1. Introduction
Intraocular fibrotic diseases, such as proliferative vitreoretinopathy (PVR),
late-stage neovascular age-related macular degeneration (nAMD), and proliferative
diabetic retinopathy (PDR), can occur in response to ocular trauma, ischemia,
inflammation, and degenerative disease. These diseases usually lead to severe
visual impairment or irreversible vision loss in the majority of patients. The
formation of subretinal, epiretinal, or intravitreal fibrotic membranes due to
extensive wound healing responses has been identified as an important step in the
pathogenesis of these fibrotic diseases. This process orchestrates the
interaction of several cellular components, including RPE cells, glial cells,
fibroblasts, and inflammatory cells [1], with many inflammatory cytokines and
growth factors, resulting in overproduction or remodeling of the extracellular
matrix (ECM). Among all these involved cell types, RPE cells have been recognized
to play a critical role in the development of retinal fibrosis [2, 3].
Epithelial to mesenchymal transition (EMT) is a biological process that refers
to the phenomenon in which epithelial cells are converted into mesenchymal cells
under specific physiological and pathological conditions. In this process,
epithelial cells lose their cell-cell junctions and apical-basal polarity,
reorganize their actin cytoskeleton, and gain cell motility and invasion
properties. This is characterized by the downregulation of epithelial molecular
markers, such as occludin, zonula occludens-1 (ZO-1), and E-cadherin, and
upregulation of mesenchymal molecular markers, such as -SMA and
fibronectin (FN). Accumulating evidence has revealed the pivotal role of EMT in
wound healing, tissue reconstruction, and fibrosis, including the formation of
fibrotic lesions on the retina [4, 5]. In the development of PVR and at the
end-stage of nAMD, with the stimulation of various growth factors and cytokines,
the RPE cells, which are a monolayer of highly polarized hexagonal cells under
physiological conditions, undergo EMT. This enables them to lose their epithelial
characteristics, transdifferentiate into fibroblast-like cells, and migrate to
the subretinal space, resulting in the formation of a fibrotic membrane after
rhegmatogenous retinal detachment in PVR or choroidal neovascularization in AMD
[6, 7].
ER stress has long been studied for its role in neurodegenerative diseases and
diabetes, including AMD and PDR. During the past 10 years, accumulating evidence
has indicated the potential role of ER stress in a variety of fibrotic diseases,
including fibrosis of the liver, heart, kidney, and lung [8, 9]. The ER is a
multifunctional cellular organelle that is essential for maintaining protein
homeostasis. Accumulation of misfolded/unfolded proteins in the ER lumen leads to
activation of the unfolded protein response (UPR), a process known as ER stress,
with the release of a critical ER stress regulator, glucose-regulated protein
78/binding immunoglobulin protein (GRP78/BiP). Recently, several studies reported
the association between ER stress and EMT in regulating fibrosis development in
various cell types, such as pulmonary epithelial cells, peritoneal mesothelial
cells, and human lens epithelial cells [10, 11, 12]. However, the effects of ER stress
on EMT in RPE cells have not been studied.
In this study, we conducted a series of experiments to investigate the impact of
ER stress on the process of EMT in the RPE cell.
2. Materials and methods
2.1 Retinal pigment epithelial cell culture
Human RPE cells were isolated from postmortem human fetal eyes (gestational age,
16‒18 weeks) obtained from Advanced Bioscience Resources Inc. (Alameda, CA, USA)
and cultured as previously described [13]. Briefly, cells were cultured in DMEM
containing 2 mM L-glutamine, 100 U/mL penicillin,
100 g/mL streptomycin, and 10% heat-inactivated fetal
bovine serum (FBS). Cultured human RPE cells were used from passage 2 to 4. Cells
were confirmed to be RPE by their typical morphology, their immunoreactivity for
cytokeratin (95%), and the lack of immunoreactivity for macrophage or
endothelial cell markers.
2.2 Western blotting
RPE cells were lysed in lysis buffer (RIPA buffer, Thermo Fisher, USA), and the
extracted cell lysates were separated using Tris-HCl/Tris-Glycine gels (Ready
Gel; Bio-Rad, Hercules, CA, USA). The proteins were then transferred to
polyvinylidene fluoride microporous membranes (Millipore, Billerica, MA, USA).
