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The Journal of Immunology, 2000, 165: 4372-4378.
Copyright © 2000 by The American Association of Immunologists

CXCR4 Receptor Expression on Human Retinal Pigment Epithelial Cells from the Blood-Retina Barrier Leads to Chemokine Secretion and Migration in Response to Stromal Cell-Derived Factor 1{alpha}1

Isabel J. Crane2, Carol A. Wallace, Susan McKillop-Smith and John V. Forrester

Department of Ophthalmology, University of Aberdeen Medical School, Aberdeen, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Retinal pigment epithelial (RPE) cells form part of the blood-retina barrier and have recently been shown to produce various chemokines in response to proinflammatory cytokines. As the scope of chemokine action has been shown to extend beyond the regulation of leukocyte migration, we have investigated the expression of chemokine receptors on RPE cells to determine whether they could be a target for chemokine signaling. RT-PCR analysis indicated that the predominant receptor expressed on RPE cells was CXCR4. The level of CXCR4 mRNA expression, but not cell surface expression, increased on stimulation with IL-1ß or TNF-{alpha}. CXCR4 protein could be detected on the surface of 16% of the RPE cells using flow cytometry. Calcium mobilization in response to the CXCR4 ligand stromal cell-derived factor 1{alpha} (SDF-1{alpha}) indicated that the CXCR4 receptors were functional. Incubation with SDF-1{alpha} resulted in secretion of monocyte chemoattractant protein-1, IL-8, and growth-related oncogene {alpha}. RPE cells also migrated in response to SDF-1{alpha}. As SDF-1{alpha} expression by RPE cells was detected constitutively, we postulate that SDF-1–CXCR4 interactions may modulate the affects of chronic inflammation and subretinal neovascularization at the RPE site of the blood-retina barrier.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The retinal pigment epithelium forms part of the blood-retina barrier, with tight junctions, under normal circumstances, preventing the potentially sight-threatening influx of cells into the retina from the fenestrated choroid. The retinal pigment epithelium also has a major role in the physiological renewal of photoreceptor outer segments and in the provision of a transport and storage system for nutrients essential to the photoreceptor layer. The ability of the retinal pigment epithelial (RPE)3 cell to phagocytose photoreceptor outer segments gives it some properties in common with macrophages, including an extensive phagolysosomal system. Under some conditions, RPE cells may express proteins typical of macrophages and myeloid cells such as Fc receptors, CD68, and inducible NO synthase (1). RPE cells have been shown to produce a wide variety of cytokines in vitro particularly after stimulation with proinflammatory cytokines such as IL-1, TNF-{alpha}, and IFN-{gamma}, including chemokines such as RANTES, IL-8, and monocyte chemoattractant protein-1 (MCP-1) (2, 3, 4).

Chemokines are a family of small proteins mainly thought of as proinflammatory, as they are inducible in inflammatory conditions and act primarily as chemoattractants and activators of specific leukocytes at sites of inflammation (5). However, more recently some chemokines, such as stromal cell-derived factor (SDF) 1, have been shown to have fundamental roles in leukocyte trafficking of immature blood cells and naive lymphocytes, regulation of proliferation and mobilization of hematopoietic cells, regulation of angiogenesis, and fetal development (6).

Chemokine receptors are seven-transmembrane-spanning, G protein-coupled receptors (7), a range of which has been found in particular on lymphoid and myeloid cells and more recently on epithelial cells (8, 9, 10). Cell types such as macrophages have been shown to express receptors for chemokines in addition to producing chemokines, e.g., macrophage-inflammatory protein-1{alpha} (MIP-1{alpha}) (11, 12), potentially leading to autocrine stimulation. We were interested in determining whether this was the case for RPE cells in which increased chemokine production during inflammation is likely to enhance breakdown of the blood-retina barrier and destruction of the retina.

