissue was superfused with bicarbonate buffer (pH 7.4) beneath the coverslip at a minimum rate (0.5–0.7 ml/min) so as not to disrupt the chemotactic gradient established adjacent to the agarose pellet. The image was recorded for 120 min. Leukocyte rolling velocity, flux, adhesion, and emigration were recorded for 5 min at various intervals under normal time, and in between, by time-lapse video recording. The velocity of chemotaxis for each leukocyte was calculated from time-lapse recordings. Leukocyte responses induced by this directional inflammatory cue were compared between WT and CD44-deficient mice.

The cremaster was removed quickly after intravital microscopy and placed in 10% neutral buffered Formalin (Sigma-Aldrich, St. Louis, MO). The tissue was embedded in paraffin, sectioned (3 μm), and stained with H&E. Section analysis was performed under air and oil emersion to identify the types of leukocytes infiltrating the tissue.

Hyaluronan was localized in tissue, as previously described (21). Cremaster tissue was collected from mice treated with MIP-2 only, or mice that had been pretreated with hyaluronidase type IV (Sigma-Aldrich) made in PBS, pH 6.7 (Sigma-Aldrich). The tissue was embedded in paraffin, sectioned (3 μm), and incubated with biotinylated hyaluronan-binding protein (HABP, 10 μg/ml in PBS; Seikagaku Kogyo, Tokyo, Japan) overnight at 4°C. After washing, they were then incubated with peroxidase-labeled ABC avidin-biotin complex and developed with Sigma Fast diaminobenzidine chromogen solution (Sigma-Aldrich).

Rooster comb hyaluronan (Sigma-Aldrich) was immobilized on plastic by placing 100 μl of hyaluronan in PBS (2.5 mg/ml) on a 35-mm cell culture dish (Corning Glass, Acton, MA), which was then left uncovered and incubated overnight at 37°C to allow the solution to dry. VCAM-1 was immobilized by incubating 100 μl of VCAM-1 in ddH2O (5 μg/ml; R&D Systems) overnight at 4°C. In this case, plates were kept moist to prevent drying. All plates were blocked with 1% BSA (Sigma-Aldrich) in PBS for 1 h at 37°C before use. The parallel plate flow chamber assay was performed, as previously described (16, 22). Briefly, hyaluronan- or VCAM-1-coated plates were placed into a polycarbonate chamber with parallel plate geometry. Freshly collected murine whole blood was diluted 1/10 in HBSS (Invitrogen Life Technologies, Burlington, Canada). Using a syringe pump (Harvard Apparatus, St. Laurent, Canada), diluted blood or BW5147 T lymphoma cells transfected to express the TCR (23) at 5 × 105 cells/ml in DMEM supplemented with 10% FCS, 1 mM sodium pyruvate, and 2 mM l-glutamine (Invitrogen Life Technologies) were then perfused across the plate at a constant rate to mimic physiological shear forces encountered in the vasculature (1 dyne/cm2). BW S147 T cells were used as a positive control for CD44-hyaluronan interactions. After 5 min, leukocyte/substratum interactions were visualized using phase contrast microscopy on an inverted microscope fitted with a digital video camera. All experiments were recorded for later analysis. Rolling cells were defined as those traveling slower than free-flowing cells. Adherent cells were defined as those remaining stationary for at least 10 s. To verify the specificity of the interaction of BW5147 cells with hyaluronan, BW5147 cells resuspended at 5 × 105 cells/ml in DMEM, as described above, were incubated with 10 μg/ml KM201 (anti-CD44; American Type Culture Collection, Manassas, VA) for 30 min at 4°C before perfusion over hyaluronan.

