During eye development in the axolotl (Ambystoma mexicanum Shaw), morphogenetic movements bring together tissues from head epidermis, neuroectoderm and neural crest. The stages 0 to 14 of axolotl eye development were expanded from Rabi’s (1898) stages 1 to 10 and correlated with Harrison’s (1969) stages. At the onset of neurulation (stage 13 of Harrison), the head epidermis is already determined to form skin, and the neuroectoderm is committed to form brain, because these tissues develop autonomously in 60% Leibovitz L-15 culture medium. However, a sequence of mutual tissue interactions is necessary to stimulate eye development. When head epidermis and neuroectoderm were cocultured, eyes developed, containing retinas with photoreceptors (stage 8) and lenses with secondary lens fibres (stage 8). The first event needed in this case appears to be the secretion of a growth factor from the head epidermis which stimulates retina development from the neuroectoderm. When neuroectoderm cultures were exposed to nondialysable extracts (30μgml−1) of an adult epidermis derivative, the bovine cornea, pigmented retinas (stage 6) and at higher concentrations (3000 µg ml−1) neural retinas developed (stage 6). In turn, lens formation is stimulated in the head epidermis by a retina-derived growth factor. A mutation that causes adult eyelessness (e eyeless, nonlethal, recessive) affects the earliest event in eye development (stage la), while a mutation that causes arrest of eye development (mi microphthalmic, lethal, recessive) acts in a later event (stage 8). Two possibilities have been considered in the case of mutation e: either the head epidermis does not secrete sufficient amounts of active growth factor, or the presumptive retina itself is defective. The latter statement turned out to be correct, because mutant e neural plates rarely developed early retina stages (stage 5) in organ culture when combined with wild-type head epidermis. On the other hand, wild-type neural plates formed advanced retinas (stage 8) in all cases when combined with mutant e head epidermis. As expected, no retina or lens developed when both neural plate and head epidermis were from mutant e donors. The heterozygous presence of genes e and r (renal insufficiency, lethal, recessive) produces duplications of the presumptive retina at the optic stalk. This observation is consistent with the notion that the mutation e, assisted by the r locus, causes a primary failure in the presumptive retinal region.

The eye tissues of the axolotl (Ambystoma mexicanum Shaw) are derived from distant embryonic regions, which are brought together by the morphogenetic movements (Reyer, 1977). These embryonic regions involve first the head epidermis and neuroectoderm and later neural crest pigment cells (Frost, Epp & Robinson, 1984) and neural crest mesoderm. At the late gastrula stage, the prechordal plate mesoderm secretes a neuralizing protein factor (Tiedemann & Born, 1978) which stimulates the cells of the overlying neural plate, thus separating the already determined optic fields (Mangold, 1929; Adelmann, 1937). In these fields the presumptive dorsal retinal cells are derived from the contralateral body side (Jacobson & Hirose, 1978). The optic fields are located in the anterolateral neural plate (Fischel, 1921; Manchot, 1929; Woerdeman, 1929; Adelmann, 1929) but they rapidly move to the neural folds (Spemann, 1938; Jacobson, 1962; Brun, 1981) where they contact the head epidermis. The latter tissue is necessary for retina development (Filatow, 1926; Detwiler & Van Dyke, 1954; Rollhâuser-Terhorst, 1981) to the rhodopsin-containing photoreceptor stage (Papermaster & Schneider, 1982). The nature of the retina-stimulating effect of the head epidermis has been unknown. Using neural plate cultures we could stimulate retina development with a macromolecular extract of bovine cornea, a head-epidermis derivative. The retina in turn stimulates the development of a lens (Le Cron, 1907; Liedke, 1955; Karkinen-Jaaskelainen, 1978) with lens-specific proteins, the crystallins (Brahma & McDevitt, 1974; Brahma & Bours, 1976), by secreting protein growth factors (Arruti & Courtois, 1978; Beebe, Feagans & Jebens, 1980; Courty et al. 1985).

Two mutations (Malacinski & Brothers, 1974; Malacinski, 1978) are known to disrupt eye development in the axolotl; mi (microphthalmic) (Humphrey & Chung, 1977) acts late in eye development, and causes degeneration of already differentiated cells derived from the head ectoderm at hatching time, while e (eyeless) (Humphrey, 1969, 1975; Briggs, 1973) acts early in eye development, suppressing growth of the presumptive eye tissues and some hypothalamic cells (Van Deusen, 1973). Both mutations are recessive. While the first is lethal, the latter produces secondary failures in the pituitary gland and the gonads, thus causing sterility. One purpose of our study has been to describe the normal stages 0 to 14 of axolotl eye development, expanded from Rabi’s (1898) stages 1 to 10, and to correlate them with the whole body developmental stages (Harrison, 1969; Bordzilovskaya & Detlaf, 1975; Schreckenberg & Jacobson, 1975). Next we wanted to determine the morphological stages at which eye development was disturbed by the mutations mi and e.

In the mutant e the question arose whether the primary failure resides in the presumptive retinal cells themselves or whether perhaps the head epidermis does not produce sufficient amounts of active retina-stimulating factor. We used mutant e-wild-type tissue combinations in organ culture to answer this question.

