Synthesis and Surface Expression of Hyaluronan by Dendritic Cells

Transkrypt

Synthesis and Surface Expression of
Hyaluronan by Dendritic Cells and Its
Potential Role in Antigen Presentation
This information is current as
of March 2, 2017.
Mark E. Mummert, Diana Mummert, Dale Edelbaum,
Francis Hui, Hiroyuki Matsue and Akira Takashima
J Immunol 2002; 169:4322-4331; ;
doi: 10.4049/jimmunol.169.8.4322
http://www.jimmunol.org/content/169/8/4322
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References
The Journal of Immunology
Synthesis and Surface Expression of Hyaluronan by Dendritic
Cells and Its Potential Role in Antigen Presentation1
Mark E. Mummert, Diana Mummert, Dale Edelbaum, Francis Hui, Hiroyuki Matsue, and
Akira Takashima2
H
yaluronan (HA)3 is a linear glycosaminoglycan composed of multiple copies of the disaccharide units of
glucuronic acid (GlcA) and N-acetylglucosamine
(GlcNAc; [␤-1,4-GlcA-␤-1,3-GlcNAc-]n). With ⬎10,000 disaccharide repeats (n ⫽ ⬎10,000), a HA molecule can be easily
⬎4000 kDa (with each disaccharide unit being ⬃400 Da) in its
molecular mass, showing an extended molecular size of ⬎10 ␮m.
About 50% of the HA in the body is found in the skin, where HA
serves as a major structural component of the dermal extracellular
matrix and the intercellular space in epidermis (1, 2).
Despite its chemical simplicity, HA exhibits diverse biological
functions, presumably reflecting its unique ability to bind to many
different proteins via a highly conserved domain known as the link
module. These proteins include CD44 (3, 4), link protein (5), aggrecan (6), versican (7), hyaluronectin (8), and neurocan (9). The
Department of Dermatology, University of Texas Southwestern Medical Center, Dallas, TX 75390
Received for publication April 18, 2002. Accepted for publication August 7, 2002.
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 National Institutes of Health Grants RO1AI46755,
RO1AR35068, RO1AR43777, RO1AI43232, and RO3AR47402.
2
Address correspondence and reprint requests to Dr. Akira Takashima, Department
of Dermatology, University of Texas Southwestern Medical Center, 5323 Harry
Hines Boulevard, Dallas, TX 75390-9069. E-mail address: akira.takashima@
utsouthwestern.edu
3
Abbreviations used in this paper: HA, hyaluronan; BM, bone marrow; bPG, bovine
proteoglycan; DC, dendritic cell; DNFB, dinitrofluorobenzene, FN, fibronectin; GlcA,
glucuronic acid; GlcNAc, N-acetylglucosamine; HAS, HA synthase; Hyal, hyaluronidase; LC, Langerhans cell; LYVE-1, lymphatic vessel endothelial HA receptor-1;
PEG, polyethylene glycol; PI, propidium iodide; RP, random peptide.
Copyright © 2002 by The American Association of Immunologists, Inc.
lymphatic vessel endothelial HA receptor-1 (LYVE-1) is a recently identified HA receptor that is expressed selectively on the
surface of lymph vessel endothelium (10, 11); this receptor also
contains a link module. The receptor for HA-mediated motility is
distinct from other HA-binding proteins in its binding to HA via a
non-Link module sequence (reviewed in Ref. 12).
Unlike conventional glycosaminoglycans that are synthesized in
the Golgi network, HA is synthesized at the inner face of the
plasma membrane, and the growing polymer is extruded through
the membrane to the outer surface as it is being synthesized. Three
HA synthases (termed HAS1 through HAS3) polymerize the disaccharide units by alternate addition of GlcA and GlcNAc to the
growing chain of HA polymers using UDP-GlcA and UDPGlcNAc as substrates. These enzymes differ from each other in
catalytic activities (HAS3 HAS2 HAS1) as well as in the sizes of
the final products. HAS1 and HAS2 polymerize long stretches of
GlcA-GlcNAc disaccharide chains, whereas HAS3 polymerizes
relatively short stretches (⬍300 kDa) (13, 14). HA polymers are
degraded into monosaccharides by three enzymatic reactions. Hyaluronidases (Hyal) degrade HA polysaccharides to oligosaccharides, which are further digested into GlcA and GlcNAc by ␤-Dglucuronidase and ␤-N-acetyl-D-hexosaminidase, respectively
(15). Four distinct Hyal have been identified: Hyal-1 through
Hyal-3 and the sperm-specific Hyal PH-20 (16 –19). These enzymes appear to exhibit different catalytic profiles; e.g., Hyal-1
cleaves HA polymers into small oligosaccharides, whereas Hyal-2
cleaves them into intermediate size fragments of ⬃20 kDa.
HA had long been recognized to serve primarily as a filling
material of extracellular spaces. This classic view was rapidly revised in 1990, when several groups reported that CD44 acts as a
cell surface receptor of HA (3, 4). Identification of two additional
0022-1767/02/$02.00
Hyaluronan (HA) is a large glycosaminoglycan consisting of repeating disaccharide units of glucuronic acid and N-acetylglucosamine. HA is known to act as a filling material of extracellular matrices and as an adhesive substrate for cellular migration.
Here we report that dendritic cells (DC) express mRNAs for HA synthases and hyaluronidases, actively synthesize HA, and display
HA on their surfaces. Interestingly, HA expression levels on DC were not significantly altered by their maturation states. With
respect to physiological function, three specific HA inhibitors, i.e., bovine proteoglycan, a 12-mer HA-binding peptide (GAH
WQFNALTVR) termed Pep-1, and an oligomeric Pep-1 formulation, all interfered with DC-induced activation of CD4ⴙ T cells
isolated from DO11.10 TCR transgenic mice. For example, Pep-1 oligomer efficiently inhibited DC-dependent cluster formation,
IL-2 and IFN-␥ production, and proliferation by DO11.10 T cells in vitro without affecting the viabilities of DC or T cells, DC
function to uptake exogenous proteins, or DC-T cell conjugate formation at earlier time points. These observations suggest a
paracrine mechanism by which DC-associated HA facilitates some of the late changes in T cell activation. Although T cells
constitutively expressed mRNAs for HA synthases and hyaluronidases, their surface HA expression became detectable only after
activation. Oligomeric Pep-1 and bovine proteoglycan both inhibited mitogen-triggered T cell activation in the absence of DC,
suggesting an autocrine mechanism by which HA expressed by T cells assists their own activation processes. Finally, adoptively
transferred DO11.10 T cells showed progressive mitosis when stimulated with Ag-pulsed DC in living animals, and this clonal
expansion was inhibited significantly by administration of Pep-1 oligomer. Our findings may introduce a new concept that
relatively simple carbohydrate moieties expressed on DC and perhaps T cells play an important immunomodulatory role during
Ag presentation. The Journal of Immunology, 2002, 169: 4322– 4331.
Materials and Methods
Animals and cells
Female BALB/c mice and DO11.10 transgenic mice (6 –10 wk old) were
used in this study. The phenotypic and functional features of XS52 and
XS106 DC lines were described previously (30, 31). Splenic DC and bone
marrow (BM)-derived DC were prepared from BALB/c mice as previously
described (32, 33). The BW5147 thymoma, B16-F10 melanoma, and
KM81 B cell hybridoma lines were purchased from American Type Culture Collection (Manassas, VA). CD4⫹ T cells were isolated from spleens
of DO11.10 transgenic mice (BALB/c background) by positive selection
with Dynabeads (Dynal, Lake Success, NY), followed by depletion of IA⫹
cells with MACS microbeads (Miltenyi Biotec, Auburn, CA). CD3⫹ T
cells were isolated from spleens of BALB/c mice using a T cell isolation
column (R&D Systems, Minneapolis, MN), followed by depletion of IA⫹
cells. All animal experiments were approved by the institutional review
board at University of Texas Southwestern Medical Center and conducted
according to the guidelines from the National Institutes of Health.
Reagents and Abs
OVA peptide 323–339 was synthesized at the Biopolymers Facility, University of Texas Southwestern Medical Center. OVA protein was purchased from Sigma-Aldrich (St. Louis, MO). LPS (Escherichia coli 026:
B6) and Con A were obtained from Sigma-Aldrich and Amersham
Pharmacia Biotech (Piscataway, NJ), respectively. The mAb KJ1.26,
which recognizes the transgenic TCR expressed by DO11.10 T cells, and
its isotype-matched control IgG were purchased from Caltag (Burlingame,
CA). Other mAbs were purchased from BD PharMingen (San Diego, CA).