Murine neutrophils were isolated from bone marrow. Blood was collected from anesthetized mice by cardiac puncture with a heparinized syringe. Mice were then euthanized, and the femurs and tibias were removed. The ends of the bones were resected, and the bone marrow was removed by perfusion of 5 ml of ice-cold PBS. The bone marrow was then suspended by drawing it through a 20-gauge needle. Marrow cells were then pelleted in a centrifuge (250 × g, 4°C, 12 min) and resuspended in 2 ml of PBS. The cell solution was placed over a discontinuous Percoll gradient consisting of a stock Percoll solution (90 ml of Percoll, 10 ml of 10× HBSS) diluted to 72, 64, and 52% in HBSS. The cell solution was spun at 1100 × g, 4°C, for 30 min. Purified murine neutrophils localized to a band between the 72 and 64% layers. This band was removed with a transfer pipette, washed in PBS, and suspended in murine plasma at 1.0 × 107 cells/ml. The heparinized blood was centrifuged at 500 × g, 4°C, for 15 min. The clear layer of plasma was removed using a transfer pipette and stored at 4°C until needed.

The under-agarose assay was performed, as described previously, with minor modifications (12, 24). The 35 mm × 10-mm Falcon petridishes (BD Biosciences, Mississauga, Canada) were filled with 3 ml of a 0.45% agarose solution containing 50% H2CO3-buffered HBSS (Sigma-Aldrich) and 50% RPMI 1640 (Invitrogen Life Technologies) culture medium containing 20% heat-inactivated FCS (Sigma-Aldrich). After the agarose solidified, three wells 3.5 mm in diameter and 2.4 mm apart were cut in a straight line into the gel. The gels were equilibrated for 1 h in a 37°C/5% CO2 incubator. A total of 12.5 pM (10 μl of a 1 mg/ml solution) murine rMIP-2 was added to the central well of the gels, and 1.0 × 105 neutrophils were loaded into the outer wells of the gel. Gels were incubated for 3 h in a 37°C/5% CO2 incubator. During this period of time, the neutrophils migrated toward the chemoattractant-containing well. Results were recorded at ×20 magnification using a video camera attached to a Zeiss Axiovert 135 microscope.

Mouse primary lung endothelial cells were isolated from 5- to 7-day-old CD44−/− and control B6/129 WT mice, according to the protocols described (25). Using this protocol with Tie2-GFP mice and flow cytometry, we verified that ∼93–98% of the isolated cells were GFP positive, confirming that the majority of the purified cells were of endothelial cell origin (26). Freshly isolated mouse endothelial cells were cultured in microvascular endothelial cell medium-2 (Clonetics EGM-2MV BulletKit; Cambrex Bio Science, Walkersville, MD) in 35-mm petridishes precoated with mouse laminin (20 μg/ml; Upstate Biotechnology, Lake Placid, NY). After reaching confluence in 5–6 days, the cells were lysed and used for Western blotting.

Endothelial cell lysates were prepared using Laemmli buffer with 10% 2-ME, 10 μg/ml leupeptin, and 10 μg/ml aprotinin. The proteins were separated by electrophoresis in 10% SDS-polyacrylamide gels, transferred to a polyvinylidene difluoride Hybond-P transfer membrane (Amersham Biosciences, Little Chalfont, U.K.), and blotted using a specific rat mAb against mouse CD44 (at 1 μg/ml; Pierce, Woburn, MA). After washing, the membrane was incubated with a secondary, HRP-conjugated goat anti-rat IgG (Pierce) and treated with ECL reagents (ECL kit from Amersham Biosciences). The blotted bands were detected with high performance autoradiography films from Amersham Biosciences.

All data are shown as means ± SEM. Student’s t test was used to determine the significance of differences between population means.

Fig. 1,A is a representative flow cytometry plot demonstrating that neutrophils from WT mice express ample CD44, whereas neutrophils from CD44−/− mice do not express CD44 to a greater degree than the isotype control primary Ab or the secondary Ab alone. We next investigated the presence/absence of CD44 on endothelium of WT and CD44−/− mice. Murine endothelia were isolated, grown to confluence, and Western blotted for the presence of CD44, as described in Materials and Methods. As shown in Fig. 1 B, a single band at 79–80 kDa was seen in the WT, but absent from the CD44−/− endothelia. Thus, we confirmed the presence of CD44 in WT neutrophils and endothelium, but not in CD44-deficient mice.