Eye stages

Axolotls were mated overnight. They were either wild type or heterozygous for e, r or mi. Spawnings were raised in 10% (v/v) Steinberg’s saline. Stages were monitored according to Bordzilovskaya & Detlaf (1975) and Harrison (1969). At all stages, individuals or their eyes were fixed in Bouin’s fixative (Conn et al. 1960) for 2·5 h. To avoid pain, old embryos, larval and adult axolotls were anaesthetized in 0·1% (w/v) ethane-zn-aminobenzoate methanesulphonate (Sigma) prior to fixation. Tissues were dehydrated in a graded ethanol series, amylacetate, toluene, embedded in paraplast and sectioned at 7μm. Rehydrated sections were stained with haematoxylin-eosin (Conn et al. 1960) and mounted in Permount (Fisher). Staging of eye cross sections was according to Rabl (1898) (Fig. 1) as far as they were treated there. Both Harrison’s and Rabi’s stages had to be extended. In mutants (Figs 2, 3) and organ-cultured eyes, development was not always uniform, and stages were assigned according to the most developed structures.

Fig. 1.

Stages 0 to 13 (opposite) and 14 (above) of eye development in wild-type axolotls. Sectioned at 7μin paraplast and stained with haematoxylin-eosin (except stage 13 sectioned at 1 μm in Araldite and stained with pararosaniline-methylene blue). See Results for explanation of the annotations and a description of the stages. At stage 0, the neural fold of the head region is curved downward and the most anterior portion appears on the ventral side in cross section. Stage 14 is from an axolotl simultaneously heterozygous for genes e and r. Vision was normal, although several retinal vesicles (arrows) had formed at the optic stalk. Scale bars, 200μm; 0 to 3, 4 to 10, 11 to 12, 12 to 14 equal magnifications.

Fig. 1.

Stages 0 to 13 (opposite) and 14 (above) of eye development in wild-type axolotls. Sectioned at 7μin paraplast and stained with haematoxylin-eosin (except stage 13 sectioned at 1 μm in Araldite and stained with pararosaniline-methylene blue). See Results for explanation of the annotations and a description of the stages. At stage 0, the neural fold of the head region is curved downward and the most anterior portion appears on the ventral side in cross section. Stage 14 is from an axolotl simultaneously heterozygous for genes e and r. Vision was normal, although several retinal vesicles (arrows) had formed at the optic stalk. Scale bars, 200μm; 0 to 3, 4 to 10, 11 to 12, 12 to 14 equal magnifications.

Fig. 2.

Eye development in vivo in axolotls homozygous for gene e. (A) Retarded optic vesicle (ov) (stage la, but smaller) in a 4-day embryo (Harrison stage 30). By that time, wild-type eyes would reach stage 2. (B) Small lens vesicle (Zv) (stage 5) next to disorganized neural retina (nr) mass (abnormal stage lb to 4) in a 10-day embryo at hatching (Harrison stage 42). Wild-type eyes would be at stage 7. (C) A pair of retinal rudiments with hypothalamus rudiment (Ay) in between at time of hatching. Tapetum nigrum (tn) pigmented (stage 6), neural retina (nr) remaining a flat epithelium (resembling stage lb). (D) Eyeless orbital area of 56-day-old larva (Harrison stage 47). A small necrotic retinal cell cluster (arrow) may still be discernible. Wild-type eyes would be at stage 11. Scale bars, 100μM.

Fig. 2.

Eye development in vivo in axolotls homozygous for gene e. (A) Retarded optic vesicle (ov) (stage la, but smaller) in a 4-day embryo (Harrison stage 30). By that time, wild-type eyes would reach stage 2. (B) Small lens vesicle (Zv) (stage 5) next to disorganized neural retina (nr) mass (abnormal stage lb to 4) in a 10-day embryo at hatching (Harrison stage 42). Wild-type eyes would be at stage 7. (C) A pair of retinal rudiments with hypothalamus rudiment (Ay) in between at time of hatching. Tapetum nigrum (tn) pigmented (stage 6), neural retina (nr) remaining a flat epithelium (resembling stage lb). (D) Eyeless orbital area of 56-day-old larva (Harrison stage 47). A small necrotic retinal cell cluster (arrow) may still be discernible. Wild-type eyes would be at stage 11. Scale bars, 100μM.

Fig. 3.

Eye development in axolotl homozygous for gene mi. Small eye at same stage as wild-type eyes (stage 9, but smaller) in a 20-day-old larva (Harrison stage 43). Cell number is reduced by cell death in cornea, lens, retina, brain areas, and gills. The lens is outside the optic cup, because of the drastic space reduction in the vitreous chamber. Annotations as in Fig. 1. Scale bars, 100 μm.

Fig. 3.

Eye development in axolotl homozygous for gene mi. Small eye at same stage as wild-type eyes (stage 9, but smaller) in a 20-day-old larva (Harrison stage 43). Cell number is reduced by cell death in cornea, lens, retina, brain areas, and gills. The lens is outside the optic cup, because of the drastic space reduction in the vitreous chamber. Annotations as in Fig. 1. Scale bars, 100 μm.

Bovine cornea extracts

Adult bovine corneas were homogenized in 0·005 M-sodium phosphate buffer, pH7·2, and centrifuged at 12000g, 5°C, for 30min. The supernatant was lyophilized. Samples of cornea powder were sterilized in 70% (v/v) ethanol for 15 min at 23°C, and dialysed (Spectrum, relative molecular mass cut-off 12–14×103) overnight against deionized water, pH5·l, 5°C. The sterile contents of the dialysis bag were diluted with sterile water to reach 40 % of the desired volume and were mixed with 60 % sterile full-strength Leibovitz L-15 medium (Grand Island) with glutamine and 100 mg ml−1 gentamicin sulphate (Sigma).

