Patterning the Vertebrate Neuraxismateriais.dbio.uevora.pt/BD/Morfogenese/segment_encefalo.pdf · 2020. 5. 19. · plane of the ectoderm (7, 8). Whereas the molecular identity of

Patterning the Vertebrate Neuraxis

Andrew Lumsden and Robb Krumlauf

Neuraxial patterning is a continuous process that extends over a protracted period of development. During gastrulation a crude anteroposterior pattern, detectable by molecular markers, is conferred on the neuroectoderm by signals from the endomesoderm that are largely inseparable from those of neural induction itself. This coarse-grained pattern is subsequently reinforced and refined by diverse, locally acting mechanisms. Segmentation and long-range signaling from organizing centers are prominent among the emerging principles governing regional pattern.

1 he central nervous system (CNS) arises from the neural plate, a cytologically homogeneous sheet of epithelial cells that forms the dorsal surface of the gastrula-stage embryo. The peripheral nervous system arises from ectodermal placodes and neural crest cells that form at the lateral fringes of the plate. The neural plate subsequently rolls up on its anteroposterior (AP) axis to form a tube, the expanded anterior end of which then partitions into a series of vesicles representing the anlagen of fore-, mid-, and hindbrain. Posteriorly, the long, uniformly narrow tube forms the spinal cord. These early morphological features of the neuraxis, accompanied by position-specific expression of developmental control genes, dictate the overall plan of the CNS and predict its regional specializations. Within each region, a large diversity of neuronal cell types is then generated, each with distinct identities in terms of morphology, ax-onal trajectory, synaptic specificities, neurotransmitters, and so on. The intricate spatial order of differentiated neurons, essential to the subsequent formation of functional circuits, is crucially dependent on correct regional specification.

Signals from adjacent tissues are involved at all stages of neuraxial patterning. Neural-inducing factors and modifiers produced during gastrulation by the (endo)mesoderm establish an initial crude AP pattern in the overlying neural plate. Although the precise nature of this early patterning information remains unclear, the inductive signals that confer forebrain identity appear to differ qualitatively from those that operate more posteriorly. The coarse-grained pattern that emerges at the end of gastrulation

A. Lumsden is in the Medical Research Council Brain Development Programme, Department of Developmental Neurobiology, United Medical and Dental Schools, Guy's Hospital, London SE1 9RT, UK. E-mail: [email protected]. R. Krumlauf is in the Division of Developmental Neurobiology, National Institute for Medical Research, The Ridgeway, Mill Hill, London NW17 1AA, UK. E-mail: [email protected]

is progressively refined, resulting in a precise regional variation in cell identity.

Patterning of cell types appears to be organized on a Cartesian grid of positional information, the coordinates of which correspond with the AP and dorsoventral (DV) axes of the neural tube: analyses of cell fate after experimental rotation of the neural plate (1,2) have indicated that regional fate is determined along the AP axis before and independently of fate restriction on the DV axis. We will confine our discussion to the assignment of AP regional identity, as patterning on the DV axis has been comprehensively reviewed elsewhere (3, 4). We will focus on two well-studied regions, the hindbrain and the midbrain, to illustrate distinct but not mutually exclusive modes of local patterning: segmentation plays a prominent role in the hindbrain, whereas a discrete signaling region at the isthmus sets up the AP polarity of the midbrain. The fore-brain has been less intensively studied, but interesting parallels with hindbrain development are considered.

Early Role of the Mesoderm in Regionalization

Investigation of the earliest developmental events has focused on the amphibian embryo, in which it was first seen that neural fate is imparted to competent ectoderm by signals emanating from the dorsal blastopore lip. Spemann noted that the early dorsal lip, grafted heterotopically, could induce an entire neuraxis but the later dorsal lip could induce only posterior CNS (5). Although the details of how the dorsal lip (Spemann's organizer) confers polarity on the neuraxis are still unknown, emphasis has been placed on the posterior-to-anterior (P-to-A) progression of mesoderm involution during gastrulation and a two-step, activation-transformation action of its signals on the dorsal ectoderm. Activating signals from early-involuting mesoderm are

thought to induce a default state of anterior neural differentiation, which is then modified to a more posterior character by transforming signals from later-involuting mesoderm (6). The level of transforming signal impinging on any one level of the neuraxis might confer its local identity (Fig. 1). Inducing signals could pass to the ectoderm either vertically, from underlying cells, or tangentially, from organizer cells still in the plane of the ectoderm (7, 8).

Whereas the molecular identity of activating signals remains uncertain, three secreted proteins expressed by the organizer— follistatin, noggin, and chordin—are capable of inducing the expression of anterior neural plate markers in naive ectodermal cells (9). These candidate activators share no obvious structural features, but each can bind directly to and antagonize the actions of members of the-BMP family of signaling molecules (10, 11). Induction of anterior neural plate may thus involve inhibition of the neural inhibitors BMP2 and BMP4, which are present in the presumptive neurectoderm.

Candidate transforming signals include basic fibroblast growth factor (bFGF) (12) and retinoic acid (RA) (13), both of which can posteriorize anterior neural tissue but have little neuralizing capacity on their own. However, signaling through the bFGF receptor is not necessary for this process; when blocked in transgenic Xeno-pus embryos by the expression of a truncated, dominant-negative form of FGFR1, the posterior CNS forms normally (14). Strongly indicating a posteriorizing role for RA, however, the expression of a con-stitutively active retinoid receptor results in a posteriorized axis, whereas dominant-negative retinoid receptor expression results in an anteriorized axis (15). The posteriorizing role of RA will be considered again in the context of local patterning in the hindbrain.

In addition to signals that may influence pattern along the entire neuraxis, there is also evidence for head-specific induction pathways. The Lim-1 and Otx-2 homeobox genes are expressed in the organizer region (Hensen's node of amniote embryos), and their inactivation in mice deletes all head structures, including prosencephalon, mesencephalon, and anterior hindbrain, whereas the posterior hindbrain and spinal cord are unaltered (16-18). The requirement for these genes appears to distinguish early organizer functions from later ones. Both genes are later expressed in the prechordal plate (16, 19), emphasizing the importance of this structure in endowing the overlying forebrain with unique characteristics. A recently identified secreted protein, cerberus, has potent forebrain-inducing activity (20).

SCIENCE • VOL. 274 • 15 NOVEMBER 1996 1109

mailto:[email protected]

mailto:[email protected]

Cerberus is abundantly expressed in the deep-layer cells of the organizer that con- stitute the leading edge of the gastrulating endomesoderm. Both the maintained expression and inducing activity of cerberus would seem to depend on coexpression of other organizer factors. It seems likely that in vivo forebrain-inducing activity lies in the prechordal plate and the endomesoderm immediately anterior to it, where expression of cerberus and chordin overlap (20). The different inductive activities of early and later dorsal lips, first recognized by Spemann, are now being dissected at molecular and genetic levels; it appears that the acquisition of coarse-grained pattern along the neuraxis is controlled by mechanisms that differ between the anterior-most (prechordal) and the more posterior (epi- chordal) regions of the neuraxis.

Hind brain-* Segmented Region of the Neuraxis

Pronounced axial variation involving a comparatively small repertoire of cell types makes the hindbrain an attractive and ac- cessible system for the study of local CNS pattern. Furthermore, early development of the hindbrain is characterized by metamer- ism, suggesting the early allocation of defined sets of precursor cells and the existence of precise boundaries to both cellular assemblies and realms of gene action. In the chick embryo, the segmented pattern of the hindbrain emerges upon neural tube closure as a series of bulges-rhombomeres-and is virtually complete at the onset of neurogenesis. Segmentation of the vertebrate hindbrain bears a superficial resemblance to segmentation of the Drosophila embryo: rhombomeres form by internal subdivision rather than by budding from a growth zone, and

they have a pair-wise organization (21 ). Neuronal pattern. Two patterns of

metameric cellular organization can be dis- tinguished in the hindbrain, one involving neurons of the reticular formation and the other involving motor neurons (Fig. 2). Eight identified types of reticular neuron are repeated through sequential rhombomeres such that each contains a more or less complete set (22). Motor neurons also develop in each rhombomere, but have rhombomere-specific identities (23, 24). The segmental disposition of branchiomotor nuclei in the early hindbrain has a close anatomical (23) and functional (25) correspondence with target structures associated with the segmented series of branchial arches that lie beneath it. Later in develo~ment. the segmental origins of these cells become obscured as certain reticular cells become more numerous in particular rhombomeres (26) and the motor nuclei condense and migrate to new positions. Fate-mapping studies have also revealed metameric origins for the adult sensory nuclei (27). In the hindbrain, segmentation is involved in specifying the pattern of developing structures, but not in deploying them in the adult.

Compamt- l ike properties of rhombomeres . Developmental compartments provide a way of allocating blocks of cells with distinct properties (28). The contain- ment of polyclonal assemblages of neuroep- ithelial cells within rhombomeres has been shown by lineage-tracing studies in chick (29). Compartmental restriction of cell mingling persists while the epithelium is predominantly germinative (30); later, young neurons may escape the restriction once they have acquired their ultimate positional specification. Rhombomeric domains of the germinative (ventricular) zone

Fig. 1. Acquisition of AP pattem during neural induction in Xenopus. In the late gas- tlula, shown in hemisection, involuted cells have reached the anterior pole of the presumptive CNS. Radial signals (white arrows) from the leading-edge endoderm (yellow) and the mesoderm (orange) induce neural fate in the overlying ectoderm (blue). Forebrain (dark pink) is induced by leading-edge endoderm and mesoderm. More posterior levels of the ectoderm are activated (light pink) and transformed by a graded posteriorizing a c t i i (green). The yellow arrow shows the route of planar signals.

Transformation \

Posterior 1

Dorsal mesoderm

Dorsal lip of blastopore

Antenor

Endoderm /

remain lineage-restricted up to late stages, when neurogenesis is nearing completion (31 1.

Rhombomeres partition from one another according to an adhesion differential that displays a two-segment repeat (Fig. 3) (32) Consistent with an expected tendency of neighboring cell groups to separate, enlarged intercellular space is the earliest specialization of rhombomere boundaries (33). Complementing this adhesive differential is an alternating periodicity to the expression domains of the Eph-like receptor tyrosine kinases and their ligands (Fig. 4) (34-36). Perturbing Rtk-1 (Sek-1) function in zebrafish and Xenopus embryos by expression of a dominant-negative form of the mouse Sek-1 receptor results in failure to establish sharp inter-rhombomere boundaries (37). These ligand-receptor partners may thus mediate repulsive interactions that serve to

Fig. 2. Pattems of cell organization in the 3-day embryonic chick hindbrain. Superimposed on the rhombomere pattem (rl to r7) are the reticular neurons (left side) and the motor neurons (right side). Reticular neurons are classified (and colored blue or green) according to axonal trajectory. Mo- tor neurons (in the right side basal plate, B) are classified as somatomotor (yellow), Innervating extrinsic eye muscles (IV, troclear; VI, abducens); branchiomotor (orange), innervating branchial muscles in the first arch (V, trigeminal), second arch (WI, facial), and thlrd arch (IX, glossopharyn- geal); and vestibuloacoustic efferents (red), which share the Vllth nerve exit point (dotted circle) with the faclal motor neurons in the alar plate (A). F, floor plate.

SCIENCE VOL. 274 15 NOVEMBER 1996

sharpen rhombomere borders. They may also provide a potential mechanism where- by cells in adjacent rhombomeres interact with each other to establish additional cell states at the inter-rhombomere boundaries (2 1, 33. 38-4 1 ). Finallv. the inter-rhom- . , . , . bomere boundaries become colonized by axons, perhaps on account of both the local expression of growth-promoting. molecules (23) and the availability of extracellular space (33).

Hox genes enc0d.e positional value along the AP axis. Prime candidates for conferrine " rhombomere identity are the clustered homeobox-containing genes of the Hox family (42), homologs of the HOM-C genes that encode parasegment identity in Drosophila. Expression of genes at the 3' ends of the Hox clusters precedes rhombomere formation and becomes progressively restricted (43) such that expression boundaries coin- cide with the interfaces between rhombomeres (44). Their expression patterns form an ordered and nested set of domains along the neuraxis, with a two-rhombomere periodicity. Superimposed on this pattern are rhombomere-specific variations in expression levels (Fig. 4).

Considering the distribution of transcripts and the general synergy among Hox genes detected in mouse null mutants, it is

the transformation of r2 to an r4 identity (45, 46). However, loss of Hoxa-1 function results in the deletion of r5, reduction of 1-4, and loss of specific neuronal nuclei (42), abnormalities that are not obviously consistent with conferring specific identity on an existing repetitive ground plan; but it remains possible that Hox genes could have dual roles, both in segmentation and segment identification.

Positional values appear to be conferred on rhombomeres by Hox expression, but it is unclear how the Hox eenes become acti- " vated at appropriate levels of the neuraxis. Candidates for this role include kreiskr. a b-Zip member of the c-maf proto-oncogene family (47) expressed in r5 and r6, and Krox-20, a zinc finger gene that is expressed in two stripes in the neural plate that become 1-3 and r5 (48). In kr-1- mouse embryos, the neural tube posterior to the r3/r4 boundary appears unsegmented, a defect that is attributable to the loss of 1-5 and r6 as identifiable territories (49, 50). Targeted disruption of Krox-20 results in the elimination of r3 and r5 and the formation of a partially fused r2/r4/r6 territory (51). This phenotype suggests that Krox-20 may be responsible for generating single-compart- ment periodicity from cues established by

possible that the identity of individual rhombomeres could be defined by the cooperative action of Hox proteins (42). They may also have singular effects: ectopic expression of Hoxa-1, for example, results in

W d

-1 Stage 8 to 9: Odd and even gene expression

Stage 9 to 10: Cell lineage restriction by differential affinity 1

Stage 13 on: Rhornbornere boundary specialization

Fig. 3. Stages leading to cell compartmentation and rhombomere boundary (yellow) formation in the chick embryo. An adhesion differential between adjacent domains (blue, green) segregates cells at the interfaces. Although the molecular basis of the differential adhesion is unknown. it fol- lows the same two-segment repeat as displayed by Krox-20 and other genes shown in Fig. 4. Mol- ecules with boundary-restricted expression (yellow) include Plzf (39), Fgf-3 (40). vimentin (38), low-PSA-NCAM, and laminin (23). Stages numbers refer to normal chick development.

upstream genes. Absence of the r5 stripe of Krox-20 expression and the more anterior expression of group 4 Hox genes in kr-I- mice is consistent with their regulation by kreiskr, although direct interaction has yet to be shown. In contrast, Krox-20 is a direct modulator of the r3/r5 activity of both Hoxa-2 (52) and Hoxb-2 (53).

A major gap in our understanding of hindbrain segmentation is the lack of candidate segmentation genes. Despite the conserved role of HodHOM genes in specifying segmental identity, the upstream pathway appears not to be conserved. How- ever, segmentation is a generic property of metazoan organization that has evolved many times (54), making it likely that Hod HOM genes have been coupled independently to segmentation.

Retinoid signaling and AP position. In addition to the putative role of kreiskr and Krox-20 in locally regulating Hox expression, RA has strong candidacy as an overall mediator of nested Hox expression, consistent with its posteriorizing effect on CNS regionalization. Excess RA causes both an anterior shift of Hox gene expression and an A-to-P transformation of regional fate (55) that includes the ordered transformation of anterior rhombomeres to a more posterior

kreisler K~ox-20 Sek- 1 Sek-2 Sek-3 Sek-4 Ebk

Elk4 Elf-2 Elk-L3 Hoxa- 1 Hoxb- 1 Hoxa-2 Hoxb-2 Hoxa-3 H~xb-3 Hoxd-3 Hoxd-4 Hoxb-4 Hoxa-4

Fgf-3 follistatin CRABP-1 RARa RARP

Fw. 4. Summary of the correlation of gene expression with specific rtiom- bomeres, compiled from analysis in mouse and chick embryos. (Top) Odd-numbered rhombomeres are indicated in blue and even-numbered rhombomeres in green, with each segment designated as rl to r7. The rod-like notochord and overlying floorplate are indicated in black. The vertical yellow lines indicate the boundaries between rhombomeres. The patterns of gene expression are depicted in arbitrary colors with the darkest dors indicating the highest levels of expression. Related genes are indicated by the same color for convenience: Hox homeobox genes (orange), Eph family tyrosine kinase receptors (blue), Eph receptor ligands (green), retinoid- or signaling-related series (magenta). and earty expressed transcription factors (dark purple).


type (56, 57). Conversely, RA-deficient quail embryos have a small hindbrain, lacking posterior rhombomeres (58). In addition, the suppression of RA signaling by expression of a dominant-negative retinoid receptor also results in anteriorization (15). Furthermore, Hox genes have the molecular machinery for responding directly to retinoid signaling (55).

Consistent with a direct role for the organizer, Hensen's node is a rich source of RA and produces increasing amounts during regression (59); thus, the nested expression of Hox genes could be controlled either by a P-to-A gradient of RA diffusing directly from the node, or by an increasing exposure to RA of cells that pass through the node, in A-to-P succession (60). However, it has yet to be shown that RA normally forms a graded signal of either kind or, indeed, that a gradient is necessary; the activity of retinoids could be locally modified by coactivators and corepressors of retinoid signaling (61 ).

Rhombomere autonomy and plasticity. Transplantation experiments in avian embryos reveal a direct correlation between commitment to rhombomere-specific fate and Hox expression: Grafts of neural plate- stage tissue acquire the complement of Hox transcripts and neuroanatomical features of their new location (62), whereas grafts of emerging rhombomeres maintain both their identity and specific Hox expression (2,63). By contrast, transplantation of rhombomeres into the post-otic region results in the activation of posterior Hox gene expression (64), suggesting that their commitment is not irreversible. However, they cannot easily be shifted from an even- to an odd-numbered fate, suggesting that commitment to "odd" or "even" may be an early step in segmentation.

In addition, the even-numbered rhombomeres appear to influence the fate of odd-numbered rhombomeres, thus providing a secondary mechanism for establishing positional differences. Inter-rhombomere interactions control cell survival in the neural crest of r3 and r5, the maintenance of Krox-20 expression in r3, and the repression of follistatin in r3 (6.5).

Midbrain-the Role of the lsthmic Signaling Region

In the midbrain, beyond the anterior limit of Hox gene expression, local AP pattern is generated within an unsegmented field through the activity of a long-range signaling region, the isthmic constriction at the junction of mesencephalic and rhombence- phalic vesicles (Fig. 5).

Establishment of midbrain polmity by En- grailed. Signals from the isthmus regulate expression of two Engraikd genes (66) in a gradient that decreases both anteriorly, through the mesencephalic vesicle, and posteriorly, through rl (Fig. 5). Knockout experiments have shown that En-1 has a critical role in the early specification of the entire region of its expression, whereas En-2 function is restricted to cerebellar morphogenesis (66). However, the En-1 mutant phenotype, agenesis of the tectum (dorsal midbrain) and cerebellum (anterior hindbrain), is completely rescued by insertion of the En-2 complementary DNA into the En-1 locus (66,67), demonstrating that the contrasting phenotypes of En-1 and En-2 mutations reflect differences in the temporal and spatial expression of the respective proteins and not a divergence in their biochemical activity.

En expression is the earliest known marker for mesencephalic polarity, later manifested in a pronounced variation in cytoarchitecture and the acquisition of different sets of afferent inputs from the retina: the wsterior tectum receives axons from the nasal retina, whereas the anterior tectum becomes innervated bv temwral retina. The molecular basis of this discrimina- tion may involve ligands for Eph-related receptor tyrosine kinases, RAGS (68) and ELF-1 (69), that are expressed in decreasing P-to-A gradients-reflecting the earlier pattern of En-and that may function as growth inhibitors of Mek-4 receptor-bear- ing temporal axons.

Trans~lantation studies in avian embw- os have shown that En expression correlates with later morphology. Thus, when the mesencephalic vesicle is reversed on the AP axis at E2, the En gradient readjusts to its

Fig. 5. Early midbrain patterning. In an early neural tube stage embryo. Fgf-8 (green) is expressed in a ring of cells at the isthmus, the constriction between the mesencephalic vesicle (M), and rhombomere 1 (rl). Wnt- 1 (yellow) is expressed in a ring of cells immediately rostra1 to Fgf-8 and along the dorsal midline. Both En- 1 and En-2 (blue) are expressed in gradients that decrease anteriorly and posteriorly from the isthmus. Sonic hedgehog (Shh) expression, at the ventral midline, is shown in red. T, telencephalon; D, diencephalon; SC, spinal cord; N, notochord.

original polarity, and both the graded cytoarchitecture and pattern of retinotectal pro- jections develop normally (66). When reversed at E3, however, the En gradient does not adjust, and both cytoarchitecture and retinotectal projection are subsequently in- verted. This association has been strength- ened by experiments in which En is misexpressed in the anterior tectum through use of a retroviral vector: nasal axons arborize ectopically in the anterior tectum, whereas temporal retinal axons fail even to enter the midbrain (70). Furthermore, the altered retinotectal specificity after En misexpression in the anterior midbrain is associated with ectopic up-regulation of RAGS and ELF-I, defeating their normal P-to-A expression gradient and effectively converting temporal axon-specific anterior tectum into nasal axon-specific posterior tectum (71). Expression of these effector genes, downstream of En, suggests that the normal graded expression of En may polarize the dorsal mesencephalon.

Reguhon of Engrailed expression. Grad- ed mesencephalic expression of En appears to be regulated by signaling from the posterior border of the mesencephalic field. When grafted to the caudal forebrain, the posterior border (isthmus) induces En expression and the formation of a complete optic tectum from the surrounding tissue (72). Two secreted signal molecules, Wnt-1 and FGF8, have been implicated in the isthmic control of En expression. Wnt-1, a homolog of the segment polarity gene wing- less (a regulator of Engrailed in Drosophila), is expressed in the midbrain region of the neural late and later in a ring of cells that " lies just anterior to the isthmus. As for their cognates in flies, Wnt-1 and En expression appears to be mutually interdependent: in Wnt-1 -I- mice, En is expressed normally at first but is then progressively lost (73) along with the dorsal midbrain. Thus, although Wnt-1 is critically involved in the maintenance of En expression, it is not a candidate for inducing En expression or for directly setting up midbrain polarity. However, another secreted factor. FGF8. ex~ressed in a , A

circumferential ring immediately posterior to that of Wnt-I, has midbrain-inducing and -polarizing abilities (74). When a bead coated with recombinant FGF8 is implanted in the posterior diencephalon of chick embryos, expression of Fgf8, Wnt-I, and En-2 is induced in the surrounding cells. These cells later display the character of a complete ectopic midbrain, whose AP polarity is reversed with respect to that of the "host" midbrain. Thus, neuroectodermal Fsf8 expression may be sufficient to establish both midbrain pattern and polarity. Fgf8 is expressed earlier in axial mesoderm cells that lie beneath the presumptive isth-


I ~ I C reg1011 of the ~ leu ra l plate (75, 76 ) anii tha t h,lve the cap'lclty to lniluce En expres- s ~ o n (77, 78) ; meoilermal FGF8 1s thus i~np l~c ;~ te i i ;I:, II homeogenet~c ~niiucer In t h ~ s local cci~ltrol of neural pattern.

O the r Ilkel\- targets of FGF8 are the palreii hox genes, I'ax-2, l'ux-5, and Pas-8, which m,ly he reiluireii, singly or together, tor s p e c ~ f ~ c a t ~ c i n cif the isthmu\. In P u s - 5 ' - nice (79) and zehraf~sh treated ~ v ~ t h func- t ~ o n - b l o c k ~ n ~ ant~hoiiies to p,lx(:f[h]), a prebumeii homolog of Pax-5 (80 ) , the ~ s t h - I ~ L I S IS iieleteii. In the zebraf~sh experiments, the expressLon of both Wnt-1 and En-2 was ,~lso represseil, suggestlng their direct posi- tlve regulat~on b\- p,lx(zf[h]). Inileeil, con- enbus P a x - b ~ n i l ~ n g sites have been iilentl- tied a ~ t h ~ n a n enh'lncer recion of the En-2 gene: when these sltes are mutateii, the r n ~ i l h r a ~ n / h i ~ l i i b r i ~ i ~ ~ iiomain cit reporter expresslon is lost (81 ).