The model of acute inflammation that was used involved intrascrotal administration of MIP-2 chemokine for 4 h. To identify the type of cells that had been recruited to the site of inflammation at this time point, H&E staining was performed on each tissue after the experiment, as described in Materials and Methods. As can be seen in Fig. 2,A, there was no neutrophil extravasation in WT tissues intrascrotally injected with saline. In contrast, neutrophils could clearly be seen emigrating from the postcapillary venules (V) of WT mice given MIP-2 injection (Fig. 2,B), while very few neutrophils could be seen emigrating from the venules of CD44−/− mice (Fig. 2,C). This emigration pattern was also noted when a MIP-2 gel was placed 350 μm from a postcapillary venule and chemotaxis was examined. Neutrophils could clearly be seen emigrating across the vessel endothelium and migrating in one direction (bottom) toward MIP-2 gel (Fig. 2,D), while very few neutrophils could be seen in CD44−/− mice (Fig. 2, E and F). However, those few CD44−/− neutrophils that did emigrate outside the vasculature moved through the interstitium very effectively (Fig. 2, C, E, and F). Leukocyte analysis of the tissues confirmed that the cellular infiltrate consisted of 99% neutrophils, characterized by their multilobed nuclei and 1% lymphocytes. No macrophages could be identified in the tissues studied.

To further understand how CD44 deficiency could result in a very significant reduction in the number of extravasating neutrophils, a systematic in vivo analysis of the cremasteric microcirculation was performed. Venules from CD44−/− and WT mice were chosen that ranged in diameter from 28 to 34 μm. None of the basal parameters was different between CD44−/− mice and WT mice. To rule out any effects of differences in circulating leukocyte numbers, leukocyte counts were performed on all mice used in this study. These analyses revealed that circulating leukocyte counts did not differ significantly between CD44−/− (5.3 ± 0.5 × 106/ml) and WT (5.7 ± 0.7 × 106/ml) mice (Fig. 3,A). Interestingly, as demonstrated in Fig. 3,B, leukocyte rolling flux was not affected by 4-h intrascrotal MIP-2. Although activated lymphocytes have been shown to roll via CD44 (27), the general population of rolling leukocytes was not reduced in CD44−/− mice relative to WT mice. In fact, the number of rolling leukocytes was increased by ∼30% in the mutant mice. Rolling velocities were not significantly different between CD44−/− and the WT mice (Fig. 3 C).

To investigate whether hyaluronan could be an active participant in leukocyte rolling in vivo, WT mice were treated with 20 U/g hyaluronidase for 2 h (before administration of MIP-2 intrascrotally), a protocol previously shown to deplete hyaluronan from endothelium (19). The effectiveness of this treatment was verified by investigating the presence of hyaluronan in the tissues after the experiment. Tissue sections from MIP-2-treated mice or mice treated with both MIP-2 and hyaluronidase were stained with HABP. Hyaluronan was localized within the connective tissue of the muscle and surrounding blood vessels (Fig. 4). When compared with tissues from untreated mice (Fig. 4, A and B), staining was less intense in tissues taken from mice treated with hyaluronidase (Fig. 4, C and D). This confirmed the effectiveness of the enzymatic procedure in removing hyaluronan from the tissue.

Despite the removal of hyaluronan, we still found leukocyte rolling flux not to be significantly affected in the hyaluronidase-treated mice (40.1 ± 12.8 cells/min), suggesting no involvement of hyaluronan in mediating leukocyte rolling in our model of acute inflammation (Fig. 3, B and C).

