Available online at www.sciencedirect.com
Cellular Signalling 20 (2008) 1159 – 1168
www.elsevier.com/locate/cellsig
Endogenous Gas6 and Ca 2+ -channel activation modulate phagocytosis by
retinal pigment epithelium
Mike O. Karl a,⁎, Wolfram Kroeger b , Soenke Wimmers a , Vladimir M. Milenkovic c ,
Monika Valtink a,1 , Katrin Engelmann a,2 , Olaf Strauss a,c
b
a
University Eye Hospital Hamburg, Martinistrasse 52, 20246 Hamburg, Germany
Institut fuer Klinische Physiologie, Charite-Universitaetsmedizin Berlin, Campus Benjamin Franklin, Berlin, Germany
c
Experimental Ophthalmology, University Eye Clinic, Franz-Josef-Strauss-Allee 11, Regensburg, 93053, Germany
Received 15 September 2007; received in revised form 27 January 2008; accepted 6 February 2008
Available online 16 February 2008
Abstract
Mutation or loss of MerTK as well as deficiency of αvβ5-integrins, gives rise to retinal-degeneration due to inefficient phagocytosis of
photoreceptor outer-segment fragments by the retinal pigment epithelium (RPE). This study shows that Gas6 expressed endogenously by human RPE
promotes phagocytosis. The RPE expresses Gas6 more highly in vivo and in serum-reduced conditions in vitro than in high-serum conditions,
suggesting a negative-feedback control. An antibody-blockage approach revealed that Gas6-expressing RPE phagocytizes photoreceptor outersegment fragments due to stimulation of MerTK by endogenous Gas6 in vitro. MerTK- and Gas6-antibodies reduced phagocytosis. Blocking L-type
Ca2+-channels with nifedipine inhibited MerTK dependent phagocytosis in vitro. Application of integrin inhibitory, soluble, RGD-containing
peptides or soluble vitronectin reduced L-type Ca2+-channel currents in RPE. Herbimycin A, which reduces phosphorylation of integrin receptorassociated proteins and decreases L-type Ca2+-channel currents in RPE, eliminates the inhibiting vitronectin effect and abolishes phagocytosis. Thus,
Gas6-promoted phagocytosis was inhibited by L-type Ca2+-channel blockage, which in turn may be activated by integrin receptor stimulation. These
results suggest that L-type Ca2+-channels could be regulated downstream of both MerTK and αvβ5-integrin, indicating that the binding and uptake
mechanisms of phagocytosis are part of a converging pathway.
© 2008 Elsevier Inc. All rights reserved.
Keywords: Ca2+-channels; Gas6; Integrin; MerTK; Phagocytosis; Retinal pigment epithelium
1. Introduction
In the retina, the renewal of photoreceptor outer-segments is
a vital task of the contiguous retinal pigment epithelium (RPE).
Phagocytizing thousands of disposed outer-segment fragments
per day makes RPE cells some of the most active phagocytes in
⁎ Corresponding author. Present address: University of Washington, Department
of Biological Structure, 1959 NE Pacific Street, HSB I-400, UW Mail Box 357420,
Seattle, WA 98195, USA. Tel.: +1 206 543 8043.
E-mail address: mail@mikeology.de (M.O. Karl).
1
Present address: Institute of Anatomy, Medical Faculty “Carl Gustav Carus",
University of Technology Dresden, Fetscherstr. 74, 01307 Dresden, Germany.
2
Present address: Department of Ophthalmology, Städtisches Klinikum
Chemnitz gGmbH, Flemmingstr. 2, Chemnitz, 09116 and CRTD / DFG-Center
for Regenerative Therapies Dresden - Cluster of Excellence, Biotechnology
Center, Tatzberg 47/49, 01307 Dresden, Germany.
0898-6568/$ - see front matter © 2008 Elsevier Inc. All rights reserved.
doi:10.1016/j.cellsig.2008.02.005
the human body. Impaired phagocytosis causes inherited retinal
dystrophies [1–3] and contributes to age-related retina degeneration [4–8].
Specific recognition and processing of photoreceptor outersegment fragments during phagocytosis by RPE is regulated by
at least two major pathways, which are highly similar to the
mechanism used by other phagocytes for clearance of apoptotic
cells [3,9,10]. First, αvβ5-integrin receptors are implicated in
photoreceptor outer-segment binding, initiating signaling to
focal adhesion kinase (FAK), Src-kinase and Mer tyrosine
kinase (MerTK) [4,11,12]. Second, the receptor protein-tyrosine
kinase, MerTK, is crucial for phagocytic uptake by RPE [2,13].
Disruption of integrin or MerTK results in deficient clearance
of photoreceptor outer-segment fragments. RPE deficient for
β5-integrin or integrin receptor ligand MFG-E8 retained basal
uptake levels of phagocytic activity in vivo, but lacked the burst
that followed circadian photoreceptor outer-segment fragment
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M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
shedding in wild-type RPE. Rhythmic activation of FAK and
MerTK that mediates wild-type phagocytosis was also
completely absent under both genetic conditions [14,15]. In
contrast, MerTK-deficient RPE still binds photoreceptor outersegments but fails to internalize sufficiently [16–19]. How these
signals and participating modulators regulate phagocytosis
remains incompletely understood in terms of post-receptor
signaling [20–23]. Related pathophysiologic mechanisms are
poorly understood and potential treatments are lacking [24,25].
Increases of [Ca2+]i have been correlated with initiation as well
as termination of phagocytosis [7,26,27], and a pathophysiologic calcium homeostasis has been shown in MerTK-deficient
RPE (Royal College of Surgeons rat RPE). Reported abnormalities of MerTK-deficient RPE, such as increased L-type calcium channel activity [24], are likely secondary to the loss of
MerTK function. RPE cells express L-type Ca2+-channels and
these contribute to changes in [Ca2+]i, which are involved in the
regulation of various RPE functions [23,28].
This study identifies Gas6 and MerTK expression by human
RPE in vivo and in vitro. Phagocytosis was inhibitable by
antibodies to Gas6 or MerTK. Therefore, RPE is a potential
endogenous source of functional Gas6. Furthermore, this study
provides evidence for a role of L-type Ca2+-channels in phagocytosis to transduce signals from particle binding to integrin
receptors and to support respective crosstalk with MerTK to
trigger particle engulfment.
2. Materials and methods
2.1. Cultured cells
Experiments were performed with adult human SV40-transfected RPE, adult
human ARPE-19 and primary human RPE. SV40-RPE, previously characterized
for phagocytosis [21], were cultured with medium F99 (Ham's F12/Medium 199;
Invitrogen) supplemented with 10% (v/v) FCS (Biochrom), 1 mM Na-pyruvate
(Biochrom), 1 µg/ml bovine insulin (Sigma), 50 µg/ml gentamycin and 2.5 µg/ml
amphotericin B; otherwise as indicated. Cells were grown to subconfluence (33 °C,
5% CO2, 95% air) and passaged (1:4) using trypsin/EDTA (0.05%/0.02%;
Invitrogen). Eventually, cells were cryopreserved in liquid nitrogen (10% (v/v)
DMSO in FCS as cryoprotectant). For experiments, 104 cells (≤passage 9)/well
were seeded onto gelatine-coated 96-well plates and cultured (37 °C, 5% CO2, 95%
air). Medium was changed after 24 h and cells were used for experiments after 48 h.