Whereas l s t h ~ r l ~ c grirfts lniiuce tectal ile- velopment in the cauilal iiiencephalon, the same grafts t o the iiorsal hlnilbraln iniiuce cerebellar iie\ elopment (82 ) , ilemonstrat- ing that the competence of rh i~mhence- p h a l ~ c t i s s ~ ~ e to responti to ~ s t h r n ~ c signals il~ffers from that of mesencephal~c anii caudal i l ~ e n c e ~ h a l ~ c regions. FGF8 a l o ~ l e ap- pe'lrs to he i~lsuff ic~ent for inilucing ectoplc En-2 expresslcin or cerehellar iievelopment in the h ~ n i l h r a ~ n (74 ) , ~mplicatlng a i i i i~ t i i~n- al s~cnn l~n i ! molecules at the isthmus.

Although s~gnals from the isthmus are involveii in D,ltternlnc hoth the iior\al mesencephalon and the ilorsal anterlor rho~rlhencelihali)n. the coIlstrlctlon iloes not corre\ponJ precisel\- with the rn~ilbraln/ hiniihr,l~n junctlon. Separating structurally anil f ~ ~ n c t ~ o n a l l \ - i i ~ s t ~ n c t tectal anii isthmo- cerehellar reglons of the brain, this junctlon for~ns s o ~ n e i i~s tance ,Interlor to the con- strlctlon anii registers w ~ t h the posterlor Illnit of Otx-2 expre\slon 111 the early mes- enceph;rl~c vesicle (83) . T h e posterior- most. Otx-2-11ee;rtlve reelon of the vesicle 1s fated to j o ~ n r l and r? In the fc~rmation of the cerebellum (84) . Thus. ~t cannot be i ~ s h ~ ~ ~ n e i i that obv~ous morphological tea- tureh of the ne~lr'll tube. such as the con- t r lcr lons het~veen ves~cles, necessarily cor- reyoni l in ;I preiiict;lble way to f ~ ~ t u r e sub- iiivihions of the brain.

Forebrain-Is Segmentation Involved?

In contrdbt to h~ni lbra ln and miiihrain patterning, \\here restricted patterns of gene e sp res ion have heen tightl\- linkeil e ~ t h e r to segmentation or to the activity of a sig- nallng regLon, our uniierstaniling of earl\- forebra~n p~ltternlng IS virtually li~rliteil t o the gene expres\lon patterns. M(n t notahle among these (85 ) are the Emx, Illx, d11ii

Nkx homeobox genes, the palred hox gene Pax-6, the Lv~ngeii h e l ~ x genes BF-1 and BF-2, the Brachyury homolog Tbr-1, and the secreted factor-encoii~ng gene Wnt-3. Some of these genes are expressed in the ven t r~c~ l l a r zone, suggesting a role in regio11- al specit~cation, whereas the expresslon of others (Dlx and Tbr-1) is restricted to the ~rlantle zone, suggestlng a role In the control of i l~fferent la t io~~. In the former category, Ernx anil Otx genes are expressed in the forebrain and miiihrain In a nested arrax- relrllnlscent of that of the Hox genes more posteriorly, although w ~ t h reverseii A P sym- metr\- (86) .

Largely on the bas15 of i l e sc r~p t~ve molecular stuiiies, it has heen propiix'il that the forehra~n 1s h u ~ l t ~ iecerneal , like the hinilhraln, from a serles of metameric unlts or prci~omeres (87) . Exper~~rlenta l ev~i ience tor compartmentation is l l~r l~te i l to the ill- encephalon \\here cell lineage restriction hounilarles, aligned ~ v i t h prominent axon tracts, i le f~ne four neuromeres (88) . H o ~ v - ever, the sign~ficance of i l lencephal~c neu- rcimeres is brought into questlon by a n anal- y s~s of r e t r o v ~ r a l l ~ nlarkeii c lo~les (89 ) , which has shown that s~b l lng cells can oc- cupy multiple nuclei throughout the A P extent of the iliencephalon. Further anteriorly, in the telencephalon, the patchwork expresslon of putatlve iievelopmental control genes il~spla\-s n o ev~i ience of repeti- tlon, the essence of metarnerlsm. Nor iioes the earl\. cellular oreanization of the telencephalon slipport the notion of a segmental o r~g in ; rather, this reglon appears to he subilivliieii longituiiinally into two subre- glons, the anlagen of cortex (pallium) anii striaturn (9L7). These suhregio~ls express i i~f- ferent regulatory genes (Ems-112, Pax-6, anil Tbr-1 ilorsally; 111s-112 ventrall\-) and appear to be segregated h\- iiifferential aii- h e s ~ o n (91) . W ~ t h i n the dorsal ( co r t~ca l ) subregion, cells nlierate extens~vel\- In the A P i i i r ec t lo~~ , so that clones cross functional bouniiaries anii sibling cells contribute to w~ilely separated structures (92 ) . Supporting the vlew that the telencephalon is a single flelil, ~ v h ~ c h hecomes suhiiiviiieii longituiiinally, BF-1 is expresseii In the prospective telence~ihalic i l oma~n before the telence- phaIic/ii~encephalic houniiary appears. In B F - 1 ' mice, the cerebral hemispheres are severely ilirninisheil and ventral telence- Ilhalic markers are not expressed (93) .

Cell nlarki~lg and transplantat~on exper- llnents are required to test the postulate that egmentat lon 1s ~nvolveil In forebrain regio~lalization. Alternativel\-, or ailil~tlonally, forebrain pattern coulil ilepenil on an as yet uniiiscovereii signal~ng reglon. Whether or not a segmenteii transverse organizat~on ex- ~ s t s , a major co~ls t ra i~l t on understanding forebrain pattern has been our uncertalnt\-

regaril~ng ~ t s t opo log~c ;~ l coorii i~lates, p,lr- t~cular l \ - w ~ t h respect t o the trajectory of t he l o n g ~ t u i l ~ n a l axl\ . Here, ho~vevcr , iie- tailed i l e sc r~p t~ons of 1)V-rcstr~cteii gene express~on patterns ( 9 4 ) have more pre- clsely i ief~neii t he topology of the i l oma~ns ~vhose m e c h a n ~ s m of form,ltion we are s e e k ~ n g . T h e rapid accumulation of molec- ~ ~ l a r ilata haa provokeii exc~ tc i l specula- tlcin: with the c o m b ~ n e d app i~ca t ion of experimental emhr\-olog~cal and genetic methoiis, Lve can expect t h ~ s exci tc~l lent soon to be rel~eveil h\- enlightenment.

Spinal Cord-Late Role of the Mesoderm in Regionalization

Although s ~ ~ p e r f ~ c ~ a l l \ - un~fo rm, there ;Ire suhtle varlatlcins In cellular composltlon ;11o11g the A P a x 1 of the s p ~ n a l cord. Motor neurons are arranged in i i~sco~ l t i~ luous lon- g ~ t u i l ~ n a l column\ that occup\- iilffcrent LIV anil meil~olateral posltlons at il~fferent, plu- risegrnental levels of the neuraxls. Thus , the neurons that form the lateral motor col- umns a t l~ rnh (brachial anii lumb;lr) levels are iiisti~lct from those that torru at cervical ;111ii t ho rac~c levels, 110t o111y in the i i lent~ty of the11 peripheral targets but also in the expressLon of ii~fferent c o m h ~ n a t i o ~ l i of LIM-homeohox genes that ;Ire thought to confer targeting spec~f~c i ty (3 , 95 ) . Recent stuiiies of the spinal coril have focuseil o n the co~l t ro l of ~ t s prono~lnceii 1)V pattern ( 3 ) , ~vhereas classical s t~~ i i i e s of A P regionalization and the i~lt luence of paraxla1 me- soilerm (96) st111 neeil to be put Into a molecular context. However, gene, that 11e 5' in the Hox clusters have sharp ho~lnii- arieh of expression along the spin;rl neural tuhe, suggestlng, hy an,rloey w ~ t h the hlnii- t.ra~n, that they nlight uniierlie thi\ regional iiivers~ty. Transpcisition of pro\pective hra- chial and thoracic regions leacis to the respec~f~caticin of Hox and Llm gene expresslon, and motor neuron sub type ilevelop accoriling to the11 new posl t lon (3 ) . T h e most likely source of signals that effect the acilu~sition of this reg~onal iiientit\- 15 the Ilarax~al mesoderm (64) . Mesoiiermal co11- trol has also been irnpl~cateil In the speci- f ~ c a t i o n of primary motor neurons of the zebrafish: transplantation of single c e l l to new A P pos~t ions with respect to the adjacent somlte r eu l t s In r e s p e c i f ~ c a t ~ o ~ ~ of hoth the Llm gene coiie of the motor I I ~ L I - ron and its subsequent axon trajector\- ;~n i i target spec~f i c~ t \ - (3 ) .

Many aspects of cell p,rttern ;Ire con- herved hetween the hlnilhrain m i i the p l - nal cord, particularly ~ v ~ t h regard to the 1)V a x ~ s , ~vhe re ventral (So111c Hedgehog) and iiorsal (RMP) signali~lg systems appear to be i i ient~cal in the t ~ v o r e g i o ~ ~ \ (3 ) . It 1s also apparent that these L>V slgnalb act on cells

SCIENCE VOL 274 I5 UOVEXIRER 1996

that have already acquired a stahle and heritahle indication of their position on the A P axis, and therehy an A P posit ion-specific competence to respond (2). T h e time at which A P fate hecomes restricted differs, however, hetween spinal cord and hind-hrain. Whereas the regional A P identity of the spinal neural tuhe appears to he uncommitted for some time after closure (3, 97), hindhrain pattern is fixed and independent of position relative to the cranial mesoderm from as soon as the rhomhomeres hecome defined (2). T h e difference may stem from a phylogenetically ancient distinction hetween head and hody with respect to patterning strategy. In the head, the paraxial mesoderm is patently unsegmented and may contrihute little to patterning (98), whereas the neural crest predominates, furnishing the ectomesenchymal cells that construct the segmented hranchial skeleton and pattern the cranial nerves and muscles. Thus, in the hindhrain/hranchial region, segmen-tally restricted positional information originates in the neural tuhe and is imposed on the surrounding mesoderm. A relatively rigid set of positional values within the hindhrain region may have evolved hoth for correct deployment of its emigrant neural crest cells and in compensation for the lack of patterning information in the mesoderm. In the hody, hy contrast, the mesoderm imposes its A P positional information on the neural tuhe (99). This is seen hoth for cell pat tern within the tuhe (3, 100) and for the pattern of motor roots and dorsal root ganglia, whose overtly segmented disposition is controlled, apparently exclusively, hy the A P polarity of the somitic sclerotome (101).

Conclusions

Considerable advances have recently heen made toward understanding the mechanisms involved in neuraxial regionalization, particularly with respect to the earliest events, during gastrulation, when the molecular identity of activating and transforming signals is heing revealed. Especially promising is the evidence, from dominant gain- and loss-of-function experiments with retinoid receptors (15), that RA acts as a concentration-dependent poste-riorizing signal in vivo and is required for the correct spatial restriction of anterior markers. The action of RA in regionalizing the entire posterior CNS, as studied in early Xenopus emhryos, is mediated, at least in part, hy its direct action on the spatial regulation of Hox genes, hest known from the amniote hindhrain. A synthesis of data from these diverse experimental systems is needed to advance our understanding of this crucial molecule, as are experiments directed at elucidating its regulation and mode of action, whether as a

gradient from a single source, the organizer, or as discrete local signals from axial or paraxial mesoderm (or hoth).

O n a wider level, expression studies and functional analyses of developmental control genes in different vertehrate systems have revealed the existence of a neuraxial ground pattern that is highly conserved. It will he important to discover the genetic and cellular mechanisms involved in the suhsequent elahoration of this ground pattern that produce the very different hrains of fish and mammals. Further understanding of C N S regionalization will depend on discovering how region-specifying genes confer a particular potential , or set of potentials, with respect to the ultimate selection of regionally appropriate cell identity.

REFERENCES AND NOTES

1. C.-O. Jacobson, Zool. Bidr. Upps. 36, 73 (1964). 2. H. Simon, A. Hornbruch, A. Lumsden, Curr. Biol. 5,

205(1995). 3. Reviewed by T. M. Jessell and A. Lumsden, in Neu

ronal Development, W. M. Cowan, T. M. Jessell, S. L. Zipursky, Eds. (Oxford Univ. Press, Oxford, 1996), in press.

4. Reviewed by Y. Tanabe and T. M. Jessell, Science 274, 1115(1996).

5. H. Spemann, Embryonic Development and Induction (Hafner, New York, 1938).

6. P. D. Nieuwkoop, J. Exp. Zool. 120, 1 (1952). 7. A. Ruiz i Altaba, Trends Neurosci. 17, 233 (1994). 8. T. Doniach, C. R. Phillips, J. C. Gerhart, Science

257,542(1992) . 9. Reviewed by R. M. Harland, in (3), in press.

10. S. Piccolo, Y. Sasai, B. Lu, E. De Robertis, Cell 86, 589(1996).

11. L. Zimmerman, J. De Jesus-Escobar, R. Harland, ibid., p. 599.

12. W. G. Cox and A. Hemmati-Brivanlou, Development 121 , 4349 (1995).

13. N. Papalopulu and C. Kintner, ibid. 122, 3409 (1996).

14. K. L. Kroll and E. Amaya, ibid., p. 3173. 15. B. Blumberg et al., ibid., in press. 16. W. Shawlot and R. R. Behringer, Nature 374, 425

(1995). 17. D. Acampora et al., Development 121 , 3279

(1995). 18. S. L. Ang etal., ibid. 122, 247 (1996). 19. L. Bally-Cuif, M. Gulisano, V. Broccoli, E. Boncinelli,

Mech. Dev. 49 ,49(1995) . 20. T. Bouwmeester, S.-H. Kim, Y. Sasai, B. Lu, E. De

Robertis, Nature 382, 595 (1996). 2 1 . A. Lumsden, Trends Neurosci. 13, 329 (1990). 22. J. D. Clarke and A. Lumsden, Development 118,

151 (1993). 23. A. Lumsden and R. Keynes, Nature 337, 424

(1989). 24. H. Simon and A. Lumsden, Neuron 1 1 , 209 (1993). 25. G. Fortin, F. Kato, A. Lumsden, J. Champagnat,

J. Physiol. 486, 735(1995). 26. J. Glover and G. Petursdottir, J. Neurobiol. 22, 352

(1991). 27. F. Marin and L. Puelles, Eur. J. Neurosci. 7, 1714

(1995). 28. P. A. Lawrence and G. Struhl, Cell 85, 951 (1996). 29. S. Fraser, R. Keynes, A. Lumsden, Nature 344, 431

(1990). 30. E. Birgbauer and S. E. Fraser, Development 120,

1347(1994). 3 1 . R. Wingate and A. Lumsden, ibid. 122, 2143

(1996). 32. S. Guthrie and A. Lumsden, ibid. 112, 221 (1991). 33. I. Heyman, A. Kent, A. Lumsden, Dev. Dyn. 198,

241 (1993). 34. N. Becker et al., Mech. Dev. 47, 3 (1994).

35. A. D. Bergeman, H. J. Cheng, R. Brambilla, R. Klein, J. G. Flanagan, Mol. Cell. Biol. 15, 4921 (1995).

36. A. Flenniken, N. Gale, G. Yancopoulos, D. Wilkinson, Dev. Biol. 179, 382 (1996).

37. Q. Xu, G. Alldus, N. Holder, D. G. Wilkinson, Development 121, 4005 0995).

38. I. Heyman, A. Faissner, A. Lumsden, Dev. Dyn. 204, 301 (1995).

39. M. Cook et al., Proc. Natl. Acad. Sci. U.S.A. 92, 2249(1995).

40. R. Mahmood, P. Kiefer, S. Guthrie, C. Dickson, I. Mason, Development 121 , 1399 (1995).

4 1 . S. Martinez, E. Geijo, M. V. Sanchez-Vives, L. Puelles, R. Gallego, ibid. 116, 1069 (1992).

42. Reviewed in R. Krumlauf, Cell 78, 191 (1994). 43. M. Studer, H. Popperl, H. Marshall, A. Kuroiwa, R.

Krumlauf, Science 265, 1728 (1994). 44. D. G. Wilkinson, S. Bhatt, M. Cook, E. Boncinelli, R.

Krumlauf, Nature 341 , 405 (1989). 45. M. Zhang etal., Development 120, 2431 (1994). 46. D. Alexandre etal., ibid. 122, 735 (1996). 47. S. P. Cordes and G. S. Barsh, Cell 79, 1025 (1994). 48. D. G. Wilkinson, S. Bhatt, P. Chavrier, R. Bravo, P.

Charnay, Nature 337, 461 (1989). 49. M. A. Frohman, G. R. Martin, S. Cordes, L. P. Hal-

amek, G. S. Barsh, Development 117', 925 (1993). 50. I. J. McKay et al., ibid. 120, 2199 (1994). 5 1 . S. Schneider-Maunouryefa/.,Ce//75, 1199(1993). 52. S. Nonchev et al., Development 122, 543 (1996). 53. M. H .Shamefa / . ,Ce / /72 , 183(1993). 54. S. A. Newman, BioEssays 15, 277 (1993). 55. Reviewed in H. Marshall, A. Morrison, M. Studer, P.

Popperl, R. Krumlauf, FASEBJ. 10, 969 (1996). 56. H. Marshall etal., Nature 360, 737 (1992). 57. J. Hill, J. D. W. Clarke, N. Vargesson, T. Jowett, N.

Holder, Mech. Dev. 50, 3 (1995). 58. M. Maden, E. Gale, I. Kostetskiii, M. Zile, Curr. Biol.

6,417(1996) . 59. Y. P. Chen, L. Huang, A. F. Russo, M. Solursh,

Proc. Natl. Acad. Sci. U.S.A. 89, 10056 (1992). 60. A. Simeone et al., Mech. Dev. 5 1 , 83 (1995). 6 1 . D. J. Mangelsdorf and R. M. Evans, Cell 83, 841

(1995). 62. A. Grapin-Botton, M.-A. Bonnin, L. Ariza-Mc-

Naughton, R. Krumlauf, N. M. LeDouarin, Development 121 , 2707 (1995).

63. S. Guthrie et al., Nature 356, 157 (1992). 64. N. Itasaki, J. Sharpe, A. Morrison, R. Krumlauf,

Neuron 16, 487(1996). 65. Reviewed by A. Lumsden and A. Graham, Semin.

Cell Dev. Biol. 7, 169(1996). 66. Reviewed in A. L. Joyner, Trends Genet. 12, 15

(1996). 67. M. Hanks, W. Wurst, L. Anson-Cartwright, A. B.

Auerbach, A. L. Joyner, Science 269, 679 (1995). 68. U. Drescherefa/., Cell 82, 359(1995). 69. H.-J. Cheng, M. Nakamoto, A. D. Bergemann, J. G.

Flanagan, ibid., p. 371 . 70. N. Itasaki and H. Nakamura, Neuron 16, 55 (1996). 7 1 . C. Logan etal., Curr. Biol. 6, 1006 (1996). 72. S. Martinez, M. Wassef, R.-M. Alvarado-Mallart,

Neuron 6, 971 (1991). 73. A. P. McMahon, A. L. Joyner, A. Bradley, J. A.

McMahon,Ce//69, 581 (1992). 74. P. H. Crossley, S. Martinez, G. R. Martin, Nature

380,66(1996) . 75. M. Heikinheimo, A. Lawshe, G. Shackleford, D. B.

Wilson, C. A. MacArthur, Mech. Dev. 48, 129 (1994).

76. P. H. Crossley and G. R. Martin, Development 121 , 439(1995).

77. A. Hemmati-Brivanlou, R. Stewart, R. M. Harland, Science 250, 800(1990).

78. S. L. Ang and J. Rossant, Development 118, 139 (1993).

79. P. Urbanek, Z. Q. Wang, I. Fetka, E. F. Wagner, M. Busslinger, Cell 79, 901 (1994).

80. S. Krauss, M. Maden, N. Holder, S. W. Wilson, Nature 360, 87(1992).

8 1 . D.-L. Song, G. Chalepakis, P. Gruss, A. L. Joyner, Development 122, 627 (1996).

82. S. Martinez, F. Marin, M. A. Nieto, L. Puelles, Mech. Dev. 51,289(1995) .

83. S. Millet, E. Bloch-Gallego, A. Simeone, R.-M. Alvarado-Mallart, Development 122, 3785 (1996).

1114 SCIENCE • VOL.274 • 15 NOVEMBER 1996

84 M Hallonet M.-A Telllet N. M. Le Douar~n, 89 J A. Golden and C. L Cepko. Developmerit 122. J Rubensten, Developme!it 121 3923 (1 9951 !hid 108. 19 11 9901 65 11 996) 95. T. Tsuchda et a1 . Cell 79, 957 119941

85 Rev~ewed n A. Bang and M Goulding. Curr Opln , 90 G F~shell, C A. Mason. M. E. Hatten, Nature 362. 96 Rev~ewed n M. Jacobson. Develoi3niental Neuro- Neurob~ol 6 . 25 (1 9961 636 11 9931 h~ology (Plenum, New York. 1991 ) .

86. A. S~meone. D Acampora, M Gul~sano, A Stor- 91 M Gotz, A. W~zenmann. A Lumsden, J Pr~ce. 97. J Elsen and S. Pke Netiron 6 767 (19911 na~uolo, E Boncnel~ , Nature 358. 687 11992) Neuron 16, 551 (1 9961 08 D Noden, Develop!iient 103 121 (1 9381

87 J L R Rubenste~n, S. Mart~nez. K. Shmamura, L. 92. C Walsh and C L Cepko. Scierice 255, 434 99 C Stern. K Jaques, T. Llm S. Fraser F?. Keynes Pueles. Scierice 266. 578 11 9941 (1 9921 ihld. 113, 239 119911.

88 M C Flgdor and C D. Stern. Nattire 363. 630 93 S Xuan et a / , Neurori 14. 1141 119951. 100. J S Elsen. Scierice 252. 567 (1991). (1 9931. 94. K. Shlmamilra. D. Hart~gan. S Mart~nez, L Pueles. 101 R. Keynes and C Stern, Nature 310. 736 (1934)

Diversity and Pattern in the eve loping Spinal Cord

Yasuto Tanabe and Thomas M. Jessell

The generation of distinct neuronal cell types in appropriate numbers and at precise positions underlies the assembly of neural circuits that encode animal behavior. Despite the complexity of the vertebrate central nervous system, advances have been made in defining the principles that control the diversification and patterning of its component cells. A combination of molecular genetic, biochemical, and embryological assays has begun to reveal the identity and mechanism of action of molecules that induce and pattern neural tissue and the role of transcription factors in establishing generic and specific neuronal fates. Some of these advances are discussed here, focusing on the spinal cord as a model system for analyzing the molecular control of central nervous system development in vertebrates.

A l l neural tlrnction-trom \imple sensory re\ncinse\ ;inii motor commani i to elaho- rate cogn i t~ve behaviors-depend on the assembly o t neuronal circuits, a process 1111- t ~ a t e d during emhryon~c development. A n early and t(rniiarnent,ll step in th15 process is the gene ra t~c~n o t i i~stinct classes of neurons a t precise locations Lv~thin a pr iml t~ve neural eplthelllrm. Over the past decaiie, Inany ijt the tnechanisrns that control the ~ d e n t ~ t y o t speclflc neural cell type\ have heen iie- t~ne i i , In l x g e part through the application of molecular genetlcs In invertehrate organ- isnls such as Drosol~hll~t and Cuenorh~tbd~t~s elegun.5 hut also through cellular anii 1310- chemical approaches in vertebrates. Collec- t~velv . the studv of these d~ver se svstelns has , ,

prcjvlded considerable inslght into the rela- tlve contrlhutlons of e n v ~ r o n ~ ~ ~ e n t a l ,len,ll- - ing and lineage restrlctlon In neural iievel- on1nent ;111ii ha\ revealeii the i i ient~tv o t many of the extracellular signaling factor\ and ~ntracellular protein\ that iiirect cell bate.