Organ culture

Jelly coats of axolotl embryos at late gastrula (Harrison stage 12) were oxidized with 10 mg I−1 KMnO7 in deionized water at 23°C, 1 h. Embryos with their jelly coats were briefly immersed in 70% (v/v) ethanol using a wide-mouth pipette, and washed in two changes of autoclaved deionized water, pH 7·8, and one change of sterile-filtered 50% (v/v) modified amphibian Ringer’s solution (100%: 0·lM-NaCl, 0·002M-KC1, 0·002M-CaCl2, 0·001 M-MgSO4, 0·005M-Hepes (Sigma), 50 mg I−1 gentamicin sulphate, pH 7·8). Under sterile conditions, embryos were de jellied with forceps and transferred to a dish with fresh 50 % modified Ringer’s solution. When they had reached the earliest stage of neurulation (Harrison stage 13), the vitelline membrane was removed with forceps. The right body side of the embryo was left intact for phenotypic identification at hatching time. On the left body side the anterolateral neural plate with adhering prechordal mesoderm was excised with tungsten needles to serve as a source of the retina, or presumptive head epidermis was peeled off as a source of the lens (Fig. 4). Any cell type contamination across the neural plate margin resulting from jagged cuts will lead to false results and must be carefully avoided. Excised tissues were transferred using a Pasteur pipette to the central well of plastic organ culture dishes (Falcon) with 60 % (v/v) Leibovitz L-15 culture medium supplemented with glutamine and 50 mg I−1 of gentamicin sulphate. The culture dishes were humidified by the surrounding sterile water rings (Fig. 4). Tissues were cultured for 9 days, either as tissue sandwiches, alone as controls, or in the presence of bovine cornea extract. Cultured tissues and donor embryos, the latter raised in 50% modified Ringer’s solution, were fixed at hatching age after 9 days, sectioned and stained as described above. After staging according to Rabl (1898), explants were grouped into eye-positive (stages 1 to 9) and eye-negative (stages 0 or -, not identifiable), independently for pigmented retina, neural retina, and lens. Results were also evaluated using a four-table χ2 test for independence.

Fig. 4.

Organ culture design. Embryonic tissues were explanted at the early neural plate stage (Harrison stage 13) and cultured for 9 days to reach the hatching age (Harrison stage 42). (A) Left anterolateral neural plate neuroectoderm (dotted) developed into brain tissue, and (C) left anterolateral epidermis ectoderm (hatched) developed into skin tissue. When the two tissues were cultured together as a sandwich (B), and were both derived from wild-type donors, eyes with pigmented and neural retina and lens developed. The right body side of the donor embryo was left intact to determine the phenotype at hatching.

Fig. 4.

Organ culture design. Embryonic tissues were explanted at the early neural plate stage (Harrison stage 13) and cultured for 9 days to reach the hatching age (Harrison stage 42). (A) Left anterolateral neural plate neuroectoderm (dotted) developed into brain tissue, and (C) left anterolateral epidermis ectoderm (hatched) developed into skin tissue. When the two tissues were cultured together as a sandwich (B), and were both derived from wild-type donors, eyes with pigmented and neural retina and lens developed. The right body side of the donor embryo was left intact to determine the phenotype at hatching.

Stages of wild-type axolotl eye development

Stages 1 to 10 are those of Rabi’s drawings (1898), stages 0 and 11 to 14 are new, having never been published before. Stages 1 to 10 are described in more detail, especially concerning the retina and accessory tissues. Eye development (25 °C) (Fig. 1) is correlated with whole body stages 23 to beyond 48 of Harrison (1969).

Stage 0 (late tail bud, 7 to 11 somites: Harrison stages 23 to 26, 60 to 75 h). Pair of optic pouches (op) on lateroventral presumptive diencephalon wall. Presumptive brain ventricle closing at neural folds (nf).

Stage la (branchial swelling with clefts, 11 to 16 somites: Harrison stages 26 to 29, 70 to 85 h). Pair of optic vesicles (ov) with central optocoel (oc) on lateroventral diencephalon in contact with head epidermis (he). Optocoel continuous with neurocoel (nc) at optic constriction or presumptive optic stalk.

Stage lb (dorsal fin rudiment, 16 to 20 somites: Harrison stages 29 to 32, 80 to 100 h). Optic vesicles expanded dorsad, optocoel flattened by pressure of brain and head epidermis. Optic constriction lengthened to form optic stalk (st). Wall of optic vesicle facing brain flattened to form tapetum nigrum (tn), wall facing head epidermis thickened to form presumptive neural retina (nr). Head epidermis flat, two cell layers thick.

Stage 2 (body axis straight, dorsal fin growing: Harrison stages 32 to 36, 90 to 130 h). Planar lens placode (Ip) formed by thickened internal layer of head epidermis, external layer not thickened. Optocoel completely collapsed by thickened presumptive neural retina, except in lengthened optic stalk.

Stage 3 (branchial branches: Harrison stages 36 to 39,120 to 155 h). Lens placode contracted to form lens hemisphere (Ih). Outer layer of head epidermis not thickened. Periphery of flat optic vesicle contracted and presumptive neural retina concave, engulfing lens hemisphere. Choroid fissure (cf) ventral of optic stalk wide open. Ventral neural retina rudimentary.