The cell line ARPE-19 (ATTC) was cultured according to the provider's
instructions and medium was changed twice/week. Cells were trypsinized and
seeded on glass coverslips prior to patch-clamp investigation.
Human RPE cells were isolated and cultured by previously published methods
[21,29]. In brief, RPE cells were isolated from human donor eyes used for cornea
preparation (tenets of the Declaration of Helsinki). The eyes anterior segment,
vitreous and retina were removed. The choroid with the attached RPE layer was
separated from the sclera and incubated in 2 ml F99 with collagenase IA/IV
(Sigma; each 0.5 mg/ml) for 1 h (37 °C, 5% CO2). Subsequently, the choroids were
shaken in order to loosen RPE cells, F99 with 10% FCS was added, the cell suspension was passed through a 70 µm cell-strainer (Falcon) and cells were counted
(Beckmann Coulter Counter Z2). Cells were seeded at 104 cells/50 µl/96-well,
cultured (37 °C, 5% CO2) with medium changes 3-times/week and used for
experiments at confluence.
2.2. Immunocyto- and histochemistry
Immunocytochemistry with goat-antibodies to human MerTK and Gas6
(Santa Cruz Biotechnology) was done on tissue sections and cells cultured on
chamber-slides (Nunc Labtek). Cells were washed three times with PBS and
fixed with 50 mM glycine buffer (pH 2.0) in 70% ethanol (− 20 °C, 10 min).
Formalin fixed human eyes were mechanically processed and embedded in
paraffin. Eyes were sectioned; immunolabeled and counterstained with Lillie's
modified Mayer's hematoxylin. After blocking in PBS with 10% FCS monolayers
or sections were stained with a primary antibody for 30 min and processed using an
alkaline-phosphatase detection kit (LSAB+ system-AP, DAKO). A negative
control reaction with non-immune IgG as primary antibody was performed.
Labeled cells were visualized with a standard light microscope (Leica).
2.3. Western blotting
Whole human retina, fresh human RPE, cultured human ARPE19 and human SV40-RPE were lysed (Tris–HCl (pH 7.5) 2.5 mM, NaCl 5 mM, NP-40 1%,
Na-desoxylat 0.5%, Complete mini (Roche) 1 Tablette, PepstatinA 10 µg/ml) and
homogenized. Samples (2 µg) were separated by polyacrylamide (12%) gel
electrophoresis in the Mini-V 8.10 system (Biometra). After blocking in 5% nonfat milk in TBS-T solution (50 mM Tris–HCl (pH 7.5), 150 mM NaCl, 0.1%
Tween 20) for 2 h at room temperature, protein blots were incubated with
primary antibody dissolved in TBS-T at 4 °C overnight. Immunoblotting was
performed with two different polyclonal antibodies recognizing an epitope on the
C-terminus of human Gas6 (goat anti-human, R&D, DY885, 1:100 and goat antihuman, C-20, Santa Cruz, SC-1935, 1:500) and peroxidase-conjugated
secondary antibodies (DakoCytomation) using a Lumi-LightPLUS detection kit
(Roche). Fluorescence was detected by the ChemiImager 5500 system (Alpha
Innotech). Images were optimized for color and contrast with Adobe Photoshop.
Negative controls were performed in the absence of the primary antibody.
2.4. RT-PCR
RT-PCR was conducted on RNA isolated from ARPE19 cell line, fresh
human retina and RPE cells. Total RNA was extracted using the NucleoSpin
RNAII kit (Macherey-Nagel GmbH, Düren, Germany) while reverse transcription was carried out using the RevertAid M-MuLV reverse transcriptase
(Fermentas, St. Leon-Rot, Germany). PCR primers (sense 1, hgas6-5′CGGGCGTGGGGGCCTCAA-3′, and, antisense 1, hgas6-5′-CCGCGATTTTCATGACAGCA-3′) were designed to amplify human Gas6 cDNA sequence
(GenBank accession number NM_000820) in order to determine Gas6 message.
And another primer pair was designed as previously published by Marcandalli
et al. [34] (sense 2 hgas6-5′-CGGGCGTGGGGGCCTCAA-3′, and, antisense 2,
hgas6-5′-TGGTGGCCTCCGGCAAAGA-3′). PCR products were separated on
a 1% agarose gel. The expected PCR product should have the size of 425 bp
without the 129 bp insertion (Gas6 isoform two) or the size of 554 bp with
insertion (isoform one). Quality of the cDNA was tested in an independent PCR
reaction with primers specific for human β-actin transcripts (GenBank accession
number NM_001101) (sense, hβ-actin-5′-GTGGGGCGCCCCAGGCACCA,
and antisense, hβ-actin-5′-CTCCTTAATGTCACGCACGATT), with an
expected PCR product of 540 bp (data not shown). DNA molecular weight
marker was 2-Log DNA Ladder (New England Biolabs, Frankfurt am Main,
Germany) with fragments in kb (from top to bottom 10, 8, 6, 5, 4, 3, 2, 1.5, 1.2,
1, 0.9, 0.8, 0.7, 0.6, 0.5, 0.4, 0.3, 0.2, 0.1). The identity of the amplification
product was confirmed by sequencing.
2.5. Patch-clamp investigation
Patch-clamp recordings were made in the perforated-patch configuration
with K+-free solutions. The bath solution consisted of 130 mM NaCl, 3 mM
TEACl, 10 mM BaCl2, 0.3 mM CaCl2, 0.6 mM MgCl2, 14 mM NaHCO3, 1 mM
Na2HPO4, 33 mM HEPES, and 6 mM glucose (pH 7.2 with Tris) and the pipette
solution contained 100 mM CsCl, 10 mM NaCl, 0.5 mM CaCl2, 2 mM MgCl2,
5.5 mM EGTA, 10 mM HEPES (pH 7.2 with Tris) and 150 µg/ml nystatin.
Currents were measured using an EPC-9 patch-clamp amplifier in conjunction
with TIDA software (HEKA) for electrical stimulation, data storage and
analysis. Stock solutions in citrate buffer were prepared and added to the bath
solution to the final concentration (RGD-peptides and vitronectin, Biochrom).
2.6. Isolation and labeling of photoreceptor outer-segments
Outer-segments were isolated from fresh porcine retinas as previously described [30]. In brief, retinas were pooled and carefully homogenized on ice in
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
phosphate buffer containing Complete™ protease inhibitor (Roche; 1 tablet/50 ml
homogenate). The homogenate was then subjected to density-gradient centrifugation (140,000 ×g, 70 min, 4 °C, Beckman XL80) using a sucrose-gradient (20, 27,
33, 50 and 60% (w/v)). The orange-colored photoreceptor outer-segment fragment
containing fraction was collected, diluted 1:5 with a 20 mM Tris buffer (pH 7.2) and
pelleted by centrifugation (13,000 ×g, 10 min). Protein content of this fraction was
determined (Bradford assay), and particle number was counted using a Coulter
Counter Z2. Photoreceptor outer-segment fragments from 103 pooled retinas were
aliquotted and stored in phosphate buffer pH 7.2, supplemented with 100 mM NaCl
and 2.5% sucrose at −80 °C according to Finnemann et al. [4].