In a separate complementary series of in vitro experiments, we examined whether neutrophils could roll via CD44. Hyaluronan, the CD44 ligand previously demonstrated to support activated lymphocyte rolling under flow conditions (27), was immobilized in 35-mm tissue culture dishes. Under flow conditions (shear forces as low as 1 dyne/cm2), hyaluronan was not capable of tethering or supporting neutrophil rolling or, for that matter, any of the general pool of leukocytes. This was not due to poor binding by hyaluronan to the coverslip, because the BW5147 murine T cell line rolled (Fig. 5,A) and adhered very well on hyaluronan and could be inhibited by anti-CD44 Ab (Fig. 5 B). Moreover, as a positive control leukocytes rolled on immobilized VCAM-1 (data not shown).

We next investigated whether the presence or absence of CD44 affected neutrophil adhesion in response to MIP-2. Neutrophil adhesion was significantly increased compared with control 4 h following intrascrotal injection of MIP-2. In contrast, this increase in adhesion was not observed in CD44−/− mice (Fig. 6 A). On average, the number of leukocytes adherent to postcapillary venules after 4-h exposure to MIP-2 chemokine was 18.9 ± 2.3 cells in the WT mice compared with 6.9 ± 2.9 cells in CD44−/− mice or ∼2.7-fold decrease in adhesion. Pretreatment of WT mice with an i.p. injection of hyaluronidase 2 h before intrascrotal injection of MIP-2 also reduced neutrophil adhesion (9.6 ± 2.8 cells), clearly demonstrating for the first time that CD44 and hyaluronan contribute to neutrophil adhesion in vivo.

Similar effects on neutrophil emigration were also observed. Very few cells were observed in the tissue of untreated WT mice (0.4 ± 0.4 cells/field; Fig. 6 B). MIP-2 induced a profound increase in neutrophil emigration over 4 h (103.7 ± 19.6 cells/field). Emigration was also observed in CD44−/− mice, but it was less than half of that observed in WT mice (44.7 ± 10.2 cells/field). Again, pretreatment with hyaluronidase also prevented leukocyte emigration (22.7 ± 8.9 cells/field). Intraperitoneal administration of hyaluronidase did not reduce adhesion or emigration in CD44−/− mice, implying potentially a similar mechanism of action (data not shown).

To determine whether CD44 deficiency impaired the ability of emigrated neutrophils to migrate through the tissue, a separate series of experiments was performed in which MIP-2 was administered intrascrotally to WT or CD44−/− mice and then leukocyte migration away from venules was analyzed 4 h later. As observed in Fig. 6,B, MIP-2 induced profound leukocyte emigration into the tissue in WT mice. These cells migrated through the tissue and away from the blood vessel, with the largest number of cells observed within the first 25 μm from the vessel and progressively fewer cells further out (Fig. 7,A). In comparison, fewer migrated cells were observed in each quadrant in CD44−/− mice, but this may simply reflect the overall reduction in total leukocyte emigration rather than a defect in cell migration per se. Indeed, when the data were reanalyzed to reflect the percentage of total emigrated cells within each quadrant, except for a small difference in the closest quadrant to the blood vessel, migration away from the blood vessel was identical in WT and CD44−/− mice (Fig. 7 B).

CD44 has previously been implicated in neutrophil and cancer cell chemotaxis in vitro using various artificial substrata (14, 28). Intrascrotal administration of MIP-2 does not reflect true chemotaxis, as there is no direction to the chemotactic stimulus. Therefore, to determine whether CD44 deficiency impairs the ability of neutrophils to chemotax to MIP-2 in vivo, a final series of experiments was performed in which an agarose gel containing MIP-2 was placed 350 μm from a postcapillary venule. After 2 h, directional migration of emigrated leukocytes toward the gel was assessed. Too few emigrated cells were observed in CD44−/− mice for an accurate analysis of migration distance (data not shown). However, when migration velocity toward MIP-2 was assessed for individual cells, this was observed to be identical in WT and CD44−/− mice (Fig. 7 C). In contrast, using an in vitro under-agarose assay of neutrophil chemotaxis (29), we observed a distinct inability of CD44−/− neutrophils to chemotax toward MIP-2 (data not shown). This apparent contradiction with the in vivo data could be attributed to the coverslip lacking the complex milieu of extracellular matrix proteins that would be present in the in vivo scenario and would allow for compensation for CD44.

