The Actin Regulatory Protein HS1 Is Required for Antigen Uptake and Presentation by Dendritic Cells
The hematopoietic actin regulatory protein hematopoietic lineage cell-specific protein 1 (HS1) is required for cell spreading and signaling in lymphocytes, but the scope of HS1 function in Ag presentation has not been addressed. We show that dendritic cells (DCs) from HS1(-/-) mice differentiate normally and display normal LPS-induced upregulation of surface markers and cytokines. Consistent with their normal expression of MHC and costimulatory molecules, HS1(-/-) DCs present OVA peptide efficiently to CD4(+) T cells. However, presentation of OVA protein is defective. Similarly, MHC class I-dependent presentation of VSV8 peptide to CD8(+) T cells occurs normally, but cross-presentation of GRP94/VSV8 complexes is defective. Analysis of Ag uptake pathways shows that HS1 is required for receptor-mediated endocytosis, but not for phagocytosis or macropinocytosis. HS1 interacts with dynamin 2, a protein involved in scission of endocytic vesicles. However, HS1(-/-) DCs showed decreased numbers of endocytic invaginations, whereas dynamin-inhibited cells showed accumulation of these endocytic intermediates. Taken together, these studies show that HS1 promotes an early step in the endocytic pathway that is required for efficient Ag presentation of exogenous Ag by DCs.
The Journal of Immunology
The Actin Regulatory Protein HS1 Is Required for Antigen
Uptake and Presentation by Dendritic Cells
Deborah A. Klos Dehring,*
Edward K. Williamson,* Shuixing Li,* Fiona Clarke,* Stefania Gallucci,
and Janis K. Burkhardt*
The hematopoietic actin regulatory protein hematopoietic lineage cell-speciﬁc protein 1 (HS1) is required for cell spreading and
signaling in lymphocytes, but the scope of HS1 function in Ag presentation has not been addressed. We show that dendritic cells
(DCs) from HS1
mice differentiate normally and display normal LPS-induced upregulation of surface markers and cytokines.
Consistent with their normal expression of MHC and costimulatory molecules, HS1
DCs present OVA peptide efﬁciently to
T cells. However, presentation of OVA protein is defective. Similarly, MHC class I-dependent presentation of VSV8 peptide
T cells occurs normally, but cross-presentation of GRP94/VSV8 complexes is defective. Analysis of Ag uptake pathways
shows that HS1 is required for receptor-mediated endocytosis, but not for phagocytosis or macropinocytosis. HS1 interacts with
dynamin 2, a protein involved in scission of endocytic vesicles. However, HS1
DCs showed decreased numbers of endocytic
invaginations, whereas dynamin-inhibited cells showed accumulation of these endocytic intermediates. Taken together, these
studies show that HS1 promotes an early step in the endocytic pathway that is required for efﬁcient Ag presentation of exogenous
Ag by DCs. The Journal of Immunology, 2011, 187: 5952–5963.
endritic cells (DCs) are highly specialized for presenta-
tion of Ags to naive T cells. DCs survey peripheral tis-
sues, ingesting large volumes of material by receptor-
mediated endocytosis, phagocytosis, and macropinocytosis (1–
5). In the presence of inﬂammatory signals, DCs undergo a mat-
uration process that results in diminished endocytosis of Ag,
enhanced acidiﬁcation of Ag processing compartments, redis-
tribution of MHC molecules to the cell surface, upregulation
of costimulatory molecules, and increased cell motility. As they
mature, DCs migrate to lymphoid organs, where they present
peptides derived from non-self Ags to T cells, initiating an
adaptive immune response.
Many of these aspects of DC function rely on actin and its
regulatory proteins. During endocytosis, actin polymerization pro-
duces forces that promote internalization of plasma membrane
vesicles. This is particularly obvious for phagocytosis and mac-
ropinocytosis, which involve large actin-rich cell surface protru-
sions (6–8). However, receptor-mediated endocytosis is also
dependent on actin ﬁlaments, which work together with clathrin
and other proteins such as dynamin 2 to drive the internalization of
plasma membrane vesicles (9, 10). After vesicle internalization,
the actin cytoskeleton serves as a highway to transport vesicles to
compartments where Ag is processed, loaded onto MHC mole-
cules, and transported back to the cell surface for recognition by
T cells (1, 2, 11).
Macropinocytosis and phagocytosis depend on the Rho family
GTPases CDC42 and Rac (12–14), and diminished uptake through
these pathways in mature DCs has been linked to downregulation
of CDC42 function (13). Notably, however, receptor-mediated
endocytosis is not dependent on Rho GTPases, nor is it down-
regulated in mature DCs (13). In keeping with this ﬁnding, recent
analysis has shown that mature DCs take up Ags efﬁciently via
receptor-mediated endocytosis (15), a process that may be very
important for presentation of Ags by lymphoid-resident DCs (16).
Receptor-mediated Ag uptake by both immature and mature DCs
is likely to be particularly important at low Ag dose. In addition to
playing an important role in normal immune responses, the ability
of DCs to take up material by receptor-mediated endocytosis has
been widely exploited to target these cells for therapeutic purposes
(e.g., Refs. 17–21).
Defects in actin regulatory proteins have far-reaching effects
on DC function. Mutations in Wiskott–Aldrich syndrome protein
(WASp) and its binding partner WASp interacting protein (WIP)
lead to defects in Ag uptake, migration, and immunological syn-
apse formation (22–27). We have recently found that WASp and
WIP interact in DCs with a third protein, hematopoietic lineage
cell-speciﬁc protein 1 (HS1; also called HCLS1, LckBP1) (28).
HS1 is the hematopoietic lineage-speciﬁc homolog of the more
*Department of Pathology and Laboratory Medicine, The Children’s Hospital of
Philadelphia and Perelman School of Medicine, University of Pennsylvania, Phila-
delphia, PA 19104; and
Division of Rheumatology, Department of Pediatrics, The
Children’s Hospital of Philadelphia and Perelman School of Medicine, University of
Pennsylvania, Philadelphia, PA 19104
Y.H. and C.B. contributed equally to this work.
Current address: Department of Cell and Molecular Biology, Feinberg School of
Medicine, Northwestern University, Chicago, IL.
Current address: Department of Microbiology and Immunology, Temple University
School of Medicine, Philadelphia, PA.
Received for publication April 22, 2011. Accepted for publication September 29,
This work was supported by National Institutes of Health Grants T32AI05542806A1
and F32AI08006901 (to D.A.K.D.) and R21AI088376 (to J.K.B. and Y.A.).
Address correspondence and reprint requests to Dr. Janis K. Burkhardt, Department
of Pathology and Laboratory Medicine, Children’s Hospital of Philadelphia, 3615
Civic Center Boulevard, 816D Abramson Research Center, Philadelphia, PA 19104.