2.7. Phagocytosis assays
Photoreceptor outer-segment fragments were labeled with 100 µg SNAFL®-2
(Molecular Probes)/mg photoreceptor outer-segment protein as described
previously [21,30]. The dual-wavelength dye emits light of different wavelengths
depending on environmental pH. This allows differentiation between photoreceptor outer-segment fragment binding to, and uptake into, the cell. Binding is
detected by elevating extracellular pH, using an extracellular mounting solution
with pH 9. Bound photoreceptor outer-segment fragments display a pale yelloworange fluorescence, whereas ingested display a green fluorescence due to the
acidic pH of phagolysosomes. For assaying phagocytosis, cultures were incubated in an unsupplemented medium F99 for 12 h, otherwise as indicated. After
applying respective experimental conditions, photoreceptor outer-segment fragments were added at a concentration of 5×106/100 µl/well (1.5×107 OS/cm2). Cell
cultures were incubated at 37 °C and rinsed twice with 50 µl PBS (with Ca2+/Mg2+)
per well at the indicated time points before measurements. DAPI nucleic counterstain was applied and fluorescence was measured using a fluorescent multi-well
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plate-reader (Cytofluor®4000; Applied Biosystems) at 360 ± 40 nm excitation and
460± 40 nm emission. Cells were then rinsed twice with 50 µl PBS containing 10%
glycerol (pH 9.0), and another 50 µl fresh solution per well were left during
measurement. Binding of photoreceptor outer-segment fragments was detected by
exciting at 560 ± 10 nm and recording at 640± 20 nm wavelength. Uptake of
photoreceptor outer-segment fragments was detected by exciting at 485± 10 nm
and recording at 580± 25 nm wavelengths. In addition, a flow cytometric technique
recording photoreceptor outer-segment fragment uptake only as described in the
literature was used [30,31]. Therefore, following fluorescence plate-reader
measurements, as described above, cells were rinsed three times in PBS,
trypsinized, pooled from 8 parallel 96-wells, washed twice in PBS and resuspended
as a single cell suspension stored in Cellfix (4 °C, Becton Dickinson). The mean
cellular fluorescence with excitation at 488–515 nm and emission at 550–610 nm
was assayed on a FACScan® (Becton Dickinson) between 1 and 72 h after cell
fixation. Typically, 104 cells were counted per sample using a live gate to exclude
cell fragments, free photoreceptor outer-segment fragments and other unwanted
debris. Cellular autofluorescence was determined for each test by analyzing
cells from wells to which no photoreceptor outer-segment fragments had been
added.
Mean cellular autofluorescence was subtracted from experimental values
and photoreceptor outer-segment fragment counts were normalized for different
cell densities using the fluorescence of DAPI. Neither HA nor NIF treatment in
this study led to a significant change in cell density based on DAPI fluorescence
analysis. Records calculated in relation to the variable F99 resulted in the
phagocytosis rate, if indicated. Measurements were performed in three replicates
using eight 96-well plates for each variable unless indicated otherwise. Mean ±
S.E.M. and significance were calculated using ANOVA; differences of p ≤ 0.05
were considered statistically significant.
Fig. 1. Detection of MerTK and its ligand Gas6 in human retina cryosections. A, C) Parallel sample controls with non-immune IgG as primary antibody gave only a
slight background staining. B, D) Immunolabeling showed MerTK localizing to the RPE and PR. E) Gas6 was detected at the RPE layer. (NFL nerve fiber layer, GCL
ganglion cell layer, IPL inner plexiform layer, INL inner nuclear layer, OPL outer plexiform layer, ONL outer nuclear layer, PR photoreceptor layer, Ø artificial
fracture).
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3. Results
Disruption of all three receptor tyrosine kinases of the TAM
family (Tyro3, Axl, and MerTK) or MerTK alone, but not single
or combined loss of Axl and Tyro3, results in retina degeneration [32]. Absence or mutation of MerTK leads to deficient
photoreceptor outer-segment fragment clearance and subsequent retinal dysfunction [1,2,13]. The potent MerTK-ligand,
Gas6 (growth arrest-specific gene 6), was initially cloned from
serum-starved fibroblasts and shares about 44% sequence identity with protein-S and both share the same complex, multidomain structure [9,33,34]. Both ligands are secreted proteins
present in serum and responsible for the well-known serum
stimulation of phagocytosis by RPE in vitro [17,18,35]. But it
has still been controversial which ligands might bind and
activate MerTK in vivo to stimulate phagocytosis in the various
phagocytes and whether species differences exist [9]. A recent
report suggested that only protein-S but not Gas6 is relevant in
mice [36], but there is evidence for Gas6 expression in fresh
RPE of rats and humans [37,38]. Interestingly, it has been
shown that human protein-S neither binds nor activates human
MerTK [9]. Therefore, it is of great interest to study MerTK and
Gas6 in human tissue specifically.
Initially we examined the localization of MerTK receptor and
one of its ligands, Gas6, in the human adult retina. Immunostaining of retina sections with an antibody to MerTK demonstrated localization to the photoreceptor layer and apical portion
of the RPE monolayer (Fig. 1B and D) supporting previous
reports in rats and mice [36,37]. Interestingly, Gas6 was detected
at the photoreceptor and RPE layer (Fig. 1E). As an intrinsic
positive control, Gas6 expression was observed in thrombocytes
or other hematopoietic cells [39] in retinal blood vessels (not
shown). Negative controls gave only a background staining
(Fig. 1A and C).
Two different alternatively spliced isoforms of Gas6 have
been described [34,40]. Isoform one contains a 43 amino acid
Fig. 2. RT-PCR, immunocytochemical and immunoblot detection of Gas6 shows expression by RPE in vivo and in vitro. A) The expression of Gas6 in fresh human
RPE cells, retina and ARPE19 cells. PCR of human tissue cDNA using primers designed to span the region of the long form insert, produces one band of 439 bp
corresponding to expression of isoform two. B) Scheme of human Gas6 isoforms. The blackened box is the insert of 43 amino acids (129 bp) for isoform one. Isoform
two is without insert. C,D) Human SV40-RPE cultured 12 h serum-free and immunostained with respective antibody. C) Parallel control staining with non-immune
IgG confirms signals are specific (n N 3, bar = 20 μm). D) Gas6 was stained at 10 µg/ml antibody concentration. E) An immunoblot detected the C-terminal domain of
Gas6 in fresh human retina, fresh human RPE and in two RPE cell lines (SV40-RPE and ARPE-19) cultured 72 h under low serum (1%) containing conditions; and
F) induction of Gas6 expression upon serum reduction (24 h after seeding in 10% FCS) by culturing for 72 h in 1% FCS and with fresh culture media containing 10%
FCS as control.
