Hepcidin is a hormone central to the regulation of iron homeostasis in the body. It is believed to be produced exclusively by the liver. Ferroportin, an iron exporter, is the receptor for hepcidin. This transporter/receptor is expressed in Müller cells, photoreceptor cells and the RPE (retinal pigment epithelium) within the retina. Since the retina is protected by the retinal–blood barriers, we asked whether ferroportin in the retina is regulated by hepcidin in the circulation or whether the retina produces hepcidin for regulation of its own iron homeostasis. Here we show that hepcidin is expressed robustly in Müller cells, photoreceptor cells and RPE cells, closely resembling the expression pattern of ferroportin. We also show that bacterial LPS (lipopolysaccharide) is a regulator of hepcidin expression in Müller cells and the RPE, both in vitro and in vivo, and that the regulation occurs at the transcriptional level. The action of LPS on hepcidin expression is mediated by the TLR4 (Toll-like receptor-4). The upregulation of hepcidin by LPS occurs independent of Hfe (human leukocyte antigen-like protein involved in Fe homeostasis). The increase in hepcidin levels in retinal cells in response to LPS treatment is associated with a decrease in ferroportin levels. The LPS-induced upregulation of hepcidin and consequent down-regulation of ferroportin is associated with increased oxidative stress and apoptosis within the retina in vivo. We conclude that retinal iron homeostasis may be regulated in an autonomous manner by hepcidin generated within the retina and that chronic bacterial infection/inflammation of the retina may disrupt iron homeostasis and retinal function.

INTRODUCTION

Iron, an essential nutrient obligatory for vital cellular functions, can induce oxidative stress and cellular dysfunction when excessively accumulated. Hereditary haemochromatosis is a genetic disorder associated with iron overload [15]. The tissues commonly affected in this disease are the liver, pancreas, kidney, pituitary and heart, resulting in a myriad of diseases including liver cirrhosis, hepatocarcinoma, diabetes, cardiomyopathy, nephropathy and endocrine dysfunction [15].

Hepcidin is an important regulator of iron homeostasis. It is considered to be a circulatory hormone, secreted almost exclusively by the liver [6]. Previous studies have identified ferroportin, an iron exporter expressed in specific cell types, as the receptor for this iron-regulatory hormone [7]. The binding of hepcidin to ferroportin leads to internalization of the iron exporter with subsequent degradation in lysosomes. This results in the prevention of iron release from cells which express ferroportin. Duodenal enterocytes express ferroportin at the basolateral membrane, and the transporter is involved in the intestinal absorption of dietary iron [15]. Macrophages also express ferroportin and play a critical role in the recycling of haem–iron from aged erythrocytes. A decrease in circulating levels of hepcidin is a common feature in haemochromatosis, irrespective of whether the disease is caused by mutations in HFE (the gene coding for HFE; human leukocyte antigen-like protein involved in Fe homeostasis), Tfr2 (the gene coding for transferrin receptor 2), HJV (haemojuvelin) or hepcidin [8]. The decrease in hepcidin prevents internalization of ferroportin in duodenal enterocytes, enhancing intestinal iron absorption and thus causing iron overload. In contrast, an increase in circulating levels of hepcidin leads to iron deficiency, resulting from decreased intestinal iron absorption and decreased recycling of haem–iron [9]. It has been documented that an increase in hepcidin expression leads to iron deficiency and microcytic anaemia [10,11]. Inflammation upregulates hepcidin expression and increases circulating levels of hepcidin [11], providing a molecular basis for the anaemia associated with chronic inflammation.

Iron overload has been postulated to play a role in the pathogenesis of various neurodegenerative diseases such as Parkinson's disease [12], Alzheimer's disease [13,14] and amyotrophic lateral sclerosis [15]. This suggests that the central nervous system is susceptible to iron-induced oxidative damage. Recent studies have shown that hepcidin and its target ferroportin are expressed in the central nervous system, indicating that iron homeostasis within the brain may be regulated by hepcidin which is produced locally [16,17]. Retinas are constantly exposed to oxidative stress caused by photo-oxidation, and regulation of iron homeostasis within this tissue is important to prevent exacerbation of this oxidative stress by iron overload [18]. The RPE (retinal pigment epithelium), which constitutes the outer blood–retinal barrier, is responsible for phagocytosis of photoreceptor outer segments. These membrane segments are rich in iron, thus exposing this cell layer to increased risk for iron overload [19]. Müller cells play a critical role as a support cell to retinal neurons to maintain normal retinal function. Previous studies have shown that ferroportin is expressed in photoreceptor cells, Müller cells and RPE cells [20], suggesting that this transporter may play an obligatory role in the maintenance of iron homeostasis within the retina. We recently reported that Hfe and Tfr2, two other proteins involved in the regulation of iron homeostasis, are also expressed in the retina [21]. Despite the fact that hepcidin may be the most crucial regulator of iron homeostasis and that an exquisite regulation of iron levels within the retinal cells is obligatory for optimal visual function, the expression and function of hepcidin have not been studied in the retina. Ferroportin, the receptor for hepcidin, is expressed in the basolateral membrane of RPE and thus has access to hepcidin in the circulation. This is not true in the case of ferroportin expressed in Müller cells and photoreceptor cells. Since the retina is surrounded by retinal–blood barriers, ferroportin present in these cells may not be subject to regulation by circulating hepcidin unless the 25-amino-acid peptide can pass through the retinal–blood barriers. Therefore we were interested to know whether ferroportin expressed within the retina is regulated by hepcidin produced within the retina. The present investigation was undertaken to study the expression, function and regulation of hepcidin in mouse retina.