E-mail address: email@example.com
Abbreviations used in this article: BMDC, bone marrow-derived dendritic cell; DAB,
3,39-diaminobenzidine; DC, dendritic cell; GRP94, glucose regulated protein of
94 kDa; HS1, hematopoietic lineage cell-speciﬁc protein 1; NP-40, Nonidet P-40;
WASp, Wiskott–Aldrich syndrome protein; WIP, WASp interacting protein; WT,
Copyright Ó 2011 by The American Association of Immunologists, Inc. 0022-1767/11/$16.00
widely expressed protein cortactin (29), and we have shown that
HS1 is the only cortactin family member expressed in murine bone
marrow-derived dendritic cells (28). Like WASp, HS1 can activate
the Arp2/3 complex to drive the formation of branched actin ﬁla-
ments (30, 31). However, HS1 also binds to F-actin and stabilizes
the branched actin network generated by WASp and other proteins
(32). HS1 is a modular protein, with an N-terminal domain that
binds to the Arp2/3 complex followed by a repeat region that binds
to actin ﬁlaments (30, 31, 33, 34). The C-terminal half of the
protein functions as a signaling adapter and consists of an extended
proline-rich region and a C-terminal SH3 domain. The proline-rich
region contains sites for immunoreceptor-induced tyrosine phos-
phorylation (35–37) and provides docking sites for several SH3 and
SH2 domain-containing proteins, including Lck, Vav1, PLCg1, and
Itk (38–40). The HS1 SH3 domain mediates binding to the WIP/
WASp heterodimer (28). The SH3 domain of cortactin also binds to
dynamin 2 (41), though this interaction has not been shown for
The full scope of HS1 function within leukocyte lineages is not
known. Mutations and polymorphisms in HS1 are associated with
autoimmune disease (42, 43), though the mechanistic basis for this
association has not been determined. Aberrant HS1 expression
and/or phosphorylation in B cells are associated with chronic
lymphocytic leukemia (44–46). Deﬁciency for HS1 is associated
with disruption of actin responses and Ca
signaling events lead-
ing to IL-2 promoter activation in T cells (39, 40), with defects
in activation-induced cell death in B cells (47, 48), and with
defects in chemotaxis and cytolysis in NK cells (49).
In comparison with lymphocytes and NK cells, much less is
known about HS1 function in professional APCs. As we recently
showed, wild-type (WT) murine bone marrow-derived dendritic
cells (BMDCs) do not express cortactin, and DCs from HS1
mice lack both cortactin family members (28). These cells exhibit
defects in podosome organization and lamellipodial protrusion,
resulting in a diminished directional persistence during chemo-
taxis. Because several studies have demonstrated a role for cor-
tactin in endocytosis (50–56), we asked whether HS1 expression
is required for Ag uptake and presentation by DCs. Our results
demonstrate that HS1 expression is essential for efﬁcient Ag
processing and presentation to T cells and that this is due, at least
in part, to a requirement for HS1 in receptor-mediated endocytosis
of protein Ags.
Materials and Methods
mice (48) were fully back-crossed onto C57BL/6J or BALB/c
backgrounds. DO-11.10 mice, which express a TCR speciﬁc for chicken
, and C57BL/6J mice were obtained from The Jackson
Laboratory. O TII mice, which express a TCR speciﬁc for chicken O VA
, were obtained from Taconic. Mice were housed under pathogen-
free conditions in the Children’s Hospital of Philadelphia animal facility.
All studies involving animals were reviewed and approved by the
Children’s Hospital of Philadelphia Institutional Animal Care and Use
Primary cultures of BMDCs were prepared as previously described (57).
Brieﬂy, bone marrow progenitor cells were ﬂushed from femurs and tibia of
2- to 4-mo-old mice, and cells were cultured in IMDM supplemented with
10% FBS, recombinant mouse GM-CSF, glutamine, 2-mercaptoethanol, and
penicillin/streptomycin. Medium was replaced every 2 d, and cells were
used between 6 and 8 d of culture, when .70% of the cells are CD11c
Maturation of BMDCs was induced by culturing for 16–24 h in the presence
of 100 ng/ml LPS (Escherichia coli 055:B5; Sigma) or, where indicated,
500 nM CpG (ODN 1668) or control oligonucleotides (InvivoGen).
T cells from DO-11.10 or OTII mice were prepared
as described previously (40). Brieﬂy, CD4
T cells were enriched from
spleens and lymph nodes by negative selection with anti-CD8 and anti-
MHC class II using anti-rat IgG magnetic beads (Qiagen). The T cell
hybridoma N15, which expresses a TCR speciﬁc for VSV8/H2-Kb (58),
was obtained from Dr. E. Reinherz (Dana-Farber Cancer Institute, Boston,
MA) and cultured in RPMI 1640, 10% FCS, 0.1 mg/ml G418, and 0.2 mg/
ml hygromycin. N15 cells were incubated overnight with PMA (10 ng/ml)
prior to use in Ag presentation assays, a treatment necessary for optimal
activation (58). Maximal activation of N15 cells was measured by cross-
linking with plate-bound anti-CD3.
Analysis of BMDC maturation
For ﬂow cytometry, BMDCs were incubated with rat anti-mouse CD16/
CD32 (clone 2.4G2) to block FcgR, followed by staining with the fol-
lowing mAbs (all from BD Biosciences): allophycocyanin-conjugated
hamster anti-mouse CD11c, FITC-conjugated hamster anti-mouse CD40,
PE-conjugated rat anti-mouse mAbs to CD80, CD86, and MHC class I and
class II. Nonspeciﬁc staining was deﬁned using isotype-matched Abs.
Cells were analyzed on a FACSCalibur (BD Biosciences) using FlowJo
software (Tree Star). To measure TNF-a production, BMDC culture
supernatants were harvested at the indicated times and assayed by ELISA
Protein binding studies and Western blotting
For GST-pulldown studies, recombinant GST-tagged SH3 domain of HS1
(aa 415–486) and a variant bearing an inactivating mutation (W456Y) were
generated in bacteria as described (28). BMDCs were lysed in Nonidet P-
40 (NP-40) lysis buffer (50 mM Tris-HCl pH 8, 1% NP-40, 100 mM NaCl,
5 mM EDTA, 0.5 mM CaCl
, protease inhibitors, 1 mM Na
, and 5
mM NaF). Lysates were clariﬁed by centrifugation and exposed to gluta-
thione resin-bound GST fusion proteins for 2 h. Beads were washed in
NP-40 lysis buffer, eluted, and analyzed by Western blotting. For coim-
munoprecipitation studies, 293T cells were transiently transfected with
FLAG-tagged HS1 (FLAG.HS1 FL), an N-terminal deletion mutant
lacking amino acids 1–335, which contain the Arp2/3 complex and actin
binding regions (FLAG.HS1DN) or a variant of FLAG.HS1 DN bearing the
W456Y mutation (FLAG.HS1DN
). In addition, cells were transfected
with either enhanced GFP-tagged dynamin 2 (GFP.Dyn2 FL) or a dynamin
2 mutant lacking the proline-rich domain (GFP.Dyn2 DPRD) (41) (gifts
from Drs. H. Cao and M. McNiven, Mayo Clinic, Rochester, MN). Cells
were lysed in NP-40 lysis buffer, and HS1 was immunoprecipitated using
M2 anti-FLAG agarose (Sigma).