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
inserted sequence compared to isoform two, which is expressed
in more different tissues and was reported first. Cultured human
ARPE19 cells, fresh human RPE cells and fresh human retina
were analyzed by RT-PCR in order to determine the Gas6
isoform expression (Fig. 2A). Using a primer pair which maps a
554 bp product including the 129 bp insertion (isoform one) or
425 bp product without the insertion (isoform two; Fig. 2B) we
detected the smaller mRNA product in all three tissue preparations showing that only Gas6 transcript for isoform two is
present. These results were confirmed using the exact primer
pairs originally published to discriminate the alternative splice
variants [34].
In the next step, we confirmed Gas6 expression in human RPE using immunocytochemistry and western blotting. In
confluent SV40-RPE cells cultured 12 h serum-free prior to
immunostaining Gas6 was well detected (Fig. 2C and D). In
agreement with these results, comparative immunoblots of RPE
lysates from retina, primary human RPE, ARPE19 and SV40RPE confirmed high expression of Gas6 protein (Fig. 2E). The
applied antibodies recognize an epitope on the receptor-domain
containing C-terminus. The antibodies detected specific bands
of immunoreactive protein migrating at the expected sizes previously reported for human Gas6 protein. The slight differences
between fresh tissue und the cell cultures might be due to posttranslational modification, an intracellular precursor or degradation product. We confirmed our results by application of two
different antibodies, polyclonal Gas6 mouse (Santa Cruz, SC1935) and goat-antibodies (R&D, DY885) recognizing recombinant human Gas6, which have been previously reported to
detect human Gas6 in immunoblots or ELISA assays. Moreover, the effect of reduced-serum culture on Gas6 expression by
RPE in vitro was examined. Gas6 was originally identified by
its increased expression by fibroblasts after serum starvation
[41] and this has been confirmed in various cell types. SV40RPE cultured 72 h in either 1% or 10% fetal calf serum (FCS)
exhibited high and low Gas6 expression, respectively (Fig. 2F).
These experiments suggest that human RPE cells express Gas6
message and protein in vivo and in vitro.
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in the absence of serum. Addition of antibodies recognizing an
epitope on the C-terminus (receptor binding-site) of Gas6
blocked uptake in a concentration-dependent manner. Application of 5 and 10 µg/ml Gas6 antibodies significantly reduced
uptake to 71 ± 4% and 65 ± 5% compared to control, respectively (Fig. 3B). Photoreceptor outer-segment fragment binding
was significantly decreased only at the highest antibody
concentration tested, to 78 ± 7% (Fig. 3A). MerTK-antibodies,
recognizing an extracellular N-terminal epitope, applied at 5
and 10 µg/ml reduced photoreceptor outer-segment fragment
uptake to 70 ± 5% and 66 ± 5%, respectively (Fig. 3D). Interestingly, outer-segment fragment binding was significantly
reduced at 2.5, 5 and 10 µg/ml MerTK antibody (Fig. 3C).
3.1. MerTK and Gas6 antibodies reduce phagocytosis
In the next step we tested the functional availability of Gas6
in human RPE in vitro, and its participation as a ligand for
MerTK in the photoreceptor outer-segment fragment phagocytosis by RPE. We employed the same antibody (SC-1935),
which was previously used in an antibody-blocking approach
for other phagocytes [42]. MerTK is localized to the apical
microvilli of human RPE cells (Fig. 1B and D) [36], which also
express the potential ligand Gas6 in vivo (Fig. 1) and in vitro
(Fig. 2) [38]. We earlier established the phagocytosis assay and
demonstrated that SV40-RPE exhibit phagocytic characteristics
of primary human RPE [21]. Since we found that Gas6
expression is increased by RPE cells under serum-reduced
conditions in vitro (Fig. 2), we wanted to test its functional
availability for phagocytosis. Confluent human SV40-RPE cells
cultured 12 h serum-free phagocytized SNAFL® -2 labeled
photoreceptor outer-segment fragments during a 4 h challenge
Fig. 3. Functional blockage of Gas6 and MerTK inhibits RPE phagocytosis.
Antibodies (Ab) recognizing ligand Gas6 (A, B, E) and related receptor MerTK
(C, D, E) inhibit OS phagocytosis by Gas6 expressing SV40-RPE (precultured
for 12 h serum-free). Blockage of MerTK and Gas6 were concentrationdependent and results were specific since non-immune IgG was without effect
(data not shown). A–D) SV40-RPE, fluorescent multi-well plate-reader assay
for OS binding and uptake, (n N 3; each n represents 8 independent wells;
⁎ p b 0.05, ⁎⁎ p b 0.01, ⁎⁎⁎ p b 0.001); E) FACS analysis of OS uptake (8 wells
of one experiment were pooled per variable).
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M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
The highest reduction in binding to 62 ± 10% was observed at
5 µg/ml MerTK antibody. Control goat-IgG incubation compared to conditions without antibodies added (data not shown)
had no effect on phagocytosis as previously reported [4]. Strikingly, inhibitory effects of antibodies against Gas6 and MerTK
on photoreceptor outer-segment fragment uptake by primary
human RPE were confirmed by means of FACS measurements
(Fig. 3E).
3.2. L-type calcium channel inhibition modulates phagocytosis
We tested the L-type Ca2+-channel blocker, nifedipine, for its
effect on Gas6-dependent photoreceptor outer-segment fragment
phagocytosis by human RPE in vitro. Human RPE expresses
functional L-type Ca2+-channels, which are nifedipine inhibitable
(Fig. 5A–C). Strikingly, nifedipine inhibited phagocytosis both
dose and time dependently. Phagocytosis by endogenous Gas6expressing SV40-RPE (cultured serum-free for 12 h) was
inhibited by nifedipine significantly during a 4 h photoreceptor
outer-segment fragment challenge: In the presence of 1 µM
nifedipine, binding was reduced to 70 ± 14% and uptake to 84 ±
7%, 5 µM nifedipine reduced binding to 43 ± 18% and uptake to
54 ± 14%, and 10 µM nifedipine reduced binding to 49 ± 18% and
uptake to 44 ± 14% (Fig. 4A and B). Adding serum, a rich exogenous source of MerTK-ligands including Gas6 and Protein S
[35], stimulates photoreceptor outer-segment fragment phagocytosis by SV40-RPE cells in vitro in a concentration-dependent
manner as previously demonstrated [21,43]. Nifedipine (5 µM)
reduced-serum stimulated phagocytosis (5% FCS) in photoreceptor outer-segment fragment binding from 356 ± 2% to 267 ±
2% and uptake from 293 ± 6% to 184 ± 6% (Fig. 4C and D). FACS
measurements allowed observation of photoreceptor outersegment fragment uptake by SV40-RPE as early as 1 h after
photoreceptor outer-segment fragment challenge, and — even
more interestingly — allowed studies on primary human RPE
after a brief culture period. Nifedipine applied at 2 µM drastically
reduced SV40-RPE phagocytosis at 1 h to 20% (Fig. 4E).