MATERIALS AND METHODS

Materials

Reagents were obtained from the following sources: RNA extraction reagent TRIzol, Invitrogen-Gibco; GeneAmp RT–PCR (reverse transcription–PCR) kit, Applied Biosystems; Taq polymerase kit, TaKaRa; and Power Block, Biogenex. DMEM (Dulbecco's modified Eagle's medium)/F12 medium, Gibco, with 10% fetal bovine serum, 100 units/ml penicillin, and 100 μg/ml streptomycin was used for growing retinal cell lines. Antibodies used were obtained from the following sources: rabbit polyclonal anti-ferroportin, rabbit polyclonal anti-HNE (anti-4-hydroxynonenal), rabbit polyclonal anti-hepcidin, and hepcidin control/blocking peptide, Alpha Diagnostic International; goat anti-rabbit IgG coupled to Alexa Fluor 555, Molecular Probes; Vectashield Hardset Mounting Medium with DAPI (4′,6-diamidino-2-phenylindole), Vector Laboratories; lipofectin transfection reagent, Invitrogen; ApopTAG fluorescein in situ Apoptosis Detection Kit, Chemicon International; and ketamine, xylazine, proparacaine, and bacterial LPS (lipopolysaccharide), Sigma-Aldrich.

Animals

C57BL/6 mice (6-week-old) were used for the preparation of total RNA from neural retina and RPE/eye cup, and 3-week-old mice were used for establishment of primary RPE cell cultures. Hfe knockout mice were obtained from the Jackson laboratory (Bar Harbor, ME, U.S.A.). Albino Balb/c mice (3-week-old) were used for immunofluorescence and in situ hybridization analyses. These mice were also used for intravitreous injection of LPS to induce inflammation in the eyes. Mice were purchased from Harlan-Sprague Dawley and maintained in our facility. The experimental procedures with mice were carried out in accordance with the United States NIH guidelines and with the institutional policies governing appropriate care and use of animals in research.

Establishment of primary RPE cell cultures from mouse eyes

Three-week-old C57BL/6 mice were used for preparation of primary RPE cell cultures using a method adapted from that described for isolation of rat RPE [22]. Briefly, enucleated mouse eyes were rinsed in 5% Povidone-Iodine solution, followed by rinsing with sterile HBSS (Hanks balanced salt solution). Eyes were then placed in cold RPE cell culture medium which consisted of DMEM:F12 medium, supplemented with 25% fetal bovine serum, gentamicin (0.1 mg/ml), penicillin (100 units/ml), and streptomycin (100 μg/ml). To aid in the degradation of extracellular matrix components and enhance the dissociation of RPE from neural retina and choroid, eyes were then incubated in HBSS containing collagenase (19.5 units/ml) and testicular hyaluronidase (38 units/ml) for 40 min at 37 °C, followed by incubation in HBSS containing 0.1% trypsin (pH 8.0) for 50 min at 37 °C. Eyes were then dissected to separate RPE from neural retina. Isolated RPE cells were collected in a 15 ml centrifuge tube and centrifuged at 350 g for 10 min at 21 °C, followed by resuspension in RPE cell medium. RPE cells were then cultured at 37 °C. Purity of the culture was verified by immunodetection of RPE-65 (retinal pigment epithelial protein 65) and CRALBP (cellular ratinaldehyde binding protein), proteins known to be expressed in the RPE.

Primary cultures of Müller cells

Primary cultures of retinal Müller cells were established from retinas of C57BL/6 mice according to our previously published method [23]. Purity of the culture was verified by immunodetection of vimentin, glutamine synthetase, CRALBP and the glutamate transporter EAAT1, proteins known to be expressed in Müller cells.

Intravitreal injection

Female Balb/c mice (3 weeks) were weighed and anaesthetized by intraperitoneal injection using 15 μl (1 μl per gram body weight) of a solution of ketamine (80 mg/ml) and xylazine (12 mg/ml). Immediately before the intraperitoneal injection, 5 μl of 5% proparacaine was administered topically to the eyes. One microlitre of LPS (10 μg/μl prepared in PBS) was then injected into the vitreous body of the right eyes at the limbus, and the left eye received an equal volume of PBS. Control mice received an equal volume of PBS in both the eyes. Following intravitreal injection, eyelids were gently closed and the animals were placed in a cage for observation. Within 30 min, the animals resumed activity and showed no evidence of stress or discomfort. The next day (24 h after injections), mice were killed by CO2 inhalation. One set of the mouse eyes were harvested for cryosectioning and immunohistochemistry. Neural retina and RPE were dissected from remaining eyes for isolation of RNA.

RT–PCR

Neural retina and RPE/eye cup were prepared according to our previously published method [24] and used for preparation of total RNA. RT–PCR was carried out under optimal conditions depending on the nature of the specific PCR primer pairs listed in Table 1. HPRT1 (hypoxanthine phosphoribosyl transferase 1) was used as an internal control for the PCR reaction. The hepcidin PCR product was subcloned into the pGEM-T Easy vector and sequenced to confirm its molecular identity. The subcloned plasmid was also used for generation of sense and antisense riboprobes for in situ hybridization.