None of the above reagents, except for LPS, contained detectable amounts
of endotoxin as tested by the OCL-100 system (BioWhittaker,
Walkersville, MD).
tions), the cells were incubated for 10 min on ice in a hypotonic buffer (10
mM KCl, 1.5 mM MgCl2, 10 mM Tris-HCl, 1% 2-ME, 1 ␮g/ml leupeptin,
and 1 mM PMSF, pH 7.4) and harvested with a rubber policeman. The
cellular fractions were then freeze-thawed twice, homogenized, and centrifuged at 400 ⫻ g for 5 min to remove nuclear materials. The supernatants
were further centrifuged at 25,000 ⫻ g for 60 min to obtain pellets (membrane fraction) and supernatants (cytoplasmic fraction). High m.w. materials were precipitated from each fraction with 1% cetylpyridinium chloride in 0.02 M NaCl in the presence of carrier HA (200 ␮g/ml) and then
digested with papain (1 U/ml; Sigma-Aldrich) for 90 min at 60°C. After 10
min boiling, one-half of each sample was digested with Streptomyces Hyal
(80 U/ml; Sigma-Aldrich) for 16 h at 37°C, while the other half was mockdigested. Hyal-resistant glycosaminoglycans were precipitated as described above, and the pellets were dissolved in 4.0 M guanidine hydrochloride, ethanol precipitated, and measured for radioactivity. The amount
of newly synthesized HA in each fraction was then calculated by subtracting the Hyal-resistant radioactivity from the total radioactivity.
Surface HA expression
To examine surface HA expression, DC preparations were incubated for 30
min on ice with biotinylated Pep-1 (100 ␮g/ml), washed extensively, and
labeled with PE-conjugated streptavidin. For splenic DC and BM-derived
DC preparation, Pep-1 binding was analyzed within CD11c⫹ populations.
To test the specificity of Pep-1 binding, samples were pretreated with Hyal
or heparinase III (Sigma) before incubation with biotinylated Pep-1. The
carbohydrate specificities of the two enzymes was determined by measuring their abilities to release biotinylated HA polymers that had been immobilized onto the Amine CovaLink plates (Nalge Nunc International,
Rochester, NY). In some experiments BM-derived DC were tested for
Pep-1 binding and CD40 and CD86 expression after 16-h incubation in the
presence or the absence of 100 ng/ml LPS. CD3⫹ T cells isolated from
BALB/c mice were examined for Pep-1 binding after 16-h incubation in
the presence or the absence of 4 ␮g/ml Con A.
Preparation of HA inhibitors
Pep-1 (GAHWQFNALTVR) and a control peptide (SATPASAPYPLA),
termed random peptide (RP), were synthesized with the amidated GGGS
linker sequence at the C terminus as previously described (29). To prepare
oligomeric formulations, a 40-fold molar excess of Pep-1 or RP was incubated for 7 days with the tetravalent polyethylene glycol (PEG) derivative, bis[polyethylene bis(imidazolyl carbonyl)] at 37°C in DMSO. Triethylamine (0.5%) was added to the reaction to ensure maximum
nucleophilicity of the peptide ␣-amine. Reactions were terminated by dialysis against dH2O or PBS, and the density of substitution was estimated
by the bicinchoninic acid assay (to determine peptide content) and dry
weight analysis. Bovine proteoglycan (bPG) from nasal cartilage (ICN Biomedicals, Costa Mesa, CA) was dissolved in PBS. Again, none of the HA
inhibitors contained detectable amounts of endotoxin.
Ag presentation assays
The following primer sets were used: 5⬘-CTTTCAAGGCACTGGGC
GAC-3⬘ and 5⬘-CACCGCTTCATAGGTCATCC-3⬘ (first reaction for
HAS1),5⬘-GGAAAGCTTGACTCAGACACAAGAC-3⬘and5⬘-AGGGAA
TTCGTATAGCCACTCTCGG-3⬘ (second reaction for HAS1), 5⬘-TGG
AACACCGGAAAATGAAGAAG-3⬘ and 5⬘-GGACCGAGCCGTGTAT
TTAGTTGC-3⬘ (HAS2), 5⬘-CCATGAGGCGGGTGAAGGAGAG-3⬘ and
5⬘-ATGCGGCCACGGTAGAAAAGTTGT-3⬘ (HAS3), 5⬘-TATCCAAC
CGGCCATTCATCACTG-3⬘ and 5⬘-ATACCCCGCTTGTCACACCACT
TG-3⬘ (Hyal-1), 5⬘-CATCTTCACTGGCCGACCCTTTGT-3⬘ and 5⬘-TCG
CCACCCCAGCCCAGATAGC-3⬘ (Hyal-2), and 5⬘-CCTAGGCCTAATGA
TGGTG-3⬘ and 5⬘-GCTAGTATGGGCTTTGTGG-3⬘ (Hyal-3). PCR products after 35 cycles of amplification with an annealing temperature of 53°C
(for Hyal-3) or 57°C (for others) were separated by electrophoresis through
1.2% agarose and visualized with ethidium bromide.
Immunoregulatory activities of HA inhibitors were tested using an in vitro
Ag presentation system in which CD4⫹ T cells (2.5 ⫻ 105 cell/ml, ⬎95%
CD4⫹ cells) purified from DO11.10 TCR transgenic mice and gammairradiated splenic DC or BM-derived DC preparations (2.5 ⫻ 104 cells/ml,
⬎70% CD11c⫹ cells) isolated from BALB/c mice were cocultured in the
presence or the absence of 2 ␮g/ml OVA323–339 peptide. T cell proliferation was measured by [3H]thymidine uptake on day 3 as previously described (30, 31, 33). Viabilities of DC and T cells were examined by propidium iodide (PI) uptake by the CD11c⫹ and KJ1.26⫹ populations,
respectively. In some experiments DO11.10 T cells were labeled with
CFSE (Molecular Probes, Eugene, OR) and examined for their fluorescence profiles on day 2. Cytokine production was examined by ELISA on
day 1 (33). To study DC-T cell conjugate formation, BM-derived DC labeled with a red fluorescence dye PKH-26 (Sigma-Aldrich) and CSFElabeled DO11.10 T cells were incubated for 30 min in the presence or the
absence of OVA peptide, followed by FACS analyses in which DC-T cell
conjugates were detected as double-positive events (34). In vivo effects of
HA inhibitors were examined in adoptive transfer experiments. Briefly,
BALB/c mice received i.p. injections of CFSE-labeled DO11.10 T cells
(106 cells/mouse), OVA peptide-pulsed BM-DC (3 ⫻ 105cells/mouse), and
oligomeric Pep-1 or RP (20 ␮g/mouse), and the cells recovered from the
peritoneal cavity on day 2 were examined for CFSE profiles.
HA biosynthesis in DC
Other functional assays
To study HA synthesis, XS106 DC were labeled for 16 h with [3H]glucosamine (25 ␮Ci/ml). After collecting culture supernatants (secreted frac-
To study potential roles of T cell-associated HA, CD3⫹ T cells isolated
from BALB/c mice were stimulated by Con A or the combination of anti-
RT-PCR analyses
surface HA receptors (receptor for HA-mediated motility and
LYVE-1) has further supported this concept (10, 11, 20). More
recently, degradation products of HA ranging from 1.5 kDa (hexasaccharides) to 500 kDa have been shown to trigger the secretion
of various cytokines and chemokines by macrophages (21–24).
Thus, HA may potentially regulate diverse cellular functions. On
the other hand, due to the unavailability of specific inhibitors, the
physiological functions of HA have been mostly deduced from the
biological changes caused by HA receptor antagonists, such as
anti-CD44 mAb and CD44-Fc fusion proteins. This approach,
however, may not be legitimate, especially from the viewpoint that
CD44 also binds other molecules, including collagens, fibronectin,
chondroitin sulfates, heparin, heparan sulfate, and serglycins (25–
28). As an initial step toward studying HA function more directly,
we have recently developed a 12-mer peptide inhibitor of HA using the phage display technique; this peptide, termed Pep-1, binds
to HA in soluble, immobilized, and cell surface-associated forms
and inhibits HA-mediated cell adhesion and migration (29). Taking advantage of this unique inhibitor, we now test the hypothesis
that HA may regulate Ag presentation processes.
4323
4324
CD3 mAb and anti-CD28 mAb as described previously (33). The ability of
DC to uptake exogenous proteins was examined using FITC-conjugated
BSA (Sigma-Aldrich). Briefly, BM-derived DC were incubated for 10 min
with FITC-BSA and then examined for FITC profiles within the CD11c⫹
populations. Cell adhesion was examined as described previously (29).