Stage 4 (branchial filaments: Harrison stages 39 to 40, 140 to 190h). Lens vesicle (lv) with small central lentocoel (lc) still attached to outer layer of head epidermis or presumptive epicornea. Vitreous chamber (vc) between lens vesicle and neural retina opened. Papilla nervi optici still indistinct. Embryonic pigment granules and yolk platelets in all ocular cell types.

Stage 5 (branchial covers united: Harrison stages 40 to 41,180 to 220h). Primary lens fibres (pf) elongating on retinal side; their pigment granules and yolk platelets disappearing. Lens vesicle still attached to epicornea (ec). Some new totally pigmented cells in dorsal tapetum nigrum. Dorsal neural retina with internal plexiform layer (ip) in central two thirds. Papilla nervi optici (pn) well developed. Retina ventral of optic stalk half the length of dorsal retina. Nick between ciliary epithelium (ce) and neural retina at presumptive ora serrata (Os) . Space between optic cup and epicornea open.

Stage 6 (branchial branches elongated: Harrison stage 41, 200 to 230 h). Lens vesicle with primary lens fibres, detached from one-cell-layer thick epicornea. Lens capsule (c) thickened. Lentocoel maximal. Optic stalk forms sheath around thin optic fibre tract (ot). Dorsal tapetum nigrum completely pigmented; ventral tapetum not pigmented. Ganglion cell layer (gc) two-cell-layers thick. Internal nuclear layer (in) three-cell-layers thick.

Stage 7 (mouth open: Harrison stage 42, 220 to 260 h). Primary lens fibre hillock large, almost filling lentocoel. Nuclei of primary lens fibres degenerating, basophilic. Neural retina with rods (r) and cones. Internal plexiform layer complete in dorsal retina, external plexiform layer (ep) irregular and thin. Nerve fibre layer still indistinct. Ciliary epithelium about five cells long in cross section; external layer pigmented, internal layer not pigmented. Internal limiting membrane (il), Bruch’s membrane (Br), and Bowman’s membrane (Bo) thin. Ciliary stromal fibroblasts, iridophores, and xanthophores present. Epicornea one- to two-cell-layers thick. Sclera (5) cells on external side of tapetum nigrum. Sclerocorneal junction (sj) very thin, anterior chamber (ac) between epicornea and lens still partially open.

Stage 8 (hatched: Harrison stages 42 to 43,250 to 330h = 10 to 14 days). Lentocoel collapsed; lens epithelium (le) in direct contact with lens fibre core. Nuclei in primary lens fibres lost. Nuclei in secondary lens fibres (sf) degenerating, basophilic, except in subequatorial zone. Ventral tapetum nigrum now also completely pigmented. Ventral retina two thirds as long as dorsal retina when viewed in cross section. Central portion of ganglion cell layer flattened, one-cell-layer thick. External plexiform layer distinct. Sclerocorneal junction complete, although still very thin. Anterior chamber closed.

Stage 9 (forelimb stump with up to three digits: Harrison stages 43 to 45, 12 to 23 days). Lens epithelium flattened. Lens fibre orientation spherical. Internal plexiform layer (ip) complete in ventral retina. Ventral retina as long as dorsal retina in cross section. Nuclei of outer nuclear layer (on) of neural retina elongating and aligned. Iris (i) rudiment one to three cells long in cross section.

Stage 10 (shoulder and elbow flexible, hind limb stump: Harrison stages 45 to 46, 20 to 40 days). Lens bigger. Nerve fibre layer (nf) of neural retina distinct. Ganglion cell layer one-cell-layer thick. Nuclei of outer nuclear layer 1-5 times as long as wide. Ora serrata constricted. Ciliary (cs) and iris (is) stromata irregular or missing.

Stage 11 (hind limb with three digits, knees may be flexible: Harrison stages 47 to 48, 35 to 80 days). Nuclei in secondary lens fibres largely lost, except in sub-equatorial zone. Ciliary and lens epithelia flattened. Iris still only three cells long in cross section. Epicornea still only one- to two-cell-layers thick. Bowman’s and Bruch’s membranes thin. Dorsal sclera with orbital cartilage (ca), although scleral cells still sparse and scattered. Canal of Schlemm missing, and choroid (ch) still consisting only of a loose net of capillaries.

Stage 12 (larva 8 to 12cm long: Harrison stages ?, 50 to 150 days). Nuclei of lens epithelium at most 1·5 times flatter than high. Secondary lens fibres flattened and oriented spherically. Zonule of Zinn (Zi) weak, enclosing new posterior chamber (pc) between iris and enlarged vitreous chamber (vc). Iris in cross section elongated, about six cells long. Ciliary and iris stromata completely coat ciliary and iris (ie) epithelia on their external sides. Stromal capillaries present. Trabecular mesh (tm) in corneociliary angle present, but canal of Schlemm possibly absent. Capillaries of choroid condensed, one-capillary-layer thick, unpigmented. Tensor choroidae inside of thin sclerocorneal junction indistinct. Bruch’s membrane thicker. Retinal internal limiting membrane distinct. Retroorbital muscles (rm) proximal to orbital cartilage and sclera (s). Cartilage still incomplete on ventral sclera. Sclera with three flat cell strata. Epicornea with three cell layers. Bowman’s membrane thick, coated on its internal side with three strata of corneal stroma (cos) cells. Eyelids absent, although small rudiment of Harderian gland (Ha) appearing at dorsal presumptive conjunctiva.