CD44 is expressed on both leukocytes and endothelium. To determine whether endothelial or leukocyte CD44 was responsible for the decrease in adhesion and emigration observed in vivo, CD44 chimeric mice were generated via bone marrow transplantation. WT mice that received CD44−/− bone marrow were deficient in leukocyte CD44 (leukocyte CD44−/− mice), whereas CD44−/− mice that received WT bone marrow were deficient in endothelial CD44 (endothelial CD44−/− mice). No significant changes were observed in rolling flux between the chimeric and WT mice exposed to MIP-2 (Fig. 8,A). In addition, no changes were observed in the leukocyte rolling velocities between the chimeric and WT mice following MIP-2 exposure (data not shown), suggesting that the generation of chimeric mice did not have any general untoward effects on leukocytes or endothelium. Neutrophil adhesion was significantly reduced in mice whose endothelium lacked CD44 (8.0 ± 2.0 cells), which was similar to adherent neutrophils in CD44−/− mice (6.9 ± 2.9 cells) (Fig. 8 B). Surprisingly, mice deficient in only leukocyte CD44 had similar numbers of adherent neutrophils (14.9 ± 3.0 cells) compared with WT mice (18.5 ± 2.7 cells).

Although the leukocyte CD44−/− mice did not have significant impairment in neutrophil adhesion, Fig. 8,C illustrates that these mice clearly demonstrated a profound reduction in the ability to emigrate out of the vasculature (25.4 ± 9.3 neutrophils vs 103.8 ± 19.6 neutrophils in WT mice). Endothelial CD44−/− mice, which had shown reduced adhesion, also exhibited reduced emigration (Fig. 8 C) in response to MIP-2.

Numerous studies have indirectly implicated CD44 as an important molecule in neutrophil motility and recruitment (14, 22, 30). In this study, we have systematically examined the role of CD44 on neutrophils as a molecular mediator of the recruitment cascade. Rolling of the neutrophils in postcapillary venules was not mediated by CD44, and circulating neutrophils did not interact with the CD44 ligand, hyaluronan, in vitro. Our data also suggest a very limited role for CD44 in the migration of neutrophils through tissue in this in vivo model system. However, CD44 was found to play an important role in the adhesion of neutrophils to the endothelium, and this process required hyaluronan. Finally, the transmigration of neutrophils across the venular endothelium was dependent upon both hyaluronan and CD44. By making use of bone marrow transplantation to generate chimeric mice, we were able to demonstrate that both neutrophil CD44 and endothelial CD44 contributed to the phenotypes observed.

Previous investigators have shown that superantigen-activated T cells are induced to bind hyaluronan and extravasate to the peritoneum in a CD44- and hyaluronan-dependent manner (5). In vitro, activated T cells and T cell lines were shown to roll on hyaluronan-coated plates and on cultured endothelial cell lines that express CD44 and hyaluronan (10, 27). These data implicated both CD44 and hyaluronan on endothelial cells and CD44 on T cells, and suggested a sandwich model, whereby CD44 on the endothelial cell presents hyaluronan to the T cell, which has an active CD44 capable of binding hyaluronan. Our in vivo data would to some extent support this view for neutrophils. Both neutrophil adhesion and emigration were dependent upon CD44 as well as hyaluronan. Furthermore, using chimeric mice lacking CD44 on leukocytes or endothelium revealed that both sources of CD44 were necessary for neutrophil emigration out of the vasculature. This is entirely consistent with the sandwich model of a CD44-hyaluronan-CD44 interaction proposed previously for T cell extravasation.

