Sources and sink of retinoic acid in the embryonic chick retina: distribution of aldehyde dehydrogenase activities, CRABP-I, and sites of retinoic acid inactivation

Developmental Brain Research 127 (2001) 135–148 www.elsevier.com / locate / bres

Research report

Sources and sink of retinoic acid in the embryonic chick retina: distribution of aldehyde dehydrogenase activities, CRABP-I, and sites of retinoic acid inactivation ¨ Mey a , *, Peter McCaffery b,c , Monika Klemeit a Jorg a

¨ Biologie II, Zoologie /Tierphysiologie, RWTH Aachen, Kopernikusstraße 14, 52074 Aachen, Germany Institut f ur b Shriver Center for Mental Retardation, Waltham, MA, USA c Department of Psychiatry, Harvard Medical School, Boston, MA, USA Accepted 6 February 2001

Abstract Previous experiments in mice and zebrafish led to the hypothesis that an asymmetric distribution of the transcriptional activator retinoic acid (RA) causes ventral–dorsal polarity in the vertebrate eye anlage. A high concentration of RA in the ventral retinal neuroepithelium has been suggested to induce developmental events that finally establish topographic order in the retinotectal projection along the vertical eye axis. In the present study we have investigated potential sources and sinks of RA during embryonic development of the chick retina. At embryonic day (E)1 to E2, when the spatial determination of the eye primordia takes place, no RA synthesis by aldehyde dehydrogenases was detectable, and neither immunoreactivity for retinaldehyde dehydrogenase RALDH-2 nor for cellular retinoic acid binding protein CRABP-I was observed. These components of RA signal transduction appeared in the eye between E3 and E5. At later stages, RA-measurements with a reporter cell line showed highest synthesis in the retinal pigment epithelium (RPE) and at the ventral and dorsal poles of the retina. RA degradation occurred mostly in a horizontal region in the middle of the retina with only small differences along the nasal–temporal axis. CRABP-I immunoreactivity appeared first in differentiating retinal ganglion cells with no indication of a spatial gradient across the ventral–dorsal eye axis. RA-production depended on three NAD 1 -dependent enzyme activities, which could be competitively inhibited by citral. One enzyme, located in the dorsal retina (corresponding to mouse RALDH-1), and one enzyme in the RPE (RALDH-2) were aldehyde dehydrogenases of the same molecular weight (monomers about 55 kDa) but with different isoelectric points (6.5–6.9; 4.9–5.4). The third RA-synthesizing activity (pI 6.0–6.3) was limited to the ventral retina, and likely corresponded to mouse RALDH-3. The restricted localization of retinoid-metabolizing activities along the dorsal–ventral axis of the embryonic chick retina does support the idea that RA is involved in dorsal–ventral eye patterning. However, the late time of appearance of aldehyde dehydrogenase activities and CRABP-I points to functions in cellular differentiation that are distinct from the initiation of the dorsal–ventral polarity.  2001 Elsevier Science B.V. All rights reserved. Theme: Development and regeneration Topic: Pattern formation, compartments, and boundaries Keywords: Chick development; Retinoic acid; Retina; Aldehyde dehydrogenase; CRABP; Axial polarity

Abbreviations: C-D, chick-dorsal, aldehyde dehydrogenase activity present in the dorsal segment of the embryonic chick retina, probably homologous to mouse RALDH-1; C-RPE, chick-retinal pigment epithelium, aldehyde dehydrogenase activity present in the chick RPE, probably homologous to mouse RALDH-2; C-V, chick-ventral, aldehyde dehydrogenase activity present in the ventral segment of the embryonic chick retina, probably homologous to mouse RALDH-3; COUP-TF, chick ovalbumin upstream promoter-transcription factor; CRABP, cellular retinoic acid binding protein; CYP, cytochrome P450-linked oxidase; DMEM, Dulbecco’s modified Eagle medium; E followed by number, embryonic day; ECL, enhanced chemoluminescence; FCS, fetal calf serum; HBSS, Hank’s buffered saline solution; HH followed by number, stage of chick embryonic development according to Hamburger and Hamilton, 1951; HPLC, high pressure liquid chromatography; IEF, isoelectric focusing; PFA, paraformaldehyde; RA, retinoic acid; RALDH, retinaldehyde dehydrogenase; RAR, retinoic acid receptor; RPE, retinal pigment epithelium; RXR, retinoid X receptor; SDS–PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis *Corresponding author. Tel.: 149-241-804-852; fax: 149-241-888-8133. E-mail address: [email protected] (J. Mey). 0165-3806 / 01 / $ – see front matter  2001 Elsevier Science B.V. All rights reserved. PII: S0165-3806( 01 )00127-4

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1. Introduction The establishment of axial polarity is a critical process of early embryonic development. In the retina, molecular differences along dorsal–ventral and nasal–temporal axes arise concomitant with the growth of the eye anlage [16,47,53]. The graded expression of molecular determinants along the eye axes is believed to endow retinal neurons with spatial identity in a topographic coordinate system that subsequently is the basis for topographic development of central axonal projections [33]. While several transcription factors have been found to exert a critical influence on the determination of retinal polarity, the transcriptional activator retinoic acid (RA) is thought to play an important role by giving rise to the original dorsal–ventral polarity. This hypothesis assumes a source of RA production in the proximity of or within the ventral part of the eye anlage. The source could be the spatially restricted activity of an aldehyde dehydrogenase [20,27,32]. RA is a lipophilic molecule which easily diffuses through cellular membranes. If it is gradually metabolized or inactivated by binding proteins while spreading in the tissue, a decreasing gradient will be established with distance from the source of production. The asymmetry created by diffusing RA could, during a sensitive period, induce the expression of spatially restricted transcription factors that have different thresholds for RA-activation. It has been suggested that in the critical period axial differences in RA synthesis become fixed through the permanent expression of different RA-generating enzymes within the retina [2]. Though several lines of indirect evidence support this concept in zebrafish and mice [11,12,24], no functional experiments have been performed to confirm the role of RA in the avian retina. For the graded distribution of RA itself, quantification with HPLC and indirect measurements with a bioassay showed a higher ventral concentration of all-trans RA in the retina of E13 mouse embryos, E4 chick embryos and 15-somite zebrafish larvae [4,24,32]. Due to the relatively large amounts of tissue needed for biochemical analysis of RA, HPLC data provided only a global estimate of RA concentrations along the vertical eye axis [32]. The biological availability of RA in the tissue depends not only on the synthesis of RA but also on its metabolic inactivation. So far, one cytochrome P450-linked oxidase, named CYP26, is known to catalyze the oxidation of RA [50,51]. In mouse embryos, CYP26 mRNA is first restricted to the dorsal half of the eye anlage, then recedes from the dorsal pole and, during invagination of the secondary eye vesicle, becomes localized in a horizontal stripe in the middle of the retina [20,31]. Although CYP26 expression has been well characterized in the ectoderm of gastrula and neurula stage chick embryos and in limb development [45], nothing is known about RA degradation in the chick retina. In addition to the nuclear receptors that transmit RA