Table 1
PCR primers
Gene name NCBI Accession No. Primer sequence 
Mouse hepcidin AF297664 Forward: GCACCACCTATCTCCATCAACAGA 
  Reverse: GGTCAGGATGTGGCTCTAGGCTAT 
Rat hepcidin AF344185 Forward: GAAGGCAAGATGGCACTAAGCA 
  Reverse: TCTCGTCTGTTGCCGGAGATAG 
Human hepcidin NM_021175 Forward: ACTGTCACTCGGTCCCAGACA 
  Reverse: TCCAAGACCTATGTTCTGGGG 
Mouse Hfe NM_010424 Forward: GGCTTCTGGAGATATGGTTAT 
  Reverse: GACTCCACTGATGATTCCGATA 
Mouse Hjv NM_027126 Forward: GGCTGAGGTGGACAATCTTC 
  Reverse: GAACAAAGAGGGCCGAAAG 
Mouse TLR4 BC029856 Forward: CTGCCAAGTCTCAGCTATCT 
  Reverse: CTGCTAAGAAGGCGATACAA 
Rat TLR4 NM_019178 Forward: ACAAGAGCCGGAAAGTTATT 
  Reverse: CTGCTAAGAAGGCGATACAA 
Human TLR4 BC117422 Forward: ACAGACTTGCGGGTTCTACA 
  Reverse: CTGCTGAGAAGGCGGTACAG 
Gene name NCBI Accession No. Primer sequence 
Mouse hepcidin AF297664 Forward: GCACCACCTATCTCCATCAACAGA 
  Reverse: GGTCAGGATGTGGCTCTAGGCTAT 
Rat hepcidin AF344185 Forward: GAAGGCAAGATGGCACTAAGCA 
  Reverse: TCTCGTCTGTTGCCGGAGATAG 
Human hepcidin NM_021175 Forward: ACTGTCACTCGGTCCCAGACA 
  Reverse: TCCAAGACCTATGTTCTGGGG 
Mouse Hfe NM_010424 Forward: GGCTTCTGGAGATATGGTTAT 
  Reverse: GACTCCACTGATGATTCCGATA 
Mouse Hjv NM_027126 Forward: GGCTGAGGTGGACAATCTTC 
  Reverse: GAACAAAGAGGGCCGAAAG 
Mouse TLR4 BC029856 Forward: CTGCCAAGTCTCAGCTATCT 
  Reverse: CTGCTAAGAAGGCGATACAA 
Rat TLR4 NM_019178 Forward: ACAAGAGCCGGAAAGTTATT 
  Reverse: CTGCTAAGAAGGCGATACAA 
Human TLR4 BC117422 Forward: ACAGACTTGCGGGTTCTACA 
  Reverse: CTGCTGAGAAGGCGGTACAG 

In situ hybridization

Mouse eyes were frozen in Tissue-Tek OCT (optimal cutting temperature), and sections were made at 10 μm thickness and fixed in 4% paraformaldehyde. Treatment of tissue sections and hybridization with digoxigenin-labelled sense and antisense riboprobes were carried out as described previously [2527]. The hybridization signals were detected with anti-digoxigenin antibody conjugated to alkaline phosphatase. The colour reaction was developed with Nitro Blue Tetrazolium/5-bromo-4-chloro-3-indolyl phosphate. Cryosections were hybridized with the sense (negative control) riboprobe to determine non-specific binding. For the preparation of antisense and sense riboprobes, a 269 bp product specific for mouse hepcidin was amplified by RT–PCR and subcloned into the pGEM-T Easy vector, and the orientation of the insert was identified by sequencing. The specificity of the probe for hepcidin was confirmed by searching the GenBank database with the nucleotide sequence of the segment encompassed by the primers as the query. The probes were prepared by in vitro transcription with appropriate RNA polymerases after linearizing the plasmid with suitable restriction enzymes.

Immunofluorescence analysis

Cryosections of mouse eyes were fixed in 4% paraformaldehyde for 10 min, washed with 0.01 M PBS (pH 7.4), and blocked with 1×Power Block for 60 min. Sections were then incubated overnight at 4 °C with one of the following primary antibodies: polyclonal anti-hepcidin (1:50 dilution), polyclonal anti-ferroportin (1:100 dilution) or polyclonal anti-HNE (1:500 dilution). The specificity of the hepcidin antibody was confirmed by using the peptide-neutralized antibody. The hepcidin antibody was mixed with the antigenic peptide, and incubated for 2 h at 37 °C plus 24 h at 4 °C. The mixture was centrifuged at 3000 g for 15 min at 4 °C to pellet the immune complex. The supernatants were incubated with the sections overnight at 4 °C to serve as negative controls. Additional negative controls involved the omission of the primary antibodies. Sections were rinsed and incubated for 1 h with goat anti-rabbit IgG coupled to Alexa Fluor 555 at a dilution of 1:1000. Coverslips were mounted with Vectashield Hardset mounting medium with DAPI (a nuclear stain) and sections were examined by epifluorescence using an Axioplan-2 microscope, equipped with an HRm camera and the Axiovision imaging program (Carl Zeiss).