Scjme o t the most intrwuine behaviors ,, , ~ ,

depend cjn the clrcuits that are formed dur- int! the develijnnlent of the verrehrate hraln and bpinal cord, yet our under\tanii~ng o t neural iievelopmenr is more tragrnentary In the vertebrate central nervous system

The authors are at the Hoviard Hughes Medcal lnsttute. Department of Biochemistry and Molecular Bophyscs Center for Neurob~ology and Behavor. Coumba Un~ver- slty. New York, NY 10032, USA

( C N S ) than in other systelns ( 1 ) . Here \ve revie~v recent progre\\ In iietining ho\v cli- verse cell types in the vertebrate C N S are generate'{, toclr\ing largely on the splnal cijrii, because it is the sinlplest and mo\t conserved regLon of the vertebrate C N S (Fig. 1A) . 111 ailcditlon, phy\iological anii ana tom~ca l analves (i t neuronal c~rcuitrv In the spinal corii' have provided, from 'the tlnle of Sherrington, a \oliii celllrlar trame- \vork h r in terpre t~ng the neural hases of sensory anii motor tlrnctions (2 ) . Although the tllnctions encc~iieii In spinal cord clr- cultry are llrn~teii hy comparison to those o t 111;111v other hraln strlrctures. stuiiieh on the develop~nent of spinal nelrrons may reveal general strategies lr\eii to ebtahlish neuronal ii~ver\ity anii clrcuitry 111 more complex regions o t the C N S .

W e exarnlne the steps involved in the generation of distinct neural cell types through the use o t \ome\vhat artlticlal sub- i i~vi\ion\ o t \vhat is evidently an ~ntegrated iievelop~nental program.

Induction of the Neural Plate

T h e iievelopment o t the spinal cord, as in other reglons ( ~ t the C N S , is initiated hy the ~ni iuct ion of the neural plate. T h e cla\\ical grafting experments of Spernann anii Mangold in amphihian embryo\ (3) estah- lished that the fi jrmat~on o t neural tlssue iiepenii\ o n signal\ prov~iieii hy prospective

a x ~ a l mesoilermal cell5 in the org;lnl:er region. Until recently the lilentity and mech- a n i s ~ n cjf actlon (j t t h e e cndogeno~15 neur;il I I I ~ U C L I I L ' factor\ h;ive r e ~ n ; i ~ n e d obscure. Stlriiie\ o t nelrr;rl i n d ~ r c t ~ o n In Xenoj~u, em- h r y o no\v \lrggcst thxt 111 one 111.1jor 17;lth- \Yay o t neural ~ncluctlon, f<lctora ,rnt,igcini:e the signals med~a ted by the rr;lnstcirming gro\vth f;~ctor-p (TGFP)-like protein, hone morphogen~c protein4 (RMP4). ~ v h ~ c h re- prebsc neural ; ~ n d p ron lo te eplder~llal cell tare (4) (Fig. 2 ) .

B,MP .si,y~nilng and )~c.urcil indztction T h e 17osslhility that neural iniluction nliqht result trom the in,ictiv;ltlcin of ,I \ lgnal~ng p a t h ~ v ~ ~ y that represses neur;rl tare c~ncrgecl frc)m the ol?erv,iticin th,lt c l ~ ~ ~ o c l : ~ t ~ o n of h l ; l s t~r l i i -~t ;~~c ectocIer111 into slnqlc cells, pre\~r~llahly prevrnting ~ n t u c e l l ~ ~ l ; ~ r s~qn,rl- ing, \va, sutticient to e l i c ~ t the fcirmat~cjn o t nelrral tijsue (5). Me~llher\ o t the TGFR tamily \vere \1.1qgc5tcil to medi;ltc thi. repres lve slgn,ll on the h;r.i. (it experllllents des~gned mitially to te.t whether the TC;FP-like protcin ; ~ c t i v ~ n Lva5 req~rlreil fiir the iniilrction o t nleoclerlll (6 ) . Injection o t transcripts that cncodcil ,r do~nin , rnt neg~i- tive fc~rrn of an ,ictivin receptor hlocked mcsoilerm,il ilifirenti;ltion. Rut ectoilcrm;rl cells e x p r e l n g thls receptor i\otcirm unex- pectedly d~fterenti,rtcii into neur;ll tl\sue, \~lgge\ting that the 13lock;rdc o t , ict iv~n receptor \ign,rl tr;ln5ductlon I \ \ ~ l f t ~ c ~ e n t to trigger neural incluctlon. TLVO 11nc. o t evidence iniiic,lte that RMP4 rather than ;ic- t lvm itself 15 likely to he the cneloqeno~rs TC;FP-l~ke proteln t h ; ~ t interacts \r.~tll t l ~ i \ receptor anii repre5ie neural dlffercntia- tion. F~rs t , RMP4 I aliiely cxprc>5cd In the earlv cctoiierm ;~ni i its exnresslon 1s extin- glr~\heii trom neural plate c e l l during neural in i iuct~on ( 7 ) . Second. RMP4 hut not actlvln can prevent the cxprcs\lon of neural marker5 anii promote e p ~ d u ~ n ; ~ l ditkrenti;r- tion in ii~ssoclated ectoiierm;il cells (8). Or - ganizer-derlvcd signal> might therefore induce neural tlbsue hy 1rle;inj of cndogcncxr, proteins that hlock \iqn,tlinq ~ne,li;rteii hy RhlP proteins.

Support for thli, icde;~ h;li c o ~ n e fro111 the iiemonstration that thrcc c;~ndicl;lte neural incl~~cers expresecd hy cirfi;lnl:cr t ~ j s u c can act in this Inanncr (Firr. 2. l? ;ind (:). T h e , , . ,

e n d o g e n o ~ r ac t iv in-hini i in p r c ~ t c ~ n tol- llstatln is expre\scd hy orq;ln1:er c ~ l l s , ;lnd

SCIENCE L'OL 274 l i hOL'EX1RER 1996 1115

Review Article

Fgf8 signaling for development of the midbrain andhindbrain

Hidekiyo Harada,1,2 Tatsuya Sato3,4 and Harukazu Nakamura4*1Genetics and Development Division, Toronto Krembil Research Institute, 2Department of Physiology, Faculty of

Medicine, University of Toronto, Toronto, Ontario, Canada; 3Department of Developmental Neuroscience, GraduateSchool of Medicine, Tohoku University, Sendai 980-8575, Japan and 4Frontier Research Institute for InterdisciplinaryScience, Tohoku University, Sendai 980-8578, Japan

In this paper, we review how midbrain and hindbrain are specified. Otx2 and Gbx2 are expressed from the earlyphase of development, and their expression abuts at the midbrain hindbrain boundary (MHB), where Fgf8

expression is induced, and functions as an organizing molecule for the midbrain and hindbrain. Fgf8 inducesEn1 and Pax2 expression at the region where Otx2 is expressed to specify midbrain. Fgf8 activates Ras-ERKpathway to specify hindbrain. Downstream of ERK, Pea3 specifies isthmus (rhombomere 0, r0), and Irx2 mayspecify r1, where the cerebellum is formed.

Key words: Fgf8 signaling, hindbrain, isthmus, midbrain, Pea3, Ras-ERK.

Introduction

Vertebrate brain is formed at the rostral part of the

neural tube and consists of regionally differentiated

structures. Each region consists of specific histological

organization, and is formed by specific histogenesis.The neural tube from which whole central nervous sys-

tem (CNS) develops is a simple tube at first. So it is

very interesting how distinct region develops from a

simple structure. Regionalization begins with the pri-

mary brain vesicle formation at the rostral end of the

neural tube: forebrain (prosencephalon), midbrain

(mesencephalon) and hindbrain (rhombencephalon).

Then the forebrain is subdivided into telencephalonand diencephalon, and the hindbrain is subdivided into

metencephalon and myelencephalon. These sec-

ondary brain vesicles are the fundamental brain plan

(Fig. 1).

Specificity of the region is determined by the combi-

nation of the transcription factors expressed (Naka-

mura et al. 2008), and the organizing signal from the

local organizing centers fix preexisting transcriptionfactors or induce new transcription factors to specify

the neighboring region. New local organizing center is

generated by pre-existing transcription factors. Mid-

brain-hindbrain boundary (MHB) functions as a local

organizing center for the development of the midbrain

and hindbrain. From a very early stage of develop-

ment, Otx2 is expressed rostrally, and Gbx2 isexpressed caudally (Simeone et al. 1992; Bouillet et al.

1995). In chick embryos, the MHB region is not cov-

ered either by Otx2 or by Gbx2 at stage 8, but their

expression becomes overlapping around the MHB at

stage 9–10 (Garda et al. 2001). By mutual repressive

activity, their expression abuts at the MHB, where Fgf8

expression is induced overlapping with the Gbx2

expression (Fig. 1) (Hidalgo-S�anchez et al. 1999; Kata-hira et al. 2000). Fgf8 is the organizing molecule for

the midbrain and hindbrain. MHB is also called as the

isthmus. Here, we will review how midbrain and hind-

brain are specified, and how Fgf8 organizes midbrain

and hindbrain development.

Isthmus organizer

Heterotopic transplantation of the diencephalon to the

posterior part of the midbrain between quail and chick

embryos resulted in the fate change of the transplant

to differentiate to the midbrain tectum (Nakamura et al.

1988). But diencephalon kept its original fate when it

was transplanted to the anterior part of the midbrain.

These results indicated that developmental fate of the

*Author to whom all correspondence should be addressed.Email: [email protected] 7 March 2016; revised 22 April 2016;

accepted 22 April 2016.ª 2016 Japanese Society of Developmental Biologists

Develop. Growth Differ. (2016) 58, 437–445 doi: 10.1111/dgd.12293

The Japanese Society of Developmental Biologists

diencephalon is plastic, and that some morphogenetic

signal exists near the posterior part of the midbrain

(Nakamura et al. 1986; Nakamura & Itasaki 1992).

Transplantation of the MHB (isthmus) to the dien-

cephalon induced En2 expression in the host dien-

cephalon, and the En2-induced-region differentiatedinto the tectum (Martinez et al. 1991; Bally-Cuif et al.

1992; Bally-Cuif & Wassef 1994). A breakthrough

came from the finding that Fgf8 is expressed in the

isthmus (Heikinheimo et al. 1994; Crossley & Martin

1995), and insertion of the Fgf8 soaked bead into the

diencephalon exerted similar effects as the isthmus

transplantation (Crossley et al. 1996). Thus, the isth-

mus was accepted as the organizer, and the Fgf8 asthe organizing molecule. Subsequent gain-of-function

studies of Fgf8 in chick and mice have all suggested

that Fgf8 is a signaling molecule emanating from the

isthmus (Liu et al. 1999; Martinez et al. 1999; Shamim

et al. 1999). Wnt1 is expressed at the posterior marginof the mesencephalon, and was regarded as a candi-

date organizing molecule since Wnt1 knock-out mice

show defects in midbrain and hindbrain (McMahon &

Bradley 1990; Thomas & Capecchi 1990; McMahon

et al. 1992). Wnt1 plays essential role in keeping Fgf8

expression in the isthmus (Matsunaga et al. 2002). Lim

homeodomein protein Lmx1b and Wnt1 are expressed

at the posterior margin of the midbrain (Fig. 1), andLmx1b induces Wnt1 expression (Matsunaga et al.

2002). It was shown that Lmx1b cell-autonomously

repressed Fgf8 expression, but Wnt1 induced Fgf8

expression. Since Wnt1 is a secreted molecule, these

relationships keep Fgf8 expression in the isthmus

(Fig. 2). Wnt1 assures cell proliferation not the regional

specificity (Matsunaga et al. 2002).

Specification of the midbrain

Otx2 is expressed in the mesencephalon and prosen-

cephalon from a very early stage of development while

Gbx2 is expressed in the metencephaon (Simeone

et al. 1992; Bouillet et al. 1995). En1 (Engrailed 1) and

Pax2 covers entire mesencephalon (Asano & Gruss

1992; Rowitch & McMahon 1995; Araki & Nakamura

Fig. 2. Maintenance of isthmus organizer. Otx2 and Gbx2 are

expressed from a very early stage of development in the midbrain

and hindbrain, respectively. Fgf8 expression is induced at the

boundary of Otx2 and Gbx2 expression, overlapping with Gbx2

expression. Lmx1b and Wnt1 are expressed at the posterior bor-

der of the midbrain. Lmx1b induces Grg4 expression, which

represses Fgf8 expression. Lmx1b also induces Wnt1 expres-

sion, which induces Fgf8 expression. By these mechanisms,

Fgf8 expression is kept in the isthmus.

Fig. 1. Schematic drawing of the brain vesicles, transcription

factors expressed and the organizing centers. Primary brain vesi-

cles, forebrain (prosencephalon), midbrain (mesencephalon) and

hindbrain, are subdivided into five secondary brain vesicles,

telencephalon (Tel), diencephalon (Di), mesencephalon (midbrain),

metencephalon (Met) and myelencephalon (Myel). Otx2 and

Gbx2 are expressed from the early stage of development cover-

ing rostral to midbrain and hindbrain, respectively. Fgf8 expres-

sion is induced at the border of Otx2 and Gbx2 expression

overlapping to Gbx2 expression. Fgf8 induces and stabilizes En1

and Pax2 expression in the region where Otx2 is expressed to

specify midbrain. Fgf8 activates Ras-ERK pathway to specify

hindbrain. Fgf8 expression is also induced at the anterior part of

the neural tube to specify telencephalon. Diencephalon-midbrain

boundary is determined by repressive activity of Pax6 and En1/

Pax2, and mid-hindbrain boundary is determined by repressive

activity of Otx2 and Gbx2.

ª 2016 Japanese Society of Developmental Biologists

438 H. Harada et al.

1999; Shamim et al. 1999; Nakamura 2001). En2expression in the mesencephalon is in a gradient as

rostral low caudal high and stronger in the isthmus.

Application of Fgf8 soaked bead in the diencephalon

induced En2 expression around the bead (Crossley

et al. 1996). Misexpression of En1/En2, Pax2 or Fgf8

in diencephalon, where they are not expressed,

induced each other’s expression and the tectum was

formed there (Araki & Nakamura 1999; Okafuji et al.1999; Sato et al. 2001). It was concluded that En1/2,

Pax2 and Fgf8 are in a positive feedback loop for their

expression (Araki & Nakamura 1999; Funahashi et al.

1999; Okafuji et al. 1999). Mice lacking En1 show sev-

ere midbrain and hindbrain deletion, and die at birth,

whereas mice lacking En2 show only subtle defects

(Joyner et al. 1991; Millen et al. 1994; Wurst et al.

1994; reviewed in Joyner 1996). Pax2 and Pax5 dou-ble knockout mice lack midbrain and cerebellum (Sch-

warz et al. 1997; Urb�anek et al. 1997). In homozygous

no isthmus mutant zebrafish, which carry inactivated

Pax-2.1 genes, Pax5 expression is lost and the fish

lack the isthmic constriction, cerebellum and optic tec-

tum (Lun & Brand 1998; Pfeffer et al. 1998).

Misexpression of Otx2 in the metencephalon

repressed Gbx2 and Fgf8 expression, and the tectumdifferentiated ectopically in place of the cerebellum in

chick (Katahira et al. 2000), and the inferior colliculi

enlarged and extended caudally in mouse (Broccoli

et al. 1999). On the other hand, Gbx2 misexpression

in the mesencephalon repressed Otx2 expression. As

a consequence, the hindbrain expanded rostrally and

midbrain was reduced both in mouse and chick (Millet

et al. 1999; Katahira et al. 2000). Both in Otx2 andGbx2 misexpression, Fgf8 expression was induced at

the border of Otx2 and Gbx2 expression domain over-

lapping with Gbx2 expression (Broccoli et al. 1999;

Millet et al. 1999; Katahira et al. 2000). Otx2 null

mouse embryos lack the forebrain, midbrain and ros-

tral hindbrain and die in early embryogenesis (Acam-

pora et al. 1995; Matsuo et al. 1995; Ang et al. 1996).

Gbx2 knockout mice showed abnormalities in the isth-mus structure and caudal expansion of the midbrain

(Wassarman et al. 1997).

From these experiments, it was concluded that the

region where Otx2, En1 and Pax2 are expressed is

specified as the mesencephalon (Fig. 1) (Nakamura

et al. 2005). Further experiments showed that Pax3/7

confers tectum property to the mesencephalon (Mat-

sunaga et al. 2001).

Fgf8 signaling

Now we focus more on the Fgf8 signaling around the

isthmus. There are eight splicing isoforms of Fgf8, and

it was shown by RT-PCR that Fgf8a and Fgf8b areexpressed in the isthmus. Fgf8b contains additional 11

amino acids to Fgf8a, and has stronger transformation

activity than Fgf8a (MacArthur et al. 1995a,b). Fgf8a

misexpression by electroporation in the forebrain to

hindbrain in chick embryos did not affect molecules

expressed in the midbrain and hindbrain, but induced

En2 expression in the diencephalon (Sato et al. 2001).

Consequently, fate of the caudal diencephalon chan-ged to midbrain; tectum extended rostrally (Fig. 3A).

Fgf8a misexpression in mouse by Wnt1 regulatory ele-

ment, which assures expression at the dorsal part of

the caudal diencephalon to midbrain caused prolifera-

tion of neural precursors and expansion of the mid-

brain, but no apparent fate change of the midbrain/

hindbrain occurred (Lee et al. 1997).

Implantation of an Fgf8b soaked bead in the dien-cephalon could induce Gbx2, and repressed Otx2 in

the midbrain both in chick and mouse embryos (Liu

et al. 1999; Martinez et al. 1999). Misexpression of

Fgf8b by in ovo electroporation in chick embryos

exerted drastic effects. Otx2 expression in the mid-

brain was repressed, and Gbx2, Irx2, Pax2 and En2

expression extended to the caudal diencephalon. Con-

sequently, the cerebellum differentiated in place of thetectum (Fig. 3B–D) (Sato et al. 2001). Misexpression of

Fgf8b in transgenic mice in which Fgf8b expression is

regulated under Wnt1 regulatory element exerted simi-

lar effects on downstream gene expression (Liu et al.

1999).

Since Fgf8b has stronger transforming activity than

Fgf8a (MacArthur et al. 1995b), it was assumed that

the difference in the effects between Fgf8a and Fgf8bcould be attributed to the difference in signal intensity

(Sato et al. 2001). This assumption was supported by

quantitative analysis and by structural analysis. For

quantitative analysis, electroporation at different con-

centration of Fgf8b expression vector (1.0, 0.1, 0.01

and 0.001 lg/lL) was performed. It was shown that

electroporation with 1.0 and 0.1 lg/lL of Fgf8b

expression vector transformed the fate of the midbrainto that of the hindbrain, and that electroporation with

0.01 lg/lL of Fgf8b expression vector exerted Fgf8a

type effects; rostral expansion of the tectum (Sato

et al. 2001). Structural analysis by surface plasmon

resonance showed that Fgf8b binds more intimately to

the c isoforms of the Fgf receptor 1–3 (FGFR1-3)

owing to the additional 11 amino acids, phenylalanine

32 (F32) being most important. Mutation of F32 to ala-nine (F32A) reduced the affinity to FGFR, and the

mutation converted the action of Fgf8b to that of Fgf8a

(Olsen et al. 2006).

Competence of the tissue to organizing signals

depends on pre-existing transcription factors. Otx2


Fgf8 signaling for development of the midbrain and hindbrain 439

and Gbx2 are expressed from a very early stage in the

midbrain and hindbrain, respectively, and Fgf8 expres-

sion is induced at the boundary of their expression

overlapping with Gbx2. Fgf8 activates Ras-ERK path-way in the region where Gbx2 is expressed to specify

hindbrain (Sato & Nakamura 2004). Strong Fgf8 signal

could repress Otx2 expression and induce Gbx2

expression in the midbrain, and activates Ras-ERK

signaling to change the fate of the midbrain to that of

the hindbrain. The result of co-transfection of Otx2

and Fgf8b expression vector uncovered that Otx2

increases Fgf signal threshold required for hindbraindifferentiation. Transfection of Fgf8b vector at concen-

tration of 1.0 lg/lL or of 0.1 lg/lL vector could

convert the midbrain to the hindbrain property. But

co-transfection of 0.1 lg/lL of Fgf8b vector and

(A) (B) (C) (D)

(E) (F) (G) (H)

(I) (J) (K)

Fig. 3. Effects of Fgf8 and its signal transduction. (A) Misexpression of Fgf8a. Fate change of the diencephalon to midbrain occurs, and

the tectum at the experimental side (right) extends toward diencephalon. (B–D) Misexpression of Fgf8b. Fate change of the midbrain to

hindbrain occurs, and the cerebellum differentiates in place of the tectum (cer-ect). In the ectopically differentiated cerebellum external

granular layer (arrows) and the Purkinje cells (arrow heads) are differentiated. (E–H) Effects of Fgf8a (F), Fgf8b (G) and 1/100 diluted

Fgf8b (H). Transfection region is indicated in E. ERK is activated corresponding to the transfection site after Fgf8b misexpression (G).

Ectopical actigation of ERK is seen only in the diencephalon after misexpression of both Fgf8a and 1/100 dilution of Fgf8b, which exerts

similar gross morphological effects. (I–K) Disruption of Ras-ERK signaling by dominant negative form of Ras (RasS17N, a serine-to-aspar-

agine mutation at residue 17). ERK activity is down regulated (I), and the tectum is formed in place of the cerebellum.



1.0 lg/lL of Otx2 vector could not change the fate ofthe midbrain (Sato et al. 2001). The results explain

why the posterior part of the midbrain that receives

strong Fgf8 signal differentiates as the midbrain though

it receives strong Fgf8 signal.

Signaling pathway toward hindbrain development

It is very interesting how the Fgf8 signal is transducedto specify midbrain and hindbrain development. Fgf

signal is received by a receptor tyrosine kinase, which

activates Ras-ERK pathway in many cases (reviewed

by Katz & McCormick 1997; Rommel & Hafen 1998).

ERK activation could be detected by anti-di-phos-

phorylated ERK (dpERK) antibody (Gabay et al. 1997;

Christen & Slack 1999; Shinya et al. 2001; Corson

et al. 2003), and it was shown that ERK is activatedaround the isthmus (Fig. 3I) (Sato & Nakamura 2004).

After misexpression of Fgf8b, ERK was activated cor-

responding to the transfection of Fgf8b (Fig. 3E, G)

(Sato & Nakamura 2004). It is very interesting that sim-

ilar ERK activation pattern was obtained after Fgf8a

and 1/100 diluted-Fgf8b misexpression that exerts

Fgf8a type effects (Fig. 3F, H). Disruption of Ras-ERK

signaling pathway changed the property of the hind-brain to that of the midbrain, that is, by misexpression

of dominant negative form of Ras (dnRas) which

antagonizes Ras activity, or by misexpression of Spro-

uty2, negative regulator of Ras-ERK pathway (Fig. 3I),

in the hindbrain, the tectum differentiated in place of

the cerebellum (Fig. 3J, K) (Sato & Nakamura 2004;

Suzuki-Hirano et al. 2005). It was suggested that Irx2

is phosphorylated by ERK, and is involved in cerebellardifferentiation (Matsumoto et al. 2004). All of these

studies suggest that strong Fgf8 signal activates Ras-

ERK pathway in the region where Gbx2 is expressed

to organize hindbrain development.