Con A-activated splenic T cells and B16-F10 melanoma cells were labeled
with [35S]methionine/cysteine (ICN Biomedicals) and incubated for 30 min
at room temperature on Nunc plates coated with HA or for 60 min at 37°C
on tissue culture plates coated with bovine fibronectin (FN; SigmaAldrich). After removal of nonadherent cells, the adherent cells were solubilized in 1% SDS and counted for radioactivity. The percentages of
adherent cells were calculated by dividing the recovered counts per minute
by the total counts per minute added to each well.
Statistical analyses
Comparisons between two groups were performed with Student’s twotailed t test, and more than two groups were compared by ANOVA. Each
experiment was repeated at least twice to assess reproducibility.
Results
HA synthesis in DC
shown). To study the subcellular distributions of newly synthesized HA, we measured Hyal-sensitive radioactivities in the membrane fractions, cytoplasmic fractions, and culture supernatants.
Consistent with the current idea that HA polymer is synthesized at
the inner face of the plasma membrane and then extruded to the
cell surface, most of the newly synthesized HA molecules were
recovered from the membrane fractions (Fig. 1B). These results
suggest that DC actively synthesize HA and perhaps express the
resulting HA polymer at the outer surface.
Detection of surface HA expression on DC and T cells
Based on our previous observation that Pep-1 binds to HA molecules in soluble, immobilized, and membrane-associated forms
(29), we thought that Pep-1 might serve as a unique tool to study
HA expression on DC. Pep-1 in a biotinylated form showed
marked binding to the surfaces of XS106 DC (Fig. 2A). By contrast, a 12-mer control peptide termed RP showed no detectable
binding. Moreover, surface binding of Pep-1 was abolished almost
completely by Hyal pretreatment of XS106 DC, indicating that HA
moieties are the major target of Pep-1. Biotinylated Pep-1, but not
RP, also bound significantly to other tested DC preparations, including XS52 DC, splenic DC, and BM-derived DC (Fig. 2B).
Once again, Hyal pretreatment of each DC preparation markedly
reduced Pep-1 binding (data not shown). To determine whether
HA expression on DC might be regulated by their maturation
states, we incubated BM-derived DC in the presence or the absence of LPS. As shown in Fig. 2C, HA expression levels remained relatively unchanged after LPS treatment, although surface
expression of CD40 and CD86 was markedly elevated in LPSstimulated DC. These observations indicate that DC in both immature and mature states express detectable amounts of HA on
their surfaces.
CD3⫹ T cells isolated from BALB/c mice expressed mRNAs
for HAS1, HAS3, and Hyal-3, and Hyal-2 mRNA became detectable after Con A activation, suggesting that T cells might express
HA under certain conditions (Fig. 1A). In fact, Con A-activated T
cells showed significant surface binding of biotinylated Pep-1,
whereas only marginal, if any, binding was observed in the absence of Con A stimulation (Fig. 2D). Although the extent of binding to activated T cells was relatively modest, Hyal pretreatment
markedly diminished Pep-1 binding, indicating the specificity.
Thus, we concluded that T cells express detectable amounts of HA
on their surfaces after activation.
Because CD44 is generally considered to function as a primary
HA receptor, we next studied the distributions of HA and CD44 on
the surfaces of XS106 DC. Pep-1 (green) bound diffusely and uniformly to the cell bodies (Fig. 2E, left). This Pep-1 binding pattern
showed only occasional overlap with the binding pattern of antiCD44 mAb (red). Once again, no detectable binding was observed
for the RP control (Fig. 2E, right). These observations imply that
some of the HA molecules on DC are not colocalized with CD44.
Molecular targets of Pep-1 on DC
FIGURE 1. HA synthase and Hyal mRNA expression and HA synthesis
by DC. A, The XS52 and XS106 DC lines and BM-derived DC cultures (7
days old) were examined for mRNA expression of the indicated HA synthases and Hyal by RT-PCR. CD3⫹ T cells isolated from BALB/c mice
were also examined for mRNA profiles before and after 16-h stimulation
with Con A (4 ␮g/ml). Data shown are ethidium bromide staining profiles
of the PCR products after 35 cycles of amplification. B, XS106 DC were
examined for their capacity to synthesize HA by metabolic labeling of
[3H]glucosamine. The data shown are the amounts of Hyal-sensitive radioactivities (mean ⫾ SE; n ⫽ 5) recovered from the indicated subcellular
fractions. All data in this figure are representative of three independent
experiments showing similar results.
Heparan sulfate proteoglycans are major proteoglycans found in
the extracellular matrix (especially in basement membrane) and at
the cell surface. Although heparan sulfate polysaccharide side
chains generally consist of repeated units of GlcA and N-deacetylated/N-sulfated Glc, a unique subunit composition of GlcA and
N-unsubstituted Glc has also been identified (36, 37). Thus, we
examined whether Pep-1 might also recognize heparan sulfates
through the above HA-like epitope. Pretreatment of XS106 DC
with heparinase III, which is known to selectively degrade heparan
sulfates (38), reduced the extent of Pep-1 binding, albeit modestly
even at the highest concentration (330 U/ml; Fig. 3A). Importantly,
We first examined whether DC express mRNAs for HA synthases
(HAS1 through HAS3) and Hyal (Hyal-1 through Hyal-3), the
enzymes known to be involved in HA synthesis and degradation
(13, 35). All tested HAS and Hyal mRNAs were detected in mouse
BM-derived DC cultures (Fig. 1A). Stable DC lines (XS series)
established from mouse epidermis (30, 31) showed similar profiles, except that HAS1 or HAS2 mRNA was undetectable in the
mature DC line XS106 or the immature line XS52, respectively.
Underlying mechanisms or biological consequences of this finding
(differential expression of HAS1 and HAS2 mRNAs) remain unknown at present. Nevertheless, our observations suggest the potential of DC to synthesize and catabolize HA polymers.
When cultured in the presence of [3H]glucosamine, XS106 DC
showed marked incorporation of radioactivities, and significant
portions (15–20%) of the total cell-associated radioactivities were
found to be susceptible to Hyal-mediated digestion (data not
HA EXPRESSION AND FUNCTION IN DC
4325
Hyal pretreatment at the lowest concentration (37 U/ml) caused
more prominent effects, suggesting that HA moieties are the major
target for Pep-1 on DC. On the other hand, our finding that heparinase III showed no detectable enzymatic activity to digest HA
substrates (Fig. 3B) strongly suggests that heparan sulfates serve as
a second carbohydrate target for Pep-1 binding on DC.
Impact of HA inhibitors on DC-dependent, Ag-specific T cell
proliferation
A majority of CD4⫹ T cells purified from DO11.10 TCR transgenic mice showed a characteristic phenotype of naive T cells:
CD25⫺ (⬎95%), CD69⫺ (⬎97%), CD62Lhigh (⬎84%), CD44low
(⬎91%), and CD45RBhigh (⬎98%) (data not shown). These CD4⫹
T cells, which express the transgenic TCR ␣- and ␤-chains specific
for OVA peptide 323–339 (39), allowed us to study the potential
contribution of HA to the primary T cell activation process.
DO11.10 T cells showed robust [3H]thymidine uptake when stimulated by splenic DC in the presence of OVA323–339 peptide (Fig.
4A). By contrast, no significant T cell proliferation was detected in
the absence of DC or OVA peptide, indicating DC dependency and
Ag specificity. Purified bPG, which has been used as a specific
probe and a standard inhibitor of HA (40), completely abolished
DC-induced proliferation of DO11.10 T cell proliferation, whereas
the same reagent showed no inhibitory potential after heat inacti-
vation. It should be noted that HA, but not heparan sulfates, efficiently competed with bPG in the binding to HA-coated plates,
indicating that bPG does not cross-react with heparan sulfate (Fig.
4B). Moreover, DC-dependent, Ag-specific proliferation of
DO11.10 T cells was also inhibited by Pep-1, but not by RP control (Fig. 4C). Thus, the two tested HA inhibitors with totally different chemical structures exhibited the same pharmacological activity (i.e., to block DC-induced T cell activation), with the
implication that HA molecules associated with DC and/or T cells
may play an indispensable role in Ag presentation.