signals to the DNA, two cellular retinoic acid binding proteins (CRABP-I, -II) are known to exist. Only one, CRABP-I, is expressed in the chick nervous system [15,41]. Several hypotheses about their functions have been raised, including the transfer of RA to nuclear receptors and differential accumulation of RA under conditions of vitamin A scarcity. Alternatively, CRABP-I may be part of retinoid catabolism because oxidation of exogenous RA is faster in the presence of high CRABP levels, suggesting that it protects RA-sensitive cells from overdose or takes part in the formation of morphogenetic gradients [36,41]. The main purpose of this study was to test the gradient hypothesis by characterizing the spatial distribution of RA production and degradation in the embryonic eye cup before and after invagination of the secondary optic vesicle. In addition, we wanted to differentiate retinoid metabolism between retina and retinal pigment epithelium (RPE). We employed a bioassay, based on an RA-sensitive reporter cell line [49,52], to measure aldehyde dehydrogenase activities, RA-release and breakdown at specific locations in the chick retina. Since an asymmetric distribution of CRABP-I would corroborate a polarizing morphogenetic effect of RA, the distribution of this binding protein was also investigated.

2. Materials and methods Fertilized eggs of White Leghorn chickens were obtained from a local poultry farm, incubated from 12 h to 17 days at 378C, 65% humidity, in a forced draft incubator and staged according to Hamburger and Hamilton (HH; [8]). Comparisons between retinal tissue of different embryonic stages or different locations in the retina were based on samples normalized for protein content. Preparations are listed in Table 1.

2.1. Protein biochemistry and Western blotting For biochemical analysis, tissue was dissected in icecold Hank’s buffered saline (HBSS). Depending on the stage of development we prepared whole eye anlagen, retina with developing pigment epithelium, or pure retina and RPE (Table 1). Tissue was collected on ice, spun down to remove dissecting buffer and triturated in an equal volume of 10 mM phosphate buffer pH 7.4 with 30 mM NaCl containing a cocktail of protease inhibitors of 1 mM PMSF, 1 mM leupeptin, 1% aprotinin (4.8 U / mg) and 1 mM pepstatin. The homogenate was centrifuged for 60 min at 100,0003g to obtain a supernatant containing the cytoplasmic proteins. Protein concentrations in these extracts were determined with the BCA protein assay. Homogenates were subjected to discontinuous sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS– PAGE) under denaturing conditions. After electrophoretic

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Table 1 Preparation of eye tissue from chick embryos a Time of incubation

Embryonic stage

Tissue preparation

23–33 h 33–45 h, E2 45–52 h, E2–3 E4 E5–6

HH HH HH HH HH

E8–9

HH 33–35

E10 E12 E15 E17

HH HH HH HH

Anterior part of neural tube Whole eye anlagen Whole eye anlagen D, V halves of eye anlagen D, V halves of eyes, Vitreous and lens removed D, V, T, N, thirds of retina; D, DM, VM, V, T, TM, NM, N quarters of retina, RPE separated D, V thirds of retina, RPE separated D, V thirds of retina, RPE removed D, V thirds of retina, RPE removed D, V thirds of retina, RPE removed

7–9 10–11 12–13 23–24 27–29

36 38 41 43

a

E, embryonic day; HH, stage of development according to Ref. [8]; D, dorsal; V, ventral; T, temporal; N, nasal; DM, dorsomedial; VM, ventromedial etc.; RPE, retinal pigment epithelium.

separation, proteins were transferred to a cellulose nitrate membrane by a semi-dry blotting procedure (Fastblot 014¨ 200, Biometra, Gottingen, Germany). A mix of molecular weight markers was also run, blotted and stained with AuroDye forte (Amersham Buchler, Braunschweig, Germany). Blots were dried between filters and stored at 48C. Isoelectric focusing (IEF) of native proteins was performed in an Isobox IEF apparatus (Hoefer Scientific / Pharmacia, Freiburg, Germany) with commercially obtained agarose gels (pH 3–10; Isolab, Mechelen, Belgium) following the protocol recommended by the manufacturer. As in the course of our experiments these gels became no longer available, we switched to self-made gels using agarose-coated polyester film (GEL-Fix, Serva, Heidelberg, Germany), silanized glass plates and 1-mm-thick plastic spacers. The gel solution contained 0.8% agarose (suitable for IEF, Serva, 11402), 2% sorbitol (Merck, Darmstadt, Germany) and 3% ampholytes pI 4–7 (Serva, 42948). Electrode wicks were soaked in 0.5 M acetic acid and 0.5 M NaOH for anode and cathode, respectively. On one agarose gel, 125 mm3125 mm31 mm, usually 18 samples were loaded with 10 mg protein in 8 ml volume per lane. Running conditions were: 10 min at 1 W; removal of the sample mask; 5 min, 5 W; 45 min, 15 W; all at 1200 V maximum. Protein standards (Sigma) were used to mark pH positions in the gel. Lanes with pI markers were fixed and stained with Coomassie G250. For immunological detection, the separated proteins were transferred onto a cellulose nitrate membrane by capillary blotting. After temporary staining with Ponceau-S, blots were dried between filters and stored at 48C.

2.2. Immunochemical detection in Western blots and histological slices A rabbit polyclonal antiserum against the mouse class I aldehyde dehydrogenase RALDH-2 [21,54] was used in a working solution of 1 / 3000. Cellular retinoic acid binding protein (CRABP)-I was detected with a monoclonal anti-

body (MA3-813, mouse; Affinity Bioreagents, Golden, CO), diluted 1 / 1000 on blots. As secondary antibodies we used horseradish peroxidase-labeled goat-anti rabbit and goat-anti mouse monoclonal antibodies (affinity isolated, Sigma), dissolved 1 / 5000 in IGSS-quality gelatin (Amersham). Immunocytochemistry and detection were performed with the enhanced chemoluminescence method (ECL, Amersham) using the fast protocol supplied by the manufacturer. For immunohistochemical localization of CRABP-I, chick embryos of stages E2 (HH 10), E3 (HH 18), E4, E6.5 and E9 were fixed in 4% phosphate buffered paraformaldehyde (PFA, in 0.1 M PB, pH 7.4), cryoprotected in 30% sucrose / PFA at 48C, cut with a cryostat microtome (Leica CM3050) in transverse sections of 20 mm (E2, E3) or 30 mm (E4, E6.5, E9) and mounted on gelatin-coated slides. Standard immunohistochemical procedures were followed using the avidin–biotin–peroxidase complex (ABC) method with reagents from Vectastain elite kits (Vector laboratories, Burlingame, CA) and nickel sulfate intensification of diaminobenzidine. The anti CRABP-I primary antibody was diluted 1:200 in 0.04% TritonX100 / 1% serum and incubated overnight. To demonstrate the specificity of the immunocytochemical pattern, a complete staining sequence was run without prior incubation in primary antibody. In this case the staining was completely abolished.