Regulation of Ras-ERK pathway

Fgf8-Ras-ERK signaling is very strong, and activation ofFgf8-Ras-ERK signaling is only needed for short win-

dow of developmental process so that ERK activity

should be quickly down regulated (reviewed in Mason

et al. 2006). Indeed, Fgf8 itself induces negative regula-

tors for Ras-ERK signaling in the isthmus; Sprouty2

and Sef expression is induced through Fgf8-Ras-ERK

signaling (Echevarria et al. 2005; Suzuki-Hirano et al.

2005) and Mkp3 is induced through Fgf8-PI3K-AKTsignaling (Echevarria et al. 2005). Sprouty2 may alter-

natively be induced by nuclear-translocated Fgf8 itself

(Suzuki et al. 2012). ERK is phosphorylated (activated)

widely in the midbrain and hindbrain at around stage 8

and 9 in chick embryos, but ERK phosphorylation

region become narrower and phosphorylation level inthe hindbrain becomes low by stage 11 (Sato & Naka-

mura 2004). Sprouty2 has stronger repressive activity

to ERK than Mkp3 (Suzuki-Hirano et al. 2010), and

Sprouty2 misexpression in the midbrain/hindbrain

exerted similar effects as dn-Ras misexpression; the

tectum differentiated in place of the cerebellum

(Suzuki-Hirano et al. 2005). Suzuki-Hirano carefully

examined ERK activation in the midbrain after Fgf8b

misexpression by electroporation, which forces fate

changes of the midbrain to the hindbrain. ERK was

phosphorylated just after electroporation and kept

high till 15 h after electroporation, the ERK became

de-phosphorylated by 18 h after electroporation. What

happens if ERK is continuously phosphorylated by

repressing the activity of the negative regulators? When

Fgf8b and DN-Sprouty2 were co-transfected in themidbrain, ERK was kept phosphorylated in the mid-

brain until 18 h after electroporation, and the fate

change of the midbrain did not occur; midbrain devel-

oped as midbrain and the tectum differentiated

(Suzuki-Hirano et al. 2010). By turning off DN-Sprouty2

transcription by Tet-off system (Hilgers et al. 2005;

Watanabe et al. 2007) at 5.5 h after electroporation of

Fgf8b and DN-Sprouty2 expression vectors, ERK activ-ity in the midbrain was repressed by 18 h after electro-

poration, and the fate change of the midbrain to the

hindbrain occurred (Suzuki-Hirano et al. 2010).

Comparison of the phosphorylation level of ERK in

the mid-hindbrain region after DN-Sprouty2 and DN-

Mkp3 misexpression showed that Sprouty2 has a

stronger activity than Mkp3 (Suzuki-Hirano et al.

2010). After misexpression of Fgf8b and DN-Mkp3,ERK phosphorylation level in the midbrain at 18 h after

electroporation kept high but lower than that after

Fgf8b and DN-Sprouty2 misexpression, which is

because Sprouty2 may have been induced, and the

property of the midbrain changed to that of the isth-

mus (rhombomere 0, r0) and the trochlear neurons dif-

ferentiated (Suzuki-Hirano et al. 2010).

In normal development of the chick hindbrain, ERK isactivated in r0-r1 at stage 10, but in the r1 ERK activity

is downregulated at stage 11 (Suzuki-Hirano et al.

2010). Thus, after activation of ERK in the hindbrain,

activity may be downregulated differentially: the region

where ERK activity is weakly kept differentiate into the

isthmus (r0) and the region where ERK is strongly

downregulated acquires the property as r1 and the

cerebellum differentiates (Suzuki-Hirano et al. 2010).

Downstream of Ras-ERK pathway

We have shown that strong Fgf8 signal activates Ras-

ERK pathway to specify hindbrain development. As a



downstream of Ras-ERK pathway, Pea3 subfamily(Raible & Brand 2011), which consist of Pea3, Erm

and Er81 and belong to Ets-type transcription factors,

were suggested in mouse lung development (Liu et al.

2003). In the isthmus, Pea3 and Erm are expressed in

zebrafish, and their expression was induced by ectopi-

cally misexpressed Fgf8, and was repressed by FGFR

inhibitor SU5402 or by Sprouty4 (Roehl & N€usslein-

Volhard 2001; Raible & Brand 2011). In chickembryos, Pea3 is expressed in the mid-hindbrain

region, and its expression was regulated by Ras-ERK

pathway; disruption of Ras-ERK signaling by DN-Ras

repressed Pea3 expression (Harada et al. 2015). Gain

and loss of function of Pea3 exerted similar effects as

Fgf8b and DN-Ras misexpression on Otx2, Gbx2 and

Fgf8 expression, respectively. Pea3 misexpression

resulted in repression of Otx2 in the midbrain, and ros-tral extension of Gbx2 and Fgf8 expression. Repres-

sion of Pea3 function by misexpression of a chimeric

molecule of Engrailed repressor domain EH1 and Pea3

(eh1-Pea3) resulted in upregulation of Otx2 in the hind-

brain and repression of Gbx2 and Fgf8 in the hindbrain

(Harada et al. 2015).

From the effects on downstream gene expression, it

was expected that Pea3 would function to specify

hindbrain at the downstream of ERK, so that Pea3

misexpression would induce cerebellum in the mid-

brain (Harada et al. 2015). But the cerebellum was not

differentiated after Pea3 misexpression in the midbrain

region. Instead, isthmus structure was extended ros-

trally in case of Pea3 misexpression; trochlear neurons

differentiated rostral to the proper troclear nucleus.

Repression of Pea3 function by eh1-Pea3 resulted in

caudal extension of the midbrain; oculomotor neuronsdifferentiated caudal to the oculomotor nucleus. These

results indicated that Pea3 specifies isthmus (r0) under

Ras-ERK pathway (Harada et al. 2015). Recently it

was reported that bHLH transcription factor Atoh1 is

expressed in the rostral part of r1, and that Atoh1

expressing region would give rise to isthmic nuclei in

mouse and chick (Green et al. 2014). Experimental

results suggest that Pea3 may be responsible for thespecification of Atoh1 expressing region.

For the specification of the r1 region, Irx2 may be

essential. Fgf8b could ectopically induce Irx2 in the

midbrain, but Fgf8a could not (Sato et al. 2001).

Consequently, Fgf8a could not change the fate of the

midbrain. Co-misexpession of Fgf8a and Irx2 could

induce the ectopic cerebellum in the midbrain region.

Thus, Irx2 expression may be induced through

Fig. 4. Fgf8 signaling for the specification

of the midbrain and hindbrain. Otx2 and

Gbx2 are expressed from a very early

stage of development, and Fgf8 expres-

sion is induced at the border of Otx2 and

Gbx2 expression. Fgf8 signaling induces

En1 and Pax2 expression where Otx2 is

expressed to specify the midbrain devel-

opment. Transducing pathway for the

midbrain specification has not been eluci-

dated. Fgf8 activates Ras-ERK signaling

pathway, and specifies the hindbrain

development. Negative regulators for the

Ras-ERK signaling, Sprouty2, Sef and

Mkp3, are induced, and differential activa-

tion level of ERK is formed in the hind-

brain. The region where ERK activity is

strongly repressed is specified as r1 by

Irx2, and the region where ERK activity is

weakly repressed is specified as r0 by

Pea3. When Ras-ERK pathway is dis-

rupted, fate change of the hindbrain

occurs, and the tectum differentiates in

place of the cerebellum.



Ras-ERK signaling. Irx2 could induce Fgf8 and Gbx2

expression in the midbrain and convert the fate of

this region from the tectum to the cerebellum (Mat-

sumoto et al. 2004).

Concluding remarks

Regional specificity of the neural tube is determined by

the combination of the transcription factors expressed,and the signaling molecule organizes the fate of the

neighboring region by regulating expression of the

transcription factors. Isthmus functions as the orga-

nizer for the midbrain and hindbrain by emanating

Fgf8. Fgf8 induces and stabilizes expression of En1

and Pax2 in the region where Otx2 is expressed. Thus,

Fgf8 specifies the development of the midbrain since

the region where Otx2, En1 and Pax2 are expresseddifferentiates as the midbrain (Figs 1 and 4). Gbx2 is

crucial for the hindbrain development. Fgf8 activates

Ras-ERK signaling pathway where Gbx2 is expressed

and specifies the hindbrain development. Otx2 and

Gbx2 expression is overlapping at the border at early

stages of development, and their expression becomes

abutted at the midbrain/hindbrain boundary by their

mutual repressing activity, and Fgf8 expression isinduced at the boundary of Otx2 and Gbx2 expression

overlapping with Gbx2 expression. If the Ras-ERK

pathway is disrupted in the hindbrain, Otx2 expression

remains and the territory of the midbrain extends cau-

dally. Strong Fgf8 signaling can change the fate of the

midbrain to hindbrain by repressing Otx2 expression in

the midbrain and activates Ras-ERK pathway. Down-

stream of the Ras-ERK pathway, Pea3 specifies theisthmus (r0) region (Fig. 4). Atoh1 may be related to

specification of the r0 (Green et al. 2014). The tran-

scription factor that specifies the r1 downstream of

Ras-ERK pathway may be Irx2 (Fig. 4).

References

Acampora, D., Mazan, S., Lallemand, Y., Avantaggiato, V.,Maury, M., Simeone, A. & Brulet, P. 1995. Forebrain andmidbrain regions are deleted in Otx2�/� mutants due to adefective anterior neuroectoderm specification during gastru-lation. Development 121, 3279–3290.

Ang, S.-L., Ou, J., Rhinn, M., Daigle, N., Stevenson, L. & Ros-sant, J. 1996. A targeted mouse Otx2 mutation leads tosevere defects in gastrulation and formation of axial meso-derm and to deletion of rostral brain. Development 122,243–252.

Araki, I. & Nakamura, H. 1999. Engrailed defines the position ofdorsal di–mesencephalic boundary by repressing dien-cephalic fate. Development, 126, 5127–5135.

Asano, M. & Gruss, P. 1992. Pax-5 is expressed at the mid-brain–hindbrain boundary during mouse development. Mech.

Dev. 39, 29–39.

Bally-Cuif, L. & Wassef, M. 1994. Ectopic induction and reorgani-zation of Wnt-1 expression in quail/chick chimeras. Develop-ment 120, 3379–3394.

Bally-Cuif, L., Alvarado-Mallart, R. M., Darnell, D. K. & Wassef,M. 1992. Relationship between Wnt-1 and En-2 expressiondomains during early development of normal and ectopicmet–mesencephalon. Development 115, 999–1009.

Bouillet, P., Chazaud, C., Oulad-Abdelghani, M., Doll�e, P. &Chambon, P. 1995. Sequence and expression pattern of theStra7 (Gbx-2) homeoboxcontaining gene induced by retinoicacid in P19 embryonal carcinoma cells. Dev. Dyn. 204, 372–382.

Broccoli, V., Boncinelli, E. & Wurst, W. 1999. The caudal limit ofOtx2 expression positions the isthmic organizer. Nature 401,164–168.

Christen, B. & Slack, J. M. 1999. Spatial response to fibroblastgrowth factor signalling in Xenopus embryos. Development

126, 119–125.Corson, L. B., Yamanaka, Y., Lai, K. M. & Rossant, J. 2003.

Spatial and temporal patterns of ERK signaling duringmouse embryogenesis. Development 130, 4527–4537.

Crossley, P. H. & Martin, G. R. 1995. The mouse Fgf8 geneencodes a family of polypeptides and is expressed in regionsthat direct outgrowth and patterning in the developingembryo. Development 121, 439–451.

Crossley, P. H., Martinez, S. & Martin, G. R. 1996. Midbraindevelopment induced by FGF8 in the chick embryo. Nature380, 66–68.

Echevarria, D., Martinez, S., Marques, S., Lucas-Teixeira, V. &Belo, J. A. 2005. Mkp3 is a negative feedback modulator ofFgf8 signaling in the mammalian isthmic organizer. Dev. Biol.277, 114–128.

Funahashi, J., Okafuji, T., Ohuchi, H., Noji, S., Tanaka, H. &Nakamura, H. 1999. Role of Pax5 in the regulation of amid-hindbrain organizer’s activity. Dev. Growth Differ. 41,59–72.

Gabay, L., Seger, R. & Shilo, B. Z. 1997. In situ activation patternof Drosophila EGF receptor pathway during development.Science 277, 1103–1106.

Garda, A. L., Echevarr�ıa, D. & Mart�ınez, S. 2001. Neuroepithelialco-expression of Gbx2 and Otx2 precedes Fgf8 expressionin the isthmic organizer. Mech. Dev. 101, 111–118.

Green, M. J., Myat, A. M., Emmenegger, B. A., Wechsler-Reya,R. J., Wilson, L. J. & Wingate, R. J. 2014. Independentlyspecified Atoh1 domains define novel developmental com-partments in rhombomere 1. Development 141, 389–398.

Harada, H., Omi, M., Sato, T. & Nakamura, H. 2015. Pea3 deter-mines the isthmus region at the downstream of Fgf8-Ras-ERK signaling pathway. Dev. Growth Differ. 57, 657–666.

Heikinheimo, M., Lawsh�e, A., Shackleford, G. M., Wilson, D. B.& MacArthur, C. A. 1994. Fgf-8expression in the post-gas-trulation mouse suggests roles in the development of theface, limbs and central nervous system. Mech. Dev. 48,129–138.

Hidalgo-S�anchez, M., Simeone, A. & Alvarado-Mallart, R. M.1999. Fgf8 and Gbx2 induction concomitant with Otx2repression is correlated with midbrain–hindbrain fate of cau-dal prosencephalon. Development 126, 3191–3203.

Hilgers, V., Pourquie, O. & Dubrulle, J. 2005. In vivo analysis ofmRNA stability using the Tet-Off system in the chickenembryo. Dev. Biol. 284, 292–300.

Joyner, A. L. 1996. Engrailed, Wnt and Pax genes regulate mid-brain–hindbrain development. Trends Genet. 12, 15–20.



Joyner, A. L., Skarnes, W. C. & Rossant, J. 1991. Subtle cere-bellar phenotype in mice homozygous for a targeted deletionof the En-2 homeobox. Nature 338, 153–156.

Katahira, T., Sato, T., Sugiyama, S., Okafuji, T., Araki, I., Funa-hashi, J. & Nakamura, H. 2000. Interaction between Otx2and Gbx2 defines the organizing center for the optic tectum.Mech. Dev. 91, 43–52.

Katz, M. E. & McCormick, F. 1997. Signal transduction from mul-tiple Ras effectors. Curr. Opin. Genet. Dev. 7, 75–79.

Lee, S. M. K., Danielian, P. S., Fritzsch, B. & McMahon, A. P.1997. Evidence that FGF8 signalling from the midbrainhind-brain junction regulates growth and polarity in the developingmidbrain. Development 124, 959–969.

Liu, A., Losos, K. & Joyner, A. L. 1999. FGF8 can activate Gbx2and transform regions of the rostral mouse brain into a hind-brain fate. Development 126, 4827–4838.

Liu, Y., Jiang, H., Crawford, H. C. & Hogan, B. L. M., 2003.Role for ETS domain transcription factors Pea3/Erm inmouse lung development. Dev. Biol., 261, 10–24.

Lun, K. & Brand, M. 1998. A series of no isthmus (noi) alleles ofthe zebrafish pax2.1 gene reveals multiple signaling eventsin development of the midbrain–hindbrain boundary. Devel-opment 125, 3049–3062.

MacArthur, C. A., Lawsh�e, A., Xu, J., Santos-Ocampo, S., Heik-inheimo, M., Chellaiah, A. T. & Ornitz, D. M. 1995a. FGF-8isoforms activate receptor splice forms that are expressed inmesenchymal regions of mouse development. Development

121, 3603–3613.MacArthur, C. A., Lawsh�e, A., Shankar, D. B., Heikinheimo, M. &

Shackleford, G. M. 1995b. FGF-8 isoforms differ in NIH3T3cell transforming potential. Cell Growth Differ. 6, 817–825.

Martinez, S., Wassef, M. & Alvarado-Mallart, R. M. 1991. Induc-tion of a mesencephalic phenotype in the 2-day-old chickprosencephalon is preceded by the early expression of thehomeobox gene en. Neuron 6, 971–981.

Martinez, S., Crossley, P. H., Cobos, I., Rubenstein, J. L. &Martin, G. R. 1999. FGF8 induces formation of an ectopicisthmic organizer and isthmocerebellar development via arepressive effect on Otx2 expression. Development 126,1189–1200.

Mason, J. M., Morrison, D. J., Basson, M. A. & Licht, J. D.2006. Sprouty proteins: multifaceted negative-feedback reg-ulators of receptor tyrosine kinase signaling. Trends Cell Biol.

16, 45–54.Matsumoto, K., Nishihara, S., Kamimura, M., Shiraishi, T., Oto-

guro, T., Uehara, M., Maeda, Y., Ogura, K., Lumsden, A. &Ogura, T. 2004. The prepattern transcription factor Irx2, atarget of the FGF8/MAP kinase cascade, is involved in cere-bellum formation. Nat. Neurosci. 7, 605–612.

Matsunaga, E., Araki, I. & Nakamura, H. 2001. Role of Pax3/7 inthe tectum regionalization. Development 128, 4069–4077.

Matsunaga, E., Katahira, T. & Nakamura, H. 2002. Role ofLmx1b and Wnt1 in mesencephalon and metencephalondevelopment. Development 129, 5269–5277.

Matsuo, I., Kuratani, S., Kimura, C., Takeda, N. & Aizawa, S.1995. Mouse Otx2 functions in the formation and patterningof rostral head. Genes Dev. 9, 2646–2658.

McMahon, A. P. & Bradley, A. 1990. The Wnt-1 (int-1) pro-tooncogene is required for development of a large region ofthe mouse brain. Cell 62, 1073–1085.

McMahon, A. P., Joyner, A. L., Bradley, A. & McMahon, J. A.1992. The midbrain-hindbrain phenotype of Wnt-1-/Wnt-1-mice results from stepwise deletion of engrailed-expressingcells by 9.5 days postcoitum. Cell 69, 581–595.

Millen, K. J., Wurst, W., Herrup, K. & Joyner, A. L. 1994. Abnor-mal embryonic cerebellar development and patterning ofpostnatal foliation in two mouse Engrailed-2 mutants. Devel-opment 120, 695–706.

Millet, S., Campbell, K., Epstein, D. J., Losos, K., Harris, E. &Joyner, A. L. 1999. A role for Gbx2 in repression of Otx2and positioning the mid/hindbrain organizer. Nature 401,161–164.

Nakamura, H. 2001. Regionalisation and polarity formation of theoptic tectum. Prog. Neurobiol. 65, 473–488.

Nakamura, H. & Itasaki, N. 1992. Expression of en in the prosen-cephalon heterotopically transplanted into the mesen-cephalon. Dev. Growth Differ. 34, 387–391.

Nakamura, H., Nakano, K. E., Igawa, H. H., Takagi, S. & Fuji-sawa, H. 1986. Plasticity and rigidity of differentiation ofbrain vesicles studied in quail-chick chimeras. Cell Differ. 19,187–193.

Nakamura, H., Takagi, S., Tsuji, T., Matsui, K. A. & Fujisawa, H.1988. The Prosencephalon has the capacity to differentiateinto the optic tectum: analysis by chick-specific monoclonalantibodies in quail–chick-chimeric brains. Dev. Growth Differ.

30, 717–725.Nakamura, H., Katahira, T., Matsunaga, E. & Sato, T. 2005. Isth-

mus organizer for midbrain and hindbrain development.Brain Res. Rev. 49, 120–126.

Nakamura, H., Sato, T. & Suzuki-Hirano, A. 2008. Isthmus orga-nizer for mesencephalon and metencephalon. Dev. Growth

Differ. 50, S113–S118.Okafuji, T., Funahashi, J. & Nakamura, H. 1999. Roles of Pax-2

in initiation of the chick tectal development. Dev. Brain Res.

116, 41–49.Olsen, S. K., Li, J. Y. H., Bromleigh, C., Eliseenkova, A. V.,

Ibrahimi, O. A., Lao, Z., Zhang, F., Linhardt, R. J., Joyner,A. L. & Mohammadi, M. 2006. Structural basis by whichalternative splicing modulates the organizer activity of FGF8in the brain. Genes Dev. 20, 185–198.

Pfeffer, P. L., Gerster, T., Lun, K., Brand, M. & Busslinger, M.1998. Characterization of three novel members of thezebrafish Pax2r5r8 family: dependency of Pax5 and Pax8expression of the Pax2.1 (noi) function. Development 125,3063–3074.

Raible, F. & Brand, M. 2011. Tight transcriptional control ofthe ETS domain factors Erm and Pea3 by Fgf signalingduring early zebrafish development. Mech. Dev. 107, 105–117.

Roehl, H. & N€usslein-Volhard, C. 2001. Zebrafish pea3 and ermare general targets of FGF8 signaling. Curr. Biol. 11, 503–507.

Rommel, C. & Hafen, E. 1998. Ras-a versatile cellular switch.Curr. Opin. Genet. Dev. 8, 412–418.

Rowitch, D. H. & McMahon, A. P. 1995. Pax-2 expression in themurine neural plate precedes and encompasses the expres-sion domains of Wnt-1 and En-1. Mech. Dev. 52, 3–8.

Sato, T. & Nakamura, H. 2004. The Fgf8 signal causes cerebellardifferentiation by activating Ras-ERK signaling pathway.Development 131, 4275–4285.

Sato, T., Araki, I. & Nakamura, H. 2001. Inductive signal and tis-sue responsiveness difining the tectum and the cerebellum.Development 128, 2466–2469.

Schwarz, M., Alvarez-Bolado, G., Urb�anek, P., Busslinger, M. P.& Gruss, P. 1997. Conserved biological function betweenPax-2 and Pax-5 in midbrain and cerebellum development:evidence from targeted mutations. Proc. Natl Acad. Sci.

USA 94, 14518–14523.



Shamim, H., Mahmood, R., Logan, C., Doherty, P., Lumsden, A.& Mason, I. 1999. Sequential roles for Fgf4, En1 and Fgf8 inspecification and regionalisation of the midbrain. Develop-

ment 126, 945–959.Shinya, M., Koshida, S., Sawada, A., Kuroiwa, A. & Takeda, H.

2001. Fgf signalling through MAPK cascade is required fordevelopment of the subpallial telencephalon in zebrafishembryos. Development 128, 4153–4164.

Simeone, A., Acampora, D., Gulisano, M., Stornaiuolo, A. &Boncinelli, E. 1992. Nested expression domains of fourhomeobox genes in developing rostral brain. Nature 358,687–690.

Suzuki, A., Harada, H. & Nakamura, H. 2012. Nuclear transloca-tion of FGF8 and its implication to induce Sprouty2. Dev.

Growth Differ. 54, 463–473.Suzuki-Hirano, A., Sato, T. & Nakamura, H. 2005. Regulation of

isthmic Fgf8 signaling by Sprouty2. Development 132, 257–265.

Suzuki-Hirano, A., Harada, H., Sato, T. & Nakamura, H. 2010.Activation of Ras-ERK pathway by Fgf8 and its down regula-tion by Sprouty2 for the isthmus organizing activity. Dev.

Biol. 337, 284–293.

Thomas, K. R. & Capecchi, M. R. 1990. Targeted disruption ofthe murine int-1 proto-oncogene resulting in severe abnor-malities in midbrain and cerebellar development. Nature 346,847–850.

Urb�anek, P., Fekta, I., Meisler, M. H. & Busslinger, M. 1997.Cooperation of Pax-2 and Pax-5 in midbrain and cerebellumdevelopment. Proc. Natl Acad. Sci. USA 94, 5703–5708.

Wassarman, K. M., Lewandoski, M., Campbell, K., Joyner, A. L.,Rubenstein, J. L. R., Martinez, S. & Martin, G. R. 1997.Specification of the anterior hindbrain and establishment of anormal mid/hindbrain organizer is dependent on Gbx2 genefunction. Development 124, 2923–2934.