The blots were blocked with 5% milk for 1 h at room temperature and incubated
with the corresponding primary antibodies (Supplementary Table 1)
overnight at 4 C. Blots were washed and incubated with horseradish
peroxidase-labeled secondary antibodies (Vector Laboratories, Inc. Burlingame,
CA, USA) for 1 h at room temperature. After addition of chemiluminescent
detection solution (ECL Western Blotting Substrate; Thermo Fisher, Waltham, MA,
USA), images were developed using an enhanced chemiluminescence detection system
(ChemiDoc™ XRS+ System, Bio-Rad). Glyceraldehyde-3-phosphate
dehydrogenase (GAPDH) served as the protein-loading control. Densitometry was
performed based on three independent experiments using Image J software (Image J
1.50i bundled with 64-bit java 1.8.0)
(http://imagej.nih.gov.libproxy1.usc.edu/ij/; National Institutes of Health,
Bethesda, MD, USA).
2.3 RNA isolation and real-time RT-qPCR
Total RNA was isolated using a RNeasy mini kit (QIAGEN, Valencia, CA, USA), and
cDNA was synthesized from total RNA using the ImProm-IITM Reverse Transcription
System (Promega, Madison, WI, USA). Real-time PCR was performed in duplicate
using a commercial SYBR Green master mix (Roche, Basel, Switzerland) with a
LightCycler 480 (Roche) according to the manufacturer’s protocols. The cycling
conditions for the RT-qPCR were as follows: pre-incubation at 95 C for
5 min, followed by 45 cycles of denaturing at 95 C for 10 s, annealing
at 55 C for 10 s, and extension at 72 C for 10 s. The primer
sequences are shown in Supplementary Table 2. Relative changes in mRNA
expression were determined by calculating the CT (cycle
threshold) values. mRNA levels were normalized relative to GAPDH mRNA and
reported as fold-change over controls.
2.4 Immunofluorescence staining and immunohistochemistry
RPE cells were cultured in 4-chambered culture slides (Falcon, Thermo Fisher).
The cells were rinsed in phosphate-buffered saline (PBS) for
3 min, fixed with 4% paraformaldehyde for 10 min,
and permeabilized with 0.1% Tween 20 (Bio-Rad) in PBS. After 30-min blocking
with 5% normal goat serum, the cells were incubated with the primary antibodies
(Supplementary Table 1) overnight at 4 C and then incubated
with the corresponding secondary antibodies for 30 min at room temperature. All
antibodies were diluted in PBS. The slides were mounted with mounting medium
containing DAPI (Vector Laboratories) and examined with a KEYENCE fluorescence
microscope (BZ-X700, San Diego, CA, USA).
2.5 MTT assay
Human RPE cells were cultured in DMEM containing 10% FBS in a 96-well plate and
then treated with different concentrations of TM (10‒200 ng/mL) for 24 h. Cell
viability was then assessed using a
3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) assay
according to the manufacturer’s instructions (Sigma, St Louis, MO, USA). The
absorbance of each well was read by a microplate manager (Bio-Rad Laboratories,
Inc. USA) at 550 nm.
2.6 Transwell cell migration assay
A transwell cell migration assay with human RPE cells was performed using a
modified Boyden chamber model (Transwell apparatus, 8.0-m pore size,
Falcon). For detecting RPE cell migration in the Transwell, the polyethylene
terephthalate membrane insert was coated with fibronectin (25 g/mL)
for 1 h. RPE cells were pre-treated with TM (100 ng/mL) or PBS (Ctrl) for 24 h
and were then trypsinized and resuspended in 0.5% FBS DMEM to a concentration of
5.0 10 cells/mL. A 100-L RPE cell suspension (final
concentration: 5 10 cells/well) was plated into the upper
chambers, and the lower chambers were filled with 0.6 mL of 0.5% FBS DMEM. After
a 1-h incubation period (37 C, 95% air/5% CO), PDGF-BB (25
ng/mL) or PBS was added to the lower chambers to induce chemotaxis. After 5 h of
incubation, Transwell inserts were washed with PBS and fixed with pre-chilled
methanol (10 min at 4 C). Cells were then stained with hematoxylin, and
all non-migrant cells were removed from the upper faces of the Transwell
membranes with a cotton swab. Migration was quantified by counting the number of
stained cells/10 in images taken with a scanning microscope (Aperio
ScanScope, Leica, Wetzlar, Germany). Four randomly chosen fields were counted per
membrane. Each experiment was performed in duplicate and repeated at least three
times.