2.3. Assays for biological activity of RA The reporter cell line [49] consists of F9 teratocarcinoma cells transfected with the b-galactosidase gene under control of the RA responsive element from the RA receptor b (RARb). Tests of the reporter cells with RA stereoisomers showed them to be most sensitive to alltrans RA with a detection threshold of 10 212 M. For 13-cis RA, about 100-fold higher concentrations and for 9-cis RA, about 300-fold higher concentrations are necessary to match the color reaction elicited by all-trans RA

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[25]. The cells were grown in CO 2 -buffered DMEM, supplemented with 10% FCS, penicillin / streptomycin, and 0.8 g / l geneticin. For the RA assay, reporter cells were plated into gelatin-coated 96-well microtiter plates, grown to confluence, and cultured for about 12 h with samples diluted in cell culture medium. To visualize the induction of b-galactosidase, the cells were fixed with 1% glutaraldehyde, washed thoroughly with 0.1 M PBS, and incubated for 3 to 8 h at 378C with 45 ml / well of 0.1 M PBS containing 3.3 mM K 4 Fe(CN) 6 33H 2 O, 3.3 mM K 3 Fe(CN) 6 33H 2 O, 6 mM MgCl 2 , and 2 mg / ml 5-bromo4-chloro-3-indolyl-b-D-galactopyranoside (X-Gal, dissolved in dimethyl formamide as 40 mg / ml aliquots). The intensity of the blue reaction product was measured with an ELISA reader (El x 800, BIO-TEK, Winooski, VT) at 630 nm. The F9 reporter cell line was used to investigate RAsynthesizing enzyme activities. As described previously [26], cell homogenates were equalized for protein content and separated by IEF gel electrophoresis (see above). Then, parallel lanes of the gel were cut into 24 or 32 consecutive slices 2.25 mm apart. These IEF fractions were distributed into the wells of microtiter plates, where proteins were eluted and assayed for RA synthesis from 50 nM all-trans retinaldehyde in the presence of 0.6 mg / ml dithiothreitol and 0.8 mg / ml NAD 1 . After 3 h of incubation at 378C in darkness, 25 ml / well of reaction products were removed and tested for RA content with the reporter cells in 75 ml medium / well. For the characterization of RA synthesizing enzymes, parallel assays were carried out in the absence of NAD 1 or using 100 mM citral as enzyme inhibitor. In addition to enzyme activities, we measured release and degradation of RA by brain, retina and pigment epithelium from E8–E9 chick embryos. For the detection of spatial differences the retinas were cut into quarters moving either from dorsal to the ventral side or from temporal to nasal. The tissues were dissected in ice-cold HBSS and collected in DMEM / 10% FCS on ice. In 100 ml medium the dorsal, dorsomedial, ventromedial, ventral, nasal, nasomedial, temporomedial, temporal quarter from

one retina, or the RPE from one eye were separately collected. For comparison in the same assay, we tested one optic tectum, one forebrain hemisphere, one brain stem hemisection, and meninges from one side of the brain, which were each collected in 300 ml medium. To investigate RA-release into the medium, tissues were incubated for 7 h at 378C in darkness in closed Eppendorf vials without the addition of cofactor or other substrate. The activity of RA-metabolizing enzymes was tested by adding 1 nM all-trans RA. As enzyme inhibitors we used 10 mM ketoconazole (blocking cytochrome P450 oxidases) or 0.1 mM citral (blocking RA-synthesis). Assays for measuring RA synthesis always contained ketoconazole, assays for RA degradation were carried out in the presence of citral. In control experiments both inhibitors were used. After incubation the samples were centrifuged (5 min, 10,0003 g) to remove tissue flakes, and 50 ml of supernatant were tested for RA-activity with the reporter cell line (plus 100 ml medium per well of a microtiter plate, procedure as described above). For direct visualization of RA-activity in the retina we used a reporter mouse strain, which contains a hsplacZ transgene (b-galactosidase) under the control of an RA response element [39,48]. Eyes from E17 reporter mice were dissected, corneas and scleras were stripped from the eye cups. Retinas with attaching vitreous body and lens were fixed for 30 min in 0.2% glutaraldehyde, 2 mM MgCl 2 in PBS, washed three times for 10 min in PBS and then reacted in X-Gal buffer as described above for the reporter cells.

2.4. RA-application in ovo In vivo, RA was applied either locally [11] or by injection onto the chorioallantoic membrane (Table 2). For local application in young embryos, anion exchange resin beads (AG1-X8 and AG1-X2, 200–400 dry mesh size, BioRad) were soaked for 1 h in 10 26 M all-trans RA dissolved in 10% DMSO / DMEM, washed once in DMEM and implanted adjacent to the eye anlagen of E2 and E3 chick embryos. All procedures with RA were performed

Table 2 RA-application and analysis of RALDH-2 immunoreactivity Time of RA-application in ovo

Method of application, concentration of all-trans RA

Evaluation with immunoblots (duration of RA-exposure)

E2 (HH 9–11)

Anion exchange beads, soaked with 1 mM RA Anion exchange beads, soaked with 1 mM RA Injection onto the CAM a , 5 ml 10 mM RA Injection onto the CAM, 5–10 ml 10 mM RA Injection onto the CAM, 10 ml 25 mM RA

E5 (3 days)

E3 (HH 17–18) E6 E7 E16 a

CAM, chorioallantoic membrane.

E5 (2 days) E9 (3 days) E9 (2 days) E17 (1 day)

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under amber light. Since the ion exchange beads release RA continuously within 8 (AG1-X8) to 24 h (AG1-X2) they were used within 45 min after washing. In older embryos 5–10 ml of 10–25 mM all-trans RA, suspension in 10% DMSO / DMEM, were injected onto a blood vessel of the chorioallantoic membrane.

2.5. Statistical analysis Data on RA-release and RA-degradation were analyzed statistically using the Jmp software (SAS Institute, Cary, NC). For the evaluation of RA-production, amounts of RA released were calculated for tissue samples from localized areas of the retina and then normalized using the total retinal RA-release in every experiment. Since data were distributed normally (Shapiro–Wilk W test), we performed ANOVA (analysis of variance), followed by an a posteriori contrast test to compare values from the central retina with the edges. In addition, the RA data of retinal quarters were compared to the value expected for an even RA-release across the retinal surface (two-tailed t-test). To evaluate RA-degradation, we calculated the percentage of RA removed from the medium by each tissue sample. The inhibition of enzymatic RA-degradation by ketoconazole was then tested with the Wilcoxon signed rank test.