Watanabe, T., Saito, D., Tanabe, K., Suetsugu, R., Nakaya, Y.,Nakagawa, S. & Takahashi, Y. 2007. Tet-on inducible sys-tem combined with in ovo electroporation dissects multipleroles of genes in somitogenesis of chicken embryos. Dev.

Biol. 305, 625–636.Wurst, W., Auerbach, A. B. & Joyner, A. L. 1994. Multiple devel-

opmental defects in Engrailed-1 mutant mice: an early mid-hindbrain deletion and patterning defects in forelimbs andsternum. Development 120, 2065–2075.



Vol.:(0123456789)1 3

Cellular and Molecular Life Sciences https://doi.org/10.1007/s00018-018-2974-x

REVIEW

Hindbrain induction and patterning during early vertebrate development

Dale Frank1 · Dalit Sela‑Donenfeld2

Received: 9 May 2018 / Revised: 19 November 2018 / Accepted: 21 November 2018 © Springer Nature Switzerland AG 2018

AbstractThe hindbrain is a key relay hub of the central nervous system (CNS), linking the bilaterally symmetric half-sides of lower and upper CNS centers via an extensive network of neural pathways. Dedicated neural assemblies within the hindbrain control many physiological processes, including respiration, blood pressure, motor coordination and different sensations. During early development, the hindbrain forms metameric segmented units known as rhombomeres along the antero-posterior (AP) axis of the nervous system. These compartmentalized units are highly conserved during vertebrate evolution and act as the template for adult brainstem structure and function. TALE and HOX homeodomain family transcription factors play a key role in the initial induction of the hindbrain and its specification into rhombomeric cell fate identities along the AP axis. Signaling pathways, such as canonical-Wnt, FGF and retinoic acid, play multiple roles to initially induce the hindbrain and regulate Hox gene-family expression to control rhombomeric identity. Additional transcription factors including Krox20, Kreisler and others act both upstream and downstream to Hox genes, modulating their expression and protein activity. In this review, we will examine the earliest embryonic signaling pathways that induce the hindbrain and subsequent rhombomeric segmentation via Hox and other gene expression. We will examine how these signaling pathways and transcription factors interact to activate downstream targets that organize the segmented AP pattern of the embryonic vertebrate hindbrain.

Keywords Hindbrain · Neural specification and patterning · Hox proteins · Meis and Pbx proteins · FGF, Wnt and retinoic acid signaling · Rhombomere patterning

Introduction

The central nervous system (CNS) has been a classic model system to study pattern formation during early vertebrate development. Vertebrate CNS morphology is strikingly con-served, from fish to mammals. Dorsal ectodermal cells on

the outer surface of the embryo are induced to neural fates by neighboring dorsal mesoderm cells. This induced neural plate tissue subsequently thickens, elevating at the embryo’s two lateral edges to form the neural folds. These two aris-ing neural folds merge at the dorsal midline of the embryo, creating the neural tube. The neural tube is asymmetric; the wider, thicker walled anterior region forms the brain, while the narrower, posterior part forms the spinal cord.

The brain subdivides into three regions with distinct antero-posterior (AP) characteristics: the most anterior forebrain (telencephalon and diencephalon), the midbrain (mesencephalon) and most caudally, the hindbrain (rhom-bencephalon). These regions are the morphological basis of distinct functional units of the brain. In the forebrain, the telencephalon gives rise to the cerebral cortex, basal gan-glia and hippocampus. The diencephalon gives rise to the thalamus, hypothalamus and pineal gland. In the lateral dien-cephalon regions, optic vesicles form the eye structure. The midbrain will generate centers of sensory and motor control, whereas the hindbrain gives rise to the cerebellum, pons and

Cellular and Molecular Life Sciences

Dale Frank and Dalit Sela-Donenfeld have contributed equally to this work.

* Dale Frank [email protected]

* Dalit Sela-Donenfeld [email protected]

1 Department of Biochemistry, Faculty of Medicine, The Rappaport Family Institute for Research in the Medical Sciences, Technion-Israel Institute of Technology, 31096 Haifa, Israel

2 Koret School of Veterinary Medicine, The Robert H Smith Faculty of Agriculture, Food and Environment, The Hebrew University of Jerusalem, 76100 Rehovot, Israel

http://orcid.org/0000-0002-1857-6820

http://orcid.org/0000-0001-7498-061X

http://crossmark.crossref.org/dialog/?doi=10.1007/s00018-018-2974-x&domain=pdf

D. Frank, D. Sela-Donenfeld

1 3

medulla [1]. The hindbrain controls crucial physiological processes such as motor activity, respiration, sleep and blood circulation. It also receives processes and transmits multi-ple sensory inputs including the auditory, precerebellar and vestibular systems. Different streams of neural crest cells are generated in the hindbrain to give rise to cranial sensory ganglia, Schwann cells, cardiac connective tissue and most of the cranial skeleton [2].

The embryonic hindbrain is divided into seven or eight segmented regions called rhombomeres (r). In the most ros-tral hindbrain, r1 borders the midbrain in a region called the midbrain–hindbrain boundary, whereas at the most caudal end, r7/r8 borders the most-anterior spinal cord. Each rhom-bomere has unique gene expression patterns that promote the region-specific fates, differentiation of neurons and pro-duction of distinct neural crest streams. Rhombomeric units are highly conserved during vertebrate evolution, acting as templates for the adult hindbrain structure and function, by giving rise to the pons, medulla oblongata and different cer-ebellar cell layers. The most caudal region of the central nervous system, the spinal cord, forms posterior to the hind-brain, extending to the rear of the body.

This elegant CNS morphogenesis is achieved in the vertebrate embryo through a series of inductive events. In amphibians, Mangold and Spemann in the 1920s found that grafting a dorsal mesodermal lip removed from the blasto-pore of an early gastrula stage embryo induced a secondary nervous system in the naïve ventral ectoderm of the trans-planted host [3]. This region is called “the organizer”, and the phenomenon of this experiment was termed “embry-onic induction”. These experiments were confirmed in other vertebrates in analogous fish and bird Spemann–Mangold organizer regions [4, 5]. More recent genetic studies suggest that a neural inducing organizer activity also exists in mam-mals. The mammalian organizer center may be composed of two regions acting in differing time periods. The earlier anterior visceral endoderm establishes initial embryonic AP polarity, while the later node region induces neural tissue [6].

Seminal experiments of Nieuwkoop, Eyal-Giladi, Toivonen and Saxen suggested that there are two induc-tive steps involved in neural induction and patterning [7–9]. In the first step, the “organizer” initially induces a general neural tissue having an anterior forebrain fate in a process called “activation”. The second “transformation” step is a caudalizing-posterior induction, which re-specifies the “acti-vated” neuro-ectoderm into more posterior CNS cell fates such as hindbrain and spinal cord [7].

Three different neural inducing proteins (noggin, chor-din, follistatin) were initially identified and characterized in amphibians [10–12]. These proteins all induce ante-rior-pan neural tissue. While structurally different, these proteins share one common activity, the ability to inhibit

BMP protein signaling activity [12–14] by blocking BMP ligand–receptor interactions [14–16]. Thus, BMP antago-nism serves as the initial “activation” signal inducing ecto-derm to rostral neural fates. Since BMP signaling actively drives ectoderm cells to epidermal/non-neural cell fates, BMP signaling inhibition suffices to convert ventral epider-mal ectoderm to a more dorsal neural fate [17–19].

Parallel to the discovery of the “activation” signal of BMP antagonism, neural caudalizing “transformation” molecules were also identified. Using Xenopus and chick experimental embryology techniques, in addition to zebrafish and mouse genetics, three signaling pathways were identified that cau-dalize rostral neural tissue. These include the basic fibro-blast growth factor (bFGF), retinoic acid (RA) and canonical Wnt signaling pathways [20–35]. For each pathway, multiple ligands, receptors and antagonists are expressed in different temporal windows and embryonic locales during the neural patterning process. The same pathways seem to have variant spatial and temporal roles in specifying multiple posterior cell fates in the developing vertebrate nervous system. These caudalizing factors induce an initial posterior neural “ground state”, which undergoes fine-tuning into distinct locales such as hindbrain and spinal cord along the neural AP axis.

While the sequence of morphological events that lead to the partition of the CNS into sub-domains is well known, the genetic networks that govern the specification and con-nectivity of different cell types along the CNS to yield a functional nervous system are only partially resolved (reviewed in [36–39]). This review examines the conserved transcriptional networks and morphogens that orchestrate the intricate regulation of early hindbrain specification and patterning in different vertebrates. It will cover both the ear-liest stages of hindbrain development/induction, as well as later stages of rhombomere specification. By addressing the complex interactive dynamics between signaling pathways, the earliest activation of Hox and TALE homeodomain pro-teins, as well as the later expression of non-homeodomain transcription factors, this review provides a unique com-prehensive synopsis of the central processes of hindbrain development in relevant vertebrate systems.

The transcription factor blueprint of the hindbrain

The Hox genes are one of the most ancient regulators of body formation in metazoans, being crucial for AP axis formation in the developing vertebrate embryo. Hox pro-teins partner up with another ancient family of homeodo-main proteins, the TALE class proteins, Meis and Pbx. By their joint interactive activities during the earliest stages of neural development, Meis, Pbx and Hox proteins act to induce and specify the hindbrain. Vertebrates typically have


1 3

thirteen Hox gene paralog groups (PG) expressed along the AP axis. The most anterior extent of Hox expression along the AP axis is the hindbrain, and the PG1–4 group genes are regionally expressed in the hindbrain, and required for its formation. Acting with the Hox genes to induce the hind-brain are the Meis/Pbx proteins. Meis/Pbx expression can precede Hox gene expression. In some systems, Meis/Pbx was shown to be required for the earliest activation of Hox gene expression in the hindbrain. In addition, various dimers or trimer of Meis/Pbx/Hox proteins directly activate target genes in the developing hindbrain. This review will address how these transcription factors interact with signaling path-ways to regulate the earliest formation of the hindbrain.

Homeodomain proteins

TALE class homeodomain proteins: Meis, Prep and Pbx proteins

The correct temporal and spatial expression of Hox par-alogous group (PG1–4) proteins is crucial for establishing the initial segmentation of the hindbrain. Hindbrain induc-tion is also dependent on Three-Amino acid Loop Exten-sion (TALE) homeodomain proteins, which belong to the MEIS (Meis1–3 proteins), PREP and PBC (Pbx1–4 proteins) groups. TALE proteins are atypical homeodomain-contain-ing transcription factors having three additional amino acids

between the first and second helix of the homeodomain [40]. During hindbrain development in zebrafish and Xenopus, meis/pbx genes are activated very early, preceding hox gene expression in the presumptive neural plate. Meis/Pbx and Hox proteins interact at two distinct levels. Early in zebrafish and Xenopus development, Meis/Pbx proteins are required to initially activate PG1–4 hox gene expression in the hindbrain [41–44]. Later, Meis (Prep)/Pbx/Hox protein combinations bind target genes to activate their transcription. Some of these target genes are also hox and meis genes themselves, but non-hox gene targets also lie downstream to Meis/Pbx/Hox (Fig. 1) [45, 46].

Studies in Xenopus and zebrafish embryos revealed the requirement of Meis1 and Meis3 for proper hindbrain for-mation [41–44, 47–50]. The expression of meis3 initiates early, at late gastrula stages in the presumptive hindbrain region. At later stages, its expression becomes localized to the r2–r4 region [42, 43, 47, 48]. In contrast, expression of the meis1 and meis2 genes is more general, expanding into more anterior neural regions than meis3 in multiple verte-brate species, such as mouse, chick, Xenopus and zebrafish [43, 51–56]. In Xenopus and zebrafish, loss of Meis/Pbx function, obtained by either dominant-negative proteins, antisense oligonucleotide morpholino (MO) knockdown, or genetic mutation, triggers a loss of the entire hindbrain region, accompanied by a loss in expression of a many hind-brain markers, including hox PG1-4 genes [41–44, 47]. This is concomitant with a posterior expansion and enlargement

Fig. 1 Pbx/Meis proteins lie upstream of Hox gene expression in the early hindbrain. A Schematic representation of Pbx and Meis regula-tory activity in the early hindbrain. Pbx/Meis proteins are induced in the neural plate by Wnt signaling prior to Hox gene expression. Pbx and Meis activate (individually and additively) the transcription of PG1–4 Hox genes, by binding to their Pbx/Meis-responsive elements. In parallel, Pbx/Meis also activate FGF and RA signaling to promote Hox gene expression. This is mediated by their binding to responsive elements in the Raldh2 gene, the main RA-synthesizing enzyme, as well as by inducing FGF gene expression that in turn inhibits the RA-degrading enzyme Cyp26. Following the initial activation of Hox gene expression, Meis/Pbx synergize with HoxA1 protein to induce expression of other Hox genes (not shown). B Meis3 regulates

early homeobox gene expression (from Elkouby et al. [44]). Meis3 knockdown inhibits early HoxD1, HoxA2 and Gbx2 gene expression. Embryos were injected into one blastomere at the two-cell stage with Meis3-MO (10–12.5 ng/blastomere; b, d, f). Gene expression was examined at late gastrula, stage 12.5. All embryos are viewed dor-sally, and oriented with anterior at the top, posterior at the bottom. The dashed line in b, d, f indicates the dorsal midline; the XMeis3-MO-injected side is on the right. Early expression of HoxD1, HoxA2 and Gbx2 is inhibited on the Meis3-MO-injected side (100%, n = 16; 88.2%, n = 34; 66.4%, n = 33, respectively). In neurula-stage Meis3-morphant sibling embryos, a typical inhibition of posterior neural cell types was observed


1 3

of more anterior forebrain structures. Meis3 also suffices for hindbrain induction, as its overexpression induces ectopic hindbrain formation while repressing forebrain formation in both Xenopus and zebrafish embryos and explants [42–44, 47, 48, 57]. In mouse and chick, Meis/Prep and Pbx proteins also are required for activation of hindbrain enhancers in the hoxb1 and hoxb2 genes [58–61].

Slightly later in zebrafish development, Meis3 syner-gizes with HoxB1b (HoxA1 in other vertebrates) protein to induce expression of various other hindbrain markers such as hoxb1a (hoxb1) and krox20 [48]. Similarly, in Xeno-pus, HoxD1 and Meis3 co-expression enhances hoxb3 and krox20 expression [62]. In mice and zebrafish, the krox20 promoter has a functional r3-specific Hox/Meis/Pbx binding site, although it does not appear to be a direct Meis3-target in Xenopus [44, 63, 64]. Moreover, hoxd1 is a direct-target gene of Meis3 in Xenopus that acts downstream of Meis3 to induce hindbrain cell fates [44, 57, 62]. In Xenopus, the Zic1 and Pax3 transcription factors are required upstream of Meis3. Knockdown of Zic1 or Pax3 prevents early meis3 gene expression, leading to a loss of hindbrain cell fates [57]. Ectopic Meis3 can rescue Zic1, but not Pax3 knock-down phenotypes.

Together with Meis proteins, the PBC family proteins are central for hindbrain induction. Pbx4 directly interacts with Meis1 and Meis3 proteins in zebrafish, and perturba-tion of either Pbx2 or Pbx4 activities eliminates r2–r6 cell fates in zebrafish [41, 43, 48, 65, 66]. Pbx4 is expressed in the presumptive hindbrain region, where its early expression closely overlaps the meis3 and hoxb1b (hoxa1) genes [48]. In Xenopus, Meis1 and Pbx1 proteins interact to regulate neural cell fate specification, where Pbx1 knockdown also disrupts hindbrain formation [50]. Pbx1 is expressed in the presumptive forebrain, hindbrain and neural crest regions [53]. In Xenopus, Pbx1 and HoxD1 proteins strongly activate a heterologous mouse hoxb1 enhancer, but Meis3 had no additive effect with either protein when tested separately or together [62, 67]. A Pbx protein partner that enhances Meis3 activity in Xenopus has not been identified. In chick and mice, Meis2 and Pbx proteins were found to synergize and to activate a krox20 enhancer element in r3 [63]. Interestingly, in zebrafish, ectopic Meis1 expression rescued hindbrain for-mation in the absence of zygotic Pbx4 protein. This suggests a potential Pbx-independent mechanism of action, although maternal Pbx protein involvement was not fully ruled out (Fig. 1), [43].

Hox proteins

PG1 Hox proteins are key factors controlling early hind-brain specification. The PG1 Hox proteins are homologs of the Drosophila labial gene and include the HoxA1, HoxB1 and HoxD1 proteins. In all vertebrates, expression of the

PG1 hox genes precedes all other hox genes in the presump-tive hindbrain region at early gastrula stages and persists through neurula stages [68–77]. PG1 proteins are essential for correct hindbrain induction and segmentation. Com-bined knockdown of HoxB1a (Mouse Hoxb1) and HoxB1b (Mouse Hoxa1) proteins in zebrafish, or hoxA1 and hoxB1 gene deletion in mouse embryos led to significant hindbrain perturbation [75, 76, 78, 79]. In Xenopus embryos, triple knockdown of all the PG1 genes, hoxa1, hoxb1 and hoxd1 caused a complete loss of r2–6, and the entire hindbrain resembled the Hox non-expressing r1 region [77]. This phe-notype is reminiscent of Pbx2 loss-of-function in zebrafish embryos, arguing for cooperative function of Pbx2 and PG1 Hox proteins in the specification of r2–6 regions [66]. Sup-porting this assumption, HoxA1 hexapeptide mutant proteins that fail to interact with Pbx proteins cause severe hindbrain phenotypes in mice [80]. Moreover, the combined PG1 loss-of-function phenotype is synergistically stronger than that of each of the individual inhibitions, and in Xenopus, PG1 knockdown could be rescued by overexpression of HoxD1 protein alone. This suggests at least partial functional redun-dancy between PG1 members [77]. To gain insight into the transcriptional network regulated by HoxA1, microarray analysis was performed on the prospective r3–5 region of hoxa1 null and wild-type mouse embryos [81]. Around 300 genes were differentially expressed between the samples. While many of these genes were previously identified to play a role in hindbrain development, new target genes were also found to be downregulated in hoxa1-nulls, such as FGF receptor 3 (fgfr3), zic1, hnf1b, foxd3 and lhx5, suggesting that HoxA1 protein acts in a genetic cascade upstream to many target-genes that control wider aspects of hindbrain development than previously thought. HoxA1 mutations in humans have brainstem function perturbations, and it was suggested that this could be related to improper embryonic hindbrain development [82].

Early PG1 protein expression is a prerequisite for the proper sequential expression of later, more posteriorly expressed PG2–4 Hox genes [62, 77]. In PG2 mutant mice, development of the r3/r4 region is disrupted with poor border formation between r2/r3. Cells in r2 express only hoxa2, the most anterior of all Hox genes, and hoxa2 loss-of-function mutations cause r2 to r1 fate changes [83–87]. Notably, Hox PG1 and PG2 genes are both expressed in r4, but with temporal differences. Single hoxb2 mutant mice had no hindbrain segmentation defects, but in hoxa2/hoxb2 double-mutant mice, the r2/3 and r3/r4 borders were lost, suggesting that the action of both PG2 genes is synergis-tic [88]. Interestingly, recent sequence analysis of hoxa2 in fugu uncovered the presence of two orthologues, hoxa2a and hoxa2b. Each orthologue contains distinct cis-regula-tory elements to drive hoxa2 expression in neural crest or rhombomeres, respectively. This study suggests that these


1 3

regulatory regions are conserved throughout vertebrate evo-lution to mediate differential hoxa2 expression and activity during development [89]. A negative regulatory mechanism exists between PG3 and PG2 groups. Studies in mice and chick embryos showed that Hoxb3 protein directly binds to hoxb1 regulatory regions and represses hoxb1 expression posterior to r4 [90]. In multiple PG3 mutants, r5/6 identity was disrupted and r4-specific hoxb1 expression was ectopi-cally activated in r5/6 [91]. In PG4 mutants, in contrast, hindbrain development was normal [92].

A recent study compared the binding targets of HoxA1 proteins in zebrafish and mouse finding that they share many common hindbrain target genes. Many of these targets also shared occupancy with Meis and Pbx proteins [93]. HoxA1 also was found to bind enhancer regions of the meis2 and meis3 genes [94], suggesting that HoxA1 may be regulating the early expression of these meis genes. This coupled to studies in Xenopus showing Meis3 regulation of PG1 Hox gene expression suggests that there is a mutual co-depend-ence between Meis and Hox PG1 proteins to regulate the earliest stages of vertebrate hindbrain specification.

TALE and Hox proteins cross‑talk in the hindbrain

As mentioned previously, during early development, Meis, Pbx and Hox proteins interact at two distinct levels. Initially, Meis–Prep/Pbx and PG1 Hox proteins may reciprocally co-activate each other’s gene expression in the hindbrain. Later, Meis/Pbx/Hox protein combinations bind target genes to further activate hox gene expression in the hindbrain [42]. In zebrafish embryos, ChIP studies showed that Meis/Pbx proteins specifically bind the hoxb1a and hoxb2a promot-ers in their respective tissues of expression [95]. These promoters were also enriched for histone H4 acetylation. In embryos ectopically expressing dominant negative Pbx proteins, Meis/Pbx activity was inhibited and histone acety-lation was highly reduced. Furthermore, Meis/Pbx protein complex formation removes histone deacetylase (HDAC) from Hox-regulated promoters. Additionally, Meis proteins recruit CBP/p300 histone acetylase to hox promoters. Thus, Meis proteins function as direct transcriptional activators of the hoxb1a target gene by controlling the accessibility of HDAC/CBP proteins to its promoter [95]. More recent stud-ies in zebrafish embryos show that TALE protein complexes actively poise hoxb1a promoters for expression by chroma-tin modification, as early as blastula stages. Later expres-sion of the hoxb1a promoter at gastrula stage is triggered by Hoxb1b protein binding to these TALE protein complexes [96]. Pbx–Hox and Meis–Prep binding sites have also been used to define a shared sequence syntax system for iden-tifying functional hindbrain-specific enhancer elements in zebrafish [97]. In rhombomeric segments, Hox gene expres-sion is positively controlled by auto- and cross-regulatory

binding of Meis/Pbx/Hox proteins to enhancer elements. For example, the hoxb1 gene enhancer that drives expression in r4 harbors distinct Meis/Pbx and Hox/Pbx binding sites. In mice, Hoxa1 together with Meis/Pbx proteins initially activates this enhancer, but later, Hoxb1 itself, together with Hoxb2, Meis3 and Pbx2/4 proteins, is required for expres-sion maintenance in r4 [58, 65]. Interestingly, additional Hox proteins of the PG3 group, but not Pbx/Meis proteins, negatively regulate hoxb1 such that it remains restricted to r4 [90, 91].

Many additional examples of such interdependent, cross-regulatory loops are known. For instance, hoxd1, hoxb2 and hoxa2 all require Meis/Pbx for their expression in zebrafish, Xenopus, chick and mouse [43, 44, 49, 59, 66, 98, 99]. In mice, Hoxb2 expression is directly activated by HoxB1 in r4, and HoxB2 protein then drives hoxb1 expression. Thus, HoxB2 indirectly controls its own expression via HoxB1 [59–61, 88, 100]. Hoxa2 expression in r4 is also regulated by conserved vertebrate enhancer elements that bind Meis/Pbx and HoxB1/HoxA2 proteins [98, 99, 101]. In more anterior r2–r3, Meis3/Pbx proteins are required for the early neural expression of hoxa2 in Xenopus [44]. Studies in mouse and chick suggest that r4 activation of hoxa2 requires Meis3/Pbx proteins; however, it is still an open question for the r2 hoxa2 enhancer regions, which contain Sox binding sites and perhaps is not regulated by Meis/Pbx proteins [98, 99]. In mouse and chick, more posteriorly, hoxa3 expression in r5/6 is controlled by an element binding Meis/Pbx and HoxA3/HoxB3 proteins [102]. In r6/7, the expression of hoxb4 and hoxd4 in mice is also regulated by Meis1/Pbx as well as Hoxb4/Hoxd4-responsive elements after initial induction by retinoic acid signaling [103–105]. Finally, add-ing an additional level of complexity, all rhombomeres fail to develop properly upon Meis3 knockdown in Xenopus, despite the fact that meis3 expression is restricted to r2–4 [42, 44, 47]. The elimination of early r4 fgf3 expression in Meis3-depleted embryos likely triggers these non-autono-mous effects [57]. Similarly, r4-specific fgf3 expression is eliminated in zebrafish pbx/hox mutants [66], and this trig-gers a loss of hindbrain cell fates from r2–r6. In Xenopus or zebrafish, any direct activation of Meis/Pbx/Hox enhancer sequences in more posterior rhombomeres could be medi-ated by more ubiquitously expressed Meis or Prep proteins (Fig. 1).