2.7 Cell scratch assay
Human RPE cells were cultured in DMEM containing 10% FBS in a 12-well cell
culture plate. After the cells were cultured to confluence, a straight scratch
was made with a 20-L pipette tip. Scratched cells were then washed with
PBS once, immediately, and treated with TGF-2 (10 ng/mL), with or
without TM (100 ng/mL), for 15 h. Images were taken with a microscope (BZ-X700,
KEYENCE). Each experiment was performed in duplicate and repeated at least three
times.
2.8 TUNEL assay
Apoptosis was detected by the terminal deoxynucleotidyl transferase
(TdT)-mediated dUTP-biotin nick end-labeling (TUNEL) assay according to the
manufacturer’s instructions (Promega). Images were taken with a fluorescence
microscope (BZ-X700, KEYENCE).
2.9 Statistical analyses
All experiments were performed at least three times, and the results are
presented as mean SEM. The statistical
significance of differences between groups was analyzed using the Tukey HSD test
or a two-tailed Student’s t-test. For comparison with the control group,
Bonferroni and Holm tests or two-tailed Student’s t-tests were applied.
One-way analysis of variance with Bonferroni correction for multiple comparisons
was performed where applicable. Differences were considered significant at
p 0.05. Statistical analyses were
performed using JMP® version 7.0.1 (SAS Institute, Cary, NC,
USA).
3. Results
3.1 ER stress inhibits EMT of RPE cells
To explore the effects of ER stress on EMT of RPE cells, we used two well-known
ER stress inducers, tunicamycin (TM) and thapsigargin (TG), to induce ER stress
in RPE cells. Then we examined the expression of an epithelial marker, occludin,
and mesenchymal markers, alpha-smooth muscle actin (-SMA) and
fibronectin (FN), in RPE cells. As shown in Fig. 1, treatment with TM and TG (for
24 h) elicited ER stress in RPE cells in a dose-dependent manner, which was
indicated by the gradual increase in the expression of ER stress marker GRP78
(p 0.05). Exposure of RPE cells to different concentrations of TM
and TG (for 24 h) reduced the expression of -SMA and FN, and increased
the expression of occludin significantly (p 0.05). These phenomena
were strongly associated with the expression of GRP78 (Fig. 1). These results
suggested that ER stress inhibited EMT in RPE cells.
Fig. 1.
ER stress inhibits epithelial to mesenchymal transition in a
dose-dependent manner in human RPE cells. The human RPE cells were treated with
different concentrations of TM (0‒300 ng/mL) (A) or TG (0‒400 nM) (B)for 24 h, and the expression of -smooth muscle actin (SMA),
fibronectin (FN), occludin, and Grp78 was analyzed by western blot. GAPDH was
used as the internal control. Data are presented as mean SEM. n =
4/group. *p 0.05, **p 0.01 vs. Ctrl. Densitometry
results of western blotting are shown on the right of each blot; GAPDH was used
as a protein-loading control.