3. Results

3.1. Retinoic acid-producing enzymes in the embryonic chick retina To investigate RA-synthesizing enzymes, cytosolic proteins from neural retina and RPE were separated in isoelectric focusing gels under non-denaturing conditions. RA production catalyzed by the different IEF fractions was then measured with a bioassay based on an RA-sensitive reporter cell line [11,26]. At mid-developmental stages (between E7 and E14) we found one enzyme activity with almost neutral pI (6.5–6.9) in the dorsal retina (designated C-D for chick, dorsal) and one activity with a slightly more acidic pI of about 6.0–6.3 in the ventral retina (C-V). RA-production by both enzymes was dependent on the presence of NAD 1 as co-substrate (Fig. 1a and b). A third, even more acidic activity (pI 4.9–5.4) was also NAD 1 dependent. After complete removal of the RPE, this enzyme was not found in the neural retina. In preparations that contained membrane proteins no additional enzyme activities appeared. Since RA-production could also be blocked with the competitive inhibitor citral (Fig. 1c) the enzymes responsible for the conversion of retinal to RA are likely to be aldehyde dehydrogenases. A crossreacting polyclonal antibody against the mouse class I aldehyde dehydrogenase RALDH-2 [1,21] allowed us to further characterize the chick enzymes. In Western blots after SDS–PAGE the antibody bound to one protein

Fig. 1. RA-synthesis in the embryonic retina depends on NAD 1 and is inhibited by citral. Cytosolic proteins from chick retinas were separated by IEF, and the charge-separated fractions were tested for RA-synthesis in the presence (a) or in the absence (b) of 1.2 mM NAD 1 with a reporter cell assay. (a, b) Spatial distribution of enzyme activities in the E12 chick retina. Both, the enzyme activity in the dorsal retina (C-D, pI 6.5–6.8) and the enzyme activity in the ventral retina with a slightly more acidic pI (C-V, pI 6.3) required the presence of NAD 1 as co-substrate. As the RPE was removed from these preparations, the asterisk (*) marks the IEF fraction where an RPE-associated enzyme activity was expected in the gel. (c) Soluble proteins from E9 chick retinas including RPE were separated by charge and tested for RA-synthesis. RA-synthesis was inhibited with 100 mM citral. In addition to C-D and C-V, fractions at the acidic end of the gel, pI 4.9, contain an RPE-associated enzyme activity (data are averages of two independent experiments, agarose gels differed from preparations shown in (a) and (b)). V marks the positions of sample application in the IEF-gels.

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of apparent molecular weight 55 kDa, which was present in the retina at embryonic stages E4–E17, even though the immunoreactivity decreased during late embryonic development (.E15; Fig. 2a). The RALDH-antigen was much stronger in the dorsal than in the ventral retina. However, it appeared also in the RPE with high intensity (or in the chorioid layer, partly attached to the RPE). At E4, where no surgical separation of RPE and neural retina was possible, we found no difference in RALDH-immunoreactivity between dorsal and ventral eye anlage, probably due to the RPE signal. Protein separation with IEF according to electric charge allowed us to differentiate the 55 kDa RALDH-immunoreactive band into several proteins with different isoelectric points (Fig. 2b). At E6 and older embryonic stages, when a separation of RPE from neural retina was feasible, a RALDH-immunoreactive molecule with acidic pI (4.9–

5.4) turned out to be restricted to the RPE (or chorioid; C-RPE). In the retina, the RALDH-positive signal had a neutral pI and was present in protein preparations from the dorsal retina, but not in the ventral retina. When sliced along the temporal–nasal axis, including the dorsal pole, the antigen seemed slightly stronger in the temporal third (see also Fig. 3a). Spatial distribution and electric charge indicate that this aldehyde dehydrogenase is identical with the C-D enzyme activity (Fig. 2b and c). At both locations in the IEF gels the neutral C-D and the acidic C-RPE signals appeared as ladders of several bands separated by small differences in electric charge. After separating equal amounts of proteins from E5 (HH 27), E3 (HH 13–16) and E2 (HH 10–11) eye anlagen and HH 7–9 heads (about 70 embryos per experiment; Table 1) we detected the acidic immunoreactivity (C-RPE) with the development of RPE after E3 and before E5. The dorsal retinal

Fig. 2. RALDH-immunoreactivity and enzyme activity during embryonic development of the chick retina. (a) Cytosolic protein fractions from dorsal and ventral halves of E6–E17 chick retinas were separated by SDS–PAGE, 10 mg protein / lane. Immunoblots were visualized with ECL. RALDHimmunoreactivity at 55 kDa was always stronger in the dorsal than in the ventral half of the retina, and it decreased with embryonic age. D, dorsal; V, ventral. (b) IEF and immunoblotting of RALDH in embryonic retina and RPE / chorioid layer. Proteins, 4 mg / lane, were separated by electric charge and processed as in (a). Lanes E2, HH 10–12 eye anlagen; E5, E5 retina and RPE / chorioid layer, not separated; E9, E9 retina freed from RPE; E9 RPE; pI, marker proteins indicating pH positions in the IEF gel; s.a., area of sample application. After separation by charge, the antibody distinguished acidic immunoreactive bands associated with chick RPE (E5, E9-RPE) from a basic RALDH signal in the retina of embryonic stages E5 and E9. Neither antigen was visible in E2 eyes. (c) IEF-fractions from HH 10–11 eye anlagen and E5 retina / RPE, 20 mg protein / lane, were tested for RA synthesis with the zymography bioassay, same pI scale as in (b). Enzyme activity at E5 corresponded with peaks of immunoreactivity but was absent in E2 eye anlagen.

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zymography assays were performed. In correspondence with the appearance of immunoreactive signals we found enzymatic activity in E4 and E5 eye cups, in older retina and RPE, but not in eye anlagen at E2 (Fig. 2c). The enzyme activity which at later stages was found to be restricted to the ventral retina (C-V) appeared in E5 eye preparation as a low but broad peak (Fig. 2c, pI 6.0–6.3). Since RA might exert a negative feedback on the expression of RA-synthesizing enzymes [37] and might even trigger a spatial polarity in the distribution of aldehyde dehydrogenases [2], we tested whether exogenous RA had an influence on RALDH-immunoreactivity. Following treatment of chick embryos of various developmental stages (Table 2) with all-trans RA in ovo, the amounts of aldehyde dehydrogenase were assessed on the protein level. Retinoid application at early stages frequently caused malformations of face and eyes. In 42% (36 / 85) of the cases treated at HH 10 or earlier one eye was smaller than normal. In addition, we observed irregularities of eye pigmentation. No morphological effects were visible after treatment at E6 or later. From embryos where the severity of teratogenic effects did not prevent the preparation of retina tissue, immunoblots with RALDHantiserum were performed with optic tectum, retina and RPE. In no case did RA treatment of chick embryos alter the distribution of RALDH-immunoreactivity (Fig. 3). Western blots of IEF-gels, which allowed us to differentiate the retinal from the RPE enzyme, revealed no influence of RA on the ocular distribution of these proteins (Fig. 3a).