Non‑homeodomain transcription regulators

Three early expressed transcription factor proteins, vari-ant hepatocyte nuclear factor-1 (Vhnf1), Krox20, and the Mafb/Kreisler/Valentino proteins, are crucial for regulating the earliest expression of hox genes. These transcription factors are thus essential for the establishment of correct


1 3

rhombomeric AP identity in the early hindbrain. Genetic disruption of these proteins severely perturbs the forma-tion of r3–r6. Vhnf1 is the earliest gene expressed in r5/r6. In zebrafish, the Vhnf1 protein induces both krox20 (r5) and mafb (r5/6) expression, which subsequently determines correct rhombomeric identities. Loss of Vhnf1 reduces r5/6 domains while the hoxb1a-expressing r4 region is expanded [106–108]. In mice, a Vhnf1-binding site was identified in a regulatory element of the kreisler gene. Mutating this site results in loss of kreisler induction, indicating that Vhnf1 is essential for rhombomere-specific kreisler expression in the future r5/r6 domain [109] (Fig. 2).

Krox20 is expressed in rhombomeres 3 and 5, and is para-mount for r3/r5 regional identity and cell fate, as directly evident by various knockout or knockdown experiments in all vertebrates [107, 110–112]. In addition to Vhnf1, induc-tion of krox20 is triggered by a positive input from Pbx/Meis and HoxB1 [63, 113–115]. In chick, mice and zebrafish, krox20 promoter/enhancer sites have functional r3-specific Hoxb1/Meis/Pbx binding sites, although in Xenopus it does not appear to be a direct Meis3-target [44, 63, 64]. Follow-ing the upregulation of krox20 via these multiple regula-tors, it acts in a positive auto-regulatory loop to maintain its own expression. A recent study revealed that one of the krox20 regulatory regions is required not only to initiate its

expression but also to enable the auto-regulatory binding site to function [116]. This region acts with additional cis-regulatory elements, some of which are positioned further away from the krox20 loci. These multiple regulatory modes ensure a proper level of krox20 auto-regulation in r3 and r5. Notably, another member of Egr family, Egr4, was recently identified in Xenopus to be expressed in the posterior hind-brain [117]. Knockdown experiments uncovered a role for Egr4 in krox20 and mafb upregulation in r5 or r5/r6, respec-tively, indicating that in the frog, Egr4 mediates the effect of Vhnf1 to activate these two genes in the posterior hindbrain (Fig. 2).

Initially, Hoxb1 activates krox20 by binding to its enhancer. Later, when Hoxb1 expression levels increase and are restricted to r4, it represses krox20 expression. Recipro-cally, Vhnf1 and Krox20 proteins repress hoxb1 to limit its expression to r4. These dual-negative regulatory interactions lead to the establishment of well-defined rhombomeric iden-tities [107, 108, 112]. Moreover, Krox20 protein directly activates the expression of the ephA4 receptor gene in r3/r5, which in turn sharpens the rhombomeric borders by prevent-ing cell intermixing between segments [118–120]. Interest-ingly, two negative regulators of krox20 gene expression, Nab and Nlz (also known as Neurl1a), have been identified in zebrafish, mice and chick hindbrains [121–123]. These

Fig. 2 Upstream and downstream regulation of Krox20 in the hind-brain. A Induction of Krox20 gene expression in r3/r5 is initiated by FGF signaling. FGF induces Vhnf1 in r5/r6 that upregulates MafB, leading to Krox20 induction in r5. In r3, FGF induces Krox20 expres-sion in a Vhnf1-independent manner. Krox20 expression is restricted to the correct rhombomeres by Pax6, which is also induced by FGF. Pax6, which is expressed in r3/r5 is upregulated, and acts as a nega-tive regulator of Krox20 expression via the induction of Nab1. NLZ is another Krox20-negative regulator that is also co-expressed in r3/r5. In parallel, Hoxb1 is expressed in r4 and represses Krox20 expression in r4. Following the upregulation of Krox20 in r3/r5, Krox20 regu-lates PG-2 Hox gene expression in r3/r5. It also induces the expres-sion of its negative modulator Nab1. In addition, Krox20 represses

Hoxb1 expression in r3/r5, leading to accurate r3/r4/r5 identities. In early stages of development, Krox20 transiently activates Hoxb1 (not shown). B FGF acts upstream of Krox20 in the chick hindbrain. (a) Krox20 is expressed in control embryos, but is missing in (b) hind-brains treated with the FGF receptor inhibitor SU5402 (from Weis-inger et al. [142]). C Pax6 is a negative modulator of Krox20 that restricts its expression to r3/r5 in the chick hindbrain. (a) Hind-brains electroporated with control plasmid show normal Krox20 expression. (b) Overexpression (OE) of Pax6 leads to a reduction in Krox20 expression (arrows). (c) Expression of a dominant-negative (DN) Pax6 results in the expansion of Krox20 (arrows) to additional domains (from Kayam et al. [124])


1 3

factors co-localize with krox20, but repress its transcription, restricting its expression to the odd-numbered rhombomeres. Krox20 itself is involved in mediating its negative transcrip-tional regulation by acting as a SUMO ligase that simulates Nab protein activity [123]. Moreover, Hoxb1 was found to upregulate nlz and thus indirectly repress krox20 expression [115]. The paired-rule gene pax6 is also co-expressed with krox20 in r3/r5 but negatively regulates it, leading to the stabilization of krox20 expression borders in r3/r5 [124]. Pax6 protein negatively modulates krox20 gene expression via its ability to induce nab1 gene expression; nab1 protein subsequently binds to krox20 regulatory elements to repress its expression (Fig. 2).

Mafb/Kreisler/Valentino proteins are expressed in r5/r6, and are obligatory for their specification. In mafb mutants, r5/6 identity is lost and r4 is expanded [125, 126]. Mafb and Vhnf1 mutually regulate gene expression in a positive feedback loop. Mafb protein regulates ephrinB2a expres-sion, which represses r4, and further promotes r5/6 iden-tities [108, 127]. Vhnf1 together with Mafb and Krox20 also activates hoxa3 and hoxb3 expression, which is cru-cial to specify r5/6 identity [108, 128, 129]. In zebrafish embryos, krox20 transcriptional activation also requires the Vhnf1 protein, which synergizes with FGF activity to induce mafb/kreisler/valentino and krox20 gene expression. Vhnf1 protein in turn represses hoxb1 gene expression, limiting its expression to r4, thus enabling formation of more posterior rhombomeric fates [107, 108]. Subsequently, Vhnf1 together with Mafb and Krox20 activate hoxa3 and hoxb3 expression in r5/6 to specify r5/6 identity [108, 128] (Fig. 2).

Signaling pathways induce the hindbrain

FGF signaling

FGF signaling plays a key role in hindbrain induction and patterning. Pioneer studies in the chick embryo revealed that caudal epiblast cells fated to give rise to hindbrain charac-ter are located adjacent and lateral to the anterior primitive streak of stage 4 embryos [130, 131]. As cells in and around the streak express different FGFs [132, 133], a possibility was raised that FGFs participate in the induction of the hind-brain. Exposure of neural plate explants from different stages and axial levels to FGFs confirmed that FGF signaling is required, although not sufficient, to induce cells of hindbrain character [134].

In zebrafish embryos, the FGF3 and FGF8 genes are expressed in the presumptive hindbrain primordia at 80–90% epiboly, before the onset of rhombomere seg-mentation [135, 136]. At segmentation initiation, FGF3/8 transcripts are expressed in the central r4 region, with r3/r5 overlap. When morphological segments have formed,

FGF3 continues to be expressed in r4, but FGF8 expres-sion is extinguished, with newly shifted expression to the more anterior isthmus region. FGF3/8 activities appear to be functionally redundant since a strong hindbrain phenotype is only seen in zebrafish embryos co-injected with morpholino oligonucleotides (MO) to both genes. FGF3/8 knockdown severely inhibited formation of all rhombomeres, except r4. In FGF3/8 morphant embryos, initial hoxB1 expression in r4 was normal but declined with time, suggesting that neigh-boring rhombomeres are necessary to maintain its expres-sion. However, in FGF3/8 morphants, neurons derived for r1–3 and r5–7 are disrupted, but r4-derived neurons formed fairly typically. The possibility that FGFs are acting redun-dantly in the hindbrain is also reinforced in mice, where otic-placode induction is severely perturbed in FGF3 null embryos, but hindbrain segmentation remains normal [137].

The expression of FGF3 in hindbrain primordia is con-served in vertebrates [133, 135, 136, 138–141]. Yet, in con-trast to the r4-restricted expression in Xenopus and zebrafish, in chick and mice FGF3 is also evident in r2 and r6 [105, 120, 140–142]. It was suggested that early FGF3/8 signaling in r4 forms a primary hindbrain-inducing center since trans-planted r4 cells or ectopic expression of FGF3/8 induces expression of r5/6-specific markers in chick and zebrafish embryos [135, 143]. In Xenopus, FGF8 is also involved in hindbrain induction; one of its splice forms, FGF8a, medi-ates hindbrain specification since its knockdown severely perturbs formation of hindbrain and other posterior neural fates [25]. Noticeably, studies in chick and mice demon-strate that much after rhombomere specification takes place, FGF3 is downregulated from r2/r4/r6 but maintained in all rhombomere boundaries [120, 144]. Rhombomere bounda-ries display unique cellular and molecular properties that are different from rhombomere bodies [145–149]. In chick, the expression of different boundary markers requires FGF3 [144]. Recent findings suggested that rhombomere bound-aries serve as pools of neural-stem-like cells that express Sox2 and nestin and contribute neurons to the hindbrain at stages when rhombomere cells are actively differentiating [150]. Whether FGF signaling is also required for the devel-opment and/or maintenance of these neural stem cells in the hindbrain awaits further studies.

A tight cross-talk exists between different transcription factors and FGF signaling in the hindbrain. These interac-tions play critical early roles in hindbrain induction and seg-mental patterning. For instance, the activity of Meis proteins is required to induce FGF3 expression as Meis3 knockdown in Xenopus resulted in elimination of early FGF3 expression in r4 [57]. As a consequence, the entire hindbrain failed to form due to the loss in FGF signaling [42, 151], and Meis3 protein cannot induce hindbrain marker expression in the absence of FGF signaling [24, 151]. Moreover, in an attempt to screen for Hoxb1 target genes in zebrafish, a novel gene,


1 3

ppp1r14al, was identified that is induced by Hoxb1 in r4. Ppp1r14al in turn regulates FGF3 expression in r4, indi-cating that it is also essential for the establishment of the earliest hindbrain signaling center in r4 [46]. The PG1–3 Hox groups, Vhnf1, Krox20 and Kreisler/MafB/Valentino proteins all act downstream of FGF signaling to induce hindbrain segments. For instance, FGF3 signal indirectly activates hoxa2, hoxb2 and hoxb3 expression via induction of krox20 gene expression [128, 129, 132, 143, 152]. FGF signaling also up-regulates vhnf1 gene expression, which in zebrafish controls caudal hindbrain specification by syner-gizing with FGF activity to induce mafb/kreisler and krox20 expression. Upon FGF signaling activation, Mafb and Vhnf1 proteins are upregulated and bind to specific enhancer ele-ments in the krox20 promoter. This transcriptional regulation is required for initial krox20 expression in r3/r5, but not for its later maintenance [107, 113]. In addition, Vhnf1 protein also appears to activate fgf3 expression and to repress hoxB1 gene expression, presumably to exclusively limit its expres-sion to r4. Thus, in the hindbrain, the FGF-inducing center acts at a pivotal position, being downstream to Meis/Pbx/Vhnf1/Hox PG1 proteins, but upstream to PG2–3 Hox and Vhnf1/Krox20/Kreisler gene activity.

Studies in the chick hindbrain demonstrated that FGF signaling activates the MAP kinase signal transduction pro-tein ERK1/2 that in turn induces the expression of the Ets-family transcription factor pea3. This signaling cascade is required for krox20 induction in the early hindbrain [142, 153, 154]. FGF3 was also found to induce the expression of pax6 in r3/r5, that in turn negatively regulates krox20 expression, via the induction of nab1 [124]. Thus, by regu-lating both krox20 and pax6, which mutually repress each other, FGF3 acts as a guardian to sharply define rhombomere borders. Moreover, Sprouty4 protein, which is also induced throughout the hindbrain by FGF, acts in a negative-feed-back loop to define sharp rhombomeric domains. Enhanced FGF signaling by sprouty4 knockdown triggers premature and ectopic krox20 gene expression fusing r3 with r5 that eliminate r4 cell fates [115]. Thus, a multitude of molecular interactions are modulated via the FGF pathway to provide an accurate positional identity of hindbrain rhombomeres.

Wnt signaling

Many Wnt ligands, such as Wnt-1, -3, -3a, -4, -8, -8b and -10, are expressed in the developing CNS [33, 155–161]. Their role in posterior neural development was first shown in Wnt1 and Wnt3a knockout mouse embryos, in which midbrain, hindbrain and spinal cord structures were poorly developed [162–164]. Wnt3 null mouse embryos had pos-terior truncations, with expanded expression of the otx2 forebrain marker, and a concomitant loss of expression

of the hindbrain hoxb1 marker [165]. Similar results were seen in chick embryos implanted with beads soaked with a soluble form of the Frz receptor (mFrz8-CRD) that antag-onizes endogenous Wnt ligand signaling. These embryos had an expansion of forebrain markers (otx2 and pax6) and a down-regulation of hindbrain (gbx2) markers [166]. In cultured chick neural explants, mFrz8-CRD also elimi-nated expression of the gbx2, krox20 and hoxb4 hindbrain markers [166, 167]. Similar results were also confirmed in Xenopus embryos, where expression of dominant-negative Wnt proteins that inhibit canonical Wnt activity anteriorized embryos [34]. Like Wnt3−/− mouse embryos, these embryos also exhibited neural tube closure defects and expression of the xanf1 and otx2 forebrain markers was posteriorly expanded, while expression of the hindbrain krox20 marker was reduced [34]. Specific Wnt3a-MO targeting suggested a role in hindbrain development in both zebrafish and Xenopus embryos [44, 168–170]. Targeted mesodermal Wnt3a knock-down in Xenopus ablated hindbrain formation [44]. Wnt3a morphant embryos exhibited neural convergent and exten-sion defects, having the typical caudal expansion of fore-brain markers, with depletion of hindbrain markers. Moreo-ver, during late gastrula stages, expression of homeoproteins that regulate early hindbrain induction such as meis3, hoxd1 and gbx2 was reduced [44]. Wnt3a MO pheno-copied the neural phenotypes of the general canonical Wnt inhibitor Dkk1 [44], further supporting a role for Wnt3a as the pri-mary posterior neural inducer in vertebrates. In contrast, zebrafish mutated for both Wnt1/Wnt10b did not have abnor-mal hindbrain morphology [171, 172], while knockdown of Wnt3a or Wnt8b did affect hindbrain patterning in zebrafish [172], indicating a potential redundancy in the function of different Wnt ligands for normal hindbrain patterning.

Complementing these loss-of-function studies, Wnt gain-of-function activity induces hindbrain neural cell fates. The Xenopus animal cap (AC) explant system has provided a great tool for examining the role of Wnt signaling in neural patterning. BMP4 antagonism in AC explants induces neural tissue. Such neuralized explants mimic the initial state of the newly induced embryonic neural plate. AC explants express pan-neural and anterior neural markers, and will develop as anterior forebrain/cement gland in the absence of additional caudalizing signals [10]. Neuralized AC explants overex-pressing different Wnt ligands or downstream effectors, such as β-catenin or inducible constitutively active Tcf, robustly induced expression of hindbrain markers, while strongly repressing anterior neural marker expression [33, 35, 44, 173–175]. In addition, expression of the earliest hindbrain specifying homeoproteins, meis3, hoxd1, hoxa2 and gbx2, along with caudalizing FGF3 and FGF8 genes, were also induced in gastrula-stage-neuralized AC explants overex-pressing either Wnt3a or β-catenin [44, 176]. Chick fore-brain explants were also caudally transformed by Wnt3a


1 3

added to the culture medium (supplemented with FGF), as evident by the induced expression of gbx2 and krox20, instead of otx2 [166]. Canonical Wnt induction of posterior neural cell fates was shown in both frogs and zebrafish to occur specifically during mid-late gastrula stages [35, 44, 169, 170].

Zebrafish embryos overexpressing a heat-shock protein (Hsp)–Wnt8 driver-plasmid induced at gastrula stages had an anterior shift in hindbrain markers, with forebrain mark-ers pushed to the anterior extremity [169]. Zebrafish head-less mutants lacked midbrain, eye and forebrain tissues and weakly expressed anf1, six3 and rx3 forebrain markers. Concomitantly, krox20 expression was expanded anteriorly [177]. Headless was identified as a point mutation in the Wnt downstream negative-effector Tcf3 gene, and mutant TCF3 protein was unable to translocate to the nucleus or to bind DNA, thus causing a loss-of-function phenotype [177]. In cells where the canonical Wnt pathway is not activated, Tcf3 represses expression of Wnt-pathway target genes. The loss of Tcf3-mediated repression in the headless mutants reflects an overactive canonical Wnt pathway in the embryos [177]. Xenopus embryos overexpressing an inducible β-catenin pro-tein activated at gastrula stages, or a CMV-promoter driving zygotic Wnt3a expression both induced caudalized embryos with ectopic expansion of hindbrain markers anteriorly, and down-regulation of anterior markers [35, 44]. This anterior transformation in morphology and gene expression pattern to more caudal fates was also evident in Dkk1 null mouse embryos [178] and chick embryos implanted with Wnt3a-soaked beads [166].

The Wnt ligands that induce hindbrain from the parax-ial mesoderm may vary between species. In Xenopus, this ligand is Wnt3a, since its specific knockdown inhibits hind-brain formation despite relatively normal wnt8 mesodermal expression levels [44]. In zebrafish, paraxial mesoderm expressed Wnt8a protein is required for hindbrain forma-tion [169]. In mice and chick, Wnt3 and Wnt8 ligands are also expressed during early development. Whether their hindbrain inducing effects are mediated via the mesoderm or directly in the neural plate still needs to be determined.

RA signaling

RA signaling is also seminal for hindbrain patterning. Yet, unlike FGF and Wnt, its availability is largely dependent on diet, as it is produced from vitamin A (reviewed in [179, 180]). The sensitivity of the hindbrain to small perturbations in RA levels has made it an excellent model to study how morphogen gradients govern pattern formation. Studies in frog [32, 74, 181], chick [104, 182], quail [183, 184], mouse [79, 185, 186], rat [187, 188] and zebrafish [189–192] have generated a vast amount of knowledge on the manner by

which RA controls hindbrain regional identity. RA signal-ing is unique among morphogens as it relies not only on its source of production, but on its site of degradation, creating a posteriorhigh–anteriorlow activity gradient.

Initially, the existence of a RA gradient was debatable since depletion of endogenous RA was rescued by applying uniform and high doses of RA throughout the embryo, which resulted in fairly normal hindbrain [180, 193–195]. These findings suggest that a RA gradient is generated not only through diffusion from its posterior source but also from its anterior inactivation. Indeed, there are two main groups of metabolic enzymes that coordinate RA levels. RALDHs convert retinaldehyde into RA, whereas CYP26s inactivate RA via oxidation [196–199]. RA is initially synthesized in the presomitic mesoderm by RALDH, and it diffuses into the adjacent hindbrain in a posteriorhigh–anteriorlow gradi-ent. CYP26 is expressed in the anterior hindbrain, where it leads to RA degradation, further reinforcing the decrease in RA signaling in the rostral embryo [193–195, 200–205]. RA transduces its effects by binding to RA receptors (RARs) and retinoid X receptor (RXRs) [206–210]. Unlike FGF and Wnt, RA uses a nuclear, rather than a membranal, receptor. Upon binding to RA, the receptors act as transcriptional reg-ulators by directly binding specific RA-responsive elements (RAREs) that are positioned at the 3′ or 5′ ends of different hindbrain patterning genes, such as hoxa1, hoxb1, hoxb4 and vhnf1 [79, 104, 108, 211–219]. For instance, in r6/r7, the initial expression of hoxb4 and hoxd4 is activated through RAREs [103–105]. Interestingly, the early transcription of the hoxb1 gene is initiated by a conserved 3′ RARE, but later, when hoxb1 expression is restricted to r4, its repres-sion in r3 and r5 is mediated by a 5′ RARE [220, 221]. Furthermore, the rarβ gene itself contains a HoxB4/HoxD4-responsive element [105], providing a positive feed-forward loop to maintain RA levels and PG4 Hox expression in r6/7.

In addition to controlling the expression of hindbrain segmentation genes in a graded manner, RA regulates its own metabolism in the same manner. RA downregulates the expression of Raldh genes in a negative feedback loop, while upregulating the expression of Cyp26 in a positive feedback loop [191, 194, 195, 202, 222–225]. Yet, RA also induces its own synthesis via a feed-forward mechanism. Experiments in mice and Xenopus embryos revealed that Hoxa1–Pbx1/2–Meis2 directly binds a specific regulatory element that is required for maintaining Raldh2 expres-sion levels. As RA induces the expression of Hoxa1, this study revealed an indirect autoregulation of RA synthesis via Hoxa1 [226]. Moreover, RA positively regulates the expression of the intracellular RA-binding proteins, Crabps, which mediate RA transfer to Cyp26 to trigger its degrada-tion [194]. Intriguingly, Crabps were also found to promote the delivery of RA to its receptors, thus eliciting RA sign-aling [191, 227, 228]. The duality of RA regulation on its


1 3

own metabolism requires future research to reveal how its autoregulation is controlled in the hindbrain.

Interestingly, while expression of the RA-degrading enzyme Cyp26a1 gene is tightly regulated by the levels of RA [194], two other members of the cyp26 family, cyp26b1 and cyp26c1, are not directly regulated by RA signaling and display an unexpected segmental expression with lower lev-els in r3/r5 than in r2/r4/r6 [193, 195, 229]. A recent study in zebrafish found that this segmental expression plays a key role in sharpening r3–r5 segmental gene expression [229]. During hindbrain segmentation, some r3/r5 (Krox20+) cells are initially found in r4, but later switch into r4 (Hoxb1+) identity. This study found that the krox20-intermingled cells are exposed to lower RA levels in r4 due to the elevated activity of cyp26b1 and cyp26c1, which results in downreg-ulation of krox20 and upregulation of hoxb1. The coupling between segmental gene expression and dynamic levels of RA provides the first evidence describing how a signal like RA that is thought to act mainly in a graded caudal–rostral axis orchestrates boundary sharpening via regulating levels of gene expression in alternating rhombomeres [230].