Retinal cell lines and primary retinal cell cultures were grown on 12 mm coverslips for 24 h in 24-well cell culture plates at 37 °C in a 5% CO2 incubator. Medium was removed and the cells were fixed in ice-cold methanol for 10 min after air drying. Cells were then washed with 0.01 M PBS (pH 7.4) and blocked with 1×Power Block for 120 min. Cells were incubated overnight at 4 °C with polyclonal anti-hepcidin (1:50 dilution), polyclonal anti-ferroportin (1:100 dilution), or polyclonal anti-HNE (1:500 dilution). After incubation of the cells with primary antibody, the same protocol was followed as used for staining the tissues. Negative control cells were treated likewise, but in the absence of the primary antibodies.

Generation of hepcidin-specific promoter–reporter constructs

The human hepcidin promoter–EGFP (enhanced green fluorescent protein) construct was generated by first subcloning the 2 kb hepcidin promoter (obtained by PCR using human genomic DNA as the template) into the pGEM-T Easy vector and then the HindIII/XhoI-digested promoter was inserted into pUIIR3-EGFP vector. The primers used for PCR were: 5′-ATACTCGAGACTCTCACCCAGGCTGGG-3′ (sense) and 5′-AAGCTTCATCGTGCCGTCTGTCTGGCT-3′ (antisense). After transfection with the hepcidin promoter–EGFP construct, cells were treated with LPS for 4 h and EGFP expression was monitored by epifluorescence under the fluorescence microscope.

In situ detection of DNA fragmentation by the TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling) assay

The TUNEL assay was performed using the ApopTAG fluorescein in situ apoptosis detection kit. After staining, the tissue sections were viewed by epifluorescence using standard fluorescence excitation and emission filters. Each section was scanned systematically from the temporal to the nasal ora serrata for fluorescent cells indicative of apoptosis. To distinguish between structures that autofluoresced versus those that were TUNEL-positive, all slides were examined first with the rhodamine filter and then with the FITC filter. Autofluorescent structures were visible under both filters, whereas TUNEL-positive cells were detectable only with the FITC filter.

Primary cultures of RPE from wild-type and Hfe−/− mouse retinas

Wild-type and Hfe−/− mice were obtained from the same litter originating from the mating of heterozygous mice. Genotyping was performed according to the protocol supplied by The Jackson Laboratory. Mice (3-week-old) were used to establish primary cultures of RPE.

RESULTS

Expression of hepcidin in retina

To investigate whether hepcidin is expressed in the retina, RT–PCR was done with RNA isolated from neural retina and RPE/eye cup of normal mouse. We found evidence for the expression of hepcidin mRNA in RPE/eye cup as well as in the neural retina (Figure 1A). The PCR product was subcloned in pGEM-T Easy vector and sequenced to confirm the molecular identity of the product using BLAST analysis. We further analysed the expression pattern of hepcidin mRNA and protein in the retina by in situ hybridization and immunofluorescence. In situ hybridization revealed the expression of hepcidin mRNA in the inner nuclear layer, the inner segment of photoreceptor cells and the RPE cell layer (Figure 1B). The signals observed were specific because no positive signals were detected with a sense riboprobe. Immunofluorescence analysis of hepcidin protein showed positive signals throughout the retina (Figure 1C). The cells in the inner nuclear layer, which include Müller cells, were immunopositive. The expression was also evident in photoreceptor cells and the RPE (Figure 1C). Negative controls with omission of the primary antibody or with peptide-neutralized antibody did not yield positive signals, indicating the specificity of the signals obtained with the antibody. The expression pattern of hepcidin protein corroborates the expression pattern of hepcidin mRNA. The inner nuclear layer contains, among other cell types, the nuclei for Müller cells, the inner segment of photoreceptor cells enriched in RNA and the RPE cell layer containing the nuclei for the RPE. Thus, there is a good correlation between the expression pattern of hepcidin mRNA and that of hepcidin protein.

Expression of hepcidin in mouse retina

Figure 1
Expression of hepcidin in mouse retina

(A) RT–PCR analysis of hepcidin mRNA in neural retina and RPE/eye cup from mouse eyes. (B) Analysis of hepcidin mRNA in mouse retina by in situ hybridization. Right panel shows a representative section of retina hybridized with a digoxigenin-labelled antisense riboprobe specific for hepcidin. Left panel is a negative control with a sense probe. GCL, ganglion cell layer; INL, inner nuclear layer; ONL, outer nuclear layer. (C) Immunofluorescence localization of hepcidin protein in mouse retina. Left panel, negative control without the primary antibody. Middle panel, antibody specificity using the hepcidin antibody which had been neutralized with the antigenic peptide. Right panel, the pattern of expression of hepcidin in retina. (D) RT–PCR analysis of hepcidin mRNA in primary RPE cells and primary Müller cells from mouse eyes. HPRT1 was used as an internal control. (E) Immunofluorescence showing hepcidin protein expression in primary Müller cells, primary RPE cells, and their respective cell lines rMC1 and ARPE19. Right panel indicates negative control with the omission of the primary antibody.