For Western blotting, proteins were separated by SDS-PAGE, transferred
to nitrocellulose, and blocked in 3% BSA in PBS. Blots were probed with
primary Abs as indicated. Rabbit anti-mouse HS1 was described previously
(28, 40). Goat anti-dynamin 2 (C-18) and rabbit anti-WIP (H-224) were
from Santa Cruz, rabbit anti-WASp was from Upstate, rabbit anti-GFP was
from Invitrogen, and mouse anti-GAPDH (6C5) was from EMD Chem-
icals. Primary Abs were detected with secondary Abs coupled to IR800
(Rockland) or Alexa Fluor 680 (Invitrogen) and visualized using an Od-
yssey Imager (Licor). Quantitation was performed within the linear range.
Macropinocytosis and phagocytosis assays
To measure macropinocytosis, BMDCs were washed and resuspended at
8 3 10
/ml in IMDM containing 0.5% BSA. Cells were mixed with pre-
warmed FITC–dextran (70,000 m.w.; Invitrogen) or Lucifer yellow (In-
vitrogen) at 1 mg/ml ﬁnal concentration and incubated at 37˚C for the
indicated times. After washing three times with cold FACS buffer (5%
FBS, 1 mM EDTA in PBS), internalized tracers were measured by ﬂow
cytometry. Time points were performed in duplicate, and mean ﬂuores-
cence intensity values were calculated.
To analyze phagocytosis, BMDCs were plated on coverslips in 6-well
plates and cultured overnight. Media was replaced with ice-cold media
containing 25 mg/ml FITC–zymosan (Invitrogen) and incubated for 1 h at
4˚C. After washing three times with cold media, prewarmed media was
added to initiate uptake. Cells were incubated at 37˚C for the indicated
times and then ﬁxed in 3% paraformaldehyde/PBS. Cells were perme-
abilized with 0.3% Triton X-100, blocked with 0.05% saponin/1.25% ﬁsh
skin gelatin in Tris buffered saline, and labeled with Alexa Fluor 594–
phalloidin or with anti-LAMP1 (1D4B) followed by Alexa Fluor 647 anti-
Rat Ig. The 1D4B mAb developed by Dr. J.T. August was obtained from
the Developmental Studies Hybridoma Bank, developed under the auspices
of the National Institute of Child Health and Human Development and
maintained by The University of Iowa Department of Biology (Iowa City,
IA). Cells were imaged using a Leica SP2 confocal microscope. Zymosan
particles were scored as internalized if they were surrounded by phalloidin
or LAMP1 staining.
The Journal of Immunology 5953
Ag presentation assays
VSV8 peptide (RGYVYQGL) from the vesicular stomatitis virus N protein
peptide (ISQAVHAAHAEINEAGR) from chicken OVA
were synthesized at the University of Chicago peptide facility, puriﬁed by
HPLC, and veriﬁed by mass spectrometry. OVA protein was purchased
from Worthington and further puriﬁed by size exclusion chromatography
to ensure removal of contaminating peptides and LPS. For phagocytosis
studies, latex beads (polystyrene 3-mm microspheres; Polysciences) were
freshly coated by overnight incubation with the indicated ratios of OVA
and BSA, maintaining a concentration of 10 mg/ml total protein. Com-
plexes of the VSV8 peptide and the chaperone glucose regulated protein of
94 kDa (GRP94) were generated as described in Ref. 57. The N-terminal
portion of GRP94 (N1-355), is sufﬁcient to mediate peptide binding,
receptor-mediated uptake by DCs, and cross-presentation of peptide Ags to
T cells (57). N1-355 was expressed in SF9 cells and puriﬁed by afﬁnity and
size exclusion chromatography under LPS-free conditions as detailed in
Vogen et al. (59). Peptide–GRP94 complex was prepared as previously
described (57, 59). Brieﬂy, N1-355 and VSV8 were mixed in buffer A (20
mM HEPES, pH 7.2, 150 mM NaCl, 10 mM KCl, 1 mM MgCl
), heat shocked for 10 min at 50˚C and then incubated at room
temperature for 30 min. The complex was separated from unbound peptide
by size exclusion chromatography.
To me asure Ag presentation on MHC class II molecules, BMDCs were
harvested and replated at 5 3 10
cells/well in ﬂat-bottom 96-well plates
in quadruplicat e. Cells were then pulsed with either OVA protein or
peptide at th e indicated concentrations for 4 h. To control for
contaminating peptide in the OVA protein, some DCs were pre-ﬁxed with
paraformaldehyde prior to addition of protein. Where indicated, mannose
receptors were blocked by incubation with 3 mg/ml mannan from Sac-
charomyces cerevisiae (Sigma) for 1 h prior to addition of soluble OVA.
For presentation of phagocytos ed OVA, cells were incubated for 4 h with
OVA/BSA-coated beads at a ratio of 60 beads per cell. Ag-pulsed DCs
were washed and cocultured with 2 3 10
T cells for 24 h. Cells and
supernatants were lysed together, and IL-2 was measured by ELISA
(R&D Systems). Presentation on MHC class I molecul es was performed
similarly, except that BMDCs were pulsed with N1-355–VSV8 complex
or free VSV8 peptide for 3–4 h, and then an equal number of N15 hy-
bridoma cells was added to each well. IL-2 released after 24 h was
measured by ELISA.
Biochemical analysis of receptor-mediated endocytosis
Seventy-ﬁve to one hundred micrograms of GRP94 N1-355, mouse trans-
ferrin, or OVA (US Biological) were each labeled with 1.0 mCi [
(Amersham) in Tris buffered saline, pH 7.2, using IODOBEADS (Amer-
sham) according to the manufacturer’s recommendations. The unbound
I]iodine was removed on Biogel P10 columns (Bio-Rad, Hercules,
CA). Final sp. act. was 2000–6000 cpm/ng protein. To assay receptor-
mediated endocytosis, BMDCs were harvested, washed with HBSS,
0.1% BSA, and adjusted to 1 3 10
/ml. Cells were then loaded with
I]transferrin, or [
I]OVA (10 mg/ml) for 1 h at 4˚C.
After washing with ice-cold HBSS/BSA, cells were resuspended in the
same buffer, and aliquots of 1 3 10
cells each were warmed to 37˚C for
the indicated times before returning to 4˚C. Surface-bound transferrin
was stripped by mixing cells with an equal volume of acid (0.25 M acetic
acid, 0.5 M NaCl, pH 2.3) for 6–10 s, followed by immediate neutrali-
zation with 50 ml 1 M sodium acetate, pH 8. Surface-bound GRP94 or
OVA was stripped by treatment with 1.25 mg/ml pronase (Roche) in
DMEM for 45 min at 4˚C. After stripping, cells were washed, and cell-
associated radioactivity was measured on a gamma-counter.