3.2 Induction of ER stress inhibits TGF-2-induced EMT in
RPE cells
TGF-2, the major TGF- isoform in the posterior eye, is a
critical inducer of EMT in RPE cells [14, 15]. To investigate whether ER stress
can inhibit TGF-2-induced EMT of RPE cells, we examined the expression
of epithelial markers and mesenchymal markers in RPE cells at both the mRNA and
protein levels. As shown in Fig. 2A and Supplemental Fig. 1, treatment with
TGF-2 (10 ng/mL) significantly upregulated the expression of
-SMA and FN, and decreased the expression of occludin at both protein
and mRNA levels. This effect was significantly inhibited by a 4-h pre-treatment
with 200 ng/mL and 300 ng/mL TM. In accordance with the results of western
blotting, immunofluorescence showed that -SMA (Fig. 2B) and FN (Fig. 2C) staining were markedly attenuated by 24-h TM treatment; however,
immunostaining of ZO-1 with TM treatment (100 ng/mL) didn’t appear to be
obviously stronger than that of the control (Fig. 2D). Moreover, the staining of
-SMA (Fig. 2B) and FN (Fig. 2C) was strongly enhanced by treatment with
TGF-2, which was considerably inhibited by TM treatment. Interestingly,
not only was ZO-1 expression reduced, but the distribution of ZO-1 staining was
changed by TGF-2 treatment (Fig. 2D). Moreover, the morphology of RPE
cells changed to a spindle-like shape after TGF-2 treatment, according
to the distribution of FN, -SMA, and ZO-1, whereas with TM treatment,
RPE cells maintained a hexagonal shape, as shown by the staining of ZO-1 (Fig. 2D). Taken together, these results implied that induction of ER stress could
inhibit TGF-2-induced EMT of RPE cells.
Fig. 2.
ER stress inhibits TGF-2-induced epithelial to
mesenchymal transition in RPE cells. (A) Human RPE cells were pre-treated with
or without different concentrations of TM for 4 h and then treated with
TGF-2 (10 ng/mL) for another 48 h. The expression of -smooth
muscle actin (SMA), fibronectin (FN), occludin, and Grp78 proteins was analyzed
by western blot. GAPDH was used as internal control. Data are presented as mean
SEM. n = 6/group. NS, not significant. *p 0.05, **p 0.01 vs. Ctrl. (B) Human RPE cells were treated with 0 or 100 ng/mL TM with
or without TGF-2 (10 ng/mL) treatment for 24 h. The immunofluorescence
staining for FN is shown in green. Nuclei are stained blue. (C) Human RPE cells
were treated with 0 or 100 ng/mL TM with or without TGF-2 (10 ng/mL)
treatment for 24 h. The immunofluorescence staining for -SMA is shown
in green. Nuclei are stained blue. (D) Human RPE cells were treated with 0 or 100
ng/mL TM with or without TGF-2 (10 ng/mL) treatment for 24 h. The
immunofluorescence staining for zonula occludens-1 (ZO-1) is shown in green.
Nuclei are stained blue. Scale bar: 50 m for FN and -SMA, 20
m for ZO-1.
3.3 Effects of ER stress on TGF- signaling
The TGF- signaling pathway is one of the best-recognized pathways that
promotes EMT in various cell types, including RPE cells [16, 17]. Therefore, we
further examined the effects of ER stress on the expression and phosphorylation
of the TGF- receptor 2, SMAD2, and SMAD3, induced by TGF- in
RPE cells. As shown in Fig. 3, TGF- (10 ng/mL, 30 min)
significantly induced the phosphorylation of TGF- receptor 2 and SMAD2/3
(p 0.01), which were markedly inhibited by pre-treatment with TM for
16 h (p 0.01). Interestingly, TM treatment alone could also inhibit
the expression of TGF- receptor 2 and SMAD2/3 at the protein level
(p 0.05). The results of quantitative polymerase chain reaction
(RT-qPCR) further validated the effects of ER stress on the expression of
TGF- receptor 2 and SMAD3; however, the expression of SMAD2 was not
inhibited by TM treatment at the mRNA level (Supplementary Fig. 2).
These results suggest that ER stress could inhibit the EMT of RPE cells by
inactivating TGF- signaling.
Fig. 3.
Effect of ER stress on TGF- signaling in human RPE
cells. The human RPE cells were pre-treated with 0 or 100 ng/mL TM for 24 h and
then treated with or without 10 ng/mL TGF-2 for another 30 min. The
expressions of TGF- R2, p-TGF- R2, SMAD2/3, p-SMAD2/3, and
GRP78 proteins were analyzed by western blot. GAPDH was used as internal control.
NS, not significant. *p 0.05, **p 0.01. Data are
presented as mean SEM.