Improvement of pharmacological activity of Pep-1 by
multimerization
While testing the impact on DC-dependent T cell activation, we
noticed that the extent of inhibition achieved with Pep-1 was rather
modest (⬍40%) even at a relatively high concentration of 300 ␮M
(Fig. 4C). To improve its biological avidity, we conjugated Pep-1
molecules to a tetravalent PEG derivative. The resulting preparations showed an estimated molecular ratio of Pep-1/PEG of 2.3–
2.9 (data not shown). We first compared this oligomeric Pep-1
formulation with the original monomeric Pep-1 for their pharmacological activities using two standard assays (29). First, the PEGconjugated Pep-1 inhibited the adhesion of CD44-expressing B16F10 melanoma cells to HA-coated plates almost completely
FIGURE 2. Surface HA expression by DC. A,
XS106 DC were examined for surface binding of
biotinylated Pep-1 or RP (filled histograms) in the
presence or the absence of Hyal pretreatment. Open
histograms, Baseline staining levels. B, The indicated DC preparations were examined for surface
binding of biotinylated Pep-1 (filled histograms) and
RP (open histograms). C, After 16-h incubation in
the presence or the absence of LPS (100 ng/ml),
BM-derived DC were examined for binding of biotinylated Pep-1 (filled histograms) or RP (open histograms). The same samples were also stained for
mAb against CD40 or CD86 (filled histograms) or
isotype-matched control IgG (open histograms). D,
After 16-h incubation in the presence or the absence
of Con A (4 ␮g/ml), CD3⫹ T cells were incubated
with biotinylated Pep-1 (filled histograms) or RP
(open histograms) in the presence or the absence of
Hyal pretreatment. E, XS106 DC were double-labeled with biotinylated Pep-1 or RP, followed by
FITC-streptavidin (green fluorescence signals) and
with PE-conjugated anti-CD44 mAb (red fluorescence signals). Data shown are randomly selected
fields (original magnification, ⫻400). All data in this
figure are representative of at least two independent
experiments showing similar results.
4326
(⬎90%) at 30 ␮M in terms of the molarity of the 12-mer Pep-1
sequence (Fig. 5A). By contrast, the original Pep-1 caused only
partial (75%) inhibition even at 300 ␮M. Secondly, the original
Pep-1 at 300 ␮M blocked the binding of soluble HA to CD44expressing BW5147 thymoma cells only marginally, whereas
complete inhibition was achieved with the PEG-conjugated Pep-1
at the same concentration (Fig. 5B). The PEG-conjugated RP
showed no detectable inhibitory activity. Thus, PEG-Pep-1 formulation exhibits a significantly improved pharmacological activity
over the original Pep-1 preparation to inhibit the function of HA.
Effects of PEG-Pep-1 on DC-induced, Ag-specific T cell
proliferation
We next added PEG-Pep-1 to the cocultures of splenic DC and
DO11.10 T cells to test its impact on Ag presentation. As shown
in Fig. 5C, PEG-Pep-1 inhibited T cell proliferation almost completely (80 –100%) at 50 –150 ␮M, whereas the original Pep-1
caused only modest (30%) inhibition at 300 ␮M. Likewise, PEGPep-1, but not PEG-RP, efficiently inhibited DO11.10 T cell proliferation triggered by BM-derived DC (a second DC preparation)
at a relatively low concentration of 60 ␮M (Fig. 5D). Moreover,
tetravalent PEG alone showed no inhibitory effect on DC-induced
DO11.10 T cell proliferation, formally excluding the possibility
that free arms of PEG molecules might affect DC-T cell interaction
nonspecifically (Fig. 5E). Taken together, our observations with
the multimeric Pep-1 formulation further support our conclusion
that HA is functionally involved in DC-induced T cell activation
processes.
In an attempt to determine the relevant cellular target(s) of PEGPep-1, we pretreated BM-derived DC or DO11.10 T cells with
PEG-Pep-1 or PEG-RP, washed the cells extensively, and then
cocultured them in the presence of OVA peptide. Nontreated
DO11.10 T cells showed similar levels of [3H]thymidine uptake
when stimulated with DC pretreated with PEG-Pep-1 (141,000 ⫾
6,400 cpm) vs PEG-RP (114,000 ⫾ 18,000 cpm). Conversely,
nontreated DC induced similar degrees of proliferation by
DO11.10 T cells pretreated with PEG-Pep-1 (140,000 ⫾ 25,000
cpm) vs PEG-RP (142,000 ⫾ 1,700 cpm). Our failure to block the
function of HA by pretreatment is not totally unexpected, because
we have observed previously that Pep-1 binds to HA with a relatively low affinity (Kd ⫽ 1.7 ␮M) and in a nonsaturated fashion
(29). These features most likely reflect the chemical property of
FIGURE 4. Impact of two currently available HA inhibitors on DCdependent T cell proliferation. A, DO11.10 T cells were cultured with
splenic DC and/or OVA peptide in the presence of intact bPG (250 ␮g/ml),
heat-denatured bPG, or PBS alone. Data shown are the mean ⫾ SD (n ⫽
3) [3H]thymidine uptake. B, bPG (50 ␮g/ml) was incubated for 16 h with
the indicated doses of HA (F) or heparan sulfate (E) and then added to
HA-coated plates. The data shown are the mean ⫾ SD (n ⫽ 3) percent
inhibition of bPG binding as measured with anti-bPG mAb. C, DO11.10 T
cells were cultured with splenic DC at the indicated numbers and with
OVA peptide in the presence of Pep-1 (300 ␮M; F), RP (E), or PBS alone
(䡺). Data shown are the mean ⫾ SD (n ⫽ 3) [3H]thymidine uptake. All
data in this figure are representative of three independent experiments
showing similar results. Statistically significant differences are indicated by
asterisks (ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01).
HA, which is a long, homogenous polymer consisting of repeating
disaccharide, thus providing no discrete or isolated binding sites
for Pep-1. Alternatively, PEG-Pep-1 pretreatment may have
caused no inhibition simply due to rapid turnover of surface HA
molecules. Nevertheless, we interpreted our observations to imply
that Pep-1, even in a multimeric form, must be present continuously to block the function of HA.
An obvious question concerned the identity of the putative
counter-receptor for HA. CD44 is generally thought to serve as a
primary surface receptor of HA, and CD44 expression is detectable
on the majority of DO11.10 T cells. Anti-CD44 mAb (KM81)
significantly inhibited DC-induced activation of DO11.10 T cells
in a dose-dependent manner, whereas an isotype-matched control
IgG caused no inhibition (Fig. 6A, left panel). On the other hand,
the extent of inhibition achieved with anti-CD44 mAb was relatively modest (⬍40%) even at the highest tested concentration of
70 ␮g/ml. By contrast, the same anti-CD44 mAb completely
blocked the adhesion of activated T cells to HA-coated plates at a
much lower concentration of 7 ␮g/ml (Fig. 6A, right panel). In this
regard we have observed previously that adhesion of activated T
cells to HA-coated plates is blocked almost completely by the
monomeric Pep-1 (29). To assess the target specificity of the above
inhibitors, we crossed two different adhesive substrates, HA and
FN. As shown in Fig. 6B, adhesion of B16-F10 melanoma cells to
HA-coated plates, but not FN-coated plates, was inhibited by PEGPep-1. By contrast, PEG-RP or PEG alone showed no inhibitory
activity. Anti-CD44 mAb inhibited the adhesion of activated T
cells to HA-coated plates without affecting their binding to FNcoated plates (Fig. 6C). Having confirmed the target specificity of
anti-CD44 mAb and PEG-Pep-1, we interpret our data (shown in
Fig. 6A) to suggest that CD44 is fully responsible for the adhesive
interaction of activated T cells with HA substrates, whereas CD44
is one, but not the only, relevant receptor mediating HA-dependent
FIGURE 3. Pep-1 binding to heparin sulfate moieties on DC. A, XS106
cells were pretreated with either Hyal (top) or heparinase III (bottom) at the
indicated concentrations. The samples were then examined for binding of
biotinylated Pep-1 (filled histograms). Open histograms, Baseline staining
with biotinylated RP control. Numbers indicate the mean fluorescence intensities with Pep-1. B, Biotinylated HA polymers immobilized on Nunc
plates were incubated with Hyal (F) or heparinase III (E) at the indicated
concentrations. The amount of HA remaining on the plates was examined
using streptavidin-conjugated alkaline phosphatase. Data shown are the
mean ⫾ SD (n ⫽ 3) percent digestion.
T cell activation. Alternatively, T cells may employ different CD44
isoforms to recognize HA moieties naturally expressed on DC vs
those artificially immobilized on plates. Nevertheless, our data indicate that DC-induced T cell activation can be inhibited almost
completely by interfering with HA function by specific inhibitors
(e.g., bPG and multimeric Pep-1) or partially by blocking CD44
function with anti-CD44 mAb.