We investigated a range of chemokine receptors for expression by RPE cells and found that the receptor predominantly expressed by RPE cells is CXCR4 (fusin/LESTR). CXCR4 is a specific receptor that, so far, is only known to bind one chemokine ligand, the C-X-C chemokine, SDF-1 (13). SDF-1 occurs in two alternative splicing variants, SDF-1{alpha} and SDF-1ß, of which SDF-1{alpha} is more abundant (14). Although many chemokine systems recruit cells to sites of inflammation, the SDF-1 and CXCR4 pair is thought to have functions atypical of chemokines. They are thought to be important in the basal trafficking of lymphocytes (15, 16) with CXCR4 expressed strongly on CD45RA+ naive T lymphocytes and SDF-1 constitutively expressed in a wide range of tissues (17). Other functions include roles in B cell lymphopoiesis, bone marrow myelopoiesis, cardiac ventricular septum formation (18), architecture of the cerebellum (19) and organ vascularization (20). Both SDF-1 and CXCR4 have a wide tissue distribution, with CXCR4 found at higher levels than other chemokine receptors in both intestinal and alveolar epithelial cells (8, 9, 10) and endothelial cells (21, 22). In the brain, CXCR4 receptors have been localized to the endothelia at the blood-brain barrier, as well as to microglia, neurons, and astrocytes (23, 24). Interestingly, in intestinal epithelial cells, stimulation by SDF-1{alpha} has been reported to up-regulate production of other chemokines, IL-8, and growth-related oncogene (GRO) {alpha} (8).

The functionality and significance of CXCR4 expression on the RPE cells of the blood-retina barrier were examined.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell isolation and culture

RPE cells were isolated and cultured from eyes obtained from six different donors as previously described (25) and maintained at 37°C and 5%CO2/95% air. Two donors were male and four female, with an average age of 27 years (range, 3–44 years). They had died from various causes unrelated to eye disease. The cultures consisted of monolayers of polygonal cells (26) that were shown to be purely epithelial in nature by 100% positive staining for vimentin (Dako, High Wycombe, U.K.) and cytokeratins 8 and 18 (CAM 5.2; Becton Dickinson, San Jose, CA), which are associated with replicating RPE cells (27). THP-1, a monocytic leukemia-derived cell line, from European Collection of Cell Cultures (Centre for Applied Microbiology and Research, Salisbury, U.K.) was cultured in RPMI 1640 supplemented with 0.05 mM 2-ME, 4 mM L-glutamine, and 10% heat-inactivated FBS at 37°C and 5% CO2/95% air. The concentration of THP-1 cells was maintained between 2 and 9 x 105 cells/ml.

Treatment of cells with cytokines

For analysis of chemokine production and flow cytometry, healthy, dividing cells from passages 4 to 7 were seeded at 2 x 105 cells/ml into 25-cm2 flasks (Nunc, Naperville, IL) and cultured to 95% confluence in complete medium (Glasgow MEM supplemented with 0.225% (v/v) sodium bicarbonate, 10 mM HEPES, 10% (v/v) tryptose phosphate broth, 1 mM sodium pyruvate, MEM nonessential amino acids at single strength, 4 mM L-glutamine, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 10% FBS). Medium was aspirated and the cultures were washed gently three times with HBSS containing calcium and magnesium. The medium was replaced with serum-free complete medium for 16 h before cytokines were added and culture continued for up to 24 h. Control cultures were treated in the same way, but no cytokines were added. Conditioned medium was collected, centrifuged at 300 x g for 10 min, and stored at –80°C until assay. Protein content of the cultures was estimated using a protein-Coomassie brilliant blue binding method (28) to check that cell growth had been comparable in each of the flasks. The cytokines over this time scale did not affect RPE proliferation.

RPE cells for RNA extraction were seeded at passages 4–7 into 75-cm2 flasks in complete medium and cultured until 80% confluent. The medium was changed to serum-free medium as above, culture continued overnight, and then cytokines were added. Cells were harvested 6 h after cytokine addition as described below.

RNA extraction

RNA was extracted using a modification of the method of Chomczynski and Sacchi (29, 30). Briefly, medium was removed at the end of the incubation period, and cultures were washed three times with PBS minus calcium and magnesium and resuspended in 4 M guanidinium isothiocyanate, 25 mM sodium citrate (pH 7.0), 0.5% sarkosyl, and 0.1 M 2-ME. The RNA was extracted using phenol-chloroform-isoamyl alcohol, and the aqueous layer was retained. The RNA was precipitated twice with isopropanol, washed with 75% ethanol, and resuspended in distilled water. The concentration of the RNA was determined spectrophotometrically at 260 nm. PBS, ethanol, and water were treated with diethyl pyrocarbonate.