Figure 1
Expression of hepcidin in mouse retina

(A) RT–PCR analysis of hepcidin mRNA in neural retina and RPE/eye cup from mouse eyes. (B) Analysis of hepcidin mRNA in mouse retina by in situ hybridization. Right panel shows a representative section of retina hybridized with a digoxigenin-labelled antisense riboprobe specific for hepcidin. Left panel is a negative control with a sense probe. GCL, ganglion cell layer; INL, inner nuclear layer; ONL, outer nuclear layer. (C) Immunofluorescence localization of hepcidin protein in mouse retina. Left panel, negative control without the primary antibody. Middle panel, antibody specificity using the hepcidin antibody which had been neutralized with the antigenic peptide. Right panel, the pattern of expression of hepcidin in retina. (D) RT–PCR analysis of hepcidin mRNA in primary RPE cells and primary Müller cells from mouse eyes. HPRT1 was used as an internal control. (E) Immunofluorescence showing hepcidin protein expression in primary Müller cells, primary RPE cells, and their respective cell lines rMC1 and ARPE19. Right panel indicates negative control with the omission of the primary antibody.

Expression of hepcidin in specific retinal cell types

RT–PCR was performed using hepcidin-specific primers with RNA isolated from primary cultures of RPE and Müller cells. RPE and Müller cells showed significant expression of hepcidin mRNA (Figure 1D). To confirm further the expression of hepcidin in RPE and Müller cells, immunolocalization studies were done. Immunofluorescence analysis confirmed hepcidin expression not only in primary cultures of RPE and Müller cells but also in ARPE-19 (a human RPE cell line) and rMC1 (a rat Müller cell line) cells (Figure 1E).

Regulation of hepcidin expression by LPS

To assess the role of hepcidin in the retina during inflammation, primary cultures of RPE and Müller cells as well as RPE and Müller cell lines were treated with and without LPS (10 and 100 ng/ml in culture medium) for 4 h. The steady-state levels of hepcidin mRNA increased with LPS treatment in a dose-dependent manner in primary cultures of RPE and Müller cells (Figure 2A). The same was true for rMC1; interestingly, hepcidin expression did not respond to LPS treatment in ARPE-19 cells, but the LPS induction of hepcidin expression was evident in HRPE cells, also a human RPE cell line. LPS produces its effects by acting as a ligand of the TLR4 (Toll-like receptor-4). To examine the potential reasons for the differential effects of LPS on hepcidin expression in ARPE-19 cells versus HRPE cells, we monitored the expression of TLR4 in these cells. The expression of TLR4 was several-fold higher in HRPE cells than in ARPE-19 cells (Figure 2B). RNA from human colon was used as a positive control. These data suggest that ARPE-19 cells failed to respond to LPS treatment because of the markedly reduced levels of TLR4 expression. The increase in hepcidin mRNA in primary cultures of RPE and Müller cells following LPS treatment was associated with an increase in hepcidin protein levels as detected by immunofluorescence (Figures 2C and 2D).

Regulation of hepcidin expression by LPS

Figure 2
Regulation of hepcidin expression by LPS

(A) RT–PCR showing hepcidin mRNA levels in untreated and LPS treated (10 ng/ml and 100 ng/ml) retinal cells. (B) RT–PCR showing TLR4 expression levels in retinal cells. (C) Hepcidin expression in primary RPE cells treated with or without LPS. (D) Hepcidin expression in primary Müller cells treated with or without LPS. The first panels in (C) and (D) indicate negative control with the omission of the primary antibody.

Figure 2
Regulation of hepcidin expression by LPS

(A) RT–PCR showing hepcidin mRNA levels in untreated and LPS treated (10 ng/ml and 100 ng/ml) retinal cells. (B) RT–PCR showing TLR4 expression levels in retinal cells. (C) Hepcidin expression in primary RPE cells treated with or without LPS. (D) Hepcidin expression in primary Müller cells treated with or without LPS. The first panels in (C) and (D) indicate negative control with the omission of the primary antibody.

Induction of hepcidin promoter activity by LPS

The effect of LPS on hepcidin expression in the liver occurs via transcriptional regulation [26]. To examine whether a similar mechanism operates in the regulation of hepcidin expression in retinal cells, we monitored the effects of LPS on hepcidin promoter activity using EGFP as the reporter. For this, a 2 kb hepcidin promoter region was cloned into an EGFP vector, and the promoter–reporter construct was transfected into rMC1, ARPE-19 and HRPE cell lines. The next day (24 h after transfection), the cells were treated with LPS (100 ng/ml culture medium) for 4 h. The promoter activity was then monitored using a fluorescence microscope to detect the expression of the EGFP reporter. Vector-transfected cells served as the negative controls. The induction of hepcidin promoter activity by LPS was detectable in rMC1 cells and HRPE cells but not in ARPE-19 cells (Figure 3). This corroborates the findings on the effects of LPS treatment on hepcidin mRNA levels in these cell lines.

Effects of LPS on hepcidin promoter activity using EGFP reporter assay

Figure 3
Effects of LPS on hepcidin promoter activity using EGFP reporter assay

Left panels show the effects of LPS on EGFP reporter in the absence of hepcidin promoter. Right panels show the effects of LPS on hepcidin promoter–EGFP construct.

Figure 3
Effects of LPS on hepcidin promoter activity using EGFP reporter assay

Left panels show the effects of LPS on EGFP reporter in the absence of hepcidin promoter. Right panels show the effects of LPS on hepcidin promoter–EGFP construct.