Furthermore, the inhibitory effect of nifedipine on phagocytosis
(4 h) was confirmed by primary human RPE (cultured 12 h serumfree), which reduced uptake to 82 ± 4% and 77 ± 3% by 0.5 and
1 µM nifedipine, respectively (Fig. 4F).
3.3. Effects of αV-integrin ligands on L-type Ca2+ channel
activity
Fig. 4. L-type Ca2+-channel blockage inhibits phagocytosis. L-type Ca2+-channel
blocker nifedipine (NIF) led dose dependently A) to a decrease in binding and
B) in uptake of OS by Gas6 expressing SV40-RPE cells (precultured for 12 h
serum-free) in vitro during 4 h OS challenge (n N 3). C,D) Furthermore,
phagocytosis of SV40-RPE cells precultured in the presence of phagocytosis
stimulating serum concentrations (= MerTK-ligand source) was reduced
similarly (n N 3; compared to control in Fig. 4A). E) In FACS measurements
the NIF effect on phagocytosis could already be observed after 1 h by serum-free
cultured (12 h) Gas6 expressing SV40-RPE (n = 1, 8 wells were pooled for one
experiment) as well as F) by serum-free cultured (12 h) primary human RPE cells
after 4 h OS challenge (n N 3).
The influence of L-type Ca2+-channel blocker on phagocytic
activity implies a regulatory role for these channels in this
process. Because nifedipine inhibited phagocytic binding and
uptake, we wanted to test whether L-type Ca2+-channel might
be modulated by integrin receptors. Since it has been shown
that αV-integrins play a role in the initiation of phagocytosis, we investigated the effects of αV-integrin ligands on L-type
Ca2+-channel activity. In perforated patch-clamp recordings and
in the presence of Ba2+, ARPE-19 cells respond to depolarization from a holding potential of − 70 mV with fast activating and
inactivating inward-currents (Fig. 5A). These currents activated
at potentials more positive than − 30 mV and were reduced to
56.6 ± 7.4% by 10 µM nifedipine (n = 5) in the bath (Fig. 5B and
C). At voltage-step from − 70 to + 10 mV, the currents reached
the peak after 8.7 ± 2.5 ms (n = 6), were inactivated with a time
constant of 20.45 ± 3.4 ms (n = 6; fitted by single exponential fit)
and showed a maximal current density at + 10 mV of 0.89 ±
0.1 pApF− 1 (n = 9).
These currents were changed by application of αV-integrin
ligands (Fig. 5D–H). Application of soluble vitronectin
(0.03 µM) led to a significant reduction (n = 6; p b 0.01) of the
maximal current amplitude after 3 min, as measured by a
voltage-step from − 70 mV to +10 mV (Fig. 5D–F). The kinetic
of the currents were not changed in the presence of soluble
vitronectin. To verify the effects of soluble vitronectin we
performed a series of experiments using insoluble ligand. For
this purpose, cells were cultured on vitronectin-coated glass
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
coverslips. Compared to cells grown on uncoated coverslips,
cells grown on vitronectin-coated coverslips showed an increased Ba2+-current density at + 10 mV of 1.64 ± 0.35 pApF− 1
(n = 9; p b 0.05; Fig. 5H). Again, the kinetic parameters time-topeak and inactivation time constant were unchanged in the
presence of insoluble vitronectin. To provide further evidence
for participation of αV-integrins in the vitronectin-dependent
modulation of Ca2+-channels in RPE, we utilized RGD-peptides
(Fig. 5G). Application of the GRADSP-peptide (0.03 µM),
which is an inactive-control, led after 5 min to a slight reduction
of the maximal current amplitude at + 10 mV to 82 ± 5.4% of
control (n = 3). The GRGDNP-peptide, which is able to bind
with higher specificity to α5- than to αV-integrin, led to no sig-
1165
nificant reduction in the maximal current amplitude at + 10 mV
compared to the inactive-control (79 ± 5%; n = 4). In contrast
the GRGDSP-peptide, which mainly binds to αV-integrin, led to
a reduction in the maximal currents amplitude to 53.3 ± 8.2%
(n = 4; p b 0.05 compared to GRADSP-peptide). This effect is in
the same range as the effect of soluble vitronectin application.
Again, in all experiments using RGD-peptides, no changes in the
kinetic behavior of the currents were observed.
In a last series of experiments we investigated the involvement of intracellular signaling molecules to provide further
evidence for a receptor-coupled effect on Ca2+-channel activity
(Fig. 5I). Since αV-integrin stimulation [11,44] and L-type Ca2+channel activation [45] are known to involve Src-kinase activity we studied the effect of soluble vitronectin in cells incubated
with the cytoplasmic Src-tyrosine kinase inhibitor herbimycin A
(HA, 1 µM). Interestingly, in the presence of HA, application of
soluble vitronectin (0.3 µM) did not change the L-type Ca2+channel activity. After 5 min, the maximal current amplitude in
HA-treated cells was 110 ± 7.7% (n = 4) of the value that was
observed after establishing the perforated-patch configuration. In
the presence of vitronectin, the maximal current amplitude in
HA-treated cells was 102 ± 4.4% of the control value observed
prior to application.
Fig. 5. αV-integrin ligands modulate L-type channel Ba2+-currents in ARPE-19
cells. A) Cells were depolarized from a holding potential of −70 mV by
9 voltage-steps of 50 ms duration and of +10 mV increment. Ba2+-currents:
When 10 mM Ba2+ was added to the bath solution the ARPE-19 cells respond to
electrical stimulation with fast activating and inactivating inward-currents. B) In
the presence of 10 µM NIF the current activated by a voltage-step from − 70 mV
to +10 mV was reduced. C) Summary of the effect of NIF as current voltageplot: The maximal current-amplitudes were plotted against the step-potentials of
the electrical stimulation. The currents activate at potentials more positive than
−30 mV. In the presence of NIF the current-amplitudes were decreased. D–I)
Effects of αV-integrin ligation on L-type Ca2+-channel currents: D) Plot of the
maximal current amplitude (at +10 mV) plotted over the experiment time. The
current amplitude which has been measured directly after establishing the
perforated-patch configuration was set as 100%. The plot represents the mean
values of experiments without (n = 6) or with 0.03 µM soluble vitronectin (n = 6)
in the bath solution (arrow indicates the moment of application). E) Ba2+-current
measured after a voltage-step from −70 mV to +10 mV without and in the presence of soluble vitronectin (0.03 µM). F) Summary of the vitronectin effect on
L-type Ca2+-channel activity: The maximal current amplitude at +10 mV was
measured before (n = 6) and in the 5 min presence of the integrin ligand
vitronectin (0.03 µM). The current amplitude was normalized to the amplitude
that had been measured directly after establishing the perforated-patch configuration. G) Effects of different RGD integrin ligands on L-type Ca2+-channel
activity: The maximal current amplitude at +10 mV was measured in the 5 min
presence of the integrin ligand GRGDSP (0.03 µM), GRGDNP (0.03 µM) or
GRADSP (0.03 µM). The current amplitude was normalized to the amplitude which has been measured directly after establishing the perforated-patch
configuration. H) Comparison of Ba2+-current density in cells cultured on
vitronectin-coated coverslips and uncoated coverslips: Maximal current density
was measured at +10 mV and normalized to the cell size using the membrane
capacitance of each individual cell. I) Effect of Src-kinase inhibition: Plot of the
maximal current amplitude over experiment time, which was measured at
+10 mV and normalized in percent to the current amplitude measured just after
establishing the perforated-patch configuration in each experiment. In control
conditions the currents were measured in herbimycin A (HA, 1 µM) treated
cells. In the experimental conditions soluble vitronectin (0.03 µM) was applied
to HA-treated cells (n = 4 in both series).