RT-PCR

Poly(A)+ RNA from 5 µg total RNA was reverse transcribed with 200 U RNase H- Moloney murine leukemia virus reverse transcriptase (Superscript II; Life Technologies, Paisley, U.K.). Of this cDNA, 2 µl was used in the PCR. RNA samples that had not been reverse transcribed were included in parallel PCRs to control for genomic DNA, as primers were not intron spanning. Each PCR was conducted in a total volume of 25 µl containing 0.8 mM each of dATP, dCTP, dGTP, and dTTP; 2.5 µl Taq buffer; 0.25 µl Taq polymerase (Promega U.K., Southampton, U.K.); and 2.5 µl primer mix. All solutions, except those containing nucleic acid, and pipettes and tips were treated for 5 min before use in an XL-1000 spectrolinker (Spectronic Instruments, Rochester, NY). ß-Actin and SDF-1 primers were obtained from Oswel (Southampton, U.K.), and chemokine receptor primers (9) were obtained from MWG-Biotech (Milton Keynes, U.K). ß-Actin primers were 5'-GTCCTTAATGTCACGCACGATTTC-3' and 5'-GTGGGGCGCCCCAGGCACCA-3', and SDF-1 primers were 5'-GTCAGCCTGAGCTACAGATGC-3' and 5'-CACTTTAGCTTCGGGTCAATG-3'.

Thirty-three cycles of amplification were performed (PCR Express; Hybaid, Middlesex, U.K.), with each cycle consisting of a denaturation step at 94°C for 50 s, annealing at 55°C for 1 min, and polymerization at 72°C for 1 min 30 s. In the first cycle denaturation was conducted for 2 min, and in the final cycle polymerization was conducted for 5 min. The number of cycles in the amplification was optimized by serial sampling with increasing cycles of amplification to ensure that the PCR was in the geometric phase for all samples, enabling valid comparisons to be made between samples in the same experiment. After amplification, samples were run on a 1.8% agarose gel (molecular biology grade; Sigma, Poole, U.K.) in TBE (0.045 M Tris-borate, 0.001 M EDTA) containing 0.4 µg/ml ethidium bromide, and bands were visualized and photographed on a UV transilluminator (Syngene, Cambridge, U.K.). An MspI digest of pBR322 DNA (New England Biolabs, Hitchin, U.K.) was run in parallel and used to calculate the PCR product sizes using GeneGenius software (Syngene). Relative abundance of product was assessed by calculating the ratios of the chemokine or chemokine receptor band to the ß-actin band for each sample using the GeneGenius software (Syngene). For sequencing, 5 µl of amplification mix was reamplified as before and sequenced (Applied Biosystems, Foster City, CA).

Flow cytometry

Cells from three RPE lines, passages 4–7, cultured as above, were harvested after treatment with 20 mM EDTA for 30 min at 37°C, washed in FACS buffer (PBS containing 1% BSA and 10 mM sodium azide), and resuspended at 5 x 105/ml in 100 µl FACS buffer containing 10 µl primary Ab at 5 µg/ml. Primary mAbs were biotinylated anti-human CXCR4, clones 12G5 and 44716.111 (R&D Systems Europe, Oxford, U.K.). Isotype control Ab was biotinylated mouse IgG2a, anti-trinitrophenol (BD Biosciences, Oxford, UK). Cells were incubated with primary Ab for 30 min at 4°C, washed twice in FACS buffer, and resuspended in 50 µl streptavidin R-phycoerythrin (Caltag Laboratories, Burlingame, CA) for 30 min at 4°C before washing twice further and resuspending in 500 µl of FACS buffer for analysis in a FACScalibur (BD Biosciences). The positive control Ab was mouse anti-human HLA-A, B, C directly conjugated to R-phycoerythrin (Dako). Live cells were gated. The three cell lines gave similar results on three separate occasions, so these results were pooled.