Regulation of hepcidin expression by LPS in retina in vivo

To determine whether the effects of LPS on hepcidin mRNA and protein levels seen in vitro in retinal cell lines and in primary cell cultures are also demonstrable in vivo, we injected LPS into mouse eyes intravitreally. The next day (24 h after LPS injection), neural retina and RPE/eye cup were isolated for preparation of RNA, and retinal sections were prepared for analysis of oxidative stress and apoptotic cell death. RT–PCR showed an increase in hepcidin mRNA in both neural retina and RPE/eye cup in response to LPS treatment. Interestingly, the induction of hepcidin mRNA was seen not only in the eyes that were injected with LPS but also in the contralateral eyes from the same mice that were injected with PBS instead of LPS, even though the effects were considerably lower (Figure 4A). The induction of hepcidin expression by LPS was independent of Hfe or Hjv because the steady-state levels of mRNAs specific for these genes were not altered by LPS treatment in the retina. The lack of involvement of Hfe in LPS-induced expression of hepcidin was confirmed with primary RPE cell cultures established from retinas of wild-type mice and Hfe-null mice. The genotype identity was confirmed by analysis of Hfe mRNA in wild-type RPE and Hfe−/− RPE (Figure 4B). Treatment of Hfe−/− RPE cells with LPS induced hepcidin expression as is evident from an increase in mRNA levels (Figure 4C) and protein levels (Figure 4D).

Effects of intravitreous injection of LPS on the expression of iron-regulatory proteins in the retina and non-involvement of Hfe in the process

Figure 4
Effects of intravitreous injection of LPS on the expression of iron-regulatory proteins in the retina and non-involvement of Hfe in the process

(A) PBS was injected in both eyes of control mice (lane 1), LPS (10 μg) was injected in the right eye (lane 3) and PBS in the left eye in experimental mice (lane 2). RT–PCR was done for Hfe, HJV and hepcidin in neural retina and RPE/eye cup. HPRT1 was used as an internal control. (B) RT–PCR showing Hfe expression in primary RPE cells from wild-type and Hfe-null mice. (C) RT–PCR analysis of hepcidin in control and LPS-treated (10 ng/ml and 100 ng/ml) primary RPE cells from Hfe-null mice. (D) Hepcidin protein expression in control and LPS-treated (10 ng/ml and 100 ng/ml) primary RPE cells from Hfe-null mice.

Figure 4
Effects of intravitreous injection of LPS on the expression of iron-regulatory proteins in the retina and non-involvement of Hfe in the process

(A) PBS was injected in both eyes of control mice (lane 1), LPS (10 μg) was injected in the right eye (lane 3) and PBS in the left eye in experimental mice (lane 2). RT–PCR was done for Hfe, HJV and hepcidin in neural retina and RPE/eye cup. HPRT1 was used as an internal control. (B) RT–PCR showing Hfe expression in primary RPE cells from wild-type and Hfe-null mice. (C) RT–PCR analysis of hepcidin in control and LPS-treated (10 ng/ml and 100 ng/ml) primary RPE cells from Hfe-null mice. (D) Hepcidin protein expression in control and LPS-treated (10 ng/ml and 100 ng/ml) primary RPE cells from Hfe-null mice.

Consequences of LPS-induced increase in hepcidin expression on ferroportin levels in retina

Hepcidin is known to bind ferroportin and induce lysosomal degradation of the transporter [7]. To determine whether similar changes occur in steady-state levels of ferroportin in intact retina following LPS treatment, we monitored the expression of hepcidin and ferroportin by immunofluorescence in retinal sections from control mouse eyes, PBS-treated contralateral eyes from mice treated with LPS on one eye, and from LPS-treated eyes. To compare the expression levels among different tissue sections, the exposure time on camera was held constant. These studies showed that LPS treatment increased the levels of hepcidin protein and that this increase was associated with a substantial decrease in ferroportin protein levels (Figure 5). Similar changes, but to a smaller extent, were seen in PBS-injected contralateral eyes.

Immunostaining for hepcidin and ferroportin in LPS-injected mouse retinal sections

Figure 5
Immunostaining for hepcidin and ferroportin in LPS-injected mouse retinal sections

Top panel: negative control with no primary antibody. Middle panel: staining with hepcidin antibody in retinal sections from control, PBS- and LPS-injected mouse eyes. Bottom panel: staining with ferroportin antibody in retinal sections from control, PBS- and LPS-injected mouse eyes.

Figure 5
Immunostaining for hepcidin and ferroportin in LPS-injected mouse retinal sections

Top panel: negative control with no primary antibody. Middle panel: staining with hepcidin antibody in retinal sections from control, PBS- and LPS-injected mouse eyes. Bottom panel: staining with ferroportin antibody in retinal sections from control, PBS- and LPS-injected mouse eyes.