Differential influences of PEG-Pep-1 on different aspects of
DC-induced T cell activation
We first examined whether PEG-Pep-1 might inhibit T cell proliferation by simply killing DC and/or T cells. To test this possi-
FIGURE 6. Relative contributions of CD44 to HA-mediated regulation
of T cell activities. A, DO11.10 T cells were cultured with splenic DC and
OVA peptide in the presence of anti-CD44 mAb (E) or isotype-matched
control IgG (F) at the indicated concentrations (left panel). The same antiCD44 mAb preparation was examined in parallel for its ability to inhibit
the adhesion of Con A-stimulated splenic T cells onto the HA-coated plates
(right panel). Data shown are the mean ⫾ SD (n ⫽ 3) percent inhibition of
T cell proliferation and adhesion. B, B16-F10 melanoma cells were examined for their adhesion to HA-coated plates and FN-coated plates in parallel. All substrates were preincubated with PEG-Pep-1 (150 ␮M), PEGRP, or PEG alone. Data shown are the mean ⫾ SD (n ⫽ 3) percent
adhesion. C, Con A-stimulated splenic T cells were pretreated with antiCD44 mAb or isotype-matched control IgG and then examined for their
adhesion to HA-coated plates and FN-coated plates in parallel. Data shown
are the mean ⫾ SD (n ⫽ 3) percent adhesion. Statistically significant differences are indicated by asterisks (ⴱⴱ, p ⬍ 0.01).
bility, we added PEG-Pep-1, PEG-RP, or PEG alone to the cocultures of BM-derived DC and DO11.10 T cells in the presence of
OVA323–339 peptide or native OVA protein. None of the reagents
significantly affected the viability of either DC or T cells in the
cocultures containing OVA323–339 peptide (Fig. 7A, left panels).
Likewise, although PEG-Pep-1 inhibited DO11.10 T cell proliferation triggered by OVA protein (data not shown), none of the
reagents caused detectable death of DC or T cells in this second
system (Fig. 7A, right panels). To visually examine the impact of
PEG-Pep-1 on T cell division, we labeled DO11.10 T cells with a
fluorescence dye, CFSE. When stimulated by BM-derived DC in
the presence of OVA peptide, CFSE-labeled DO11.10 T cells
showed marked reduction in fluorescence intensity, reflecting their
progressive mitosis (Fig. 7B). By contrast, DC failed to trigger T
cell mitosis in the absence of OVA peptide, indicating Ag specificity. Consistent with our observations in [3H]thymidine uptake
assays, PEG-Pep-1, but not PEG-RP, completely prevented DCinduced, Ag-specific clonal expansion of DO11.10 T cells. It
should be noted that we examined CFSE profiles within the PInegative populations to further address the cytotoxicity issue.
FIGURE 5. Impact of multimeric Pep-1 on DC-dependent T cell proliferation. A, Original Pep-1 (E) and PEG-conjugated Pep-1 (‚) were compared at the indicated concentrations (in terms of the molarity of the 12mer Pep-1 sequence) for their abilities to inhibit the binding of B16-F10
melanoma cells to HA-coated plates. 䡺, Percent cell adhesion to noncoated
plates. Original RP (F) and PEG-conjugated RP (Œ) served as controls.
Data shown are the mean ⫾ SD (n ⫽ 3) percent adhesion. B, Pep-1 (left)
and RP (right) in the original formulations (top) and the PEG-conjugated
formulations (bottom) were compared at 300 ␮M for their abilities to inhibit the binding of FITC-conjugated HA to the surfaces of BW5147 thymoma cells (f). Maximal binding in the absence of added inhibitors and
the background autofluorescence level are indicated with solid lines and
broken lines, respectively. C, Original Pep-1 (E) and PEG-conjugated
Pep-1 (‚) were compared at the indicated concentrations for their abilities
to inhibit DO11.10 T cell proliferation triggered by splenic DC. Data
shown are the mean ⫾ SD (n ⫽ 3) [3H]thymidine uptake. D, DO11.10 T
cells were cultured with the indicated numbers of BM-derived DC and
OVA peptide in the presence of PEG-Pep-1 (60 ␮M; ‚), PEG-RP (Œ), or
PBS alone (䡺). f, Baseline response in the absence of OVA peptide. Data
shown are the mean ⫾ SD (n ⫽ 3) [3H]thymidine uptake. E, DO11.10 T
cells were cultured with BM-derived DC and/or OVA peptide in the presence of PEG-Pep-1 (150 ␮M), PEG-RP, or PEG alone. Data shown are the
mean ⫾ SD (n ⫽ 3) [3H]thymidine. All data in this figure are representative
of two independent experiments showing similar results. Statistically significant differences are indicated by asterisks (ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01).
4327
4328
FIGURE 7. Influences of PEG-Pep-1 on DC-induced T cell activation
changes at late time points. A, DO11.10 T cells were cultured for 24 h with
BM-derived DC in the presence of OVA323–339 peptide (left) or OVA protein. Samples were then examined for PI uptake within the CD11c⫹ populations (䡺) or the KJ1.26⫹ populations (f). Data shown are the mean ⫾
SD (n ⫽ 3) percentage of PI-negative cells. B, CFSE-labeled DO11.10 T
cells were cultured with splenic DC and/or OVA peptide in the presence of
PEG-Pep-1 (60 ␮M), PEG-RP, or PBS alone. Data shown are the CFSE
fluorescence profiles within PI-negative, viable cell populations. C,
DO11.10 T cells were cultured for 24 h with splenic DC and/or OVA
peptide in the presence of PEG-Pep-1 (60 ␮M), PEG-RP, or PBS alone.
Data shown are the mean ⫾ SD (n ⫽ 3) of the indicated cytokine concentrations detected in the culture supernatants. D, DO11.10 T cells were
cultured with BM-derived DC and/or OVA peptide in the presence of PEGPep-1 (60 ␮M), PEG-RP, or PBS alone. Phase-contrast photographs were
taken from randomly chosen fields after 16 h (original magnification,
⫻100). All data in this figure are representative of two independent experiments. Statistically significant differences are indicated by asterisks
(ⴱⴱ, p ⬍ 0.01).
Does PEG-Pep-1 affect other changes that accompany T cell
activation? As shown in Fig. 7C, DO11.10 T cells secreted significant amounts of IL-2 and IFN-␥ when cocultured with BMderived DC in the presence of OVA peptide, whereas neither cytokine was detected in the absence of DC or Ag. Importantly,
production of IL-2 and IFN-␥ was inhibited efficiently by PEGPep-1, but not by PEG-RP. DO11.10 T cells exhibited characteristic cluster formation only when cocultured with DC in the presence of OVA peptide (Fig. 7D). This morphological change was
FIGURE 8. Failure of PEG-Pep-1 to inhibit early DC-T cell conjugate
formation during Ag presentation. A, BM-derived DC were incubated for
10 min with FITC-BSA (filled histograms) or PBS (open histograms) in the
presence of PEG-Pep-1 (150 ␮M), PEG-RP, or PBS alone. B, BM-derived
DC labeled with PKH-26 and DO11.10 T cells labeled with CFSE were
incubated for 30 min in the presence or the absence of OVA peptide (top
panels). The numbers indicate the percentage of the double-positive events
representing DC-T cell aggregates. PEG-Pep-1 was added at the indicated
concentrations to the cocultures with OVA peptide to test its impact on
DC-T cell conjugate formation (bottom panels). Data shown in this figure
are representative of two independent experiments.
also inhibited by PEG-Pep-1, but not by PEG-RP. Thus, PEGPep-1 is capable of inhibiting multiple events that accompany DCinduced T cell activation.
The experiments described above examined relatively late
events during Ag presentation, i.e., T cell cluster formation at 16 h,
cytokine production at 24 h, T cell mitosis at 48 h, and [3H]thymidine uptake at 72 h. We next sought to examine the potential
influence of PEG-Pep-1 on earlier events. As shown in Fig. 8A,
PEG-Pep-1, PEG-RP, or PEG alone did not inhibit FITC-BSA
uptake by BM-derived DC. To test the impact on early formation
of DC-T cell aggregates, we labeled BM-derived DC with a red
fluorescence dye (PKH-26) and DO11.10 T cells with a green fluorescence dye CFSE. Significant numbers of DC-T cell conjugates
were detected as double-positive events when the two populations
were incubated for 30 min in the presence of OVA peptide (Fig.
8B). Importantly, PEG-Pep-1 even at relatively high concentrations (up to 75 ␮M in the experiment shown in Fig. 8B and 150
␮M in a different experiment) failed to inhibit this early event.
Thus, PEG-Pep-1 efficiently inhibits relatively late events that accompany DC-induced T cell activation, without affecting the viability of DC or T cells, the ability of DC to uptake exogenous
4329
proteins, or Ag-specific DC-T cell conjugate formation at earlier
time points.