1166
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
uptake was already reduced after 1 h photoreceptor outersegment fragment challenge (Fig. 6D).
4. Discussion
The major findings of the present work were the following:
I) human RPE cells are a potential endogenous source for
MerTK-ligand Gas6, II) blocking L-type Ca2+-channels reduces
MerTK-regulated phagocytosis and III) ligation of photoreceptor outer-segment binding integrin receptors modulates L-type
Ca2+-channel currents.
4.1. Signaling of MerTK dependent RPE phagocytosis
Fig. 6. Herbimycin A (HA) treatment reduces phagocytosis. SV40-RPE cultured
12 h serum-free and pretreated with HA showed reduced A) binding and
B) uptake of OS assayed after 4 h in vitro. The dose dependent stimulatory effect
of serum (=MerTK-ligand source) on phagocytosis was thereby abolished
(n = 3). C-D) FACS measurements of C) primary human RPE (n N 3) after 4 h OS
challenge and D) SV40-RPE phagocytosis (n = 1) after 1 h OS challenge confirmed the results presented in B (cells were cultured serum-free for 12 h before
experiments to enhance endogenous Gas6 expression; 8 independent wells were
pooled for one FACS experiment).
3.4. Potential modulators of L-type Ca2+ channels alter RPE
phagocytosis
To test whether the potential Src-kinase inhibitor herbimycin
A (1 µM, HA) also affects phagocytosis, RPE cells were preincubated with HA prior to OS challenge. Control phagocytic
challenge with photoreceptor outer-segment fragments and
serum (source of MerTK-ligands) for 4 h caused a strong
concentration-dependent increase in phagocytosis as previously
shown. Strikingly, this stimulation was not observed after HA
pretreatment (Fig. 6A and B). Serum stimulation (10% FCS)
of phagocytic binding (295 ± 6%) and uptake (242 ± 5%) by
SV40-RPE cells were almost completely abolished to 137 ± 6%
and 93 ± 10%, respectively. Importantly, likewise phagocytosis
of Gas6-expressing SV40-RPE (cultured 12 h serum-free) was
decreased (uptake 73 ± 7% compared to control, while binding
was unaltered; Fig. 6A and B). This effect was observed up to
12 h after photoreceptor outer-segment fragment challenge
(uptake 67 ± 4% and binding 80 ± 27% in one preliminary
experiment of 8 independent wells; data not shown). FACS
analysis of primary human RPE cells after a 4 h photoreceptor
outer-segment fragment challenge showed comparable results
under serum-free condition (69 ± 9% uptake) and similar results
under serum stimulation (Fig. 6C). 5% FCS stimulated uptake
significantly (160 ± 8%), but HA pretreatment prevented this
stimulatory action (114 ± 14%). In HA-treated SV40-RPE cells,
The physiological sources and roles of Gas6 for phagocytosis have been controversial [9,10,35–38]. Our results provide
evidence for endogenous Gas6 expression by human RPE in
vivo and in vitro extending previous results (Figs. 1 and 2). We
detected Gas6 transcript and protein in fresh human RPE cells
and human RPE cell lines. Growth-arrest-specific (GAS) genes
were originally identified by mRNA downregulation upon
induction of growth by serum or vice versa [33,41,46].
Therefore, reducing or withdrawing serum from RPE in vitro
led to increased Gas6 protein (Fig. 2), suggesting autocrine and
negative-feedback control. Using a direct antibody-blocking
approach previously described for other phagocytes [39,42] we
investigated the roles of Gas6 and MerTK. Our study shows that
photoreceptor outer-segment fragment phagocytosis by Gas6
expressing RPE (12 h serum-free precultured) is reduced in the
presence of Gas6-antibodies (Fig. 3). Thus, Gas6, which promotes phagocytosis via MerTK-activation, may stem from RPE.
Our experiments extend previous results obtained by genedeletion of MerTK and support previous reports that receptors
other than MerTK primarily mediate the initiating binding process [11,14]. For this process, Finnemann's group suggested a
pathway in which photoreceptor outer-segments bind via RGDmotif containing ligands to integrin receptors, which in turn
leads to phosphorylation of MerTK via a focal adhesion kinasedependent pathway [11,14,15]. This is in agreement with the
idea that MerTK-deficient RPE is still capable of photoreceptor
outer-segment binding [14,17,19]. Moreover, that integrin
receptors have been related to the initiating binding process is
supported by the observation that integrin ligand MFG-E8-null
retina in vivo failed to stimulate peak MerTK-phosphorylation
that follows light onset in wild-type retina [12,14]. Due to our
results with the MerTK antibody, it is still controversial whether
MerTK possesses a potential Gas6-independent adhesive function itself, or whether MerTK-activation is interacting with
photoreceptor outer-segment adhesion. The first case is supported by the observations that MerTK-antibodies significantly
inhibited phagocytosis at low concentrations, but Gas6 antibodies did not. These observations could not have been made in
cells lacking functional MerTK, which has dual extracellular
immunoglobin and fibronectin-type II motifs that suggest
potential adhesive capacities [47]. In other phagocytes it has
been suggested that MerTK might sustain active integrin
receptors, and both share common signals that allow them to
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
cooperate in a bi-directional way by amplifying internalization
signals [10].
4.2. Role of Ca2+-channels in RPE phagocytosis
This study provides evidence that MerTK-stimulated phagocytosis was inhibited by blockage of L-type Ca2+-channels
using nifedipine. Nifedipine reduces phagocytosis by various
other MerTK utilizing phagocytes [48,49]. In general, receptor
protein-tyrosine kinase stimulation may activate L-type Ca2+channels or vice versa. MerTK-deficient RPE cells have
increased L-type Ca2+-channel currents, and fail to ingest
proper amounts photoreceptor outer-segment fragments, but
still bind them. Therefore, L-type Ca2+-channel activity might
be modulated by an inhibitory function of MerTK on L-type
Ca2+-channel, which is lost in MerTK-deficient RPE and would
therefore lead to increased channel-activity [24]. Interestingly,
Gas6 application reduced secreted phospholipase-A2 type IIA(sPLA2-IIA)-induced Ca2+-influx through L-type Ca2+-channels in neurons [50] and RPE contains sPLA2-IIA, which
has been found to enhance phagocytosis [51]. But, given that
photoreceptor outer-segment fragment binding and uptake
were both significantly reduced, L-type Ca2+-channel inhibition
might also decrease binding and thereby reduce phagocytosis.