3.2. Sources and sink of RA in the embryonic retina

Fig. 3. (a) IEF and immunoblotting of RALDH in RA-treated E9 retina and RPE. Separation of soluble proteins by electric charge shows acidic RALDH-immunoreactive bands associated with the chick RPE (arrow C-RPE) and basic bands associated with the dorsal retina (arrow C-D). Treatment with RA 3 days before preparation of the tissue caused no change in immunoreactivity. Ten mg protein were loaded in all lanes. V, N, T, D, ventral, nasal, temporal, dorsal quarters of the neural retina; RPE, retinal pigment epithelium; pI-markers are indicated at the right hand side, where s.a. marks the position of sample application in the IEF-gel; tissue from RA-treated embryos in lanes 2, 4, 6, 8, 10; controls in lanes 1, 3, 5, 7 and 9. (b) SDS–PAGE and immunoblotting of RALDH in RA-treated E5 eye and tectum anlagen. Cytosolic protein fractions from dorsal and ventral thirds of E5 eye cups and E5 tectum anlagen were separated by SDS–PAGE, 10 mg protein / lane. Treatment with RA 3 days before preparation of the tissue caused no change in the RALDH signal. OT, optic tectum anlage; D, V, dorsal, ventral thirds of eye primordium (retina plus prospective RPE); tissue from RA-treated embryos in lanes 2, 4, 6; controls in lanes 1, 3 and 5.

enzyme C-D appeared at the same time. Representative immunoblots for E2 (HH 10–11) eye anlagen, E5 (HH 27) retina / RPE, E9 retina, and E9 RPE / chorioid are shown in Fig. 2b. With tissue samples equivalent to 20 mg protein

The tests described so far measured RA-synthesis in the presence of 1.2 mM NAD 1 and 50 nM all-trans retinaldehyde, which were added as substrates. Therefore, they do not reflect the endogenous production of RA. For this, we collected tissue from different locations in the retina, from RPE and from various brain regions in cell culture medium. After 7 h of incubation, the RA that was released from the tissue samples and had accumulated in the medium was assayed with the reporter cells. As predicted from the zymography bioassay, the endogenous RA production was high in the embryonic RPE and in the retina, but low in the optic tectum and other brain tissues. The meninges were found to be another source of RA (Fig. 4). Within the E8–9 retina, RA release was high in the dorsal and ventral quarters of the eye but low at midlevel. Across the horizontal axis of the retina, only slightly lower RA production appeared in the nasal retina, yet no pronounced gradient. Differences along the dorsal–ventral axis were highly significant (ANOVA analysis of variance, F3,36 529.92, P,0.0001) but not along the temporal axis (P.0.1). This significant regional difference was due to high RA production from dorsal and ventral quarters and low activity in the mid-horizontal retina (a posteriori

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measure for RA-inactivating enzyme activity in the tissue samples. Compared to RA synthesis, the analysis of enzymatic degradation in the E8–9 retina revealed a complementary distribution with highest decrease of RA after incubation with dorsomedial and ventromedial retinal stripes, whereas little RA was removed by dorsal and ventral quarters of the retina (Fig. 5a). To see whether oxidizing enzymes are the cause of this inactivation [31,52] we blocked RA degradation with ketoconazole. As shown in Fig. 5, RA oxidation by the middle region of the

Fig. 4. RA-production in embryonic brain and eye tissues. Tissues from E8–9 chick embryos were incubated for 7 h at 378C in 100 ml DMEM / 10% FCS, 10 mM ketoconazole (brain tissue in 300 ml) in darkness. Then the RA release into the medium was quantified with the reporter cell assay and normalized using the total RA released from one retina in each experiment. (a) Localized RA-release from retinal areas. D, dorsal; DM, dorsomedial; VM, ventromedial; V, ventral; N, nasal; NM, nasomedial; TM, temporomedial; T, temporal quarter of one retina, completely freed of RPE and vitreous body. Error bars represent 95% level of confidence of 10 independent experiments. Asterisks indicate that mean differs significantly from 25%, the expected value for equal distribution across the retinal surface (*P,0.05; ***P,0.001; two-tailed t-test). (b) Normalized RA-release from embryonic brain tissues. Ret, one retina; RPE, retinal pigment epithelium from one eye; FB, one forebrain hemisphere, pia removed; OT, one optic tectum, pia removed; BS, hemisection of one brain stem, pia removed; Men, meninges of one brain hemisphere. Error bars represent 95% level of confidence of 8–14 independent experiments.

contrast test, t-ratio 8.20, P,10 29 ). Fig. 6 presents a summary of the observed local RA release in the embryonic chick retina. In a similar set of experiments, embryonic tissues were incubated for 7 h with medium containing 1 nM all-trans RA. In control experiments we also added 10 mM ketoconazole, a general inhibitor of cytochrome P450 oxidases. The degree of RA removal from the medium that could be blocked with ketoconazole was taken as a

Fig. 5. RA-degradation in the embryonic retina and its inhibition by ketoconazole. (a) One dorsal, dorsomedial, ventromedial, and ventral quarter of an E8–9 retina was incubated for 7 h at 378C in 100 ml DMEM / 10% FCS with 1 nM all-trans RA and 0.1 mM citral in darkness. The remaining RA concentration was determined with the reporter cell assay (gray bars) and compared to controls (5100%), where 1 nM RA was incubated in the absence of retina tissue. To inhibit enzymatic oxidation of RA, 10 mM ketoconazole was added to the medium (white bars). (b) Same experiments with one RPE, one forebrain hemisphere, one optic tectum, one brainstem hemisection and meninges of one brain hemisphere from E9 chick embryos, incubated in 300 ml DMEM / 10% FCS with 1 nM all-trans RA with or without ketoconazole. Error bars represent 95% level of confidence. Asterisks indicate a significant effect of ketoconazole on RA-degradation (Wilcoxon signed rank test, *P, 0.05; **P,0.01, ***P,0.005; two-tailed t-test).