Potential function of T cell-associated HA
In vivo ability of PEG-Pep-1 to inhibit DC-induced, Ag-specific
T cell expansion
With respect to the in vivo impact of Pep-1, we have successfully
inhibited allergic contact hypersensitivity responses to dinitrofluorobenzene (DNFB) by injecting the original Pep-1 formulation
before or even after sensitization (29). Although these results may
first appear to support our working model that HA facilitates DCinduced T cell activation, it should be stated that administered
Pep-1 also blocked hapten-triggered migration of epidermal DC,
known as Langerhans cells (LC), from the epidermis at the sensitization phase and skin-directed homing of effector T cells at the
elicitation phase. Thus, our previous observations may mainly reflect the ability of Pep-1 to block HA-dependent, two-way trafficking of leukocytes from and to the skin, but not its potential
impact on the Ag presentation processes in which DNFB-pulsed
LC trigger clonal expansion of DNFB-reactive T cells. To study
the direct impact of PEG-Pep-1 on DC-induced T cell activation,
we sought to develop a new in vivo model that would not require
DC or T cell homing. When OVA peptide-pulsed DC and CFSE-
FIGURE 10. In vivo impact of PEG-Pep-1 on DC-dependent clonal expansion of T cells. A and B, BALB/c mice received i.p. injections of CFSElabeled DO11.10 T cells, OVA peptide-pulsed BM-derived DC, and the
indicated reagent. The cells recovered from the peritoneal cavity 2 days
later were examined for CFSE fluorescence profiles within the PI-negative
populations and for percentage of dividing cells (A). Results obtained from
three animals per each panel were examined for the mean ⫾ SEM percentage of dividing cells and for statistical difference (ⴱ, p ⬍ 0.05; B). All
data in this figure are representative of two independent experiments showing similar results.
labeled DO11.10 T cells were injected i.p., a significant reduction
in the fluorescence intensity was detected in cells recovered from
the peritoneal cavity 48 h later (Fig. 10A). No significant change
was observed in the CFSE profiles of DO11.10 T cells when coinjected with PBS-pulsed DC (data not shown). Importantly, a substantial reduction in T cell mitosis was detected following i.p. administration of PEG-Pep-1, but not PEG-RP. In two independent
experiments the panel receiving PEG-Pep-1 injection showed statistically significant impairment in the extent of clonal expansion
of adoptively transferred T cells compared with the control panel
receiving PBS alone and the second control panel receiving
PEG-RP (Fig. 10B). Our model, which may be seen as in vivo
cocultures of Ag-pulsed DC and T cells in the peritoneal cavity,
does not represent physiological immune responses or diseases in
living animals. However, we believe that our observation in this
model at least reflects direct influences of PEG-Pep-1 on DC-dependent, Ag-specific clonal expansion of T cells in living animals.
Discussion
FIGURE 9. HA-mediated autocrine mechanism promoting mitogentriggered T cell activation. A, CD3⫹ T cells were stimulated with anti-CD3
mAb and/or CD28 mAb in the presence of PEG-Pep-1 (60 ␮M), PEG-RP,
or PBS alone. B, CD3⫹ T cells were stimulated with Con A in the presence
of bPG (250 ␮g/ml) or PBS alone. Data shown are the mean ⫾ SD (n ⫽
3) [3H]thymidine uptake. Statistically significant differences are indicated
by asterisks (ⴱⴱ, p ⬍ 0.01).
We have demonstrated in this study that DC constitutively express
mRNAs for various enzymes involved in HA synthesis and degradation, actively synthesize HA, and uniformly express HA on
their surfaces. Moreover, we have detected HA expression on activated T cells, albeit at modest levels. These findings, which imply
an immunoregulatory role for HA, challenge the classic view of
HA as an extracellular matrix component or a filling material being
produced by fibroblasts, chondrocytes, epithelial cells, and endothelial cells. In fact, DC-induced, Ag-specific activation of T cells
was blocked significantly by all tested HA inhibitors (i.e., monomeric and multimeric Pep-1 formulations and bPG). PEG-Pep-1,
for example, efficiently inhibited DC-induced, Ag-specific T cell
clustering, cytokine production, [3H]thymidine uptake, and mitosis. Complementary observations have been reported by Galandrini et al. (43), who found that exogenous HA polymers in an
It has been an established concept that immunologically naive T
cells are activated most efficiently by DC (41, 42). From this viewpoint, our findings illustrate functional roles of HA in Ag presentation under physiological conditions. On the other hand, it remains unclear whether Pep-1 primarily acts on HA moieties
constitutively displayed on DC or whether it may also inhibit the
function of HA being expressed by T cells upon activation. To test
the latter possibility, we employed nonphysiological experimental
systems in which T cells were stimulated by mitogens in the absence of DC or Ag. When stimulated by both anti-CD3 mAb and
anti-CD28 mAb, CD3⫹ T cells showed robust proliferation (Fig.
9A). T cell responses to this mitogenic stimulation were almost
completely inhibited by PEG-Pep-1, but not by PEG-RP. Likewise, Con A-triggered proliferation of CD3⫹ T cells was almost
completely inhibited by bPG (Fig. 9B). These observations imply
a second mechanism by which HA moieties expressed by activated
T cells facilitate their own proliferation in an autocrine manner.
4330
mice expressing an antisense CD44 cDNA under the keratin-5 promoter. Interestingly, the same transgenic animals also exhibited
marked reduction in the amount of HA present in the epidermal
intercellular space and massive accumulation of HA in the superficial dermis, with the implication that CD44 expressed on keratinocytes plays a critical role in HA turnover in skin (55). Thus, it
is conceivable that anti-CD44 mAb may have inhibited LC migration indirectly by altering HA metabolism in skin, instead of directly blocking the function of LC-associated CD44. Our observations in this study suggest an even more extreme possibility, that
HA polymers expressed on DC may direct their homing to the
afferent lymphatics by binding to various putative receptors, including LYVE-1, a newly identified HA receptor expressed almost
exclusively in the lymphatic vessels (10, 11).
Degradation products of HA are known to trigger cytokine and
chemokine production by macrophages (21–24). More recently,
Termeer et al. (56, 57) have reported the potential of small HA
fragments of tetra- and hexa-saccharide to induce DC maturation
by a Toll-like receptor 4-dependent mechanism. With respect to
the physiological source of HA fragments, these investigators postulated that HA polymers in the extracellular matrices might be
digested into oligosaccharides at the sites of inflammation. Our
findings now suggest that DC (expressing mRNAs for HA synthases and Hyal) may release degradation products of HA, thereby
regulating their own maturational state in an autocrine manner.
Alternatively, T cells (expressing similar mRNA profiles) may
elaborate HA fragments during Ag presentation, triggering DC
maturation in a paracrine manner. The heparan sulfate moiety detected on DC may also play an immunoregulatory role; in fact,
heparan sulfate and its degradation products have been reported to
deliver costimulatory signals to T cells and maturation signals to
DC (58 – 60). Obviously, further studies are required to examine
the metabolic activities of DC to synthesize and degrade HA and
heparan sulfate and to characterize the immunoregulatory properties of DC-derived oligosaccharides. Nevertheless, the present
study may introduce a new concept, that Ag presentation processes
are regulated by relatively simple carbohydrates being produced
by DC.
Acknowledgments
We thank Lesa Ellinger and Pat Adcock for their assistance.
References
1. Tammi, R., J. A. Ripellino, R. U. Margolis, and M. Tammi. 1988. Localization
of epidermal hyaluronic acid using the hyaluronate binding region of cartilage
proteoglycan as a specific probe. J. Invest. Dermatol. 90:412.
2. Tammi, R., A. M. Saamanen, H. I. Maibach, and M. Tammi. 1991. Degradation
of newly synthesized high molecular mass hyaluronan in the epidermal and dermal compartments of human skin in organ culture. J. Invest. Dermatol. 97:126.
3. Aruffo, A., I. Stamenkovic, M. Melnick, C. B. Underhill, and B. Seed. 1990.
CD44 is the principal cell surface receptor for hyaluronate. Cell 61:1303.
4. Miyake, K., C. B. Underhill, J. Lesley, and P. W. Kincade. 1990. Hyaluronate can
function as a cell adhesion molecule and CD44 participates in hyaluronate recognition. J. Exp. Med. 172:69.
5. Hardingham, T. E. 1979. The role of link-protein in the structure of cartilage
proteoglycan aggregates. Biochem. J. 177:237.
6. Watanabe, H., S. C. Cheung, N. Itano, K. Kimata, and Y. Yamada. 1997. Identification of hyaluronan-binding domains of aggrecan. J. Biol. Chem. 272:28057.
7. LeBaron, R. G., D. R. Zimmermann, and E. Ruoslahti. 1992. Hyaluronate binding
properties of versican. J. Biol. Chem. 267:10003.
8. Delpech, B., and C. Halavent. 1981. Characterization and purification from human brain of a hyaluronic acid-binding glycoprotein, hyaluronectin. J. Neurochem. 36:855.