Comparably, in consequence of β5-integrin loss or overexpression burst of photoreceptor outer-segment fragment internalization is abolished or binding and uptake increased, respectively
[15,52].
Therefore, we explored the potential role of integrin receptors
in L-type Ca2+-channel regulation. The effects of αV-integrin
stimulation on L-type Ca2+-channel activity were studied by
application of RGD-motif containing soluble vitronectin, which
should displace insoluble vitronectin from its receptor, and
thus lead to de-activation of the αV-integrin [53]. Under these
conditions, L-type Ca2+-channel activity decreases. On the
other hand, cells cultured on insoluble vitronectin showed larger
L-type Ca2+-channel activity. Further verification was provided
by investigating RGD-peptides, known to interact specifically
with αV-integrin and to reduce phagocytosis by RPE in vitro
[4], which showed a significant effect on L-type Ca2+-channel
activity of RPE cells. Thus, αV-integrin stimulation activates
L-type Ca2+-channels in RPE.
L-type Ca2+-channels had previously been shown to directly
interact with Src-kinase in RPE and other cells [45,52,54], and
stimulation of αV-integrin in the initiation process of phagocytosis involves Src-kinase activity in the RPE. Herbimycin A,
a potential Src-kinase blocker, reduces phosphorylation of
integrin receptor-associated proteins [4]. L-type Ca2+-channel
activity of RPE cells treated with herbimycin A did not respond
to application of soluble vitronectin. Taken together, stimulation of αV-integrin is a potential way to activate L-type Ca2+channels in the signaling pathway of RPE phagocytosis. Integrin
receptor ligation may activate these channels in other cell types
[53], but the precise role in phagocytosis still needs to be
discovered. RPE cells display a large range of intracellular
calcium kinetics [28] and L-type Ca2+-channels may regulate
steady-state local and global Ca2+-signals [55] and participate in
1167
the regulation of [Ca2+]i oscillations [56]. In functional studies,
local changes of [Ca2+]i have been associated with photoreceptor outer-segment binding by RPE [7], comparable with
observations in other phagocytes [20,57]. Moreover, reducing
extracellular calcium or increasing [Ca2+]i may terminate RPE
phagocytosis [26,58]. Because calcium homeostasis seems to be
relevant for phagocytic activity, and since phagocytic deficiency
has been related to inherited and age-related retinal-degeneration
[6,8,15], the diversity of calcium signaling involved in phagocytosis needs to be identified in future studies.
4.3. L-type Ca2+ -channel, αv-integrin and MerTK signaling
pathways
L-type Ca2+-channels are activated by Src-kinase and inhibition of either one reduces binding and uptake of RPE phagocytosis. Previous reports localized Src-kinase actions downstream
of integrin receptors in RPE, which promote phagocytic signaling
to MerTK [11,14,15]. In macrophages, it has been shown that
MerTK is directionally and functionally linked to integrinsignaling by Src-kinase, which was found to be under MerTKand integrin-regulation [9,10]. The current work shows that Gas6
expressed endogenously by human RPE promotes phagocytosis,
which was reduced by blocking L-type Ca2+-channels. Our data
suggests that L-type Ca2+-channel regulation might be coupled to
photoreceptor outer-segment binding via integrin receptors
[53,54] as well as to uptake via MerTK [10,33,44], with Srckinase being a common denominator.
Acknowledgements
We are grateful to Dr.rer.nat. Boris Fehse (University Hamburg,
Germany) for access to and help on FACS experiments. We like
to thank Stefanie Ehmer for expert technical assistance and Paige
Etter (University of Washington, USA) for proofreading. Mike O.
Karl was supported by a grant from the Werner Otto-Foundation.
Funding: DFG STR480/8-1; STR480/8-2. The abbreviations used
are: MerTK, Mer tyrosine kinase; Gas6, Growth-arrest-specific 6;
RPE, Retinal Pigment Epithelium.
References
[1] A. Gal, Y. Li, D.A. Thompson, J. Weir, U. Orth, S.G. Jacobson, E.
Apfelstedt-Sylla, D. Vollrath, Nat. Genet. 26 (3) (2000) 270.
[2] P.M. D'Cruz, D. Yasumura, J. Weir, M.T. Matthes, H. Abderrahim, M.M.
LaVail, D. Vollrath, Hum. Mol. Genet. 9 (4) (2000) 645.
[3] S.C. Finnemann, E. Rodriguez-Boulan, J. Exp. Med. 190 (6) (1999) 861.
[4] S.C. Finnemann, V.L. Bonilha, A.D. Marmorstein, E. Rodriguez-Boulan,
Proc. Natl. Acad. Sci. U. S. A. 94 (24) (1997) 12932.
[5] E.F. Nandrot, M. Anand, M. Sircar, S.C. Finnemann, Am. J. Physiol., Cell.
Physiol. 290 (4) (2006) C1256.
[6] S. Zareparsi, M. Buraczynska, K.E. Branham, S. Shah, D. Eng, M. Li, H.
Pawar, B.M. Yashar, S.E. Moroi, P.R. Lichter, H.R. Petty, J.E. Richards, G.R.
Abecasis, V.M. Elner, A. Swaroop, Hum. Mol. Genet. 14 (11) (2005) 1449.
[7] A.L. Kindzelskii, V.M. Elner, S.G. Elner, D. Yang, B.A. Hughes, H.R.
Petty, J. Gen. Physiol. 124 (2) (2004) 139.
[8] R. Sarangarajan, S.P. Apte, Mol. Vis. 11 (2005) 482.
[9] S. Hafizi, B. Dahlback, Febs J. 273 (23) (2006) 5231.
[10] Y. Wu, N. Tibrewal, R.B. Birge, Trends Cell Biol. 16 (4) (2006) 189.
[11] S.C. Finnemann, Embo J. 22 (16) (2003) 4143.
1168
M.O. Karl et al. / Cellular Signalling 20 (2008) 1159–1168
[12] E.F. Nandrot, M. Anand, K. Atabai, S.C. Finnemann, Invest. Ophthalmol.
Vis. Sci. 47 (5) (2006) 5880.
[13] E.F. Nandrot, E.M. Dufour, A.C. Provost, M.O. Pequignot, S. Bonnel, K.
Gogat, D. Marchant, C. Rouillac, B. Sepulchre de Conde, M.T. Bihoreau,
C. Shaver, J.L. Dufier, C. Marsac, M. Lathrop, M. Menasche, M.M.
Abitbol, Neurobiol. Dis. 7 (6 Pt B) (2000) 586.
[14] E.F. Nandrot, M. Anand, D. Almeida, K. Atabai, D. Sheppard, S.C.
Finnemann, Proc. Natl. Acad. Sci. U. S. A. 104 (29) (2007) 12005.
[15] E.F. Nandrot, Y. Kim, S.E. Brodie, X. Huang, D. Sheppard, S.C.
Finnemann, J. Exp. Med. 200 (12) (2004) 1539.