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retina was prevented to a large extent by ketoconazole (Wilcoxon signed rank test, DM: P,0.005, VM: P,0.01). Incubation of 1 nM RA with ketoconazole and tissue from the (RA-releasing) RPE resulted in an increase of the RA concentration in the medium, compared to the control condition without tissue (100%). Similar to retinoid release, the pattern of inactivation of all-trans RA showed horizontal lines of equal activity with no pronounced differences between the temporal and nasal poles. RA degradation was highest in a horizontal stripe in the middle of the retina (Fig. 6b). This result was confirmed with a transgenic mouse strain, where endogenous RA induces the local expression of b-galactosidase as a reporter gene, and thus colorimetric staining for the lacZ product indicates regions of all-trans RA signaling. In the perinatal retina of this mouse, RA activity was absent in a horizontal stripe reaching from the temporal to the nasal pole (Fig. 6c). Some decline of RA activity occurred also with tissue from chick RPE, forebrain, tectum, brainstem and meninges (Fig. 5b). In all samples from brain tissue this effect was not prevented by ketoconazole, however, and was therefore not indicative of any specific catabolic activity. A significant effect of the enzyme inhibitor appeared after incubation with RPE and meninges, both tissues that released large amounts of RA. Even in the presence of 0.1 mM citral that was added in all degradation experiments the medium containing RPE tissue and ketoconazole produced an increased biological activity, suggesting that stored RA was released from the RPE.

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3.3. Retinal distribution of CRABP-I At the stage when determination of dorsal–ventral patterning of the eyes is assumed to take place (at E2, earlier than HH 18) the immunohistochemical analysis completely failed to detect CRABP-immunoreactivity in the neural tube and eye anlagen, whereas the protein was detected in the presomitic mesoderm (Fig. 7a). In accordance with earlier reports [22,35,41] we found CRABPimmunoreactivity to appear later in the nervous system, in differentiating neuroblasts. In the retina, the differentiating retinal ganglion cells and their growing axons were CRABP-positive at E3–E6.5 (stages HH 20–30; Fig. 7b– e). At stage E3 (HH 20) only very few retinal ganglion cells were stained in the center of the retina close to the vitreal surface of the neuroepithelium. As the differentiation of these cells followed a spatio-temporal gradient from a temporal / central origin to the periphery of the retina, the pattern of CRABP expression followed this course of development. Although at E6 a somewhat stronger staining with the antibody against CRABP-I appeared in the optic fiber layer and retinal ganglion cells of the ventral retina, immunoblotting experiments showed no quantitative difference of CRABP-I-immunoreactivity along the dorsal– ventral eye axis (Fig. 8). This was confirmed immunohistochemically with E9 retina wholemounts, which produced a homogeneous staining of evenly spaced cells across the entire surface of the retina (data not shown). In IEF gels (Fig. 8a) the chick CRABP-I appeared as a sharp band with pI 4.5, in SDS–PAGE (Fig. 8b) it migrated with

Fig. 6. Local RA-release and degradation in the E8–9 chick retina. In vitro measurements from retinal stripes cut along the dorsal–ventral and nasal–temporal axes were used to calculate local release and degradation of RA. Data were calculated by geometric averaging, interpolated and plotted with Sigma Plot. (a) RA concentration in 100 ml medium released from tissue equivalent to one quarter retina, (b) percent removal of 10 29 M all-trans RA from 100 ml medium incubated with tissue equivalent to one quarter retina (100, no RA left; 0, same concentration as after incubation without tissue). (c) Retina from E17 RAREhsplacZ mouse, seen from the back side. This mouse is transgenic for an RA-response element driving the b-galactosidase gene, and thus colorimetric staining for the lacZ product indicates those regions of RA signaling [39]. The horizontal stripe of CYP26 expression mentioned in the text is clearly defined in this mouse by its absence of lacZ induction [31]. This indicates that the catabolic enzyme degrades all RA in this region. On either side of the stripe both dorsal and ventral zones are strongly positive due to the expression, respectively, of the RA synthetic enzymes RALDH-1 and RALDH-3 [31,48]. Although RA synthesis by ventral RALDH-3 is twofold greater than that of RALDH-1, the colorimetric assay for lacZ is not sufficiently linear to show this difference.

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Fig. 7. CRABP-I immunohistochemistry in the embryonic chick retina. (a) Transverse section through the head with primary optic vesicles of a stage HH 11 (E2) chick embryo. No CRABP-I signal is visible, whereas staining was positive in the presomitic mesoderm (not shown). (b) HH 18 (E3) secondary eye cup. Differentiating retinal ganglion cells are CRABP-I positive in the center of the retinal neuroepithelium. (c) Higher magnification of CRABP-immunoreactivity (arrow), limited to the vitreal layer of the central retina at E4. (d) Section through optic nerve head of E6.5 retina showing CRABP-I in nerve fibers and retinal ganglion cells (arrows). (e) Higher magnification of E6.5 retina, where CRABP-immunoreactivity is only visible in retinal ganglion cells (arrow) and optic fiber layer. (f) Section through a peripheral part of an E9 retina. Two rows of CRABP-positive cells are seen between the GCL and the proliferating neuroepithelium. These cells are presumably amacrine cells of the future inner nuclear layer. (g) Higher magnification of CRABP-immunoreactive amacrine cells in the E9 retina. ne, neuroepithelium in the stage of cell proliferation; D, dorsal; V, ventral; RPE, retinal pigment epithelium; OFL, optic fiber layer; GCL, ganglion cell layer. All scale bars are 50 mm.

a relative molecular weight of ca. 15 kDa. Corroborating the immunocytochemical results, Western blots revealed increasing levels of CRABP-I during the period of retinal neurogenesis. CRABP-I was not detected at E2 and E3, was weak at E5, and strong at stages E7 through E20 (Fig. 8). After neuronal differentiation, retinal ganglion cells seemed to downregulate the protein. However, the retinal levels of CRABP-I did not decline because at E9 and at later stages the protein remained present in different cell types in the inner nuclear layer, presumably amacrine cells (Figs. 7f, g and 8b).