9. Rauch, U., L. Karthikeyan, P. Maurel, R. U. Margolis, and R. K. Margolis. 1992.
Cloning and primary structure of neurocan, a developmentally regulated, aggregating chondroitin sulfate proteoglycan of brain. J. Biol. Chem. 267:19536.
10. Banerji, S., J. Ni, S. X. Wang, S. Clasper, J. Su, R. Tammi, M. Jones, and
D. G. Jackson. 1999. LYVE-1, a new homologue of the CD44 glycoprotein, is a
lymph-specific receptor for hyaluronan. J. Cell Biol. 144:789.
11. Prevo, R., S. Banerji, D. J. Ferguson, S. Clasper, and D. G. Jackson. 2001. Mouse
LYVE-1 is an endocytic receptor for hyaluronan in lymphatic endothelium.
J. Biol. Chem. 276:19420.
immobilized form augment T cell proliferation and IL-2 production triggered by anti-CD3 mAb or phorbol ester. These findings
made in opposing directions (i.e., loss of function and gain of
function) introduce a new concept that HA polymers being synthesized and expressed by DC and T cells facilitate Ag presentation processes.
The relevant cellular target(s) or molecular mechanism(s) for
Pep-1-induced inhibition remain unclear at this time. PEG-Pep-1
and bPG both inhibited mitogen-triggered activation of T cells in
the absence of DC, revealing a putative pathway in which HA
molecules expressed by T cells may regulate their own activation
in an autocrine manner. These observations, however, should not
be interpreted to indicate that T cell-associated HA, but not DCassociated HA, plays a regulatory role in Ag presentation under
physiological conditions. Perhaps the most unexpected observation was the failure of PEG-Pep-1 to inhibit DC-T cell conjugate
formation at an early time point, despite the fact that the same
reagent almost completely inhibited several later events in Ag presentation. Activation of naive T cells has been reported to increase
not only the extent of immunoreactivity to anti-CD44 mAb, but
also the HA binding potential of CD44 molecules (44, 45). Thus,
it is tempting to speculate that DC-associated HA may not be able
to interact efficiently with CD44 expressed by naive T cells in the
absence of T cell activation signals. Conversely, DC maturation is
accompanied by increased immunoreactivity to anti-CD44 mAb
and elevated surface expression of selected CD44 variant isoforms
(46, 47). An alternative explanation is, therefore, that DC may
need additional maturational signals from T cells for their CD44dependent interaction with T cell-associated HA. Although HAS2knockout embryos die during midgestation due to severe cardiac
and vascular abnormalities (48), HAS1/HAS3 double-knockout
mice show no apparent developmental defect (48). Experiments
are underway in our laboratories to study immunological functions
of DC and T cells isolated from the HAS1/HAS3 knockout
animals.
Our detection of HA molecules that did not colocalize with
CD44 on DC is not totally surprising, because even a CD44-negative tumor cell line has been reported to elevate surface HA expression levels upon transfection with HAS2 or HAS3 cDNA (49).
Perhaps HAS molecules at the inner face of the plasma membrane
not only catalyze HA synthesis, but also serve as a membraneanchoring mechanism for newly synthesized HA polymers (13).
Our observation that anti-CD44 mAb caused only partial inhibition
of DC-induced T cell activation is also in complete agreement with
the current idea that HA binds many other surface receptors and
extracellular matrix components (see the introduction). Thus, it
seems reasonable to state that CD44 is one of multiple HA receptors involved in HA-mediated immunoregulation.
HA molecules, which are present along the migration path (e.g.,
intercellular space in epidermis and dermal matrix) for epidermal
LC, have been considered to promote CD44-dependent LC homing. This concept was originally derived from the observations that
1) CD44 expression is detectable on epidermal LC in normal skin
(50, 51); 2) LC elevate surface expression of selected CD44 isoforms upon hapten painting (47); and 3) antagonistic anti-CD44
mAb and Pep-1 both inhibit LC migration from the epidermis and
the subsequent contact hypersensitivity responses (29, 47, 52).
More recent reports, however, do not necessarily support the above
view. Schmits et al. (53) reported that CD44 knockout mice generated by conventional homologous recombination show normal
contact hypersensitivity responses to DNFB, indicating that CD44
expression by LC is not absolutely required for their hapten-triggered migration. By contrast, Kaya et al. (54) reported severely
impaired contact hypersensitivity responses to DNFB in transgenic
38.
39.
40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
glucosamine units in native heparan sulfate revealed by a monoclonal antibody.
J. Biol. Chem. 270:31303.
Rozenberg, G. I., J. Espada, L. L. de Cidre, A. M. Eijan, J. C. Calvo, and
G. E. Bertolesi. 2001. Heparan sulfate, heparin, and heparinase activity detection
on polyacrylamide gel electrophoresis using the fluorochrome tris(2,2⬘-bipyridine) ruthenium (II). Electrophoresis 22:3.
Murphy, K. M., A. B. Heimberger, and D. Y. Loh. 1990. Induction by antigen of
intrathymic apoptosis of CD4⫹CD8⫹TCRlo thymocytes in vivo. Science 250:
1720.
Sconocchia, G., J. A. Titus, and D. M. Segal. 1997. Signaling pathways regulating CD44-dependent cytolysis in natural killer cells. Blood 90:716.
Banchereau, J., and R. M. Steinman. 1998. Dendritic cells and the control of
immunity. Nature 392:245.
Banchereau, J., F. Briere, C. Caux, J. Davoust, S. Lebecque, Y.-J. Liu,
B. Pulendran, and K. Palucka. 2000. Immunobiology of dendritic cells. Annu.
Rev. Immunol. 18:767.
Galandrini, R., E. Galluzzo, N. Albi, C. E. Grossi, and A. Velardi. 1994. Hyaluronate is costimulatory for human T cell effector functions and binds to CD44
on activated T cells. J. Immunol. 153:21.
Lesley, J., and R. Hyman. 1992. CD44 can be activated to function as an hyaluronic acid receptor in normal murine T cells. Eur. J. Immunol. 22:2719.
Lesley, J., N. Howes, A. Perschl, and R. Hyman. 1994. Hyaluronan binding
function of CD44 is transiently activated on T cells during an in vivo immune
response. J. Exp. Med. 180:383.
Aiba, S., S. Nakagawa, H. Ozawa, K. Miyake, H. Yagita, and H. Tagami. 1993.
Up-regulation of ␣4 integrin on activated Langerhans cells: analysis of adhesion
molecules on Langerhans cells relating to their migration from skin to draining
lymph nodes. J. Invest. Dermatol. 100:143.
Weiss, J. M., J. Sleeman, A. C. Renkl, H. Dittmar, C. C. Termeer, S. Taxis,
N. Howells, E. Schöpf, H. Ponta, P. Herrlich, et al. 1997. An essential role for
CD44 variant isoforms in epidermal Langerhans cell and blood dendritic cell
function. J. Cell Biol. 137:1137.
Camenisch, T. D., A. P. Spicer, T. Brehm-Gibson, J. Biesterfeldt,
M. L. Augustine, A. Calabro, Jr., S. Kubalak, S. E. Klewer, and J. A. McDonald.
2000. Disruption of hyaluronan synthase-2 abrogates normal cardiac morphogenesis and hyaluronan-mediated transformation of epithelium to mesenchyme.
J. Clin. Invest. 106:349.
Simpson, M. A., C. M. Wilson, L. T. Furcht, A. P. Spicer, T. R. Oegema, Jr., and
J. B. McCarthy. 2002. Manipulation of hyaluronan synthase expression in prostate adenocarcinoma cells alters pericellular matrix retention and adhesion to
bone marrow endothelial cells. J. Biol. Chem. 277:10050.
Cumberbatch, M., R. J. Dearman, and I. Kimber. 1996. Adhesion molecule expression by epidermal Langerhans cells and lymph node dendritic cells: a comparison. Arch. Dermatol. Res. 288:739.
Osada, A., H. Nakashima, M. Furue, and K. Tamaki. 1995. Up regulations of
CD44 expression by tumor necrosis factor-␣ is neutralized by interleukin 10 in
Langerhans cells. J. Invest. Dermatol. 105:124.
Camp, R. L., A. Scheynius, C. Johansson, and E. Puré. 1993. CD44 is necessary
for optimal contact allergic responses but is not required for normal leukocyte
extravasation. J. Exp. Med. 178:497.
Schmits, R., J. Filmus, N. Gerwin, G. Senaldi, F. Kiefer, T. Kundig,
A. Wakeham, A. Shahinian, C. Catzavelos, J. Rak, et al. 1997. CD44 regulates
hematopoietic progenitor distribution, granuloma formation, and tumorigenicity.
Blood 90:2217.