Immunocytochemistry

RPE cells were also cultured as described above but on glass coverslips. Cells were fixed in acetone for 10 min, air dried, and incubated with anti-human CXCR4 (clone 12G5) or control mouse IgG2a as above for 60 min at room temperature. Coverslips were washed three times in TBS before incubation for 60 min with biotinylated rabbit anti-mouse Ab (1:100, Dako) that had been preabsorbed with 10% normal human serum for 30 min. After washing, sections were incubated with streptavidin-biotin complex/alkaline phosphatase (Dako) for 30 min according to kit protocol before addition of Fast Red substrate solution.

Calcium mobilization

Calcium mobilization in response to chemokine stimulation was measured in individual cells using a Ca2+ imaging system with a slow-scan charge-coupled device camera and SpectroMaster monochromatic illuminator attached to an upright Olympus microscope. Images were acquired and analyzed using the MERLIN ratio imaging system. The complete Ca2+ imaging system was supplied by Perkin Elmer Life Sciences (Cambridge, U.K.). RPE cells at passages 4 and 5 were split 24 h before the experiment and were no >80% confluent at the time of the experiment. Cultures in complete medium were loaded with 5 µM fura-2-acetoxymethyl ester (Molecular Probes Europe, Leiden, The Netherlands) at 37°C for 1 h in the dark before washing twice with HBSS + 1% FCS. Cultures were retained in HBSS + 1% FBS for the experiment. Cells were focused and recording commenced. SDF-1{alpha} and IL-8 (R&D Systems) were added as a 30-µl drop close to the water-immersion objective (31) to give a final concentration of 100 nM. The fluorescence ratio was determined from background-corrected fluorescent images using dual excitation at 340 and 370 nm. The system was calibrated for free ion concentrations by imaging the cells under permeabilized conditions using calcium calibration kit 1 (Molecular Probes).

ELISA

Immunoreactive chemokine, MCP-1, RANTES, GRO{alpha}, IL-8, and SDF-1{alpha} produced by RPE cells was quantified by sandwich ELISA (R&D Systems) according to the manufacturer’s protocol. The minimum amount of chemokine detectable was <5 pg/ml. Cell-free supernatants were diluted as appropriate for the chemokine and the sensitivity of the ELISA and added in duplicate. Concentrations of chemokine detected in the supernatant were adjusted for the protein content of the cultures.

Migration assays

The functional ability of CXCR4 receptors on RPE cells was also tested in an assay of migration in response to SDF-1{alpha}. RPE cells cultured as above were harvested and incubated at 1 x 106/ml in 10 ml PBS containing 50 nM calcein acetoxymethyl ester (Molecular Probes) for 45 min with end-over-end mixing at 37°C. Cells were then centrifuged, counted, and resuspended at 5 x 105/ml in PBS plus 0.05% BSA. Cells were incubated with or without 10 µg/ml anti-human CXCR4 Ab, clone 12G5 (R&D Systems) for 15 min at 37°C. PBS (356 µl) plus 0.05% BSA with or without SDF-1{alpha} (R&D Systems) was added to the wells of a black microtiter plate (Polyfiltronics, Middlesex, U.K.) inserted in a microtiter plate chemotaxis chamber (NeuroProbe, Cabin John, MD). An adhesive framed 5-µm polycarbonate filter with polyvinylpyrrolidone (NeuroProbe) was placed over the microtiter plate and the top of the chamber latched to the bottom. Cell suspension (50 µl) was then added to the upper wells, and the chamber was incubated at 37°C for 2 h in a humidifier. The cell suspension was removed from the upper wells. To ensure complete removal of any cells that had not migrated into the filter, 200 µl of PBS plus EDTA (20 mM) was added into the upper wells and the chamber was incubated for 30 min at 4°C. The EDTA solution was decanted, the plate plus filter removed from the chamber, and the top rinsed three times in distilled water. The plate plus filter was then read in a Fluorolite 1000 (Dynex Technologies, Middlesex, U.K.), excitation 485 nm and emission 535 nm. The top of the filter was checked separately and shown to have negligible residual fluorescence. Standard curves were prepared with known cell numbers so that the percentage of cells migrating could be calculated.