Stage 13 (juvenile 20 to 25cm long: Harrison stages ?, 300 to 400 days). Lens epithelial nuclei three to four times flatter than wide. Secondary lens fibre cell boundaries indistinct. Lens slightly wider than vitreous chamber. Zonule of Zinn still weak, but more extensive. Nerve fibre layer of neural retina thick near papilla nervi optici. Nuclei of internal nuclear layer three-cell-layers spanning. Rod cells of outer nuclear layer and photoreceptor layer (pr) with rod and cone outer segments three times as long as their nuclei. Cone and possibly double cone cells more internal in these layers. Melanophores frequent in iris and ciliary stromata.

Tensor choroidae muscle (te) weak. Orbital cartilage complete on ventral sciera, about three-cell-layers thick. External cell layer of epicornea flattened. Corneal stroma five-cell-strata thick. Eyelid (I) rudiment on dorsal side. Harderian gland with few large acini.

Stage 14 (adult 30 to 35 cm long: Harrison stages ?, 7 years at 18°C). Lens twice as wide as liquid-filled vitreous chamber. Zonule of Zinn strong, with several in parttorn fibre sheets. Nerve fibre layer thick in most parts of neural retina. Internal layer of ciliary epithelium unpigmented, the external pigmented. Ciliary caecae absent. Choroid still one-capillary-layer thick. Trabecular mesh in corneociliary angle extensive both dorsally and ventrally. Tensor choroidae ventrally strong. Sclera external of cartilage about five-cell-strata thick. Epicornea four-cell-layers thick. Corneal endothelium and Descemet’s membrane absent. Dorsal eyelid only slightly enlarged, ventral eyelid missing. Dorsal Harderian gland branched with many acini. Conjunctiva distinct dorsally.

Eye development in mutant e (eyeless)

In axolotls homozygous for gene e (nonlethal, recessive) retina development is arrested at the early optic vesicle stage (Rabl stage la) (Fig. 2A), although sporadic cell divisions are still observed later. At hatching, an irregular cell mass (abnormal stages lb to 4) (Fig. 2B) represents the neural retina, and the pigmented retina is represented by a few pigmented cells. In rare cases a pigmented retinal epithelium (stage 6) persists until hatching (Fig. 2C). Small normally shaped lenses (stage 6) (Fig. 2B) may form. They are not stimulated to grow by the retina and are pushed aside by the immigrating head mesenchyme. These lenses may thus become located close to the head epidermis or close to the retinal rudiment (Fig. 2B). Both retinal rudiments and lenses disintegrate and disappear (Fig. 2D) during larval life. In axolotls heterozygous for both e and r (renal insufficiency, lethal, recessive) (Humphrey, 1964) eyes develop to stage 14 (Fig. 1); however, often several optic vesicles develop into a stack of retinal cups (Fig. 1), whereby the lens may become dislocated (Epp, 1978).

Eye development in mutant mi (microphthalmic)

Early eye development in axolotls homozygous for the mutation mi (lethal, recessive) is indistinguishable from that in the wild type almost up to hatching (Rabl stage 8). Cells in the late embryonic to larval eye (Rabl stages 7 to 9), some brain areas, the gill branches and filaments begin to die, and cell growth and mitosis are halted. At that stage the eyes look dwarfed and crippled. The eye cup is often too small to harbour the lens behind the iris (Fig. 3). Cell differentiation is not impeded directly, and lens fibre cells, retinal photoreceptor cells with rod outer segments, retinal pigment epithelial cells, retinal ganglion cells with axons in the optic nerve are all present in reduced numbers. Scleral cartilage, corneal stroma, iris stroma, eyelid and Harderian gland never develop, because the larva dies before that stage, usually within 1 to 2 weeks after hatching. The larva swims but does not feed.

Necessity for mutual head epidermis-neural plate stimulation

When wild-type anterolateral neural plate neuroectoderm or head epidermis from stage-13 early neurula were cultured for 9 days in isolation in serum-free medium, they never developed retinas or lenses in several hundred cases examined. Instead, neural plates developed only into brain tissue with nerve tracts, small dense nuclei and still remaining yolk platelets. Head epidermis formed only skin (Figs 5, 6B,C) with intracellular slime vacuoles, large nuclei, and a thick outer cell surface. When those two tissues were cultured together, however, eyes developed in almost all cases (Figs 5, 6A) in addition to brain and skin tissue. Retinas with dense nuclei but without remaining yolk platelets, and often with rod outer segments, and lenses with a core of enucleated lens fibres both reached up to Rabl stage 8. All tissue explants had some mesodermal cells, which could not be removed. Lens development was never observed in the absence of the retina, while retina development frequently occurred in the absence of a lens. It may be concluded that retina development depends on the presence of an epidermal component, and lens development in turn depends on a retinal factor.

Fig. 5.