Kaya, G., I. Rodriguez, J. L. Jorcano, P. Vassalli, and I. Stamenkovic. 1999.
Cutaneous delayed-type hypersensitivity response is inhibited in transgenic mice
with keratinocyte-specific CD44 expression defect. J. Invest. Dermatol. 113:137.
Kaya, G., I. Rodriguez, J. L. Jorcano, P. Vassalli, and I. Stamenkovic. 1997.
Selective suppression of CD44 in keratinocytes of mice bearing an antisense
CD44 transgene driven by a tissue-specific promoter disrupts hyaluronate metabolism in the skin and impairs keratinocyte proliferation. Genes Dev. 11:996.
Termeer, C. C., J. Hennies, U. Voith, T. Ahrens, J. M. Weiss, P. Prehm, and
J. C. Simon. 2000. Oligosaccharides of hyaluronan are potent activators of dendritic cells. J. Immunol. 165:1863.
Termeer, C., F. Benedix, J. Sleeman, C. Fieber, U. Voith, T. Ahrens, K. Miyake,
M. Freudenberg, C. Galanos, and J. C. Simon. 2002. Oligosaccharides of hyaluronan activate dendritic cells via toll-like receptor 4. J. Exp. Med. 195:99.
Wrenshall, L. E., F. B. Cerra, A. Carlson, F. H. Bach, and J. L. Platt. 1991.
Regulation of murine splenocyte responses by heparan sulfate. J. Immunol. 147:
455.
Kodaira, Y., S. K. Nair, L. E. Wrenshall, E. Gilboa, and J. L. Platt. 2000. Phenotypic and functional maturation of dendritic cells mediated by heparan sulfate.
J. Immunol. 165:1599.
Johnson, G. B., G. J. Brunn, Y. Kodaira, and J. L. Platt. 2002. Receptor-mediated
monitoring of tissue well-being via detection of soluble heparan sulfate by Tolllike receptor 4. J. Immunol. 168:5233.
12. Cheung, W. F., T. F. Cruz, and E. A. Turley. 1999. Receptor for hyaluronanmediated motility (RHAMM), a hyaladherin that regulates cell responses to
growth factors. Biochem. Soc. Trans. 27:135.
13. Itano, N., T. Sawai, M. Yoshida, P. Lenas, Y. Yamada, M. Imagawa,
T. Shinomura, M. Hamaguchi, Y. Yoshida, Y. Ohnuki, et al. 1999. Three isoforms of mammalian hyaluronan synthases have distinct enzymatic properties.
J. Biol. Chem. 274:25085.
14. Weigel, P. H., V. C. Hascall, and M. Tammi. 1997. Hyaluronan synthases.
J. Biol. Chem. 272:13997.
15. Roden, L., P. Campbell, J. R. Fraser, T. C. Laurent, H. Pertoft, and
J. N. Thompson. 1989. Enzymic pathways of hyaluronan catabolism. Ciba
Found. Symp. 143:60.
16. Frost, G. I., T. B. Csoka, T. Wong, and R. Stern. 1997. Purification, cloning, and
expression of human plasma hyaluronidase. Biochem. Biophys. Res. Commun.
236:10.
17. Gmachl, M., S. Sagan, S. Ketter, and G. Kreil. 1993. The human sperm protein
PH-20 has hyaluronidase activity. FEBS Lett. 336:545.
18. Lepperdinger, G., B. Strobl, and G. Kreil. 1998. HYAL2, a human gene expressed
in many cells, encodes a lysosomal hyaluronidase with a novel type of specificity.
J. Biol. Chem. 273:22466.
19. Csoka, T. B., G. I. Frost, T. Wong, and R. Stern. 1997. Purification and microsequencing of hyaluronidase isozymes from human urine. FEBS Lett. 417:307.
20. Hardwick, C., K. Hoare, R. Owens, H. P. Hohn, M. Hook, D. Moore, V. Cripps,
L. Austen, D. M. Nance, and E. A. Turley. 1992. Molecular cloning of a novel
hyaluronan receptor that mediates tumor cell motility. J. Cell Biol. 117:1343.
21. Hodge-Dufour, J., P. W. Noble, M. R. Horton, C. Bao, M. Wysoka,
M. D. Burdick, R. M. Strieter, G. Trinchieri, and E. Puré. 1997. Induction of
IL-12 and chemokines by hyaluronan requires adhesion-dependent priming of
resident but not elicited macrophages. J. Immunol. 159:2492.
22. Horton, M. R., M. D. Burdick, R. M. Strieter, C. Bao, and P. W. Noble. 1998.
Regulation of hyaluronan-induced chemokine gene expression by IL-10 and
IFN-␥ in mouse macrophages. J. Immunol. 160:3023.
23. McKee, C. M., M. B. Penno, M. Cowman, M. D. Burdick, R. M. Strieter, C. Bao,
and P. W. Noble. 1996. Hyaluronan (HA) fragments induce chemokine gene
expression in alveolar macrophages: the role of HA size and CD44. J. Clin.
Invest. 98:2403.
24. Horton, M. R., C. M. McKee, C. Bao, F. Liao, J. M. Farber, J. Hodge-Dufour,
E. Pure, B. L. Oliver, T. M. Wright, and P. W. Noble. 1998. Hyaluronan fragments synergize with interferon-␥ to induce the C-X-C chemokines mig and
interferon-inducible protein-10 in mouse macrophages. J. Biol. Chem.
273:35088.
25. Knutson, J. R., J. Iida, G. B. Fields, and J. B. McCarthy. 1996. CD44/chondroitin
sulfate proteoglycan and ␣2␤1 integrin mediate human melanoma cell migration
on type IV collagen and invasion of basement membranes. Mol. Biol. Cell 7:383.
26. Jalkanen, S., and M. Jalkanen. 1992. Lymphocyte CD44 binds the COOH-terminal heparin-binding domain of fibronectin. J. Cell Biol. 116:817.
27. Sleeman, J. P., K. Kondo, J. Moll, H. Ponta, and P. Herrlich. 1997. Variant exons
v6 and v7 together expand the repertoire of glycosaminoglycans bound by CD44.
J. Biol. Chem. 272:31837.
28. Toyama-Sorimachi, N., F. Kitamura, H. Habuchi, Y. Tobita, K. Kimata, and
M. Miyasaka. 1997. Widespread expression of chondroitin sulfate-type serglycins with CD44 binding ability in hematopoietic cells. J. Biol. Chem. 272:26714.
29. Mummert, M. E., M. Mohamadzadeh, D. I. Mummert, N. Mizumoto, and
A. Takashima. 2000. Development of a peptide inhibitor of hyaluronan-mediated
leukocyte trafficking. J. Exp. Med. 192:769.
30. Xu, S., K. Ariizumi, G. Caceres-Dittmar, D. Edelbaum, K. Hashimoto,
P. R. Bergstresser, and A. Takashima. 1995. Successive generation of antigenpresenting, dendritic cell lines from murine epidermis. J. Immunol. 154:2697.
31. Matsue, H., K. Matsue, M. Walters, K. Okumura, H. Yagita, and A. Takashima.
1999. Induction of antigen-specific immunosuppression by CD95L cDNA-transfected “killer” dendritic cells. Nat. Med. 5:930.
32. Kumamoto, T., E. K. Huang, H. J. Paek, R. F. Valentini, and A. Takashima. 2002.
Induction of tumor-specific protective immunity by in situ Langerhans cell vaccine. Nat. Biotechnol. 20:64.
33. Mizumoto, N., T. Kumamoto, S. C. Robson, J. Sévigny, H. Matsue, K. Enjyoji,
and A. Takashima. 2002. CD39 is the dominant Langerhans cell-associated ectoNTPDase: modulatory roles in inflammation and immune responsiveness. Nat.
Med. 8:358.
34. Schaefer, B. C., M. F. Ware, P. Marrack, G. R. Fanger, J. W. Kappler,
G. L. Johnson, and C. R. Monks. 1999. Live cell fluorescence imaging of T cell
MEKK2: redistribution and activation in response to antigen stimulation of the T
cell receptor. Immunity 11:411.
35. Menzel, E. J., and C. Farr. 1998. Hyaluronidase and its substrate hyaluronan:
biochemistry, biological activities and therapeutic uses. Cancer Lett. 131:3.
36. Lyon, M., J. A. Deakin, and J. T. Gallagher. 1994. Liver heparan sulfate structure:
a novel molecular design. J. Biol. Chem. 269:11208.
37. van den Born, J., K. Gunnarsson, M. A. Bakker, L. Kjellen, M. Kusche-Gullberg,
M. Maccarana, J. H. Berden, and U. Lindahl. 1995. Presence of N-unsubstituted
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