Induction of hepcidin expression by LPS in retinal cells may result in the down-regulation of ferroportin, causing iron overload in these cells in vivo. To determine whether there were any consequences to the retina from such effects, we monitored the levels of oxidative stress in control and LPS-treated retinas by using an antibody capable of detecting HNE, a metabolic product of lipid peroxidation with hydroxyl radicals. Increased accumulation of free iron is expected to generate hydroxyl radicals via the Fenton reaction, which would then lead to enhanced lipid peroxidation and increased generation of HNE. Thus, HNE serves as a surrogate for detecting increased iron levels. These studies showed that the levels of HNE were significantly higher in LPS-treated retinas compared to normal retinas (Figure 6A). We then investigated whether these changes in oxidative stress lead to apoptosis in the retina. We used the TUNEL assay to detect apoptosis. There was considerably more apoptosis in retinal sections from eyes that were injected with LPS compared with retinal sections from control mice (Figure 6B). The increased TUNEL-positive signals were detectable in the outer nuclear layer and in the inner nuclear layer. PBS-injected contralateral eyes had more apoptosis in the outer nuclear layer compared to the mice that received PBS in both eyes. Interestingly, there was no evidence of increased apoptotic cell death in the RPE cell layer and in the ganglion cell layer. Ganglion cells do not express hepcidin and hence the absence of apoptosis in these cells following intravitreal injection of LPS is not surprising. To check if the RPE was resistant to LPS-induced oxidative stress in spite of the expression of hepcidin, we treated primary RPE cells with LPS for 24 h and checked for ferroportin levels, which were found to be down-regulated (Figure 6C). The down-regulation of ferroportin was associated with increased oxidative stress as monitored by immunofluorescence with anti-HNE antibody (results not shown).

Oxidative stress and apoptosis in LPS-injected mouse retina

Figure 6
Oxidative stress and apoptosis in LPS-injected mouse retina

(A) Immunostaining for HNE showing positive staining in the outer and inner nuclear layer of LPS-injected mouse retina compared to PBS-injected control retina. There was mild staining in the PBS-injected contralateral eye of treatment mice. (B) Number of apoptotic cells in each cell layer of retina in LPS-injected mouse retinal sections versus control retinal sections. (C) Immunostaining for ferroportin levels in control and LPS-treated (100 ng/ml) primary RPE cells.

Figure 6
Oxidative stress and apoptosis in LPS-injected mouse retina

(A) Immunostaining for HNE showing positive staining in the outer and inner nuclear layer of LPS-injected mouse retina compared to PBS-injected control retina. There was mild staining in the PBS-injected contralateral eye of treatment mice. (B) Number of apoptotic cells in each cell layer of retina in LPS-injected mouse retinal sections versus control retinal sections. (C) Immunostaining for ferroportin levels in control and LPS-treated (100 ng/ml) primary RPE cells.

DISCUSSION

Previous studies have shown that Hfe is expressed in the RPE basolateral membrane [21]. Since this membrane is in contact with systemic blood via choroidal circulation, the expression of Hfe in this membrane suggests that the protein might play a critical role in the sensing of systemic iron status. Hahn et al. [20] have shown that ferroportin is expressed abundantly in RPE, Müller cells and photoreceptor cells. Ferroportin is the target for hepcidin [7], the critical iron-regulatory hormone currently believed to be produced primarily by the liver [2932]. In RPE, ferroportin is expressed in the basolateral membrane; thus ferroportin in RPE may be subject to regulation by hepatic hepcidin present in the circulation. In contrast, ferroportin expressed in Müller cells and photoreceptor cells does not have access to circulating hepcidin. Regulation of ferroportin in these cells by hepatic hepcidin would require the transfer of hepcidin across the blood–retinal barriers. Alternatively, hepcidin may be produced within the retina to exert local control on ferroportin in retinal cells independent of hepatic hepcidin. There are no reports in the literature on the expression of hepcidin in the retina. In the present study, we found that hepcidin is expressed abundantly in Müller cells, photoreceptor cells and RPE cells. These are the same cell types which also express ferroportin [20]. This represents the first report on the expression of this important iron-regulatory hormone within the retina, suggesting that ferroportin expressed in cells within the retina may be subject to regulation by hepcidin produced locally within the tissue.

The expression of hepcidin in retina raises an important question. Do changes in the expression levels of hepcidin regulate ferroportin levels in retinal cells? Hepatic expression of hepcidin is induced by inflammation [28]. The expression of iron-regulatory proteins in different retinal cell types has not been studied in retinal infections. In the present study, we mimicked bacterial infection of the retina by intravitreal injection of LPS, and examined the changes in the expression of hepcidin within the retina. Our studies show that hepcidin expressed in retinal cells is subject to regulation by LPS with obligatory involvement of the TLR4. Hfe and Hjv regulate hepcidin synthesis in the liver [33,34], but are not involved in the inflammation-mediated regulation of hepcidin expression [28]. Our findings in the retina are similar to those in the liver. The increase in hepcidin expression in response to LPS in the retina is associated with a decrease in ferroportin levels in Müller cells, photoreceptor cells and RPE cells.