WT and HS1
BMDCs were grown on 35-mm tissue culture dishes. To
inhibit dynamin, some dishes were treated with 80 mM of the dynamin
inhibitor dynasore monohydrate (Sigma) for 60 min at 37˚C in serum-free
media. Cells were then washed with HBSS and ﬁxed as monolayers with
2.5% glutaraldehyde, 50 mM sodium cacodylate, pH 7.4, containing 1.25
and 1.25 mM CaCl
Cells were washed with 100 mM sodium
cacodylate, postﬁxed in 2% aqueous OsO
followed by 2% aqueous uranyl
acetate, and dehydrated in graded EtOH. Dishes were then processed using
a modiﬁcation of Grifﬁths et al. (60) to release cell monolayers and a thin
layer of polystyrene, and cell sheets were embedded in EPON-812 (Elec-
tron Microscopy Sciences). Ultrathin cross-sections of the cell sheets
were contrasted with aqueous 1.5% uranyl acetate followed by Reynold’s
lead citrate and viewed using a JEOL 1011 electron microscope operated
at 100 kV. Images were acquired with an Orius CCD camera using Digital
Micrograph software (Gatan).
To label receptor-mediated endocytic compartments, cells were allowed
to internalize 50 mg/ml mouse transferrin–HRP (Thermo Scientiﬁc) for 15
min at 37˚C. To label the ﬂuid-phase pathway, cells were allowed to in-
ternalize 15 mg/ml soluble HRP (Sigma) for 30 min at 37˚C. In both cases,
cells were washed and ﬁxed with glutaraldehyde as described previously.
After ﬁxation, cells were incubated for 30 min with 1 mg/ml 3,39-dia-
minobenzidine (DAB; Sigma) and 0.01% H
in 200 mM sodium
cacodylate, pH 7.4. Cells were then washed with 100 mM sodium caco-
dylate and incubated with 2% aqueous OsO
to generate an electron-dense
HRP–DAB reaction product and postﬁx the cells. Subsequent processing
was as described earlier.
To determine the number of endocytic invaginations, cell proﬁles were
examined at random at 360,000 or higher magniﬁcation, and structures
were counted by an individual blinded to experimental conditions. Shallow,
wide-neck pits were distinguished from non-pit regions of the membrane
on the basis of clearly deﬁned edges, relatively regular size, and, in some
cases, an obvious clathrin coat. Pits and wide-neck invaginations were
collectively deﬁned as structures where the width of the structure at the
plasma membrane was larger than its depth. Narrow-neck invaginations
were deﬁned as structures where the width of the mouth was less than the
diameter of the invagination. Structures of an unclear nature were excluded
from analysis. To determine density of these structures as a function of
plasma membrane length, low-magniﬁcation (35000) micrographs were
captured, and the total length of the plasma membrane in each cell proﬁle
was measured using ImageJ v.1.42 (National Institutes of Health). Filo-
podia were excluded from analysis, as these were difﬁcult to measure ac-
curately and were almost never observed to have endocytic invaginations.
HS1 is not required for differentiation or maturation of DCs
One of the key changes occurring during DC maturation is the
reprogramming of actin regulatory pathways (13, 61). This in-
volves changes in expression patterns of actin regulatory proteins
(62, 63) and functional downregulation of Rho family GTPases
that drive the formation of branched actin ﬁlaments. We recently
showed that DCs express HS1, a protein that promotes the for-
mation of branched actin ﬁlaments in a Rho GTPase-independent
manner (28). To ask whether HS1 expression is also downregu-
lated upon DC maturation, we cultured BMDCs from WT mice
for 24 h with LPS or CpG DNA to induce maturation and ana-
lyzed cell lysates by Western blotting for HS1. As shown in Fig.
1A and 1B, mature DCs do not downregulate HS1 expression.
Instead, these cells express ∼2-fold more HS1 than immature
DCs, indicating that HS1 expression is upregulated as part of the
We showed previously that BMDCs from HS1
both HS1 and the related protein cortactin (28). To ask whether
HS1 is required for DC development and/or maturation, BMDCs
generated from WT and HS1
mice were cultured for 24 h with
or without LPS or CpG DNA, and surface marker expression was
analyzed by ﬂow cytometry. Although the proﬁles of individual
surface markers varied somewhat between DC preparations, cul-
tures from HS1
and WT mice generated equivalent numbers
cells, and maturation-induced upregulation of MHC
molecules and costimulatory proteins was similar in the two pop-
ulations (Fig. 1C and data not shown). Secretion of TNF-a was
also comparable in the two populations of cells (Fig. 1D). Taken
together, these results show that HS1 is not required for devel-
opment or maturation of BMDCs.
HS1 is required for presentation of protein Ags
We next asked whether HS1 is required for Ag presentation. As
shown in Fig. 2A,HS1
DCs could present OVA peptide efﬁ-
ciently to CD4
T cells, as measured by IL-2 production by T cells
from DO-11.10 mice. However, if HS1
DCs were pulsed instead
with intact OVA protein, they activated T cells much less efﬁciently
than WT DCs. Although Ag insolubility precluded our ability to
reach plateau levels of T cell activation, comparison of the Ag dose-
5954 HS1-DEPENDENT Ag PRESENTATION IN DENDRITIC CELLS
response shows that HS1
DCs required 30- to 100-fold more
protein than WT DCs to achieve comparable levels of T cell acti-
vation (Fig. 2B). Similar results were obtained with OTII cells and
BMDCs on the C57BL/6 background (Fig. 2C). To analyze
Ag presentation on MHC class I molecules, we took advantage of
our experience working with the endoplasmic reticulum chaperone
GRP94. We showed previously that GRP94 binds to various pep-
tides, and the complex is taken up by DCs through a receptor-
mediated process and transported to degradative organelles (57).
GRP94-bound peptides then escape the endocytic pathway and are
loaded onto MHC class I molecules in the endoplasmic reticulum.
As shown in Fig. 2E and 2F,HS1
DCs presented VSV8 peptide
efﬁciently to the H2-K
–restricted hybridoma N15, whose TCR is
speciﬁc for VSV8. However, presentation of GRP94–VSV8 peptide
complexes to this hybridoma was 10-fold less efﬁcient than pre-
sentation by WT DCs. Taken together, these results indicate that
HS1 is required for the uptake or processing of protein Ags, and
thereby for their presentation, but is not needed for later events
leading to productive T cell activation.
HS1 is required for efﬁcient receptor-mediated endocytosis
DCs take up Ag by multiple mechanisms, including phagocytosis,
macropinocytosis, and receptor-mediated endocytosis. Because
phagocytosis and macropinocytosis are heavily dependent on actin
polymerization to mobilize large pseudopodial extensions, we ﬁrst
asked whether HS1 is required for Ag uptake through these
pathways. To assess macropinocytosis, immature WT and HS1
DCs were allowed to take up FITC–dextran (Fig. 3A) or Lucifer
yellow (Fig. 3B), for various times, and uptake was assessed by
ﬂow cytometry. In both cases, the time course of ﬂuid phase
uptake was identical in control and HS1-deﬁcient cells. To ask
whether the maturation-associated downregulation of macro-
pinocytic activity is intact in HS1
DCs, mat ure DCs were
generated by culturing overnight in the presence of LPS. As re-
ported previously (13), WT DCs showed diminished ﬂuid-phase
endocytosis upon maturation (Fig. 3B). Mature HS1
(open squares) behaved similarly, indicating that HS1 is not re-
quired for reprogramming of the macropinocytic pathway.