Although some of the neutrophil adhesion was also CD44 dependent, only endothelial CD44 and hyaluronan were necessary for this process to occur. These data are not entirely consistent with the sandwich model described above; however, neutrophils have multiple mechanisms of adhesion. Indeed, neutrophils are known to use integrins including the β2-integrin-dependent mechanisms (9, 31), but under certain conditions, β2-integrin-independent mechanisms of adhesion also have been reported (reviewed in Ref.32). Clearly, a number of these molecules could replace the need for neutrophil CD44 to induce adhesion. Whether any of the CD44-independent neutrophil-adhesive mechanisms can bind hyaluronan remains unknown.

To date, integrins have been the dominant molecules for neutrophil recruitment. The mechanism of action by which CD44 contributes to neutrophil recruitment could be as an accessory molecule for integrins. For example, CD44 may influence integrin function: cross-linking of CD44 in cancer cells has been shown to activate LFA-1 (13). In T cells, CD44 has been shown to complex with the VLA-4 integrin, and this association is important for T cell extravasation to an inflammatory site (33). It is possible that similar CD44-integrin associations occur on neutrophils. Although we have no direct evidence for a CD44-integrin association, our in vitro under-agarose chemotaxis assay provides some indirect evidence. We have previously reported that in that system on a multiprotein substratum, β2-integrin was an essential molecule for neutrophil chemotaxis (29). Quite surprisingly, CD44−/− neutrophils were also defective in migration toward the chemokine in vitro. Clearly, this suggests that the absence of neutrophil CD44 can modulate integrin-linked migration. However, how this occurs remains unclear. It should also be noted that despite the profound migratory defect in vitro, we were not able to observe a similar defect in the rate of migration in the interstitium in vivo. One difference is the fact that neutrophils that emigrate have been shown to express β1-integrins (33), which may substitute for the absence of CD44 in vivo. In vitro, of course, the neutrophils were isolated from bone marrow and did not have the opportunity to emigrate.

Alternatively, the observed inhibitory effects on neutrophil recruitment in the absence of endothelial CD44 or hyaluronan may be unrelated to direct properties of adhesion. CD44 on the activated endothelium anchors and presents hyaluronan to passing cells. Furthermore, endothelial cell lines have been shown to express hyaluronan, which is increased under inflammatory conditions (34). This system as an integral component of extracellular matrices is responsible for binding large proteoglycans such as aggrecan and versican (8). It is well known that heparin sulfate-modified proteoglycans present on the surface of the endothelium can bind and help sequester chemokines at sites of inflammation (35, 36). Indeed, some alternatively spliced forms of CD44 can be modified by heparan sulfate and have been shown to bind the chemokine, MIP-1β (36). Because both endothelial CD44 and hyaluronan are required for neutrophil adhesion and emigration, it is possible that they both function to maintain the integrity of the proteoglycan environment, which is important for sequestering chemokines at the inflammatory site.

In conclusion, our data suggest that CD44 contributes to the acute inflammatory response as it pertains to neutrophil recruitment. We could not find a role for CD44 in either neutrophil rolling or migration through the interstitium, but did observe a role for neutrophil adhesion and transendothelial emigration. Although it has been appreciated for some time that the endothelium can express CD44, this is the first functional demonstration of this molecule in neutrophil adhesion and emigration.

We thank Lori Zbytnuik and Krista McRae for generating the chimeric mice; Carol Gzwod for her excellent advice and technical assistance during histology; and Pau Serra and Fusun Turesin for their help with flow cytometry.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by the Canadian Institutes of Health and Research Group Grant. P.K. is a Canada Research Chair and Alberta Heritage Foundation for Medical Research scientist.

3

Abbreviations used in this paper: WT, wild type; HABP, hyaluronan-binding protein.

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