It is interesting that injection of LPS in one eye leads to changes in the expression of hepcidin and ferroportin in the contralateral eye. It is possible that LPS injected in one eye may have access to the contralateral eye via the circulation. Alternatively, proinflammatory mediators, released in response to LPS in one eye, may travel to the contralateral eye via the circulation and elicit their effects. It has been shown that LPS may produce its effects on hepcidin expression in the liver through IL-6 (interleukin 6) and/or TNFα (tumour necrosis factor α) in an autocrine manner [35]. Such intermediates may also be released into the circulation from the retina of the LPS-injected eye and subsequently elicit their effects on hepcidin expression in the contralateral eye. Exposure of the retina to LPS leads to increased oxidative stress as monitored by the levels of HNE. Several studies have shown that iron overload causes oxidative stress and that HNE levels increase under these conditions [3639]. The LPS-induced oxidative stress in vivo in the retina is accompanied by apoptosis in specific cell types. A significant increase in apoptotic nuclei is seen in the outer nuclear layer, which contains the nuclei of photoreceptor cells, and in the inner nuclear layer, which contains, among other cell types, the nuclei of Müller cells. Interestingly, there is no evidence of increased apoptosis in the RPE in response to LPS in vivo even though these cells express hepcidin and ferroportin. RPE is expected to respond to intravitreally injected LPS because TLR4 is present at the apical membrane of these cells [40]. Therefore the findings that there was no increase in apoptosis in this cell layer in response to LPS treatment in vivo are intriguing. It is possible that RPE may possess robust anti-oxidant mechanisms to fight against oxidative stress. Though we observed a significant increase in oxidative stress and apoptosis in specific cell types in the retina in response to LPS injection, obvious morphological changes in the retina were not readily apparent. This could be due to the fact that the retina was exposed to LPS only for 24 h. Exposure for longer periods may increase the severity of the phenotype. Iron deficiency and anaemia are observed in chronic, but not in acute, inflammation [41].

Since loss-of-function mutations in hepcidin cause a severe form of haemochromatosis, known as juvenile haemochromatosis, our findings on the expression of this protein within the retina are of clinical relevance. To our knowledge, there have been no reports on the iron status within the retina in patients with juvenile haemochromatosis. Since ferroportin, the molecular target of hepcidin, is expressed in several retinal cell types, it is expected that the levels of this iron transporter will be increased in patients with loss of hepcidin function. What this means in terms of iron homeostasis within the retina remains to be investigated. Studies by Hahn et al. [42] showed that a combined disruption of the function of ceruloplasmin and hephaestin in mice leads to iron overload in the retina, and that the retinal accumulation of iron causes a retinal phenotype similar to that of age-related macular degeneration. Since iron is a pro-oxidant, iron-induced oxidative stress may be an aetiological factor in age-related macular degeneration in humans. Chronic retinal inflammation in humans is expected to cause iron overload within the retina, but whether such effects have any relevance to the pathogenesis of age-related macular degeneration remains to be seen. It is interesting to note that genetic variations in genes involved in inflammation and oxidative stress, including TLR4 [43], have been implicated in the pathogenesis of age-related macular degeneration [4446].

The currently prevailing notion that hepcidin is produced exclusively in the liver as a hormone which has its biological effects distantly on other cells expressing ferroportin needs to be revisited in the light of more recent findings. There is evidence that hepcidin is made in non-hepatic tissues such as brain [16], heart [47] and lung [48]. Here we have shown that various cell types within the retina express this peptide. The emerging evidence that hepcidin and its molecular target ferroportin may be expressed in the same cell (e.g. RPE cell, Müller cell and photoreceptor cell) raises several interesting issues. It is currently believed that the plasma membrane ferroportin functions as the receptor for circulating hepcidin and that the binding of hepcidin to ferroportin leads to the internalization and degradation of the transporter [7]. If the same cell type expresses hepcidin and ferroportin, it is possible that hepcidin may regulate the levels of ferroportin in the cell, even prior to the recruitment of the transporter to the plasma membrane, by binding to the transporter at intracellular sites. The consequences of such binding inside the cell remain to be investigated. The binding of ferroportin to hepcidin leads to phosphorylation of specific tyrosine residues on the transporter, a process obligatory for internalization [49]. After internalization, the transporter is dephosphorylated and subsequently ubiquitinated to initiate degradation. It is not known at present whether the binding of hepcidin to the transporter at intracellular sites in retinal cells would lead to these covalent modifications of the transporter without involving other signalling components associated with the plasma membrane.

Since the ferroportin-expressing cell types within the retina also express hepcidin, ferroportin in these cells may be regulated by hepcidin generated within these cells independent of hepcidin in the circulation. This is true even for the RPE, in which ferroportin is expressed on the basolateral membrane with access to circulating hepcidin. Thus, hepatic hepcidin is not the sole determinant of ferroportin density in the plasma membrane of retinal cells. Pathologic conditions such as retinal infections and local inflammation may increase the production of hepcidin within the retina and reduce the ferroportin density in the plasma membrane in retinal cells without any observable changes in the circulating levels of hepcidin.

We thank Muthusamy Thangaraju and Santoshanand V. Thakkar for their help with the EGFP reporter assay, and Jennifer Duplantier for assistance with the intravitreous injection.

Abbreviations

     
  • CRALBP

    cellular ratinaldehyde binding protein

  •  
  • DAPI

    4′,6-diamidino-2-phenylindole

  •  
  • DMEM

    Dulbecco's modified Eagle's medium

  •  
  • EGFP

    enhanced green fluorescent protein

  •  
  • HBSS

    Hanks balanced salt solution

  •  
  • Hfe

    human leukocyte antigen-like protein involved in Fe homeostasis

  •  
  • HJV

    haemojuvelin

  •  
  • HNE

    4-hydroxynonenal

  •  
  • HPRT

    hypoxanthine phosphoribosyl transferase

  •  
  • LPS

    lipopolysaccharide

  •  
  • RPE

    retinal pigment epithelium

  •  
  • RT–PCR

    reverse transcription–PCR

  •  
  • Tfr

    transferrin receptor

  •  
  • TLR

    Toll-like receptor

  •  
  • TUNEL

    terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling

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