Visualization of the actual RA gradient in live embryos was needed to fully confirm its existence [231]. Initially, this challenge was tackled by generating transgenic mice or zebrafish embryos where GFP/LacZ expression was driven under the control of RAREs through in vivo injection of RA–GFP fused constructs [193, 210, 232, 233]. These strat-egies supported the shape of a RA gradient, which fitted well with the expression domains of RALDH and CYP26 genes. However, the non-peptidic structure of RA, as well as the failure to observe the GFP signals at early stages, prevented full in vivo validation [194, 210, 234–236]. More recently, imaging tools were developed for this purpose. In one approach, genetically encoded probes for RA (GEPRs) were fused with different GFP variants and introduced into zebrafish. Each GERP displayed a different affinity to RA. Binding of RA to different GERPs led to conformational change that was converted into changes in fluorescence reso-nance energy transfer (FRET) [237]. This strategy confirmed the concentration gradient of RA in live embryos, where the local source and sink jointly establish the highest RA concentration in the mid-trunk and the lowest in the tail and head. A more recent technology utilized fluorescence life-time imaging microscopy and phasor analysis to calculate the relative abundances of RA along the hindbrain [238]. This strategy, which is based on the endogenous fluores-cence of RA, demonstrated that intracellular free RA forms an anteriorly declining gradient similar to that previously reported with FRET or RARE-GFP/LacZ [193, 237]. This sensitive technology also enabled visualization of random fluctuations in RA levels that can vary rapidly within one hindbrain position. This study suggested that individual cells can actively control the magnitude of random fluctuations of

RA levels to preserve the required concentrations for hind-brain segmentation [231].

In general, limiting RA activity results in shortening or loss of the posterior hindbrain and caudal expansion of its anterior part. Yet, the severity of these phenotypes depends on the level in which RA signaling was modified. Complete vitamin A deficiency (VAD) was initially studied in quail embryos, causing a complete loss of r4–r8, and a posterior expansion of r3 [183]. This morphological distortion was combined with loss of posteriorly expressed genes, such as hoxb1, FGF3, and mafB, together with caudal expansion of the r3 stripe of krox20. In rats, VAD was accomplished by a gradual, rather than total reduction of vitamin A [187, 188]. These embryos displayed a correlation between titrated reduced doses of RA and a progressive expansion of anterior rhombomeres at the expense of posterior ones. Reduction of RA-signaling levels was also achieved by modulation of its metabolism. In Xenopus, ectopic Cyp26 protein anteriorized the posterior hindbrain in a dose-dependent manner [239]. Yet, only a partial duplication of anterior rhombomeres was observed in the posterior hindbrain. Similar intermediate effects were found in two zebrafish lines, neckless and no-fin, where the raldh2 gene is mutated [200, 240]. Attenuation of RARs was another means to inhibit RA. This was per-formed by the generation of RAR null mice or zebrafish, or by expressing dominant negative RARs in mice, chick and frog [32, 74, 181, 183, 203, 241–243]. In most cases, expres-sion of posterior genes was delayed and formation of the posterior rhombomeres was disrupted but not lost. Notably, stronger defects were observed upon combination of differ-ent RAR mutations or inhibitors [185–203]. Finally, the less severe hindbrain phenotypes were achieved by manipulating RA-binding sites on target genes. For example, mutations of RAREs in the 3′ ends of hoxa1 and hoxb1 delayed their ini-tial rhombomeric expression, but this was fully restored later in development [78, 79, 213]. These studies demonstrate the complex level of regulation of RA signaling in the hindbrain, which combines different sources for RA synthesis and deg-radation, the activity of several RA receptor subtypes and the binding to RAREs on different hindbrain genes to deliver the correct patterning outcome.

Cross‑talk between hindbrain signaling pathways: cooperative or antagonistic activities?

As mentioned, the role of RA, FGF and Wnt in regulating the expression of key hindbrain genes is well documented. Yet, there are marked differences between these morpho-gens in terms of their expression/activity patterns. Whereas RA signal spans the entire hindbrain in a gradient man-ner [180], expression of FGFs and Wnts is more limited:


1 3

FGF3/FGF8 is expressed in r4 (zebrafish, frog) or r2/r4/r6 (chick and mice) [133, 135, 136, 139, 141]. Wnt8a and Wnt8b are expressed in r4 and r3/r5, respectively [244, 245]. The neural patterning of Wnt3a and Wnt8a ligands, as well as FGFs, is also expressed in the dorsal–lateral mesodermal regions similar to RA, where they are secreted to induce posterior cell fates in adjacent neural tissue (Fig. 3) [44]. How the localized FGF and Wnt signals and the graded RA pathway are integrated to govern the positional identity of the hindbrain is not well understood.

Signaling cooperativity

Interactions mediating the expression of all these signals have been demonstrated.

For example, in VAD-quail embryos or in mice carrying a null mutation for the raldh2 gene, FGF3 expression is lost [183, 246]. This observation indicated that the induc-tion and/or maintenance of FGFs in hindbrain segments involve RA signaling. Moreover, FGF3-null mice have reduced wnt8a expression in r4, whereas excess FGF8 in hindbrain explants induces ectopic wnt8a expression. These

results suggest that the wnt8a expression in the hindbrain is mediated by FGF signaling [247]. Further support for the Wnt–FGF cross-talk was shown by studying sprouty1/2 genes. Sprouty proteins are negative regulators of the FGF signaling pathway and restrict Wnt8a to r4. Mice mutated for Spry1/2 showed expansion of Wnt8a into additional rhom-bomeres, probably via releasing the negative regulation of FGF signaling [248]. However, no gross hindbrain malfor-mations appeared in the Spry1/2 mutants, thus the interac-tive roles of FGF and Wnt in regulating hindbrain induction and patterning are still unclear.

Additional evidence suggests that these morphogens cooperate to induce the hindbrain. In Xenopus embryos and explants, caudalizing canonical Wnt activity regulated early neural FGF3/8 gene expression [35, 57]. Wnt3a protein acti-vates expression of the hindbrain-specific meis3 gene [44]. Meis3 protein then directly activates FGF3/8 gene expres-sion [57, 151]. In Meis3 knockdown embryos, FGF3 expres-sion in r4 is eliminated and hindbrain cell fates are lost [57]. Ectopic FGF ligand expression can partially rescue Meis3 morphant phenotypes in Xenopus embryos [57]. Yet, neither canonical Wnt nor Meis3 protein activities efficiently cau-dalize the CNS, when downstream FGF signaling is com-promised [24, 35, 151]. Moreover, Wnt activation of Meis3 protein modulates expression of RA direct target genes, such as hoxd1 [30, 44, 62]. Hoxd1 is a direct target of RA, Wnt and the Meis3 protein [30, 44, 62, 249, 250]. Meis3/RA act synergistically to optimize hoxd1 gene expression to pro-mote correct hindbrain formation [62].

Combined FGF and RA signaling activity was also found to induce hindbrain cell fate in the chick; exposure of caudal epiblast cells to FGFs led to the induction of krox20 only when caudal paraxial mesoderm was also present [134]. This study suggested that a combination of FGF and paraxial mesoderm-caudalizing activity acts directly on epiblast cells to induce hindbrain character. This activity was later identi-fied as RA [251]. The combined activity of RA and FGFs is also demonstrated in setting up the hindbrain–spinal cord border, as marked by cdx1/4 expression. In zebrafish, loss of cdx1/4 resulted in caudal expansion of hindbrain genes. This phenotype could only be rescued when both FGF and RA were inhibited, suggesting that both signals act together to coordinate the formation of the border between the hind-brain and spinal cord [252]. These observations show that CDX proteins modify the cell competence to respond to both FGFs and RA in the posterior neural tissues, including the hindbrain. Hence, both signals are required to define the precise hindbrain–spinal cord boundary.

At slightly later stages, rhombomere-derived FGFs are required for the specification of r3 and r5, as inhibi-tion of FGF downregulates krox20 expression in both seg-ments [108, 135, 136, 142, 143]. Notably, a recent study in zebrafish elucidated a novel mechanism through which the

Fig. 3 A summary of the expression domains of different signal-ing factors in the hindbrain. The spatial organization of the multiple signaling factors triggering hindbrain induction and patterning with-out temporal separation is shown. Before rhombomere specifica-tion, FGF3/8 and canonical Wnt 3a/8a are expressed in the paraxial mesoderm adjacent to the caudal hindbrain. RA is also synthesized in the same domain and acts in a caudal–rostral gradient along the hind-brain. Slightly later, FGF3/8 and Wnt8a are expressed in r4, whereas Wnt8b is expressed in r3/r5. FGF3/8 and Wnt1/3a are also secreted from the mid–hindbrain boundary


1 3

restriction of krox20 expression to r3/5 is also mediated by different levels of RA in odd versus even segments. This is mediated by the higher expression and activity of Cyp26b in r4, so that Krox20 expressing cells that cross into the r4 territory are exposed to higher levels of RA than in r3/r5, which results in the downregulation of krox20, and the upregulation of hoxb1 gene expression [229]. Yet, although defined RA levels are also necessary for krox20 expression, RA activity is more limited than FGF3; attenuation of RA signaling results in loss of the more posterior r5 stripe of krox20, whereas it remains intact in r3 or even expands pos-teriorly, depending on the degree of RA modulation [108, 183, 188]. Nevertheless, synergism between RA and FGF is required to induce krox20 through the induction of down-stream mediators such as Vhnf1 [108].

From gastrulation stages and onwards, FGF and Wnt are also expressed in the posterior mesoderm, together with RA. All these signals were found to induce posterior and sup-press anterior expression of genes involved in rhombomere specification, as detailed in the previous sections. The meso-dermal FGFs and Wnts prevent the upregulation of cyp26 in the posterior hindbrain [23, 194] and also maintain raldh2 expression in this domain. RA, on the other hand, was found to induce the expression of wnt1/3A and the downstream FGF target gene, pea3 [253]. In this way, posterior FGF and Wnt are involved in maintaining the hindbrain RA gradient, whereas RA positively regulates the expression/activity of these morphogens in more posterior domains. The tight con-nection of FGF and Wnt signaling with RA maintains stable and adaptable RA concentration levels along the hindbrain AP axis [180, 192].

Opposing signaling activities

One of the initial indications supporting a negative cross-talk between hindbrain signals came again from VAD-quail embryos, where the posterior hindbrain was abolished, but anterior regionalization occurred normally [183, 184]. The morphogen regulating patterning in the anterior hindbrain is the mid–hindbrain boundary (MHB)-derived FGF8 protein [212, 254–256]. Grafting of MHB or adding FGF8 beads into the posterior hindbrain induced ectopic expression of genes that are normally expressed anteriorly, together with downregulation of posterior genes, like hoxb1. These results were later confirmed by pharmacologic or genetic disrup-tion of RA in other species, where the posterior hindbrain became anteriorized [179, 242, 257, 258]. Conversely, either inhibition of FGF8 or enhancement of RA activity led to the expansion of anterior Hox genes [256, 259]. Knockout of FGF8 also resulted in midbrain Wnt1 expression expanding posteriorly into the isthmus and cerebellum [260]. These and other studies indicate that posterior RA and anterior FGF8

act in opposing manners to specify Hox-positive/negative domains to define the AP borders of the hindbrain.

Intriguingly, these antagonistic FGF8 and RA activities are similar to those seen in the posterior end of the embryo; FGF3/8 are expressed in neuro-mesodermal progenitor cells at the tail and act in a posteriorhigh–anteriorlow gradient to inhibit their differentiation into mesoderm or ectoderm lineages. RA, which is produced in the more rostral par-axial mesoderm, acts in an opposite anteriorhigh–posterior-low gradient to trigger the differentiation of the progenitors and to induce Hox gene distribution [134, 261–263]. These observations strongly suggest that both signals act in the same direction to pattern the posterior hindbrain, but their interaction is antagonistic in the most anterior or posterior positions along the neural tube. Further support for this con-clusion comes from a study where the distinct effects of FGF and RA were examined on different members of the HoxB group [214]. The anterior expression border of hoxb4, which normally lies in r6/7, was expanded anteriorly upon exog-enous application of RA or by grafting posterior mesoderm adjacent to anterior rhombomeres. However, the expression pattern of the more caudal gene, hoxb9, was not modified upon manipulating RA signaling. Conversely, application or inhibition of FGF did not change the expression pattern of hoxb4 but led to a dramatic expansion or loss of hoxb9 gene expression, respectively.

The complex cross-talk between RA, FGF and Wnt is also evident in the positioning of the border between the hindbrain and the spinal cord, as marked by the anterior expression of cdx1/4 at the level of somite 3. In zebrafish, loss of cdx1/4 causes a caudal shift of the hindbrain–spinal cord border via posterior expansion of hindbrain genes (such as hoxb4), and caudalization of spinal cord markers, such as hoxb8 [252, 264]. Hence, the spatial regulation of cdx1/4 expression is crucial for positioning the hindbrain–spinal cord border region. Analyzing the regulatory role of RA and FGF revealed that in RA-deficient embryos, cdx4 expression shifts dorsally while in FGF-deficient embryos cdx4 expres-sion shifts caudally. Yet, in embryos lacking both RA and FGF signals, the shift in cdx4 axial expression is rescued and it is aligned at the level of somite 3, indicating that FGF neg-atively modulated RA activity in regulating cdx4 expression [252, 265, 266]. Thus, for hindbrain patterning, the FGF and RA pathways have additive functions, but with respect to the axial position of the hindbrain/spinal cord border region, they may be antagonistic. In this way, FGF and RA inter-actions are different in each process: additive in hindbrain patterning, antagonistic in hindbrain size specification and epistatic in neural-mesodermal tissue alignment.

In Wnt-deficient embryos, cdx4 expression shifts caudally, but the simultaneous loss of both Wnt and RA activities results in a more severe caudal shift of cdx4 gene expression, in comparison to solely inhibiting Wnt


1 3

signaling, indicating that Wnt and RA act together to regu-late cdx4 axial positioning in the hindbrain–spinal cord bor-der [170, 215, 267, 268]. These studies clearly demonstrate the multifaceted network of morphogen regulation that is differentially activated or inhibited along the hindbrain AP axis. In the future, it will be important to elucidate in greater detail the spatial and temporal dynamics of these regulatory signaling pathways to fully understand how their joint activi-ties orchestrate rhombomere specification.

Acknowledgements We wish to thank Dr. Yuval Peretz for his help with the illustrations. DF was supported by grants from the Israel Sci-ence Foundation (ISF, 658/15) and the Israel Cancer Research Fund (ICRF). DSD was supported by grants from the ISF (1515/16), The Chief Scientist Office of the Ministry of Health, Israel (3-0000-15441) and United States–Israel Binational Science Foundation (2015087).

References

1. Shoja MM, Johal J, Oakes WJ, Tubbs RS (2018) Embryology and pathophysiology of the Chiari I and II malformations: a compre-hensive review. Clin Anat 31:202–215

2. Shoja MM, Ramdhan R, Jensen CJ, Chern JJ, Oakes WJ, Tubbs RS (2018) Embryology of the craniocervical junction and pos-terior cranial fossa, part II: embryogenesis of the hindbrain. Clin Anat 31:488–500

3. Spemann H, Mangold H (1924) Uber Induktion von Embryonal-anlagen durch Implantation artfremder Organisatoren. Arch mikr Anat Entw mech 100:599–638

4. Oppenheimer JM (1936) Transplantation experiments on devel-oping teleosts (Fundulus and Perca). J Exp Biol 72:409–437

5. Waddington CH (1933) Induction by the primitive streak and its derivatives in the chick. J Exp Biol 10:38–46

6. Beddington RS, Robertson EJ (1999) Axis development and early asymmetry in mammals. Cell 96:195–209

7. Nieuwkoop PD (1952) Activation and organization of the central nervous system in amphibians. III Synthesis of a new working hypothesis. J Exp Zool 120:83–108

8. Eyal-Giladi H (1954) Dynamic aspects of neural induction. Arc Biol 65:180–259

9. Toivonen S, Saxen L (1968) Morphogenetic interaction of pre-sumptive neural and mesodermal cells mixed in different ratios. Science 159:539–540

10. Lamb TM, Knecht AK, Smith WC, Stachel SE, Economides AN, Stahl N, Yancopolous GD, Harland RM (1993) Neural induction by the secreted polypeptide noggin. Science 262:713–718

11. Hemmati-Brivanlou A, Kelly OG, Melton DA (1994) Follistatin, an antagonist of activin, is expressed in the Spemann organizer and displays direct neuralizing activity. Cell 77:283–295

12. Sasai Y, Lu B, Steinbeisser H, De Robertis EM (1995) Regu-lation of neural induction by the Chd and Bmp-4 antagonistic patterning signals in Xenopus. Nature 376:333–336

13. Re’em-Kalma Y, Lamb T, Frank D (1995) Competition between noggin and bone morphogenetic protein 4 activities may regulate dorsalization during Xenopus development. Proc Natl Acad Sci USA 92:12141–12145

14. Fainsod A, Deissler K, Yelin R, Marom K, Epstein M, Pillemer G, Steinbeisser H, Blum M (1997) The dorsalizing and neural inducing gene follistatin is an antagonist of BMP-4. Mech Dev 63:39–50

15. Zimmerman LB, De Jesus-Escobar JM, Harland RM (1996) The Spemann organizer signal noggin binds and inactivates bone morphogenetic protein 4. Cell 86:599–606

16. Piccolo S, Sasai Y, Lu B, De Robertis EM (1996) Dorsoventral patterning in Xenopus: inhibition of ventral signals by direct binding of chordin to BMP-4. Cell 86:589–598

17. Hemmati-Brivanlou A, Melton DA (1994) Inhibition of activin receptor signaling promotes neuralization in Xenopus. Cell 77:273–281

18. Wilson PA, Hemmati-Brivanlou A (1995) Induction of epidermis and inhibition of neural fate by Bmp-4. Nature 376:331–333

19. Wilson SI, Rydstrom A, Trimborn T, Willert K, Nusse R, Jessell TM, Edlund T (2001) The status of Wnt signalling regulates neu-ral and epidermal fates in the chick embryo. Nature 411:325–330

20. Cox WG, Hemmati-Brivanlou A (1995) Caudalization of neural fate by tissue recombination and bFGF. Development 121:4349–4358

21. Lamb TM, Harland RM (1995) Fibroblast growth factor is a direct neural inducer, which combined with noggin generates anterior-posterior neural pattern. Development 121:3627–3636

22. Holowacz T, Sokol S (1999) FGF is required for posterior neural patterning but not for neural induction. Dev Biol 205:296–308

23. Kudoh T, Wilson SW, Dawid IB (2002) Distinct roles for Fgf, Wnt and retinoic acid in posteriorizing the neural ectoderm. Development 129:4335–4346

24. Ribisi S, Mariani FV, Aamar E, Lamb TM, Frank D, Harland RM (2000) Ras-mediated FGF signaling is required for the formation of posterior but not anterior neural tissue in Xenopus laevis. Dev Biol 227:183–196

25. Fletcher RB, Baker JC, Harland RM (2006) FGF8 spliceforms mediate early mesoderm and posterior neural tissue formation in Xenopus. Development 133:1703–1714

26. Durston AJ, Timmermans JP, Hage WJ, Hendriks HF, de Vries NJ, Heideveld M, Nieuwkoop PD (1989) Retinoic acid causes an anteroposterior transformation in the developing central nervous system. Nature 340:140–144

27. Sive HL, Draper BW, Harland RM, Weintraub H (1990) Identi-fication of a retinoic acid-sensitive period during primary axis formation in Xenopus laevis. Genes Dev 4:932–942

28. Sharpe CR (1991) Retinoic acid can mimic endogenous signals involved in transformation of the Xenopus nervous system. Neu-ron 7:239–247

29. Ruiz i Altaba A, Jessell TM (1991) Retinoic acid modifies the pattern of cell differentiation in the central nervous system of neurula stage Xenopus embryos. Development 112:945–958

30. Kolm PJ, Sive H (1995) Hindbrain patterning requires retinoid signaling. Dev Biol 192:1–16

31. Papalopulu N, Kintner C (1996) A posteriorising factor, retinoic acid, reveals that anteroposterior patterning controls the timing of neuronal differentiation in Xenopus neuroectoderm. Develop-ment 122:3409–3418

32. Blumberg B, Bolado J Jr, Moreno TA, Kintner C, Evans RM, Papalopulu N (1997) An essential role for retinoid signaling in anteroposterior neural patterning. Development 124:373–379

33. McGrew LL, Lai CJ, Moon RT (1995) Specification of the anteroposterior neural axis through synergistic interaction of the Wnt signaling cascade with noggin and follistatin. Dev Biol 172:337–342

34. McGrew LL, Hoppler S, Moon RT (1997) Wnt and FGF path-ways cooperatively pattern anteroposterior neural ectoderm in Xenopus. Mech Dev 69:105–114

35. Domingos PM, Itasaki N, Jones CM, Mercurio S, Sargent MG, Smith JC, Krumlauf R (2001) The Wnt/beta-catenin pathway posteriorizes neural tissue in Xenopus by an indirect mechanism requiring FGF signaling. Dev Biol 239:148–160


1 3

36. Schulte D, Frank D (2014) TALE transcription factors during early development of the vertebrate brain and eye. Dev Dyn 24:99–116

37. Krumlauf R (2016) Hox genes and the hindbrain: a study in seg-ments. Curr Top Dev Biol 116:581–596

38. Hernandez-Miranda LR, Müller T, Birchmeier C (2017) The dor-sal spinal cord and hindbrain: from developmental mechanisms to functional circuits. Dev Biol 432:34–42

39. Parker HJ, Krumlauf R (2017) Segmental arithmetic: summing up the Hox gene regulatory network for hindbrain development in chordates. WIRE Dev Biol. https ://doi.org/10.1002/wdev.286

40. Burglin T (1997) Analysis of TALE superclass homeobox genes (MEIS, PBC, KNOX, Iroquois, TGIF) reveals a novel domain conserved between plants and animals. Nucleic Acids Res 25:4173–4180

41. Pöpperl H, Rikhof H, Chang H, Haffter P, Kimmel CB, Moens CB (2000) lazarus is a novel pbx gene that globally mediates hox gene function in zebrafish. Mol Cell 6:255–267

42. Dibner C, Elias S, Frank D (2001) XMeis3 protein activity is required for proper hindbrain patterning in Xenopus laevis embryos. Development 128:3415–3426

43. Waskiewicz AJ, Rikhof HA, Hernandez RE, Moens CB (2001) Zebrafish Meis functions to stabilize Pbx proteins and regulate hindbrain patterning. Development 128:4139–4151

44. Elkouby YM, Elias S, Casey ES, Blythe SA, Tsabar N, Klein PS, Root H, Liu KJ, Frank D (2010) Mesodermal Wnt sign-aling organizes the neural plate via Meis3. Development 137:1531–1541

45. Rohrschneider MR, Elsen GE, Prince VE (2007) Zebrafish Hoxb1a regulates multiple downstream genes including prick-le1b. Dev Biol 309:358–372

46. Choe S, Zhang X, Hirsch N, StraubhaarJ Sagerstrom CG (2011) A screen for hoxb1-regulated genes identifies ppp1r14al as a regulator of the rhombomere 4 Fgf-signaling center. Dev Biol 358:356–367

47. Salzberg A, Elias S, Nachaliel N, Bonstein L, Henig C, Frank D (1999) A Meis family protein caudalizes neural cell fates in Xenopus. Mech Dev 80:3–13

48. Vlachakis N, Choe SK, Sagerstrom CG (2001) Meis3 syner-gizes with Pbx4 and Hoxb1b in promoting hindbrain fates in the zebrafish. Development 128:1299–1312

49. Choe S, Vlachakis N, Sagerstrom CG (2002) Meis family pro-teins are required for hindbrain development in the zebrafish. Development 129:585–595

50. Maeda R, Ishimura A, Mood K, Park EK, Buchberg AM, Daar IO (2002) Xpbx1b and Xmeis1b play a collaborative role in hind-brain and neural crest gene expression in Xenopus embryos. Proc Natl Acad Sci USA 99:5448–5453

51. Oulad-Abdelghani M, Chazaud C, Bouillet P, Sapin V, Cham-bon P, Dolle P (1997) Meis2, a novel mouse Pbx-related home-obox gene induced by retinoic acid during differentiation of P19 embryonal carcinoma cells. Dev Dyn 210:173–183