3.4 ER stress inhibits RPE cells migration
During the process of EMT and formation of subretinal fibrosis, one of the most
important functional characteristic changes of the RPE cells is the increase of
migration. Therefore, transwell migration assay and scratch assay were used to
investigate the changes in the migratory capacity of RPE cells. PDGF-BB, an
important growth factor that is expressed in the subretinal membrane in PVR and
lesions of nAMD and is associated with the migration of RPE cells, was used to
induce the cell migration [18]. As shown in Fig. 4A, PDGF-BB significantly
stimulated the migration of RPE cells, whereas TM treatment significantly
inhibited the migration of RPE cells (60%), with or without the stimulation
of PDGF-BB (p 0.01). The results of the scratch assay further
validated the inhibitory effect of ER stress on RPE cell migration, as shown in
Fig. 4B. These data suggest that TM-induced ER stress could inhibit the migration
of RPE cells.
Fig. 4.
ER stress inhibits migration of human RPE cells. (A)The human RPE cells were treated with or without 100 ng/mL TM for 24 h, after
which the cells were treated with or without 25 ng/mL PDGF for 4 h. RPE cell
migration was detected by Boyden chamber migration assay. (B) Human RPE cells
were pre-treated with 0 or 100 ng/mL TM for 8 h and were then scratched for the
scratch assay. NS, not significant. *p 0.05, **p 0.01.
Data are presented as mean SEM.
3.5 Effects of high concentrations of TM and TG on RPE cell survival
and apoptosis
To explore the effects of TM and TG on cell viability and apoptosis in RPE cells
at the concentration we used in this study, we conducted an MTT assay and a TUNEL
assay. In the MTT assay, lower concentrations of TM (up to 300 ng/mL) and TG (up
to 250 nM) revealed no cell toxicity in RPE cells; however, high concentrations
of TM (400 ng/mL) or TG (300 nM) reduced the viability of RPE
cells. In addition, TM treatment induced apoptosis of RPE cells in a
dose-dependent manner, as shown in the TUNEL assay (Supplementary Fig.
3). These data indicated that mild ER stress does not affect cell survival in
RPE cells; however, severe ER stress inhibits cell survival and induces apoptosis
of RPE cells.
4. Discussion
To the best of our knowledge, this study demonstrates for the first time that
chemical induction of ER stress by TM or TG inhibited TGF-2-induced EMT
of RPE cells. We further confirmed that ER stress inactivated TGF-
signaling and inhibited migration of RPE cells. We also revealed that mild ER
stress does not affect the viability of RPE cells; however, severe ER stress
induced by higher concentrations of TM and TG could inhibit cell survival and
induce apoptosis of RPE cells. These results provide evidence for the association
between ER stress and EMT in RPE cells and suggest a potential role of ER stress
as a modifiable target for the prevention or treatment of fibrotic retinal
diseases, such as PVR and AMD, considering that ER stress is an important
mediator in the pathogenesis of such diseases.
According to its different functional consequences, EMT can be divided into
three different subtypes. Type I EMT is associated with implantation,
embryogenesis, and organ formation; Type II EMT is known to play a critical role
in wound healing, tissue regeneration or reconstruction, and organ fibrosis;
while Type III EMT usually occurs in neoplastic cells during the cancer progress
and metastasis [19]. In the development of PVR or in late-stage AMD, RPE cells
undergo EMT and transdifferentiate into a fibroblastic phenotype characterized by
increased EMT phenotypes and the ability to proliferate and migrate to facilitate
development of subretinal fibrosis. This process is regulated by the
participation of a number of growth factors and cytokines, such as TNF, HGF,
PDGF, and TGF-. TGF- is a multifunctional growth factor and a
major inducer of EMT in the pathogenesis of fibrotic eye diseases [20, 21].