Frequency of eye stages of neural retina, pigmented retina, and lens in explants of wild-type anterolateral neural plate and, or, anterolateral head epidermis after 9 days of culture. See Fig. 4 for experimental design. Histograms on line 1 exhibit results with wild-type embryos from a heterozygous spawning (same as in Figs 6, 9), while those on line 2 are from a pure wild-type spawning (same as in Figs 7, 8). The vertical lines separate the unidentifiable negative cases (—) on the left from the positive cases (1 to 9) on the right in each histogram. Stages 1 to 3 are hard to recognize in culture. The total of cases examined is given on the left, the percentage positive is on the lower left, and the level of significance by the χ2 test of independence appears on the lower right of the histograms. The low level of significance (P<0·10) of lens frequency in the wild-type sandwiches is caused by the low number of control tissues in the heterozygous spawning: four wild-type neural plates and four wild-type epidermis, cultured alone were all negative for eye tissues.

Fig. 5.

Frequency of eye stages of neural retina, pigmented retina, and lens in explants of wild-type anterolateral neural plate and, or, anterolateral head epidermis after 9 days of culture. See Fig. 4 for experimental design. Histograms on line 1 exhibit results with wild-type embryos from a heterozygous spawning (same as in Figs 6, 9), while those on line 2 are from a pure wild-type spawning (same as in Figs 7, 8). The vertical lines separate the unidentifiable negative cases (—) on the left from the positive cases (1 to 9) on the right in each histogram. Stages 1 to 3 are hard to recognize in culture. The total of cases examined is given on the left, the percentage positive is on the lower left, and the level of significance by the χ2 test of independence appears on the lower right of the histograms. The low level of significance (P<0·10) of lens frequency in the wild-type sandwiches is caused by the low number of control tissues in the heterozygous spawning: four wild-type neural plates and four wild-type epidermis, cultured alone were all negative for eye tissues.

Fig. 6.

Mutant e and wild-type tissue explants after 9 days of culture. (A) Sandwich of wild-type neural plate and wild-type epidermis showing differentiation of an eye with pigmented retina (tn) (stage 8), neural retina (nr) (stage 7) with rod outer segments (r), and a lens with lens epithelium (le) and secondary lens fibres (sf) (stage 8). Brain tissue in the upper left, skin tissue in the lower right, and some mesodermal cells between eye and skin. (B) Wild-type neural plate cultured alone developed into brain tissue only. (C) Wild-type epidermis cultured alone formed skin. A few mesodermal cells are apparent in the centre. (D) Sandwich of wild-type neural plate and gene e epidermis with pigmented retina (tn) (stage 6), neural retina (nr) (stage 6) and lens vesicle (Iv) (stage 4). Brain tissue just above pigmented retina and skin in the upper left area. (E) Sandwich of gene e neural plate and wild-type epidermis without eye tissues, but with brain tissue in upper right, skin in lower left, and some mesodermal cells in the centre. (F) Sandwich of gene e neural plate and gene e epidermis. Eye tissues wanting, brain tissue on the left, and skin tissue on the right. Scale bars, 100μm.

Fig. 6.

Mutant e and wild-type tissue explants after 9 days of culture. (A) Sandwich of wild-type neural plate and wild-type epidermis showing differentiation of an eye with pigmented retina (tn) (stage 8), neural retina (nr) (stage 7) with rod outer segments (r), and a lens with lens epithelium (le) and secondary lens fibres (sf) (stage 8). Brain tissue in the upper left, skin tissue in the lower right, and some mesodermal cells between eye and skin. (B) Wild-type neural plate cultured alone developed into brain tissue only. (C) Wild-type epidermis cultured alone formed skin. A few mesodermal cells are apparent in the centre. (D) Sandwich of wild-type neural plate and gene e epidermis with pigmented retina (tn) (stage 6), neural retina (nr) (stage 6) and lens vesicle (Iv) (stage 4). Brain tissue just above pigmented retina and skin in the upper left area. (E) Sandwich of gene e neural plate and wild-type epidermis without eye tissues, but with brain tissue in upper right, skin in lower left, and some mesodermal cells in the centre. (F) Sandwich of gene e neural plate and gene e epidermis. Eye tissues wanting, brain tissue on the left, and skin tissue on the right. Scale bars, 100μm.

Bovine cornea extracts stimulate axolotl retina development

Embryonic axolotl head epidermis is too small for growth factor extraction. Therefore, an adult head epidermis derivative, the bovine cornea was used.

Neural plates from wild-type axolotls, when cultured in the presence of non-dialysable bovine cornea extract (3000μg ml−1), formed neural retinas (stage 6) preferentially (Figs 7, 8B). At a lower concentration (30μgml−1), the differentiation of pigmented retinas (stage 6) was prevalent (Figs 7, 8A). Overall retina development is relatively infrequent in the presence of corneal extract. However, in several hundred cases, retina development was never observed in control neural plates cultured alone. The results suggest that head epidermis derivatives, such as the cornea, produce a macromolecular factor which stimulates retinal differentiation from wild-type neural plates.

Fig. 7.

Frequency of eye stages of neural retina, pigmented retina, and lens in wildtype neural plate explants cultured in the presence of aqueous bovine cornea extracts. See Fig. 4 for experimental design, and Fig. 5 for explanation of the histograms. Controls are shown in Fig. 5.

Fig. 7.

Frequency of eye stages of neural retina, pigmented retina, and lens in wildtype neural plate explants cultured in the presence of aqueous bovine cornea extracts. See Fig. 4 for experimental design, and Fig. 5 for explanation of the histograms. Controls are shown in Fig. 5.

Fig. 8.