To test the role of HS1 in phagocytosis, WT and HS1
were incubated with FITC-conjugated zymosan particles at 4˚C,
warmed to 37˚C to allow uptake, and analyzed by immunoﬂuo-
rescence microscopy. As shown in Fig. 3C, both WT and HS1
DCs exhibited F-actin–rich phagocytic cups at early times of in-
ternalization. To assess the kinetics of phagocytosis, cells ﬁxed at
various times after warming were labeled with ﬂuorescent phal-
loidin to deﬁne cell boundaries, and the percentage of cells with
FIGURE 1. HS1
DCs differentiate normally in response to LPS stimulation. A, HS1 expression in immature and mature DCs. BMDCs from WT and
mice were stimulated with 100 ng/ml LPS, 500 nM CpG, or control oligonucleotides. After 24 h of stimulation, cell lysates were made and blotted
with anti-HS1. GAPDH was used as a loading control. B, Quantitation of HS1 expression in DCs. Bands from blots as in A were scanned, and the intensity
of HS1 was normalized to that of GAPDH. Data represent means 6 SD from at least three independent experiments. *p , 0.05, **p , 0.01 (relative to
untreated WT DCs). C, BMDCs from WT or HS1
mice were stimulated for 24 h with 100 ng/ml LPS to induce maturation. The cells were then analyzed
by ﬂow cytometry for expression of the indicated surface proteins. Shaded proﬁles, isotype control; solid lines, unstimulated immature DCs; dashed lines,
LPS-stimulated DCs. D,WT(s) and HS1
(n) BMDCs were stimulated with LPS for the indicated times, and TNF-a in the culture supernatants was
measured by ELISA. Data represent means 6 SD from replicate wells of one representative experiment.
The Journal of Immunology 5955
internalized FITC–zymosan was determined. As shown in Fig.
3D, the kinetics of zymosan internalization were unaffected by
HS1 deﬁciency. Moreover, labeling for the lysosomal protein
LAMP1 showed that internalized particles were transported to
lysosomal compartments with normal kinetics (data not shown).
Similarly, phagocytosis of apoptotic cell debris or of ﬂuorescent
Listeria were not impaired in HS1
DCs (data not shown).
Taken together, these studies show that HS1 is not required for
macropinocytosis or phagocytosis by DCs.
We next asked whether receptor-mediated endocytosis is de-
pendent on HS1. Initially, we tested endocytosis of transferrin, with
is taken up by transferrin receptors via clathrin-coated vesicles
(64). To obtain a quantitative measure of transferrin internaliza-
tion, WT and HS1
DCs were surface-labeled with [
ferrin at 4˚C, washed, and warmed to 37˚C for various times.
Residual surface-bound transferrin was then removed by washing
at low pH, and cell-associated radioactivity was determined. As
shown in Fig. 4A, HS1
DCs internalized transferrin at only 30–
40% of the efﬁciency of WT cells. The diminished uptake was
apparent both in terms of the rate of uptake and the overall amount
of transferrin taken up. Although diminished ligand uptake can
theoretically reﬂect either reduced endocytosis or enhanced ligand
recycling, starting transferrin receptor levels were similar in WT
DCs, and the initial rates of endocytosis were already
signiﬁcantly different, arguing against a signiﬁcant contribution
from differential recycling. Thus, we conclude that the diminished
overall uptake of transferrin by HS1
DCs reﬂects a defect in
initial endocytic internalization. OVA is taken up in a receptor-
mediated fashion by the mannose receptor (65, 66), and GRP94 is
taken up by CD91 and other scavenger receptors (57, 67–70). To
ask whether HS1 deﬁciency also leads to reduced endocytosis of
these proteins, we analyzed their internalization using the same
assay. As shown in Fig. 4B and 4C, uptake of both OVA and
GRP94 was signiﬁcantly reduced in HS1
DCs, indicating that
the requirement for HS1 is a general one.
As an independent means of assessing endocytosis, HS1
were incubated for 15 min in the presence of transferrin–HRP.
Cells were then ﬁxed and processed for cytochemistry and elec-
tron microscopy. As shown in Fig. 5A, WT BMDCs showed nu-
merous vesicles and tubular endosomes ﬁlled with HRP reaction
FIGURE 2. HS1 is required for presentation
of protein Ags. A and B, BMDCs cultured from
WT and HS1
mice on the BALB/c back-
ground were pulsed with the class II-restricted
peptide (A) or with the whole OVA
protein (B) at the indicated doses for 4 h. They
were then cocultured with OVA-speciﬁc DO-
11.10 T cells, and 24 h later, IL-2 levels in the
culture supernatants were measured by ELISA.
C, BMDCs cultured from WT and HS1
on the C57BL/6 background were incubated in
the presence or absence of mannan to block
mannose receptors, and then OVA protein was
added, so that any residual uptake would be
by macropinocytosis. Ag presentation to OTII
T cells was then assayed as in A and B.In
this experiment (representing two of four rep-
licates), mannan completely blocked the re-
sponse, indicating that even very high doses
of OVA are taken up almost exclusively by re-
ceptor-mediated endocytosis. s, WT DCs with-
out mannan; n, HS1
DCs without mannan;
D, WT DCs with mannan; ♦, HS1
mannan. Inset, Data from a separate experiment
(representing two of four replicates) where T
cell responses to mannan-blocked DCs were
measurable at high OVA doses. Open bars, WT
DCs with mannan; ﬁlled bars, HS1
with mannan; differences between WT and
DCs in the presence of mannan were
not statistically signiﬁcant at any dose of OVA.
D, WT and HS1
BMDCs (C57BL/6 back-
ground) were allowed to phagocytose latex
beads coated with OVA and serum albumin in
the indicated ratios, holding total protein and
bead number constant. OTII T cell responses
were then measured. E and F, WT and HS1
BMDCs were pulsed with free VSV8 peptide
(E) or VSV8 peptide prebound to GRP94 (F)at
the indicated doses for 4 h. VSV8-speciﬁc N15
T hybridoma cells were incubated with Ag-
pulsed BMDCs for 24 h, and IL-2 levels in the
supernatants were measured by ELISA. Data
represent means 6 SD from replicate wells of
one representative experiment.
5956 HS1-DEPENDENT Ag PRESENTATION IN DENDRITIC CELLS
product. By comparison, HS1
BMDCs contained far fewer
endocytic structures (Fig. 5B). Indeed, most cell proﬁles
lacked labeled endosomes; the image shown in Fig. 5B was se-
lected because it contains one rare HRP-positive organelle (ar-
rowhead). Importantly, HS1
DCs took up free HRP by ﬂuid-
phase endocytosis as efﬁciently as WT cells. As shown in Fig. 5C
and 5D, both cell types exhibited numerous macropinosomes con-
taining electron-dense reaction product. Together with the bio-
chemical analysis of ligand internalization, these data show that
HS1 is speciﬁcally required for efﬁcient uptake of ligands via
receptor-mediated endocytosis into early endocytic compartments.
On the basis of our ﬁnding that HS1 is selectively required for
receptor-mediated endocytosis in DCs, we asked whether the Ag
presentation defects in HS1
DCs are limited to Ags taken up
via this pathway. To facilitate comparisons, we targeted the same
protein, OVA, for uptake via different endocytic mechanisms.