Statistical analysis

Assays were repeated a minimum of three times using cultures from at least three different donors. Data are presented as the mean ± SEM. The statistical significance of the results was assessed using Student’s unpaired two-tailed t test for comparison of two groups or ANOVA with Newman-Keuls post test when three or more groups were compared (GraphPad, San Diego, CA).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Chemokine receptor and SDF-1 expression on RPE cells

mRNA level. CXCR4 mRNA expression was detected consistently in different RPE cell lines (Fig. 1Go) but not in samples of RNA that had not been reverse transcribed, confirming that CXCR4 detection was not a result of genomic DNA contamination. Predicted PCR product sequences were confirmed by sequencing. Low levels of CCR1, CCR2, and CCR3 could also be detected in some lines after stimulation of RPE cells with IL-1ß and/or TNF-{alpha} (Fig. 1Go) but CCR4, CCR5, CXCR-1, CXCR-2, and CXCR-3 were not detected (data not shown). CXCR4 expression was constitutive and was increased significantly (p = 0.034) by incubating the RPE cells with IL-1ß (Fig. 1Go). The ratio of CXCR4 to ß-actin was increased from 0.191 ± 0.0045 to 0.644 ± 0.028 with IL-1ß and to 0.439 ± 0.056 with TNF-{alpha}.



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  		FIGURE 1. Expression of mRNA for ß-actin and chemokine receptors as detected by PCR. Lane 1, unstimulated THP-1 cells; lanes 2–7, RPE cell line 25B: lane 2, unstimulated; lane 3, IL-1ß; lane 4, IL-1ß and IFN-{gamma}; lane 5, TNF-{alpha}; lane 6, IL-1ß and TNF-{alpha}; lane 7, TNF-{alpha} and IFN-{gamma}; and lane 8, negative control (no cDNA). Concentrations: IL-1ß, 0.25 ng/ml; TNF-{alpha}, 10 ng/ml; IFN-{gamma}, 20 ng/ml. Results shown are representative of three experiments using RPE from different donors.

 
SDF-1 mRNA expression was detected constitutively in RPE cells and was down-regulated significantly (p = 0.034) by IL-1ß (Fig. 2Go). The ratio of SDF-1 to ß-actin was decreased from 0.331 ± 0.064 to 0.070 ± 0.044 with IL-1ß and to 0.163 ± 0.032 with TNF-{alpha}.



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  		FIGURE 2. Expression of mRNA for ß-actin and SDF-1 as detected by PCR. Lanes 1–6, RPE cell line 15A. Lane 1, unstimulated; lane 2, IL-1ß; lane 3, TNF-{alpha}; lane 4, IL-1ß and TNF-{alpha}; lane 5, IL-1ß and IFN-{gamma}; and lane 6, TNF-{alpha} and IFN-{gamma}. Concentrations: IL-1ß, 0.25 ng/ml; TNF-{alpha}, 10 ng/ml; IFN-{gamma}, 20 ng/ml. Results shown are representative of three experiments using RPE from different donors.

 
Protein level. The positive control Ab, anti-human HLA-A, B, C, identified 96.92 ± 0.733% of the gated cells. Both anti-human CXCR4 mAbs detected significant cell surface CXCR4 expression on the RPE lines when compared with the negative control Ab (clone 12G5, p < 0.0001, and clone 44716, p < 0.0167). However, clone 12G5 detected more cells positive for CXCR4 (percent cells gated, 16.28 ± 2.691, no cytokine stimulation) than 44716 (7.36 ± 1.877) (Fig. 3Go). There was no significant difference (p > 0.28) in receptor expression between populations of cells either unstimulated or stimulated with IL-1ß or TNF-{alpha} (Fig. 3GoB).



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  		FIGURE 3. Surface expression of CXCR4 on RPE cells. A, Flow cytometry histogram for RPE cell line 25B incubated with isotype control Ab, mouse IgG2a, or an equivalent concentration of anti-human CXCR4, clone 12G5. M1 is the population that is positive compared with the isotype control. Results shown are representative of three experiments using RPE from three different donors in each experiment. B, Mean percentage of gated RPE cells positive ± SEM (n = 9) in unstimulated cultures ({square}) and in cultures stimulated for 24 h with 0.25 ng/ml IL-1ß () or with 10 ng/ml TNF-{alpha} (). Anti-human CXCR4 Abs were clones 12G5 and 44716.111.