4. Discussion We analyzed RA synthesis and degradation in the embryonic chick retina. After stage E4, two RA-synthesizing aldehyde dehydrogenases, which were restricted to the dorsal retina and RPE and differed by their pI, could

be characterized in Western blots and with a zymography assay based on an RA-sensitive reporter cell line. Relative molecular weights of RALDH-immunoreactive monomers were 55 kDa, the pI ranged 6.5–6.9 and 4.9–5.4 for the retinal and RPE enzyme, respectively (each with several closely spaced bands). As published previously [32], the ventral retina contained a third enzyme activity (pI 5.9– 6.3), not recognized by our RALDH-antiserum. Unexpectedly, neither the antigens nor RA-producing activities were detectable in stage HH 7–9 heads, HH 10–11 or HH 13–14 eye anlagen, i.e. at embryonic stages when determination of the eye axes takes place. All enzyme activities required NAD 1 as co-substrate and could be inhibited by citral. The following mammalian enzymes with similar properties and distributions seem to be homologous to the avian RA-synthesizing activities [48]: RALDH-1 (dorsal retina, C-D; ALDH in Ref. [7]), RALDH-2 (pigment epithelium, C-RPE; C-MV in Ref. [32]) and RALDH-3 (ventral retina, C-V; [20]), suggesting

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reactivity was visible in this population of differentiating neurons but became downregulated as these cells matured. At later embryonic stages (E7, E9), as in the adult retina [34], CRABP-antibody stained amacrine cells in the inner nuclear layer. Although at E4 CRABP-I appeared stronger in the ventral part of the ganglion cell and optic fiber layer, the distribution did not correlate with the zones of RAsynthesis and degradation. At later stages when a dorsal– middle–ventral pattern of RA became apparent, the binding protein showed a uniform distribution in the inner nuclear layer of the retina.

4.1. Is a retinoid gradient responsible for the establishment of the dorsal–ventral eye axis?

Fig. 8. Immunoblotting of CRABP-I in embryonic retina and RPE. (a) Separation of soluble proteins by electric charge show CRABP-positive bands at pI 4.5 in embryonic retinas E5 and older, but no immunoreactivity in E2 eye anlagen. Four mg protein were loaded in lanes E2 (HH 10–12 eye cups), E5 (retina / RPE), E9 (pure retina), E9 RPE, and 10 mg protein in lanes E5 (retina / RPE, middle) and E7 (pure retina). D, M, V, dorsal, middle, ventral thirds of the neural retina; RPE, retinal pigment epithelium; pI-markers are indicated at the right hand side, where s.a. marks the site of sample application in the IEF-gel. (b) SDS–PAGE and immunoblotting of CRABP-I in dorsal (D) and ventral thirds (V) of E7, E8, E10 and E20 chick retinae, 5 mg protein / lane.

a corresponding nomenclature for the chick (cRALDH-1– 3). For the first time we show a catabolic sink of RA in the chick retina. Contrary to what would be expected from a ventral–dorsal gradient model, the RA-inactivating activity was located in a horizontal zone extending from the temporal to the nasal pole of the retina. RA-inactivation was sensitive to ketoconazole, an inhibitor of cytochrome P450 oxidases. At E9, the distribution of RA synthesizing and oxidizing activities produced a pattern of high RA concentration at the dorsal and ventral poles of the retina with a trough in the middle. Differences along the temporal–nasal axis were small. This distribution was corroborated by the similar pattern of RA-driven transgene expression in a reporter mouse. Several and in part contradictory biological functions have been suggested for the cellular RA-binding proteins within the pathway of retinoid signal transduction. Nevertheless, a graded distribution of CRABP-I, which is the RA-binding protein present in the nervous system of the chick, would support the notion of a morphogenetic gradient of RA-activity in the developing eye. We therefore investigated the distribution of CRABP-immunoreactivity in relation to the pattern of retinoid metabolic activities. Like the enzymes, CRABP-I was absent in the neuroepithelium at stage E3 and younger. The protein was first expressed by differentiating retinal ganglion cells in the central retina between E3 and E4. CRABP-immuno-

Surgical operations on the early eye anlage evaluated with neuroanatomical tracing techniques revealed that the molecular polarity of the temporal–nasal retina axis is established already in the primary optic vesicle [3,47]. At the same time, molecular markers for dorsal–ventral eye polarity appear. Experimental misexpression of early expressed transcription factors causes targeting errors of retinofugal axons later in development [16,40,53]. Teratogenic effects of excess RA on eye development and the discovery of spatially restricted RA-synthesizing aldehyde dehydrogenases in the retina prompted the hypothesis that a ventral.dorsal gradient of RA may be responsible for the original establishment of this eye axis [29,30]. That idea has subsequently been supported by ventral eye defects in RAR / RXR knockout mice [12] and after inhibition of aldehyde dehydrogenases in zebrafish [24]. Experimental administration of excess RA resulted in an upregulation of the ventral transcription factor Pax[b], downregulation of the dorsal aldehyde dehydrogenase, and the induction of an ectopic eye fissure in zebrafish [11,24]. Similar aldehyde dehydrogenase activities as in mice have then been found in chick retinal development [44]. In addition, HPLC-measurements demonstrated a higher concentration of RA in the chick ventral retina at E4, which dissipated during the following 4 days of development [32]. Yet, none of these studies addressed RA-production at precisely the stage when dorsal–ventral determination of the eye anlage takes place. The data presented here indicate that no RA-synthesis occurs within the chick eye anlage during this period, within the limits of detectability of the zymography techniques employed. Hence, RAsynthesizing enzymes within the developing chick retina may not operate as the primary ventralizing agents. It is quite possible, however, that a polarizing RA gradient derives from a source external to the eye anlage. A similar situation may be found in the neural crest and their derivatives. They constitute another system whose development is dependent on retinoid signaling, but they do not synthesize RA themselves. Before differentiating to sensory neurons, sympathetic ganglia, Schwann cells etc., the neural crest cells migrate through mesodermal ter-

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ritories containing high retinoid concentrations [1,9,19]. An analogous situation in accordance with a ventral. dorsal gradient in eye development would stipulate an RA source in the surface ectoderm or mesenchyme ventral to the optic vesicle at stage HH 9–11. In situ hybridization has in fact located RALDH-2 to such a position around the maxillar–mandibular cleft in mice [37]. When the present study was concluded, Noda et al. had cloned mouse and chick RALDH-3. Their in situ hybridization analysis shows a staining of raldh-3 transcripts in the ventral head of stage HH 9–10 chick embryos, then in the surface ectoderm covering the ventral region of the eye cup (HH 12), and later in the ventral margin of the retina [44]. Despite various attempts at influencing development of the visual system by manipulating the RA concentration, only one study reports a decisive effect of RA on the neuronal connection between the eye and central targets. Following RA treatment of Xenopus larvae, rhodamine tracing revealed a strong reciprocal retino-retinal projection in addition to overshooting of retinal fibers into the hindbrain [23]. Effects that would reflect an alteration of positional identity of retinal ganglion cells have not been described. We must conclude that at this point an influence of the RA distribution on the pattern of neural retinofugal connections is not supported by experimental data.