Overexpressed TGF- is found in the vitreous of patients with PVR, which
is highly related to the severity of the disease [22, 23]. Since TGF-2
is the predominant isoform of TGF- in the posterior eye, we used
TGF-2 to induce EMT in RPE cells in our study. Consistent with previous
studies, our results confirmed that RPE cells undergo EMT when stimulated with
TGF-2, as shown by a morphological change with up-regulation of
-SMA and FN, and downregulation of occludin at both protein and mRNA
levels. There are several signaling pathways that are involved in regulating the
process of EMT. Among them, the best known is the TGF- signaling
pathway, which signals through canonical SMAD signaling as well as through some
non-SMAD-dependent pathways, such as the MAPK pathway and PI-3 kinase pathway. In
the canonical SMAD signaling pathway, signaling is activated by the interaction
of TGF- with type II and type I serine‒threonine kinase receptors,
termed TGF-RII and TGF-RI. Following ligand binding,
TGF-RI is phosphorylated by TGF-RII, which in turn
phosphorylates SMAD2/3. Activated SMAD2/3 then binds to SMAD4 and translocated
into the nucleus to regulate the expression of target genes. Numerous studies
have revealed that the SMAD signaling pathway is critically involved in EMT and
in the pathogenesis of tissue fibrosis [24, 25]. Inactivation of SMAD signaling
significantly inhibits EMT in RPE cells and fibrogenesis in many other cell types
[24, 26, 27]. Extensive studies have revealed the essential role of SMAD3 in EMT
and fibrosis [28]. RPE cells in Smad3-null mice failed to undergo EMT in
response to retinal detachment, indicating the pivotal role of SMAD3 in the EMT
of RPE cells [29]. Furthermore, Shin et al. [11] demonstrated that ER
stress preconditioning blocked activation of SMAD signaling in human peritoneal
mesothelial cells. In this study, we found that ER stress inducers inhibited the
phosphorylation of SMAD2/3 induced by TGF-2 treatment of RPE cells. In
addition, it also inhibited the expression of TGF- receptor and SMAD3
with or without TGF- stimulation. These results support the suggestion
that ER stress inhibits EMT of RPE cells, at least in part, by inactivating the
TGF/SMAD signaling pathway.
The ER is an important organelle that is essential for the synthesis of lipids,
regulation of calcium homeostasis, and most importantly, the synthesis, folding,
assembly, modification, and translocation of proteins. Under normal conditions,
unfolded or misfolded proteins are degraded via ER-associated degradation, while
under harsh environmental stresses, such as hypoxia, nutrient deprivation, or
metabolic stress, accumulation of misfolded/unfolded proteins in the ER lumen
leads to activation of ER stress and the UPR. GRP78 is one of the most important
and predominant ER chaperones that identify misfolded proteins and facilitate
protein folding in the ER. Numerous studies have verified the specific induction
of GRP78 as an indicator of ER stress [30]. In our study, both TM and TG, two
classical chemical inducers of ER stress, increased the expression of GRP78 in a
dose-dependent manner. Over the past decade, evidence has emerged to reveal an
association between ER stress and a variety of human diseases, including
metabolic disease, neurodegenerative disorders, infectious and inflammatory
diseases, fibrosis, and cancer [8]; however, the exact relationship between ER
stress and those diseases has not been fully understood. On the one hand, a
number of studies have demonstrated that ER stress can suppress EMT and fibrosis.
For example, downregulation of GRP78 promotes EMT and migration of human
hepatocellular carcinoma cells [31]. Knockdown of GRP78 induces EMT in colon
cancer cells and Mahlavu cells, as characterized by downregulation of vimentin,
upregulation of E-cadherin, and increased migratory ability [32, 33]. A more
recent study found that matrine treatment inhibited EMT in human prostate cancer
cells as well as activating the ER stress signal pathway in both vivo and vitro
[34]. Similar results were observed in Honokiol treated melanoma cells [35],
indicating a potential suppressive effect of ER stress on EMT. Furthermore,
pre-treatment with TM or TG inhibited TGF-1-induced EMT in HPMCs,
suggesting a protective role of ER stress in TGF--induced EMT [11]. On
the other hand, ER stress can also play a facilitatory role in EMT and fibrosis.
Induction of ER stress results in EMT in human lens epithelial cells and lung
epithelial cells [12, 36]. Similarly, Moon et al. [37] showed that TM or
TG treatment induces EMT in HK-2 cells via activation of the Src pathway. Taken
together, these results suggest that the role of ER stress in EMT and fibrosis
may be context-dependent meaning based on difference of microenvironment, even
though the exact causes of the discrepancies among these studies remain unclear.
We speculate that the differences in cell types, cell status, culture conditions,
and the concentration of TM or TG used in each experiment might be possible
reasons.