 
Using immunocytochemistry, CXCR4 expression by RPE cells could be detected mainly as intracellular perinuclear staining (Fig. 4Go).



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  		FIGURE 4. CXCR4 detected by immunocytochemistry on human RPE cells. A, Cells stained with control IgG2a. B, Cells stained with anti-human CXCR4. Magnification, x125.

 
SDF-1{alpha} production by the RPE cells could be detected constitutively by ELISA but only at very low levels (1.33 ± 1.0; range, 3.34–0.27 pg/ml).

RPE cell response to SDF-1{alpha}

Calcium mobilization. Calcium imaging of individual cells showed a rapid increase in intracellular calcium in cells responding to 100 nM SDF-1{alpha} (Fig. 5Go). This could not be achieved by treating the cells in the same way with 100 nM IL-8 (Fig. 5Go) and indicated that the CXCR4 detected on the RPE cell surface was functional.



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  		FIGURE 5. Ca2+ flux generated in RPE cell cultures in response to SDF-1{alpha}. The fura-2-acetoxymethyl ester-loaded cells were stimulated with 100 nM SDF-1{alpha} (upper panel) or 100 nM IL-8 (lower panel). Arrows indicate the points at which the chemokine was added to the culture. Seconds are measured from the initiation of the recording. Traces are from individual cells in the same field of view. Results shown are representative of three experiments.

 
Chemokine production. MCP-1, IL-8, and GRO{alpha} were produced by the RPE cells in response to SDF-1{alpha} and showed the same pattern of response. A low concentration of SDF-1{alpha} (0.1 ng/ml) resulted in significant (p < 0.05) production of all three chemokines by 24 h (Fig. 6Go). Incubation with 1 and 10 ng/ml SDF-1{alpha} did not significantly increase chemokine production except for GRO{alpha} (p < 0.05), whereas with 20 ng/ml SDF-1{alpha} production of all three chemokines was significant (p < 0.05) (Fig. 6Go). No significant production of RANTES could be detected in response to these SDF-1{alpha} concentrations (data not shown). Incubation of RPE cells with SDF-1{alpha}, at either 0.1 ng/ml or 10 ng/ml, and either IL-1ß or TNF-{alpha} did not significantly (p > 0.05) affect MCP-1, IL-8, or GRO{alpha} production compared with IL-1ß or TNF-{alpha} alone (Table IGo). Production of all three chemokines was much greater after incubation with IL-1ß or TNF-{alpha} alone than with SDF-1{alpha}.



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  		FIGURE 6. MCP-1, GRO{alpha}, and IL-8 in the culture supernatant of RPE cells as determined by ELISA. RPE cultures were stimulated with increasing concentrations of SDF-1{alpha} for 24 h. Results shown are the mean of five experiments using RPE from four different donors. Error bars indicate SEM.

 

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  		Table I. MCP-1, GRO{alpha}, and IL-8 in supernatants of RPE cell cultures incubated for 24 h with combinations of IL-1ß, TNF-{alpha}, and SDF-1{alpha}1

 
Migration. RPE cells demonstrated a significant migratory response to SDF-1{alpha} with concentrations of 1, 10, and 100 ng/ml SDF-1{alpha} resulting in a significant (p < 0.05) increase in the chemotactic index (Fig. 7Go). The incubation of RPE cells with anti-CXCR4 before assay of migration in response to SDF-1{alpha} blocked the migratory response with no significant increase in the chemotactic index (p = 0.35) in this situation.



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  		FIGURE 7. Migration of RPE cells in response to SDF-1{alpha}. RPE cells were calcein labeled, and migration was measured across a 5-µm polyvinylpyrrolidone-coated polycarbonate filter over 2 h in response to increasing concentrations of SDF-1{alpha}. Labeled cells were also incubated with anti-human CXCR4 (clone 12G5) at 10 µg/ml for 15 min before the migration assay. Results shown are the mean of five experiments using RPE from four different donors. Error bars indicate SEM.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This study demonstrates for the first time CXCR4 and SDF-1 mRNA expression in human RPE cells. In addition, there is expression of CXCR4 on the cell surface of the RPE cells, which is functional as shown by signaling mobilization of intracellular calcium; production of the chemokines MCP-1, IL-8, and GRO{alpha}; and migration of the RPE cells in response to SDF-1{alpha} .