4.2. Spatial distribution of enzymes of RA metabolism in the differentiating chick retina Direct quantification of all-trans RA with HPLC as well as indirect measurements with a reporter cell line showed that by the end of the first week of chick retinal development an initial ventral–dorsal RA gradient has disappeared [32]. Yet, expression and RA-synthesizing activity of aldehyde dehydrogenases continue. The function of their complicated local distribution within the plane of the retina remains a mystery. Instead of a single gradient we detected two zones of active RA production, namely in the ventral and in the dorsal retina. These were separated by a zone of low RA synthesis and high catabolic activity. Moreover, the RPE emerged as a potent source of RA. A similar pattern has recently been found in the E13–14 mouse retina [20,31], though in this altricial mammal the RPE becomes a source of RA synthesis only shortly before birth [28]. Experiments with reporter mice that possess a retinoid-dependent transgene allowed us to visualize gene activation by retinoids in vivo [39,43,48]. In the differentiating mouse retina, corresponding to E7 chick embryos and older, RA-driven gene expression is highest in the dorsal and ventral thirds with a mid-retinal horizontal stripe devoid of RA activity (Fig. 6c). Accordingly, in situ hybridization for the cytochrome P450 oxidase CYP26 revealed a horizontal zone of mRNA expression, just above the midline and reaching from the temporal to the nasal periphery of the mouse retina [31]. The complementary distribution of CYP26 and RA activity fits because this

enzyme is believed to be responsible for the inactivation of RA [50]. This distribution is in excellent agreement with the pattern of RA production and RA degradation that we describe here for the chick retina during the second week of embryonic development. As expected if RA degradation was catalyzed by the chick homolog of CYP26, the activity in mid-retinal tissue could be inhibited by ketoconazole. In this case RA is presumably oxidized to 4-OH-all-trans RA, and this form, which may still induce transcription, will be inactivated through further oxidation, catalyzed by the same enzyme. In addition to corroborating the dorsal location of a chick RALDH (C-D, [32]), we investigated the temporal regulation of this enzyme. It appeared after the secondary eye vesicles had invaginated (E3), then became restricted to the dorsal retina, and although RALDH-immunoreactivity was detectable until hatching, it decreased during the second half of embryonic development. This analysis is consistent with data derived by Northern blotting using a probe against the chick cytosolic aldehyde dehydrogenase [6]. From E7 to E19 transcript levels declined by a factor of 20–25. A second report with in situ hybridization and an antiserum against the same enzyme detected no dorsal– ventral gradient in the eye cup stage: the primary optic vesicles were not analyzed, but in the youngest stages investigated, E3.5 (in situ hybridization), E3 (HH 14, immunohistochemistry) enzyme transcript and antigen were seen along the entire extent from dorsal to ventral pole of the retina. Only later, at E5 (in situ hybridization) and E3.5 (HH 18, immunohistochemistry), was the enzyme restricted to the dorsal half of the retina [7]. In mice, CYP26 first occurs in the dorsal eye cup. It has been suggested that by keeping local RA levels low, the catabolic enzyme may allow for the expression of RALDH-1 in the dorsal retina if one assumes the gene for this enzyme to be repressed by RA [11]. For this reason we tested whether previous administration of RA changed the protein expression of RALDH-immunoreactivity in the chick eye. No regulation was apparent at stages E5, E9 or E17. This contrasts with a paper on RA-treated zebrafish larvae, where elevated levels of all-trans RA abolished ALDH-IR in the dorsal eye. Experiments with mice indicate, however, that gene regulation of RALDH-2 by RA is indirect and dependent upon other factors: administration of teratogenic doses of RA at 8.5 days of gestation (dpc) resulted in the downregulation of RALDH-2 mRNA in caudal regions of the embryo but not in the cervical mesenchyme [37]. Moreover, this regulation was observed 12 and 27 h after RA treatment, but not after 6 h, and no regulation occurred with RA administration at 7.5 and 10.5 dpc.

4.3. Functional significance of the retinal pattern of RA distribution Our description of the retinal RA distribution as well as CYP26 in situ hybridization data [31] indicate two com-

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partments of high RA concentration separated by a horizontal zone of RA degrading activity. Several genes are distributed along the dorsal / ventral axis in the retina within compartments rather than gradients. This includes, in the mouse, COUP-TFII [31] and the tyrosine receptor kinase ligand ephrin B2 (unpublished observation), both of which are RA-inducible in P19 cells. The ventral borders of these genes match closely to the border between mouse RALDH-1 and CYP26, and a function of the RA-degrading activity may be to set up gene expression boundaries for RA-regulated genes. Nevertheless, the tripartite pattern of RA activity in the embryonic retina is not reflected by known histological features in the adult chick eye, although in the mouse a coincidence with insertion sites of extraocular muscles has been noted [48]. Molecular research during the last decade unveiled many surprising similarities in pattern formation between insects and chordates, including processes of eye development. In this context, the absence of a significant functional boundary in a tissue that displays striking molecular patterns in development has been compared with territorial boundaries in Drosophila [31]. There, developmental compartments often do not correlate with morphological landmarks in the imago. A second possibility may be that the RA-degrading enzyme is transported along the axon and that it acts at a distance from the cell body. It is already known that mouse RALDH-1 [30], RALDH-2 [1] and RALDH-3 (unpublished observation) are transported along the axons. RA released from these axons may act in a paracrine fashion in the innervated brain regions. CYP26 within the axon would then act as a sink for RA along the tectal territories innervated by this population of retinal ganglion cells.

4.4. The role of CRABP-I in the embryonic retina Although mutant mice that lack both CRABP-I and -II have no specific phenotype [18], a high conservation of the primary sequence of CRABPs as well as pathological changes resulting from ectopic CRABP expression suggest an essential role of these proteins in retinoid signaling [36]. Among several hypotheses for the function of CRABPs, the idea that CRABP may assist in establishing morphogenetic RA gradients [41] has prompted us to investigate whether retinal distribution of CRABP-I correlates in any way with the patterns of RA synthesis and degradation. This seemed not to be the case. In the E2 retina no CRABP-immunoreactivity was detectable, and at later stages CRABP-I concurred with the spreading of differentiating ganglion cells, was then downregulated in these neurons as they matured. Later, CRABP-I appeared in the inner nuclear layer, probably in amacrine cells [34]. This, again, points to additional roles of RA in the retina. A different line of research, separate from the work on morphogenetic gradients, indicates that RA is a regulator of cell differentiation, including neurogenesis and differentiation of glia [5,17,38,46]. In retinal cell cultures, RA

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induces the differentiation of photoreceptors [10,13,14,42]. Our data on CRABP-I distribution in the developing chick retina support a role of retinoids in neural differentiation, especially of retinal ganglion cells and amacrine cells.

Acknowledgements The authors thank Marianne Dohms for help with immunohistochemistry and Helga Gaube for proofreading the manuscript. This work was supported by the Deutsche Forschungsgemeinschaft, grant ME 1261 / 5-1 to J.M.

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