First, cells were incubated with soluble OVA in the presence of
excess mannan to inhibit receptor-mediated endocytosis and drive
ﬂuid-phase uptake. As shown in Fig. 2C, mannan treatment
strongly inhibited Ag presentation, conﬁrming that OVA is taken
up by receptor-mediated endocytosis, even at very high doses. In
two of four replicate experiments, no residual Ag presentation
could be detected, making it impossible to assess the effects of
HS1 deﬁciency on presentation of OVA taken up by ﬂuid-phase
endocytosis. In two of four experiments, although T cell activation
in response to mannan-treated DCs was low (∼10% of that ob-
served in the absence of mannan), it was measurable at the highest
doses of OVA (Fig. 2C, inset). In such experiments, WT and
DCs activated T cells equally well at all doses of OVA
where a response could be measured. We conclude that HS1
DCs are competent to present OVA taken up by ﬂuid-phase en-
docytosis. We also tested the presentation of phagocytosed OVA
using latex beads coated with varying ratios of OVA protein and
serum albumin. As shown in Fig. 2D, the response to phagocy-
tosed OVA was similar in T cells activated by WT and HS1
DCs. Taken together, these studies show that the requirement for
HS1 in presentation of protein Ags is linked to its role in receptor-
mediated endocytosis and that HS1 is not needed for presentation
of Ags taken up by other routes.
HS1 binds to dynamin 2 and promotes formation of plasma
The HS1 homolog cortactin has been shown to interact through its
SH3 domain with several proteins that promote endocytosis, in-
cluding WASp, WIP, and dynamin 2 (41, 52, 71). In particular,
cortactin interacts physically and functionally with dynamin 2 to
drive the internalization of endocytic vesicles (51, 56, 72–75). We
have shown that HS1 binds via its SH3 domain to the actin reg-
ulatory proteins WASp and WIP (28). To ask whether HS1 in-
teracts in a similar way with dynamin 2, we probed DC lysates
with the GST-tagged HS1 SH3 domain and blotted the bound
proteins for dynamin 2. WASp and WIP were used as positive
controls. As shown in Fig. 6A, all three proteins are associated
with the HS1 SH3 domain. Mutation of tryptophan 465, a residue
required for canonical binding to proline-rich ligands, disrupted
binding, demonstrating the speciﬁcity of these interactions. To
conﬁrm that binding of HS1 and dynamin 2 takes place in intact
cells and to map the relevant region of dynamin 2, we performed
coimmunoprecipitation studies on cells coexpressing epitope-
FIGURE 3. HS1 is not required for macropinocytosis or phagocytosis.
A, Immature WT and HS1
BMDCs were incubated with 1 mg/ml
FITC–dextran for the indicated times at 37˚C, and ﬂuorescence intensity
was assessed by ﬂow cytometry. B, WT and HS1
BMDCs were cul-
tured for 24 h in the absence or presence of 100 ng/ml LPS to induce
maturation. The cells were then incubated at 37˚C in the presence of 1 mg/
ml Lucifer yellow for the indicated times, and ﬂuorescence intensity was
assessed by ﬂow cytometry. C, WT and HS1
BMDCs were incubated
with FITC–zymosan particles (green) at 4˚C and warmed to 37˚C to induce
uptake. Cells were ﬁxed and stained with Alexa Fluor 594–phalloidin to
reveal the actin distribution (red). D, The percentage of cells containing
FITC–zymosan particles over time was determined. Data represent
means 6 SD from replicate coverslips of one representative experiment.
The Journal of Immunology 5957
tagged HS1 and dynamin 2 constructs. As shown in Fig. 6B, full-
length HS1 and dynamin 2 could be coimmunoprecipitated from
cells. This interaction does not require the Arp2/3- or actin-
binding domains of HS1, as an N-terminal deletion mutant lack-
ing these regions bound dynamin 2 efﬁciently. Indeed, on a mole-
per-mole basis, this mutant showed .200-fold more binding to
dynamin 2 compared with that of full-length HS1. The enhanced
binding ability of this HS1 truncation mutant also allowed de-
tection of binding to endogenous dynamin 2 (Fig. 6B, “end”).
Mutation of W465 within the SH3 domain of the HS1 DN mutant
dramatically diminished dynamin 2 binding, conﬁrming that the
SH3 domain of HS1 mediates interaction with dynamin 2. Con-
versely, a dynamin 2 mutant lacking the C-terminal proline-rich
domain (41) failed to coimmunoprecipitate efﬁciently with either
full-length HS1 or the HS1 deletion mutant (Fig. 6B,“DPRD”).
Low levels of association of HS1 and dynamin 2 were detectable
under conditions where SH3–PRD interactions were abrogated,
suggesting that a secondary interaction outside these domains may
exist. Nonetheless, our results indicate that HS1 and dynamin 2
interact primarily through binding of the HS1 SH3 domain to the
proline-rich domain of dynamin 2.
Dynamin is known to drive the scission of plasma membrane
invaginations during receptor-mediated endocytosis, and cells ex-
pressing a dominant-negative dynamin mutant or treated with the
dynamin inhibitor dynasore generate increased numbers of deep,
wide-neck (U-shaped) and narrow-neck (O-shaped) endocytic in-
vaginations (76). To ask whether HS1 is required at the same
step in endocytic trafﬁc, we analyzed plasma membrane invagi-
nations in WT and HS1
BMDCs. Fig. 7A shows examples
of shallow pits and wide-neck invaginations (top row), as well
as narrow-neck invaginations (bottom row) found in WT cells.
The number of these structures per linear micrometer of plasma
membrane was determined as described in Materials and Meth-
ods. As anticipated, treatment of WT cells with dynasore resulted
in a signiﬁcant increase in the surface density of endocytic inva-
ginations (Fig. 7B). These consisted primarily of deep invagina-
tions with a wide neck. In contrast, HS1
DCs showed only
about half as many endocytic invaginations as WT cells, and this
FIGURE 4. HS1 is necessary for efﬁcient receptor-mediated endocyto-
sis. A–C, WT and HS1
BMDCs were incubated with [
I]OVA (B), or [
I]GRP94 (C) at 4˚C for 1 h. Cells were then
washed and warmed to 37˚C for the indicated times. At each time point,
residual surface-bound proteins were removed by acid wash (transferrin) or
pronase treatment (GRP94 and OVA). Radioactivity in the cell pellets was
then determined and uptake calculated. Data represent means 6 SD from
independent experiments, each done in duplicate or triplicate. The number
of binding sites for each protein did not differ between WT and HS1
BMDCs within experimental error. We calculate 6000–8500 binding sites
for transferrin per cell, 20,000–40,000 sites for GRP94, and ∼80,000 sites
FIGURE 5. Analysis of ﬂuid-phase and receptor-mediated endocytosis
by electron microscopy. WT (A, C) or HS1
(B, D) BMDCs were in-
cubated with transferrin–HRP (A, B) for 15 min at 37˚C to load endocytic
compartments via receptor-mediated endocytosis. Alternatively, BMDCs
were incubated with soluble HRP (C, D) for 30 min at 37˚C to load the
endocytic pathway via ﬂuid-phase endocytosis (macropinocytosis). Cells
were ﬁxed and processed with DAB to generate an electron-dense reaction
product and visualized by transmission electron microscopy. Representa-
tive images from three independent experiments are shown. Arrowheads
indicate compartments (early endosomes and macropinosomes) ﬁlled with
electron-dense HRP reaction product. Note the relative paucity of electron-
dense deposits in the HS1
cells when HRP is delivered by receptor-
mediated endocytosis (compare A and B), but not macropinocytosis (com-
pare C and D). Scale bars, 2mm. G, Golgi complex; N, nucleus; PM,
plasma membrane; V, macropinocytic vacuoles.