Of the chemokine receptors tested for by RT-PCR, mRNA for CXCR4 was expressed most strongly and constitutively, with only low levels of CCR1, CCR2, and CCR3 detected in response to stimulation by the proinflammatory cytokines IL-1ß and TNF-{alpha}. Only 16% of the cultured human RPE cells, as determined by flow cytometry, expressed cell surface CXCR4 receptors, and this was supported by the data from the Ca2+ mobilization experiments in which only 20% of cells responded (Figs. 3GoB and 5). Analysis of primary cultures of fetal macaque neurons (32), also using the anti-CXCR4 Ab clone 12G5, showed 28% positive for CXCR4, consistent with the heterogeneity in neuronal expression observed in adult macaque brain in vivo. Jordan et al. reported that just under 50% of the human colon adenocarcinoma cell line HT-29 expressed cell surface CXCR4 and that this percentage did not change whether the cells were growing or quiescent (9). It has been reported that freshly isolated human monocytes rapidly lose the ability to respond to SDF-1{alpha} by calcium mobilization but that this ability is regained concomitantly with CXCR4 mRNA expression as they differentiate (33).

Stimulation of the RPE cells with IL-1ß or TNF-{alpha}, which increases CXCR4 mRNA expression, had no effect on the percentage of cells expressing CXCR4 or on the fluorescence intensity of those cells expressing CXCR4. IL-1ß and TNF-{alpha} have been reported to have no effect on CXCR4 expression on T cells (34), but exposure of HUVEC to these cytokines resulted in a decrease followed by an increase in CXCR4 mRNA levels (22). Modulation of CXCR4 expression by other factors has also been described, e.g., up-regulation in human T cells by IL-4 and dexamethasone (34, 35), in cultured Langerhans cells by IL-4 and TGF-ß (36), and in endothelial cells by basic fibroblast growth factor or vascular endothelial growth factor (37). Down-regulation of CXCR4 expression by IFN-{gamma}, reported in human endothelial cells (22), is also suggested in RPE cells by our studies, but as yet we have not determined this unequivocally. As SDF-1{alpha} expression has been reported to be constitutive in many tissues, CXCR4 regulation is likely to be important and may be via intracellular stores of CXCR4 (38, 39). Intracellular CXCR4 can be seen in the RPE cells and has also been observed in the promyelocytic cell line HL-60 (33) and in intestinal epithelial cells (8).

The ability of RPE cells to migrate in response to SDF-1 indicates the functionality of the RPE cell CXCR4 receptors. It has been shown with germinal center B cells that it is possible for cells to express cell surface CXCR4 but not to migrate to SDF-1, and, in this case, CXCR4 appears to be less coupled to the downstream signaling pathways (40). This degree of RPE cell migration, a mean chemotactic index (ratio of specific to background migration) of 1.1:2.0, with 2–4% of input cells migrating, is characteristic for chemokines such as MCP-1, MIP-1{alpha}, and RANTES and for lymphocytes in filter chemotaxis assays (17). There are situations in vivo in which RPE cells may migrate, e.g., in development (41), wound healing, or pathological situations such as proliferative vitreoretinopathy (42) and age-related macular degeneration (43). However, RPE cell migration in response to SDF-1{alpha} may not be relevant in vivo if there is constitutive SDF-1 production by RPE cells, as suggested by RT-PCR and ELISA, as it is unlikely that a gradient of SDF-1 could be established. In addition, wound healing is reported to be inhibited by injection of SDF-1 (E. R. Fedyk et al., unpublished observations).

As further evidence of the functionality of the CXCR4 receptors in RPE cells, these cells were shown to secrete low, but biologically significant, levels of the chemokines MCP-1, IL-8, and GRO{alpha} in response to SDF-1{alpha}. It has been shown that signaling via CXCR4 can activate NF-{kappa}B (44), which is an important transcription factor in the regulation of MCP-1, IL-8, GRO{alpha}, and RANTES (45, 46, 47, 48). Lack of production of RANTES in response to SDF-1{alpha} may be due to differences in RANTES regulation via NF-{kappa}B involving I-{kappa}B-related protein (48).