[16] A.I. Goldman, P.J. O'Brien, Science 201 (4360) (1978) 1023.
[17] M.H. Chaitin, M.O. Hall, Invest. Ophthalmol. Vis. Sci. 24 (7) (1983) 812.
[18] R.B. Edwards, P.M. Flaherty, J. Cell. Physiol. 127 (2) (1986) 293.
[19] W. Feng, D. Yasumura, M.T. Matthes, M.M. LaVail, D. Vollrath, J. Biol.
Chem. 277 (19) (2002) 17016.
[20] G. Berton, C.A. Lowell, Cell. Signal. 11 (9) (1999) 621.
[21] M.O. Karl, M. Valtink, J. Bednarz, K. Engelmann, Graefe Arch. Clin. Exp.
Ophthalmol. 245 (7) (2007) 981.
[22] J. Nguyen-Legros, D. Hicks, Int. Rev. Cytol. 196 (2000) 245.
[23] O. Strauss, Physiol. Rev. 85 (3) (2005) 845.
[24] O. Strauss, F. Stumpff, S. Mergler, M. Wienrich, M. Wiederholt, Acta.
Anat. (Basel) 162 (2–3) (1998) 101.
[25] E.M. Dufour, E. Nandrot, D. Marchant, L. Van Den Berghe, S. Gadin, M.
Issilame, J.L. Dufier, C. Marsac, D. Carper, M. Menasche, M. Abitbol,
Neurobiol. Dis. 14 (2) (2003) 166.
[26] M.O. Hall, T.A. Abrams, T.W. Mittag, Exp. Eye Res. 52 (5) (1991) 591.
[27] S. Nakashima, Y. Tsunematsu, Y. Nozawa, Nippon Ganka Gakkai Zasshi
93 (1) (1989) 149.
[28] S. Wimmers, M.O. Karl, O. Strauss, Prog. Retin. Eye Res. 26 (3) (2007)
263.
[29] K. Engelmann, M. Valtink, Graefe Arch. Clin. Exp. Ophthalmol. 242 (1)
(2004) 65.
[30] M.V. Miceli, D.A. Newsome, Exp. Eye Res. 59 (3) (1994) 271.
[31] C.J. Kennedy, P.E. Rakoczy, I.J. Constable, Curr. Eye Res. 15 (9) (1996)
998.
[32] H.M. Seitz, T.D. Camenisch, G. Lemke, H.S. Earp, G.K. Matsushima,
J. Immunol. 178 (9) (2007) 5635.
[33] S. Goruppi, E. Ruaro, B. Varnum, C. Schneider, Mol. Cell. Biol. 17 (8)
(1997) 4442.
[34] P. Marcandalli, M. Gostissa, B. Varnum, S. Goruppi, C. Schneider, FEBS
Lett. 415 (1) (1997) 56.
[35] M.O. Hall, M.S. Obin, M.J. Heeb, B.L. Burgess, T.A. Abrams, Exp. Eye
Res. 81 (5) (2005) 581.
[36] D. Prasad, C.V. Rothlin, P. Burrola, T. Burstyn-Cohen, Q. Lu, P. Garcia de
Frutos, G. Lemke, Mol. Cell. Neurosci. 33 (1) (2006) 96.
[37] M.O. Hall, A.L. Prieto, M.S. Obin, T.A. Abrams, B.L. Burgess, M.J. Heeb,
B.J. Agnew, Exp. Eye Res. 73 (4) (2001) 509.
[38] N. Kociok, A.M. Joussen, Graefe Arch. Clin. Exp. Ophthalmol. 245 (1)
(2007) 101.
[39] A. Angelillo-Scherrer, P. de Frutos, C. Aparicio, E. Melis, P. Savi, F. Lupu,
J. Arnout, M. Dewerchin, M. Hoylaerts, J. Herbert, D. Collen, B. Dahlback,
P. Carmeliet, Nat. Med. 7 (2) (2001) 215.
[40] S. Goruppi, H. Yamane, P. Marcandalli, A. Garcia, C. Clogston, M.
Gostissa, B. Varnum, C. Schneider, FEBS Lett. 415 (1) (1997) 59.
[41] C. Schneider, R.M. King, L. Philipson, Cell 54 (6) (1988) 787.
[42] J.C. Todt, B. Hu, J.L. Curtis, J. Leukoc. Biol. 75 (4) (2004) 705.
[43] P.L. Mayerson, M.O. Hall, J. Cell Biol. 103 (1) (1986) 299.
[44] Y. Wu, S. Singh, M.M. Georgescu, R.B. Birge, J. Cell Sci. 118 (Pt 3)
(2005) 539.
[45] O. Strauss, F. Buss, R. Rosenthal, D. Fischer, S. Mergler, F. Stumpff, H.
Thieme, Biochem. Biophys. Res. Commun. 270 (3) (2000) 806.
[46] G. Manfioletti, C. Brancolini, G. Avanzi, C. Schneider, Mol. Cell. Biol. 13 (8)
(1993) 4976.
[47] D.K. Graham, G.W. Bowman, T.L. Dawson, W.L. Stanford, H.S. Earp, H.R.
Snodgrass, Oncogene 10 (12) (1995) 2349.
[48] N. Tokuda, T. Mano, R.B. Levy, J. Urol. 147 (1) (1992) 278.
[49] T. Mano, D.G. Puro, Invest. Ophthalmol. Vis. Sci. 31 (6) (1990) 1047.
[50] T. Yagami, K. Ueda, K. Asakura, N. Okamura, T. Sakaeda, G. Sakaguchi,
N. Itoh, Y. Hashimoto, T. Nakano, M. Fujimoto, Brain Res. 985 (2) (2003)
142.
[51] M. Kolko, J. Wang, C. Zhan, K.A. Poulsen, J.U. Prause, M.H. Nissen, S.
Heegaard, N.G. Bazan, Invest. Ophthalmol. Vis. Sci. 48 (3) (2007) 1401.
[52] E.F. Nandrot, S.C. Finnemann, Invest. Ophthalmol. Vis. Sci. 45 (5) (2004)
3675.
[53] X. Wu, J.E. Mogford, S.H. Platts, G.E. Davis, G.A. Meininger, M.J. Davis,
J. Cell. Biol. 143 (1) (1998) 241.
[54] P. Gui, X. Wu, S. Ling, S.C. Stotz, R.J. Winkfein, E. Wilson, G.E. Davis,
A.P. Braun, G.W. Zamponi, M.J. Davis, J. Biol. Chem. 281 (20) (2006)
14015.
[55] M.F. Navedo, G.C. Amberg, V.S. Votaw, L.F. Santana, Proc. Natl. Acad.
Sci. U. S. A. 102 (31) (2005) 11112.
[56] G. Liu, N. Hilliard, G.H. Hockerman, Mol. Pharmacol. 65 (5) (2004) 1269.
[57] S. Dewitt, M.B. Hallett, J. Cell Biol. 159 (1) (2002) 181.
[58] L.M. Greenberger, J.C. Besharse, Invest. Ophthalmol. Vis. Sci. 24 (11)
(1983) 1456.