Wild-type neural plate explants after 9 days of culture in the presence of aqueous bovine cornea extracts. (A) In the presence of 30μgml−1 of extract, a pigmented retina (tn) (stage 6) with visible basement membrane of Bruch (arrows) developed. The surrounding tissue appears to be brain-like. (B) In the presence of 3000 μg ml−1 of extract a neural retina (nr) (stage 5) (arrows) formed, recognizable by the early disappearance of yolk platelets and the plexiform nerve fibres in the centre. Brain tissue occupies the centre and right side. Scale bars, 100 μM.

Fig. 8.

Wild-type neural plate explants after 9 days of culture in the presence of aqueous bovine cornea extracts. (A) In the presence of 30μgml−1 of extract, a pigmented retina (tn) (stage 6) with visible basement membrane of Bruch (arrows) developed. The surrounding tissue appears to be brain-like. (B) In the presence of 3000 μg ml−1 of extract a neural retina (nr) (stage 5) (arrows) formed, recognizable by the early disappearance of yolk platelets and the plexiform nerve fibres in the centre. Brain tissue occupies the centre and right side. Scale bars, 100 μM.

Fig. 9.

Frequency of eye stages of neural retina, pigmented retina, and lens in tissue sandwiches of anterolateral neural plate and anterolateral epidermis from donors of a gene e heterozygous spawning. Explants were either phenotypically wild type or mutant e. See Fig. 4 for experimental design, and Fig. 5 for explanation of the histograms. The probability of an e-e tissue combination is 1/16, for e-E and E-e 3/16, and for E-E 9/16 (see Fig. 5, line 1).

Fig. 9.

Frequency of eye stages of neural retina, pigmented retina, and lens in tissue sandwiches of anterolateral neural plate and anterolateral epidermis from donors of a gene e heterozygous spawning. Explants were either phenotypically wild type or mutant e. See Fig. 4 for experimental design, and Fig. 5 for explanation of the histograms. The probability of an e-e tissue combination is 1/16, for e-E and E-e 3/16, and for E-E 9/16 (see Fig. 5, line 1).

Primary failure in mutant e neural plate and retina

In heterozygous spawnings, embryos homozygous for gene e have a frequency of 1/4, and the probability of an e-e tissue combination in sandwich cultures is thus only 1/16. In the rare cases where both anterolateral neural plates and head epidermis were derived from donors homozygous for gene e, only brain and skin tissues differentiated (Figs 6F, 9). This same result was obtained in several repetitions of this experiment. All combinations of wild-type neural plates with homozygous gene e epidermis developed a retina, reaching up to Rabl stage 8. However, lenses formed less frequently, reaching stage 4 (Figs 6D, 9). When homozygous gene e neural plates and wild-type epidermis were cultured together, retinas attaining stage 5 and lens vesicles reaching stage 4 developed very rarely (Figs 6E, 9). These results suggest that homozygous gene e epidermis can stimulate wild-type retina development, while retinal stem cells in homozygous mutant e neural plates exhibit an intrinsic failure in the response to the epidermal stimulation. Lens development appears to be impeded secondarily due to the primary failure in the retina.

Normal eye development in the axolotl has been redescribed in more detail in this paper, in order to be able to assess the developmental defect in mutations that focus on the eye. As eyes of other anamniotic vertebrates, the axolotl eye lacks a retinal fovea and supraretinal blood vessels. In urodeles, the corneal endothelium and Descemet’s membrane are absent. The lens is spherical and the retinal photoreceptors are more diverse and complex (Reyer, 1977) than those of mammals. In the axolotl, the choroid remains very thin and unpigmented, while in newts (genus Notophthalmus) it is pigmented and several capillary layers thick. Neither anurans nor urodeles have a vitreous body in their vitreous chamber. In the axolotl, a scleral cartilage protects the protruding eyeball, while adult newts have no cartilage in their sclera. On the other hand, newts have well-developed dorsal and ventral eyelids, while axolotls only have a dorsal eyelid rudiment. Nictitating membranes, while present in frogs (genus Rana), are absent in urodeles. Embryonic eye tissues are laden with yolk platelets in lungfish, urodeles and anurans, while they are devoid of yolk platelets in mammals and birds.

The mutation mi (Humphrey & Chung, 1977) is expressed late during axolotl eye development (stage 8), and also affects other head tissues. In contrast, mutation e is already expressed at the beginning of eye development (stage la) in the optic vesicle and apparently in some cells of the hypothalamus (Van Deusen, 1971, 1973). Disorganized mitotic divisions still continue at a reduced rate (Ulshafer & Hibbard, 1976,1979) in later stages, and some pigmented retinal cells may differentiate (Van Deusen, 1973) in rudimentary optic vesicle cell masses (Brun, 1983, 1985). A very similar eyeless mutation is also known in the rat, except that homozygous animals are fertile (Kinney et al. 1982).

Van Deusen (1973) proposed that in the axolotl e mutation the retina- and hypothalamus-forming neuroectoderm regions have an intrinsic defect. If this hypothesis is correct, the insufficient production of retina-derived growth factors would arrest lens development as a direct consequence (Cuny & Malacinski, 1983). Other accessory tissues, such as cornea, scleral cartilage, Harderian gland, and sclera, never develop in the mutant e, probably also because they would depend on the retinal growth factors. While the preoptic nuclei in the brain are histologically normal (Eagleson & Malacinski, 1983) in the mutant e, the superficial optic layer of the tectal neuropil is hardly developed (Gruberg & Harris, 1981; Harris, 1982, 1983). The mutant e tectum develops normally, however, and full vision is restored (Epp, 1972; Hibbard, Ulshafer & Ornberg, 1975; Hibbard & Ornberg, 1976; Schwenk & Hibbard, 1977; Harris, 1981), if a wild-type eye is implanted in embryos homozygous for gene e. This demonstrates that the mutant e optic tectum is not genetically defective, and only suffers from a lack of retinal innervation.