Production of the chemokines in response to such low levels of SDF-1{alpha} is of interest. In vitro SDF-1 is a highly efficient chemoattractant, attracting 10-fold more lymphocytes than other chemokines such as MCP-1, MIP-1{alpha}, and RANTES (17) but only when compared at higher concentrations (1 µg/ml). However, in vivo SDF-1 is more potent possibly because of its ability to bind to heparin with a higher affinity than MCP-1 or IL-8, which is likely to improve presentation of SDF-1 or enable it to be retained more efficiently at sites of production (17). Thus, in vivo, concentrations of SDF-1 as low as 0.1 ng/ml are likely to have a significant role that may be distinct from that in situations when higher concentrations of SDF-1 are present. In human intestinal epithelial cells, production of GRO{alpha} and IL-8 was also evident in response to SDF-1{alpha} (8), although concentrations as low as 0.1 ng/ml SDF-1{alpha} were not examined. No significant production of the chemokines was detected until 100 ng/ml of SDF-1{alpha} was used and chemokine production was still continuing to rise at 1 µg/ml SDF-1{alpha}. As IL-1ß and TNF-{alpha} stimulate higher levels of MCP-1, IL-8, and GRO{alpha} production than SDF-1, production in response to SDF-1 may only be relevant in situations in which very little or no IL-1ß and TNF-{alpha} are present such as very early in an inflammatory response or during low-grade chronic inflammatory disease, which can be a feature of human autoimmune uveitis. In inflammatory situations in which there are substantial amounts of IL-1 and TNF-{alpha} present, SDF-1 may be down-regulated, possibly because it is unnecessary or has inhibitory functions.

IL-8 and GRO{alpha} production in response to SDF-1 may be of more significance than MCP-1, as MCP-1 production by RPE cells, although stimulated by SDF-1, is also constitutive. IL-8 and GRO{alpha} are both ELR motif-containing CXC chemokines and are strongly angiogenic (49). New vessel induction can be stimulated directly via SDF-1 (22, 37). In RPE-associated neovascular disorders such as autoimmune uveitis, in which neovascularization can play a prominent part (50); age-related macular degeneration (51); or diabetic retinopathy, SDF-1 may be secreted by the RPE cells constitutively or be available from other sources, and it is possible that it may influence the course of neovascularization either directly or indirectly via IL-8 and GRO{alpha}. The link between SDF-1 and neovascularization in situations such as diabetic retinopathy (52) has also been suggested by studies in which stimulation of HUVECs with vascular endothelial growth factor or basic fibroblast growth factor up-regulated levels of CXCR4 mRNA on these cells (37). Although SDF-1, if produced constitutively, may also be present in normal situations at low levels, its presence in these neovascular disorders in the eye may, in combination with other factors, facilitate disease progression.

Finally, the significance of CXCR4 as a coreceptor for T cell-tropic HIV-1 (53, 54) cannot be overlooked. Although normally a second cofactor CD4 is needed for virus entry into cells, there are reports that in non-CD4-expressing cells other components such as the glycolipid galactosylceramide (55, 56) may act as a cofactor. It is reported for HIV-2 that CXCR4 alone may be sufficient (57). This suggests that RPE cells could possibly be a cellular target for HIV-1 and involved in AIDS-related retinopathy and related complications.


   Acknowledgments
 
We thank The Netherlands Ophthalmic Research Institute, Amsterdam, for their valued help in providing material for this study.


   Footnotes
 
1 This work was made possible by Grant 049531 from The Wellcome Trust. I.J.C. is a Wellcome Trust Research Fellow. Back

2 Address correspondence and reprint requests to Dr. Isabel J. Crane, Department of Ophthalmology, University of Aberdeen Medical School, Foresterhill, Aberdeen, AB25 2ZD, U.K. Back

3 Abbreviations used in this paper: RPE, retinal pigment epithelial; MCP-1, monocyte chemoattractant protein-1; SDF-1, stromal cell-derived factor 1; MIP, macrophage-inflammatory protein; GRO, growth-related oncogene. Back

Received for publication January 27, 2000. Accepted for publication July 19, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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