Despite ample studies showing the association of ER stress and EMT with organ
fibrosis, evidence regarding the effects of ER stress on EMT in RPE cells remains
very limited. Yoshikawa et al. [38] reported that ER stress increased
the expression of tight junctions, such as occludin, claudin-1, and ZO-1, at both
the protein and mRNA levels in APRE-19 cells. Another study showed that
overexpression of the ER protein 29 (ERp29), which is increased during ER stress,
protected against the CSE-induced reduction in ZO-1 expression in ARPE-19 cells
[39]. However, no previous studies have explored the role of ER stress on EMT
induced by TGF- in human RPE cells. Here, we reported that chemical
induction of ER stress in RPE cells, by means of TM or TG, increased the
expression of the tight junction protein, occludin, and decreased the expression
of the mesenchymal markers, -SMA and FN. Moreover, ER stress could also
inhibit TGF--induced EMT and inactivate SMAD signaling. These results
suggest the beneficial effects of ER stress on the EMT of RPE cells.
Proliferation and migration of RPE cells are key abnormal steps in the formation
of subretinal or epiretinal fibrosis membranes during the development of PVR or
nAMD [5, 40]. Previous studies have shown that, when RPE cells undergo EMT, the
dedifferentiated cells acquire an increased migratory capacity and migrate to the
vitreous cavity to contribute to the formation of the contractile membranes [1, 41, 42]. This process is facilitated by the remodeling of the ECM, involving
proteins such as fibronectin and collagen [1, 43]. While several studies have
shown the inhibitory effect of ER stress on cell migration, no study investigated
its effect on cell migration in RPE cells. In vascular smooth muscle cells, TM
pretreatment inhibited PDFG-BB-induced cell migration [44]. Downregulation of
GRP78 increased the migratory ability of colon cancer cells and Mahlava cells
[32, 33]. In our study, consistent with its inhibitory effect on EMT, ER stress
induced by TM significantly reduced cell migration and fibronectin synthesis in
RPE cells, without causing significant apoptosis. Moreover, it could also inhibit
PDGF-BB-induced cell migration. These results indicate that ER stress might
regulate ECM remodeling and RPE cell migration, not only through EMT inhibition
but also by mediating signaling pathways involved in PDGF-BB-induced migration.
5. Conclusions
In conclusion, our data demonstrated that ER stress inhibits
TGF--induced EMT in RPE cells. Our results suggest the potential role of
ER stress in the pathogenesis of intraocular fibrotic diseases, such as PVR and
AMD and may be as a new therapeutic approach to prevent such kind of disorder.
Abbreviations
ER stress, Endoplasmic reticulum stress; EMT, Epithelial-mesenchymal transition;
UPR, Unfolded protein response; RPE, Retinal pigment epithelial; PVR,
Proliferative vitreoretinopathy; AMD, Age-related macular degeneration;
TGF-, Transforming growth factor-; TM, Tunicamycin; TG,
Thapsigargin; FN, Fibronectin; -SMA, Alpha-smooth muscle actin; ZO-1,
Zonula occludens-1; GRP78, Glucose regulated p rotein 78 kD; PERK, Protein kinase
R-like ER kinase; ATF6, Activating transcriptional factor 6; ATF4, Activating
transcriptional factor 4; TGFBR2, Transforming growth factor- receptor
II; GAPDH, Glyceralehyde-3-phosphate dehydrogenase.
Author contributions
SO, SH and XX conceived and designed the experiments; SO performed the
experiments; SO, DJ analyzed data; SO, SH and XX involved in writing and
reviewing the paper, and all authors had final approval of the submitted
versions.
Ethics approval and consent to participate
Not applicable.
Acknowledgment
The research is supported by a fellowship from the second Xiangya Hospital of
Xiangya School of Medicine of Central South University for Research Abroad. The
authors thank Christine K. Spee, Eric Barron and Ernesto Barron for technical
assistance.
Funding
This research received no external funding.
Conflict of interest
The authors declare no conflict of interest.
SH is serving as one of the Guest Editor of this journal. We
declare that SH had no involvement in the peer review of this
article and has no access to information regarding its peer
review. Full responsibility for the editorial process for this article
was delegated to GP.