Implantation of a wild-type eye causes skin melanocytes to contract in white (d) strain (Keller, Löfberg & Spieth, 1982) homozygous carriers of mutation e, while parabiosis causes a spreading of melanocytes in d strain axolotls, if they are connected with an axolotl homozygous for gene e (Epp, 1972). These data suggest that the hypothalamic melanocyte-inhibiting factor is not secreted in sufficient amounts, which causes an overproduction of pituitary melanocyte-stimulating hormone (MSH) (Epp, 1972,1978). The pineal gland (Brick, 1962) and the retinal photoreceptors (Gern, Owens & Ralph, 1978) both secrete melatonin, which also stimulates skin melanocyte contraction. At any rate, the effect on skin pigmentation of mutant e seems to be a direct consequence of the eyeless condition.

The sterility (Humphrey, 1969) of axolotls homozygous for gene e is apparently due to insufficient secretion (Van Deusen, 1973) of hypothalamic gonadotropin-releasing hormone (Kubo, Watanabe, Ibata & Sano, 1979). While the implantation of a wild-type eye has no effect on mutant gonads, the implantation of a wild-type hypothalamus region in a gene e embryo restores fertility (Van Deusen, 1973; Harris, 1983). The hypothalamic cells develop adjacent to the retinal cells in the neural plate, and may thus belong to the same stem cell lineage. From the presented evidence one may assume that all pleiotropic effects of the mutation e are consequences of the primary genetic defect in the retina area, with perhaps the exception of some hypothalamic cells.

It remained still unclear, however, whether the retinal tissue does not develop, because the interacting tissues in the surroundings of the neural plate cells do not produce enough retina-stimulating growth factors. Tissue-grafting experiments (Van Deusen, 1971, 1973) between homozygous e and wild-type gastrulae have shown that the mutant e prechordal mesoderm can sustain normal eye development in wild-type embryos, while wild-type prechordal mesoderm cannot restore eye development in mutant e embryos. Hence, the putative mesoderm-derived neuralizing factor (Tiedemann & Born, 1978) is probably normal in the mutant. We found that in the mutant e paired optic vesicle rudiments always formed, which made contact with the head epidermis (Fig. 2A). This rules out a primary barrier function of the neural crest head mesenchyme in this mutant. Brun (1978, 1980) suggested that retina development might fail, because of a failure in the overlying mutant e head epidermis. It is known that retina development from cultured wild-type neural plate tissue depends on the presence of head epidermis (Filatow, 1926; Rollhäuser-Terhorst, 1981). The nature of this tissue interaction has been unknown however. Understanding the nature of this tissue interaction is crucial for the investigation of the failure in the mutant e retinal region. Unfortunately, embryonic axolotl head epidermis is too small for biochemical extractions, and we therefore resorted to extracts of adult bovine corneas, which are head epidermis derivatives. Dialysed cornea extracts stimulate development of pigmented or neural retinas, depending on the extract concentration, in cultured wild-type neural plates. In the embryo, the neural retina directly faces the head epidermis and thus receives a higher concentration than the pigmented retina behind it. The results suggest that the head epidermis stimulates retina development via a macro-molecular growth factor. In our cultures, mutant e head epidermis was fully capable of wild-type retina stimulation, while mutant e neural plates hardly developed retinas, even in the presence of wild-type epidermis. This demonstrates that the mutant e neural plate and retina are intrinsically defective, and cannot be rescued by the head-epidermis-derived growth factor.

Knowing that the presumptive retina cells are the primary target of the mutation e, we would like to know which molecular system is disrupted in these cells. At this point we can only offer some speculations. Ulshafer & Hibbard (1979) argued that the observed premature thickening and maturation of the basal lamina, that surrounds the mutant e optic vesicles, could suppress eye development. The composition of basal laminae modifies the cellular responsiveness to growth factors (Gospodarowicz, Gonzales & Fuji, 1983). Basal laminae can act as filters or adsorbants for macromolecules. Wild-type retina development can also be prevented by intracellular injection of antibodies directed against gap junction proteins (Warner, Guthrie & Gilula, 1982). Therefore, a reduction or malfunctioning of gap junctions in the mutant e presumptive retinal region could be the primary reason for the eyeless condition. A third possibility would be that a shift in embryonic to retina-specific gene expression takes place during retina development, involving two isoforms of proteins needed for cell proliferation. If the retina-specific isoform is malfunctioning or unstable, this could lead to a decrease in mitotic activity of the retina, and eventually to eyelessness. These ideas may help to design experiments that will elucidate the primary cause of eyelessness in the mutant e of the axolotl.

We thank Françoise Briggs and Leah Dvořak (Indiana University Axolotl Colony) for the heterozygous spawnings of gene e axolotls. Hervé Coët helped with the photo-reproduction. Support for these studies was provided by the Swiss National Fund for Scientific Research and the US National Science Foundation PC 83-15899.

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