5958 HS1-DEPENDENT Ag PRESENTATION IN DENDRITIC CELLS
decrease was observed for both wide-neck and narrow-neck invag-
inations. These data indicate that HS1 and dynamin affect distinct
steps of endocytosis in DCs and support a model in which an HS1-
dependent step occurs early in the formation of membrane inva-
ginations, whereas dynamin is needed to complete later steps of
membrane deformation and vesicle scission.
DC sentinel function depends on the ability of these cells to take
up a wide variety of antigenic materials. This is achieved through
a combination of phagocytosis, macropinocytosis, and receptor-
mediated endocytosis (1, 3, 4, 11). We have found that HS1
plays an important role in this process by promoting receptor-
mediated endocytosis of protein Ags. DCs from HS1
show diminished receptor-mediated endocytosis of multiple pro-
teins and impaired presentation of exogenous protein-derived Ags
on both MHC class I and MHC class II.
The requirement for HS1 in Ag trafﬁcking and presentation
appears to be restricted to receptor-mediated endocytic pathway.
DCs presented peptide Ags efﬁciently, indicating that
HS1 is not required for actin-dependent events at the DC side of
the immunological synapse (27, 77–79). Moreover, although there
is strong evidence that actin polymerization is important for mac-
ropinocytosis and phagocytosis (12, 14, 80), HS1 expression was
not required for these processes. By extension, as we ﬁnd no cor-
tactin expression in murine BMDCs (28), cortactin family mem-
bers generally are dispensable. Both macropinocytosis and phago-
FIGURE 6. The SH3 domain of HS1 interacts with the proline-rich
domain of dynamin 2. A, GST alone, the SH3 domain of HS1 fused to
GST, or the inactive SH3
mutant fused to GST was immobilized on
glutathione resin and incubated with DC lysates. Bound proteins were
analyzed by SDS-PAGE and immunoblotting with anti-dynamin 2, anti-
WASp, or anti-WIP. Total, protein expression in whole-cell lysates. B,
293T cells were transiently transfected with full-length GFP-tagged dy-
namin 2 (GFP.Dyn2 FL, black arrowheads) or a mutant lacking the pro-
line-rich domain (DPRD, gray arrowheads), together with FLAG-tagged
HS1 (FLAG.HS1 FL), an N-terminal deletion mutant of HS1 lacking the
actin regulatory region (DN), or the DN mutant bearing a point mutation in
the SH3 domain (DN
). Cells were lysed, immunoprecipitated with
anti-FLAG, and Western blots were probed as indicated. Note that the
DPRD mutant is not detected by the anti-dynamin 2 (which was generated
against this C-terminal region of dynamin 2) but is readily detectable in the
WCL using anti-GFP Ab. end, endogenous dynamin 2; *HS1 variants.
WCL, whole-cell lysate from which immunoprecipitation was performed.
FIGURE 7. HS1 promotes endocytic invagination. A, Electron micro-
graphs illustrating various stages in the process of coated vesicle forma-
tion. All images are from WT BMDCs; structures in HS1
comparable in morphology. Top row, Examples of shallow pits and wide-
necked invaginations. Bottom row, Examples of narrow-neck invagina-
tions. B, Quantitation of the frequency of the structures shown in A in
WT cells, WT cells treated with 80 mM dynasore, and HS1
proﬁles were photographed at random, and the number of structures in
each category per linear micrometer of plasma membrane was determined
as described in Materials and Methods. Values represent mean 6 SEM
from three to four independent experiments, totaling 66–68 cells and 100–
500 pits or invaginations per experimental condition. ***p , 0.001 (rel-
ative to untreated WT cells).
The Journal of Immunology 5959
cytosis are downregulated upon DC maturation (13, 65), whereas
receptor-mediated endocytosis proceeds unabated in mature DCs
(13, 15, 81). Phagocytosis and macropinocytosis depend on the
actin nucleating proteins WASp and WAVE, and the downregu-
lation of these processes is linked to diminished activation of the
Rho family GTPase CDC42 upstream of these events (13). No-
tably, HS1 differs from WASp and WAVE in that it does not de-
pend on activation by Rho GTPases, and so is not subject to
downregulation at this level. Indeed, we ﬁnd that HS1 expression
is modestly upregulated upon DC maturation. Moreover, HS1 is
enriched in podosomes, which are largely dissolved on DC mat-
uration (28), presumably increasing the pool of HS1 available to
support endocytic functions in mature DCs.
Our analysis of HS1
DCs shows a 2-fold decrease in the
maximal uptake of endocytosed proteins, yet these cells require
30- to 100-fold more protein Ag to achieve the same level of T cell
activation as that of WT DCs. The assay we used for receptor-
mediated endocytosis measures only a single, synchronized round
of protein uptake. When multiplied by the many rounds of en-
docytosis that take place within the 4-h period of the Ag pulse, the
defects we observe could readily account for at least 30- to 50-fold
reduction in overall Ag uptake. Thus, the requirement for HS1 to
facilitate endocytosis is likely to be the primary cause of the di-
minished Ag presentation that we observe in HS1-deﬁcient DCs.
The ﬁnding that Ags taken up by phagocytosis and macro-
pinocytosis are presented efﬁciently further supports this conclu-
sion. Nonetheless, it is possible that HS1 is also required for
additional intracellular steps of Ag trafﬁcking. Testing this pos-
sibility will require quantitative analysis of the fate of the endo-
cytosed pool of protein.
Although the DC literature tends to focus on phagocytosis and
macropinocytosis, receptor-mediated endocytosis is also an im-
portant means of Ag uptake. Typically, it is argued that receptor-
mediated endocytosis is particularly important at low ligand
concentrations. While this is no doubt true, our side-by-side
comparison of presentation of OVA-derived peptides after up-
take by receptor-mediated endocytosis and macropinocytosis
shows that receptor-mediated uptake is dominant even at very high
Ag doses. DCs express numerous cell surface receptors that
preferentially bind non-self molecules and take them up in a highly
efﬁcient manner. OVA is taken up by mannose receptors (65,
66) (Fig. 2C), which is important for uptake of pathogen-derived
glycoproteins (82–84). Uptake and cross-presentation of GRP94–
peptide complexes also require receptor-mediated endocytosis,
though the relevant receptor(s) remain controversial (57, 67–70).
Other receptors also play an important physiological role. Among
these are the C-type lectin DC-SIGN, which promotes the endo-
cytosis of HIV and other viruses and their presentation on MHC
class I (85–88). The multilectin receptor DEC205 plays a similar
role (89, 90). Even FcgRs, which are best known for their role in
phagocytosis, carry out receptor-mediated endocytosis of small
immune complexes (91–94). CD1b recycling occurs via receptor-
mediated endocytosis and continues in mature DCs (81), a phe-
nomenon that has been proposed to be the basis of effective
presentation of lipid Ags from mycobacteria, under conditions
where presentation of Ags on MHC class II fails (95). We show
here that HS1 is required for receptor-mediated uptake via three
distinct receptors, and it seems likely that this is a general re-
quirement. Going forward, it will be interesting to ask whether
HS1 expression has an important impact on DC function in vivo,
particularly in the context of viruses, lipid Ags, or DC vaccines
that rely on receptor-mediated uptake. Moreover, as HS1 is also
expressed in B cells, it will be interesting to ask whether BCR-
dependent uptake of Ags is also HS1 dependent.
HS1 functions both as an actin regulatory protein and an adapter
molecule, and it seems likely that both of these functions are called
into play to promote receptor-mediated endocytosis. We show that
HS1 interacts with dynamin 2 through binding of its C-terminal
SH3 domain to the proline-rich domain of dynamin 2, the same
mechanism used to mediate cortactin–dynamin 2 interaction (41).
Notably, however, the endocytic phenotype of HS1
not mirror that of cells treated with the dynamin GTPase inhibitor
dynasore. In keeping with studies in other cell types (76), we
found that treatment of DCs with dynasore led to the accumula-
tion of deep endocytic invaginations. The accumulation of wide-
necked (U-shaped) structures was predominant, though narrow-
necked (O-shaped) endocytic invaginations were also observed.
The latter structures are also seen in cells expressing dominant-
negative dynamin mutants and are thought to represent a require-
ment for dynamin in the ﬁssion of fully formed coated pits (9, 96).
The U-shaped invaginations represent an earlier endocytic inter-
mediate and are proposed to represent an additional requirement
for dynamin in driving membrane deformation (76). In contrast to
dynamin-inhibited cells, HS1-deﬁcient DCs exhibited abnormally
low numbers of both U-shaped and O-shaped endocytic invagi-
nations. The simplest interpretation of this ﬁnding is that HS1 is
required for an early step in membrane invagination, preceding
the deep invaginations formed by dynamin activity. These early
intermediates are thought to be metastable (97, 98), perhaps
explaining why no clear accumulation of shallow pits was ob-
served in HS1-deﬁcient DCs. This interpretation is consistent with
a two-step model for dynamin function, in which dynamin ﬁrst
serves as regulatory GTPase to ensure vectorial coat assembly
and enhance membrane curvature, and subsequently, in an assem-
bled state, uses GTPase-driven mechano-chemical activity to drive
vesicle scission (96). According to this model, SH3 domain-
containing proteins such as HS1 sense the status of cargo load-
ing, membrane curvature, and/or coat assembly and regulate this
In addition to binding dynamin 2, HS1 activates the Arp2/3
complex, binds actin ﬁlaments, and interacts with the actin regu-
latory proteins WASp and WIP (28, 30). Thus, it is also likely that
HS1 promotes actin-dependent aspects of endocytosis. The liter-
ature regarding the requirement for actin polymerization in en-
docytosis is conﬂicting. In S. cerevisiae, actin recruitment occurs
after clathrin coat formation and provides an essential force for
membrane deformation (99). In mammalian cells, however, results
are variable and depend on the cell type and the nature of the
endocytic structure (100–103). Recent work from the Kirchhausen
group (104–106) has shown that actin regulatory proteins are
recruited to long-lived endocytic structures and are particularly
important for internalization of large structures, such as clathrin
plaques and some virus particles. On the basis of this work, it
seems that actin polymerization provides the force needed to gen-
erate oversized clathrin-coated structures and to overcome other
obstacles, such as high membrane tension (107). Because HS1
promotes the formation of branched-actin ﬁlaments, it may play
a key role in this process. Importantly, the interactions of HS1
with the actin cytoskeleton and dynamin are likely to be coupled.
Indeed, we ﬁnd that interactions between HS1 and dynamin are
dramatically enhanced by deletion of the N-terminal actin regu-
latory region of HS1. This and similar observations regarding HS1
binding to WASp and WIP (28) are compatible with a model in
which the N terminus of HS1 partially restricts availability of the
SH3 domain, such that interaction with the actin cytoskeleton
releases the SH3 domain and promotes binding to dynamin 2
and other molecules. This model explains our inability to coim-
munoprecipitate endogenous dynamin 2 and HS1, as the pool of
5960 HS1-DEPENDENT Ag PRESENTATION IN DENDRITIC CELLS
active HS1 would be selectively associated with the insoluble
actin cytoskeleton. A positive feedback loop may also be at play,
as engagement of the dynamin proline-rich domain b y SH3
domains has been shown to promote dynamin GTPase activity
(108, 109). Such a mechanism would promote the coupled as-
sembly of actin and dynamin at endocytic invaginations.
An important unresolved question is to what extent HS1 function
is distinct from that of its more widely expressed homolog cortactin.
In non-hematopoietic cells, the weight of the evidence indicates that
cortactin plays a key role in endocytic uptake (51, 56, 72–75, 110,
111), although some studies challenge this view (112, 113). Cor-
tactin is recruited to long-lived clathrin-rich regions of the mem-
brane, and kinetic analysis shows that cortactin recruitment peaks
at a late time point, close to the time of dynamin recruitment (72,
104). Furthermore, phosphorylation of cortactin promotes its as-
sociation with dynamin (56, 74), and the tyrosine phosphorylation
sites that have been linked to regulated cortactin function are
conserved in HS1. Given all the biochemical and cell biological
similarities between cortactin and HS1, why did hematopoietic
lineages evolve a distinct family member? There are, in fact, some
important functional differences between HS1 and cortactin. The
structural features necessary for F-actin binding are distinct (31).
Moreover, HS1 is less efﬁcient than cortactin at driving Arp2/3
complex-nucleated actin polymerization (30). Actin binding pro-
motes cortactin interaction with dynamin (74), and our evidence
suggests that the same is true for HS1. Thus, the difference in
mode of actin binding between HS1 and cortactin may translate
into important differences in the mechanisms by which these pro-
teins link the actin cytoskeleton to dynamin function. Because
BMDCs from HS1
mice lack both HS1 and cortactin, they
provide an ideal experimental system in which to carry out com-
parative reconstitution studies to address this possibility.
We thank Drs. H. Cao and M. McNiven for providing dynamin constructs,
Dr. T. Svitkina and the University of Pennsylvania Electron Microscopy
Resource Laboratory for providing access to equipment for electron micros-
copy, and Dr. A. Mantegazza for help with analysis of macropinocytosed
and phagocytosed Ags. We thank Drs. T. Svitkina, M. Marks, and T. Laufer
and members of the Burkhardt laboratory for helpful discussions.
The authors have no ﬁnancial conﬂicts of interest.
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