52. Cecconi F, Proetzel G, Alvarez-Bolado G, Jay D, Gruss P (1997) Expression of Meis2, a Knotted-related murine homeobox gene, indicates a role in the differentiation of the forebrain and the somatic mesoderm. Dev Dyn 210:184–190

53. Maeda R, Mood K, Jones TL, Aruga J, Buchberg AM, Daar IO (2001) Xmeis1, a protooncogene involved in specifying neural crest cell fate in Xenopus embryos. Oncogene 20:1329–1342

54. Zerucha T, Prince VE (2001) Cloning and developmental expression of a zebrafish meis2 homeobox gene. Mech Dev 102:247–250

55. Bumsted-O’Brien KM, Hendrickson A, Haverkamp S, Ashery-Padan R, Schulte D (2007) Expression of the homeodomain tran-scription factor Meis2 in the embryonic and postnatal retina. J Comp Neurol 505:58–72

56. Sanchez-Guardado LO, Irimia M, Sanchez-Arrones L, Bur-guera D, Rodrıguez-Gallardo L, Garcia-Fernandez J, Puelles L, Ferran JL, Hidalgo-Sanchez M (2011) Distinct and redundant expression and transcriptional diversity of MEIS gene paralogs during chicken development. Dev Dyn 240:1475–1492

57. Gutkovich YE, Ofir R, Elkouby YM, Dibner C, Gefen A, Elias S, Frank D (2010) Xenopus Meis3 protein lies at a nexus down-stream to Zic1 and Pax3 proteins, regulating multiple cell-fates during early nervous system development. Dev Biol 338:50–62

58. Ferretti E, Cambronero F, Tumpel S, Longobardi E, Wiede-mann LM, Blasi F, Krumlauf R (2005) Hoxb1 enhancer and control of rhombomere 4 expression: complex interplay between PREP1- PBX1-HOXB1 binding sites. Mol Cell Biol 25:8541–8552

59. Ferretti E, Marshall H, Popperl H, Maconochie M, Krumlauf R, Blasi F (2000) Segmental expression of Hoxb2 in r4 requires two separate sites that integrate cooperative interactions between Prep1, Pbx and Hox proteins. Development 127:155–166

60. Maconochie MK, Nonchev S, Studer M, Chan SK, Popperl H, Sham MH, Mann RS, Krumlauf R (1997) Cross-regulation in the mouse HoxB complex: the expression of Hoxb2 in rhombomere 4 is regulated by Hoxb1. Genes Dev 11:1885–1895

61. Jacobs Y, Schnabel CA, Cleary ML (1999) Trimeric associa-tion of Hox and TALE homeodomain proteins mediates Hoxb2 hindbrain enhancer activity. Mol Cell Biol 19:5134–5142

62. Dibner C, Elias S, Ofir R, Souopgui J, Kolm PJ, Sive H, Pieler T, Frank D (2004) The Meis3 protein and retinoid signaling interact to pattern the Xenopus hindbrain. Dev Biol 271:75–86

63. Wassef MA, Chomette D, Pouilhe M, Stedman A, Havis E, Dinh CD, Schneider- Maunoury S, Gilardi-Hebenstreit P, Charnay P, Ghislain J (2008) Rostral hindbrain patterning involves the direct activation of a Krox20 transcriptional enhancer by Hox/Pbx and Meis factors. Development 135:3369–3378

64. Stedman A, Lecaudey V, Havis E, Anselme I, Wassef M, Gilardi- Hebenstreit P, Schneider-Maunoury S (2009) A functional interaction between Irx and Meis patterns the anterior hindbrain and activates krox20 expression in rhombomere 3. Dev Biol 327:566–577

65. Popperl H, Bienz M, Studer M, Chan SK, Aparicio S, Brenner S, Mann RS, Krumlauf R (1995) Segmental expression of Hoxb-1 is controlled by a highly conserved autoregulatory loop depend-ent upon exd/pbx. Cell 81:1031–1042

66. Waskiewicz AJ, Rikhof HA, Moens CB (2002) Eliminating zebrafish pbx proteins reveals a hindbrain ground state. Dev Cell 3:723–733

67. Chan SK, Popperl H, Krumlauf R, Mann RS (1996) An extraden-ticle-induced conformational change in a HOX protein over-comes an inhibitory function of the conserved hexapeptide motif. EMBO J 15:2476–2487

68. Wilkinson DG, Bhatt S, Cook M, Boncinelli E, Krumlauf R (1989) Segmental expression of Hox-2 homoeobox containing genes in the developing mouse hindbrain. Nature 341:405–409

69. Sundin OH, Busse HG, Rogers MB, Gudas LJ, Eichele G (1990) Region-specific expression in early chick and mouse embryos of Ghox-lab and Hox 1.6, vertebrate homeobox-containing genes related to Drosophila labial. Development 108:47–58

70. Frohman MA, Boyle M, Martin GR (1990) Isolation of the mouse Hox-2.9 gene; analysis of embryonic expression suggests that positional information along the anterior-posterior axis is specified by mesoderm. Development 110:589–607

71. Murphy P, Hill RE (1991) Expression of the mouse labial-like homeobox-containing genes, Hox 2.9 and Hox 1.6, during seg-mentation of the hindbrain. Development 111:61–74

72. Frohman MA, Martin GR (1992) Isolation and analysis of embry-onic expression of Hox-4.9, a member of the murine labial-like gene family. Mech Dev 38:55–67

https://doi.org/10.1002/wdev.286


1 3

73. Godsave S, Dekker EJ, Holling T, Pannese M, Boncinelli E, Durston A (1994) Expression patterns of Hoxb genes in the Xenopus embryo suggest roles in anteroposterior specification of the hindbrain and in dorsoventral patterning of the mesoderm. Dev Biol 166:465–476

74. Kolm PJ, Apekin V, Sive H (1997) Xenopus hindbrain patterning requires retinoid signaling. Dev Biol 192:1–16

75. Rossel M, Capecchi MR (1999) Mice mutant for both Hoxa1 and Hoxb1 show extensive remodeling of the hindbrain and defects in craniofacial development. Development 126:5027–5040

76. McClintock JM, Kheirbek MA, Prince VE (2002) Knockdown of duplicated zebrafish hoxb1 genes reveals distinct roles in hind-brain patterning and a novel mechanism of duplicate gene reten-tion. Development 129:2339–2354

77. McNulty CL, Peres JN, Bardine N, van den Akker WM, Durston AJ (2005) Knockdown of the complete Hox paralogous group 1 leads to dramatic hindbrain and neural crest defects. Develop-ment 132:2861–2871

78. Gavalas A, Studer M, Lumsden A, Rijli FM, Krumlauf R, Cham-bon P (1998) Hoxa1 and Hoxb1 synergize in patterning the hind-brain, cranial nerves and second pharyngeal arch. Development 125:1123–1136

79. Studer M, Gavalas A, Marshall H, Ariza-McNaughton L, Rijli FM, Chambon P, Krumlauf R (1998) Genetic interactions between Hoxa1 and Hoxb1 reveal new roles in regulation of early hindbrain patterning. Development 125:1025–1036

80. Remacle S, Abbas L, de Backer O, Pacico N, Gavalas A, Gofflot F, Picard JJ, Rezsohazy R (2004) Loss of function but no gain of function caused by amino acid substitutions in the hexapeptide of Hoxa1 in vivo. Mol Cell Biol 24:8567–8575

81. Makki N, Capecchi MR (2011) Identification of novel Hoxa1 downstream targets regulating hindbrain, neural crest and inner ear development. Dev Biol 357:295–304

82. Tischfield MA, Bosley TM, Salih MA, Alorainy IA, Sener EC, Nester MJ, Oystreck DT, Chan WM, Andrews C, Erickson RP, Engle EC (2015) Homozygous HOXA1 mutations disrupt human brainstem, inner ear, cardiovascular and cognitive development. Nat Genet 37:1035–1037

83. Prince V, Lumsden A (1994) Hoxa-2 expression in normal and transposed rhombomeres: independent regulation in the neural tube and neural crest. Development 120:911–923

84. Gavalas A, Davenne M, Lumsden A, Chambon P, Rijli FM (1997) Role of Hoxa-2 in axon pathfinding and rostral hindbrain patterning. Development 124:3693–3702

85. Barrow JR, Stadler HS, Capecchi MR (2000) Roles of Hoxa1 and Hoxa2 in patterning the early hindbrain of the mouse. Develop-ment 127:933–944

86. Ren S, Angran P, Rijli FM (2002) Targeted insertion results in a rhombomere 2- specific Hoxa2 knockdown and ectopic activation of Hoxa1 expression. Dev Dyn 225:305–315

87. Oury F, Murakami Y, Renaud J, Pasqualetti M, Charnay P, Ren S, Rijli FM (2006) Hoxa2- and rhombomere-dependent development of the mouse facial somatosensory map. Science 313:1408–1413

88. Davenne M, Maconochie MK, Neun R, Pattyn A, Chambon P, Krumlauf R, Rijli FM (1999) Hoxa2 and Hoxb2 control dors-oventral patterns of neuronal development in the rostral hind-brain. Neuron 22:677–691

89. McEllin JA, Alexander TB, Tümpel S, Wiedemann LM, Krum-lauf R (2016) Analyses of fugu hoxa2 genes provide evidence for subfunctionalization of neural crest cell and rhombomere cis-regulatory modules during vertebrate evolution. Dev Biol 409:530–542

90. Wong EY, Wang XA, Mak SS, Sae-Pang JJ, Ling KW, Fritzsch B, Sham MH (2011) Hoxb3 negatively regulates Hoxb1 expres-sion in mouse hindbrain patterning. Dev Biol 352:382–392

91. Gaufo GO, Thomas KR, Capecchi MR (2003) Hox3 genes coor-dinate mechanisms of genetic suppression and activation in the generation of branchial and somatic motoneurons. Development 130:5191–5201

92. Horan GS, Ramırez-Solis R, Featherstone MS, Wolgemuth DJ, Bradley A, Behringer RR (1995) Compound mutants for the par-alogous hoxa-4, hoxb-4, and hoxd-4 genes show more complete homeotic transformations and a dose-dependent increase in the number of vertebrae transformed. Genes Dev 9:1667–1677

93. De Kumar B, Parker HJ, Paulson A, Parrish ME, Zeitlinger J, Krumlauf R (2017) Hoxa1 targets signaling pathways during neural differentiation of ES cells and mouse embryogenesis. Dev Biol 432:151–164

94. De Kumar B, Parker HJ, Paulson A, Parrish ME, Pushel I, Singh NP, Zhang Y, Slaughter BD, Unruh JR, Florens L, Zeitlinger J, Krumlauf R (2017) HOXA1 and TALE proteins display cross-regulatory interactions and form a combinatorial binding code on HOXA1 targets. Genome Res 27:1501–1512

95. Choe S, Lu P, Nakamura M, Lee J, Sagerstrom CG (2009) Meis cofactors control HDAC and CBP accessibility at Hox-regulated promoters during zebrafish embryogenesis. Dev Cell 17:561–567

96. Choe SK, Ladam F, Sagerström CG (2014) TALE factors poise promoters for activation by Hox proteins. Dev Cell 28:203–211

97. Grice J, Noyvert B, Doglio L, Elgar G (2015) A simple predic-tive enhancer syntax for hindbrain patterning is conserved in vertebrate genomes. PLoS One 10:e0130413

98. Tümpel S, Cambronero F, Sims C, Krumlauf R, Wiedemann LM (2008) A regulatory module embedded in the coding region of Hoxa2 controls expression in rhombomere 2. Proc Natl Acad Sci USA 105:20077–20082

99. Lampe X, Samad OA, Guiguen A, Matis C, Remacle S, Picard JJ, Rijli FM, Rezsohazy R (2008) An ultraconserved Hox-Pbx responsive element resides in the coding sequence of Hoxa2 and is active on rhombomere 4. Nucleic Acids Res 36:3214–3225

100. Gavalas A, Ruhrberg C, Livet J, Henderson CE, Krumlauf R (2003) Neuronal defects in the hindbrain of Hoxa1, Hoxb1 and Hoxb2 mutants reflect regulatory interactions among these Hox genes. Development 130:5663–5679

101. Tümpel S, Cambronero F, Ferretti E, Blasi F, Wiedemann LM, Krumlauf R (2007) Expression of Hoxa2 in rhombomere 4 is regulated by a conserved cross-regulatory mechanism dependent upon Hoxb1. Dev Biol 302:646–660

102. Manzanares M, Bel-Vialar S, Ariza-McNaughton L, Ferretti E, Marshall H, Maconochie MM, Blasi F, Krumlauf R (2001) Independent regulation of initiation and maintenance phases of Hoxa3 expression in the vertebrate hindbrain involve auto- and cross-regulatory mechanisms. Development 128:3595–3607

103. Gould A, Morrison A, Sproat G, White RA, Krumlauf R (1997) Positive crossregulation and enhancer sharing: two mechanisms for specifying overlapping Hox expression patterns. Genes Dev 11:900–913

104. Gould A, Itasaki N, Krumlauf R (1998) Initiation of rhombo-meric Hoxb4 expression requires induction by somites and a retinoid pathway. Neuron 21:39–51

105. Serpente P, Tumpel S, Ghyselinck NB, Niederreither K, Wiede-mann LM, Dolle P, Chambon P, Krumlauf R, Gould AP (2005) Direct crossregulation between retinoic acid receptor beta and Hox genes during hindbrain segmentation. Development 132:503–513

106. Sun Z, Hopkins N (2001) vhnf1, the MODY5 and famil-ial GCKD-associated gene, regulates regional specification of the zebrafish gut, pronephros, and hindbrain. Genes Dev 15:3217–3229

107. Wiellette EL, Sive H (2003) vhnf1 and Fgf signals synergize to specify rhombomere identity in the zebrafish hindbrain. Develop-ment 130:3821–3829


1 3

108. Hernandez RE, Rikhof HA, Bachmann R, Moens CB (2004) vhnf1 integrates global RA patterning and local FGF signals to direct posterior hindbrain development in zebrafish. Develop-ment 131:4511–4520

109. Kim FA, Sing A, Kaneko T, Bieman M, Stallwood N, Sadl VS, Cordes SP (2005) The vHNF1 homeodomain protein estab-lishes early rhombomere identity by direct regulation of Kre-isler expression. Mech Dev 122:1300–1309

110. Moens CB, Cordes SP, Giorgianni MW, Barsh GS, Kimmel CB (1998) Equivalence in the genetic control of hindbrain segmen-tation in fish and mouse. Development 125:381–391

111. Voiculescu O, Taillebourg E, Pujades C, Kress C, Buart S, Charnay P, Schneider-Maunoury S (2001) Hindbrain pattern-ing: Krox20 couples segmentation and specification of regional identity. Development 128:4967–4978

112. Aragón F, Vázquez-Echeverría C, Ulloa E, Reber M, Cereghini S, Alsina B, Giraldez F, Pujades C (2005) vHnf1 regulates specification of caudal rhombomere identity in the chick hind-brain. Dev Dyn 234:567–576

113. Chomette D, Frain M, Cereghini S, Charnay P, Ghislain J (2006) Krox20 hindbrain cis-regulatory landscape: interplay between multiple long-range initiation and autoregulatory ele-ments. Development 133:1253–1262

114. Bouchoucha YX, Reingruber J, Labalette C, Wassef MA, Thie-rion E, Desmarquet-Trin Dinh C, Holcman D, Gilardi-Heben-streit P, Charnay P (2013) Dissection of a Krox20 positive feedback loop driving cell fate choices in hindbrain patterning. Mol Syst Biol 9:690

115. Labalette C, Wassef MA, Desmarquet-Trin Dinh C, Bouchou-cha YX, Le Men J, Charnay P, Gilardi-Hebenstreit P (2015) Molecular dissection of segment formation in the developing hindbrain. Development 142:185–195

116. Thierion E, Le Men J, Collombet S, Hernandez C, Coulpier F, Torbey P, Thomas-Chollier M, Noordermeer D, Charnay P, Gilardi-Hebenstreit P (2017) Krox20 hindbrain regulation incorporates multiple modes of cooperation between cis-acting elements. PLoS Genet 13:e1006903

117. Bae CJ, Jeong J, Saint-Jeannet JP (2015) A novel function for Egr4 in posterior hindbrain development. Sci Rep 5:7750

118. Theil T, Frain M, Gilardi-Hebenstreit P, Flenniken A, Charnay P, Wilkinson DG (1998) Segmental expression of the EphA4 (Sek-1) receptor tyrosine kinase in the hindbrain is under direct transcriptional control of Krox-20. Development 125:443–452

119. Cooke JE, Kemp HA, Moens CB (2005) EphA4 is required for cell adhesion and rhombomere-boundary formation in the zebrafish. Curr Biol 15:536–542

120. Sela-Donenfeld D, Kayam G, Wilkinson DG (2009) Bound-ary cells regulate a switch in the expression of FGF3 in hindbrain rhombomeres. BMC Dev Biol. https ://doi.org/10.1186/1471-213x-9-16

121. Mechta-Grigoriou F, Garel S, Charnay P (2000) Nab proteins mediate a negative feedback loop controlling Krox-20 activity in the developing hindbrain. Development 127:119–128

122. Runko AP, Sagerström CG (2003) Nlz belongs to a family of zinc-finger-containing repressors and controls segmental gene expression in the zebrafish hindbrain. Dev Biol 262:254–267

123. García-Gutiérrez P, Juárez-Vicente F, Gallardo-Chamizo F, Charnay P, García-Domínguez M (2011) The transcription fac-tor Krox20 is an E3 ligase that sumoylates its Nab coregulators. EMBO Rep 12:1018–1023

124. Kayam G, Kohl A, Magen Z, Peretz Y, Weisinger K, Bar A, Novikov O, Brodski C, Sela-Donenfeld D (2013) A novel role for Pax6 in the segmental organization of the hindbrain. Devel-opment 140:2190–2202

125. Moens CB, Yan YL, Appel B, Force AG, Kimmel CB (1996) Valentino: a zebrafish gene required for normal hindbrain seg-mentation. Development 122:3981–3990

126. Prince VE, Moens CB, Kimmel CB, Ho RK (1998) Zebrafish hox genes: expression in the hindbrain region of wild-type and mutants of the segmentation gene, valentino. Development 125:393–406

127. Cooke J, Moens C, Roth L, Durbin L, Shiomi K, Brennan C, Kimmel C, Wilson S, Holder N (2001) Eph signalling func-tions downstream of Val to regulate cell sorting and boundary formation in the caudal hindbrain. Development 128:571–580

128. Manzanares M, Cordes S, Ariza-McNaughton L, Sadl V, Maruthainar K, Barsh G, Krumlauf R (1999) Conserved and distinct roles of kreisler in regulation of the paralogous Hoxa3 and Hoxb3 genes. Development 126:759–769

129. Manzanares M, Nardelli J, Gilardi- Hebenstreit P, Marshall H, Giudicelli F, Martinez-Pastor MT, Krumlauf R, Charnay P (2002) Krox20 and Kreisler co-operate in the transcriptional control of segmental expression of Hoxb3 in the developing hindbrain. EMBO J 21:365–376

130. Rao BR (1968) The appearance and extension of neural dif-ferentiation tendencies in the neurectoderm of the early chick embryo. Wilhelm Roux Arch Entwickl Mech Org 160:187–236

131. Garcia-Martinez V, Alvarez IS, Schoenwolf GC (1993) Loca-tions of the ectodermal and nonectodermal subdivisions of the epiblast at stages 3 and 4 of avian gastrulation and neurulation. J Exp Zool 267:431–446

132. Niswander L, Martin GR (1992) Fgf-4 expression during gastrulation, myogenesis, limb and tooth development in the mouse. Development 114:755–768

133. Mahmood R, Kiefer P, Guthrie S, Dickson C, Mason I (1995) Multiple roles for FGF-3 during cranial neural development in the chicken. Development 121:1399–1410

134. Muhr J, Graziano E, Wilson S, Jessell TM, Edlund T (1999) Convergent inductive signals specify midbrain, hindbrain, and spinal cord identity in gastrula stage chick embryos. Neuron 23:689–702

135. Maves L, Jackman W, Kimmel CB (2002) FGF3 and FGF8 mediate a rhombomere 4 signaling activity in the zebrafish hindbrain. Development 129:3825–3837

136. Walshe J, Maroon H, McGonnell IM, Dickson C, Mason I (2002) Establishment of hindbrain segmental identity requires signaling by FGF3 and FGF8. Curr Biol 12:1117–1123

137. Wright TJ, Mansour SL (2003) Fgf3 and Fgf10 are required for mouse otic placode induction. Development 130:3379–3390

138. Mahmood R, Mason IJ, Morriss-Kay GM (1996) Expression of Fgf-3 in relation to hindbrain segmentation, otic pit position and pharyngeal arch morphology in normal and retinoic acid-exposed mouse embryos. Anat Embryol (Berl) 194:13–22

139. Lombardo A, Isaacs HV, Slack JM (1998) Expression and functions of FGF-3 in Xenopus development. Int J Dev Biol 42:1101–1107

140. Powles N, Marshall H, Economou A, Chiang C, Murakami A, Dickson C, Krumlauf R, Maconochie M (2004) Regulatory analysis of the mouse Fgf3 gene: control of embryonic expres-sion patterns and dependence upon sonic hedgehog (Shh) sig-nalling. Dev Dyn 230:44–56

141. Weisinger K, Wilkinson DG, Sela-Donenfeld D (2008) Inhibi-tion of BMPs by follistatin is required for FGF3 expression and segmental patterning of the hindbrain. Dev Biol 324:213–225

142. Weisinger K, Kayam G, Missulawin-Drillman T, Sela-Donen-feld D (2010) Analysis of expression and function of FGF-MAPK signaling components in the hindbrain reveals a central role for FGF3 in the regulation of Krox20, mediated by Pea3. Dev Biol 344:881–895

https://doi.org/10.1186/1471-213x-9-16

https://doi.org/10.1186/1471-213x-9-16


1 3

143. Marín F, Charnay P (2000) Hindbrain patterning: FGFs regulate Krox20 and mafB/kr expression in the otic/preotic region. Devel-opment 127:4925–4935

144. Weisinger K, Kohl A, Kayam G, Monsonego-Ornan E, Sela-Donenfeld D (2012) Expression of hindbrain boundary markers is regulated by FGF3. Biol Open 1:67–74

145. Guthrie S, Lumsden A (1991) Formation and regeneration of rhombomere boundaries in the developing chick hindbrain. Development 112:221–229

146. Martinez S, Geijo E, Sánchez-Vives MV, Puelles L, Gallego R (1992) Reduced junctional permeability at interrhombomeric boundaries. Development 116:1069–1076

147. Heyman I, Faissner A, Lumsden A (1995) Cell and matrix spe-cialisations of rhombomere boundaries. Dev Dyn 204:301–315

148. Cayuso J, Xu Q, Wilkinson DG (2015) Mechanisms of bound-ary formation by Eph receptor and ephrin signaling. Dev Biol 401:122–131

149. Terriente J, Pujades C (2015) Cell segregation in the verte-brate hindbrain: a matter of boundaries. Cell Mol Life Sci 72:3721–3730

150. Peretz Y, Eren N, Kohl A, Hen G, Yaniv K, Weisinger K, Cin-namon Y, Sela-Donenfeld D (2016) A new role of hindbrain boundaries as pools of neural stem/progenitor cells regulated by Sox2. BMC Biol 14:57

151. Aamar E, Frank D (2004) Xenopus Meis3 protein forms a hind-brain inducing center by activating FGF/MAP kinase and PCP pathways. Development 131:153–163

152. Maconochie MK, Nonchev S, Manzanares M, Marshall H, Krum-lauf R (2001) Differences in Krox20-dependent regulation of Hoxa2 and Hoxb2 during hindbrain development. Dev Biol 233:468–481

153. Lunn JS, Fishwick KJ, Halley PA, Storey KG (2007) A spatial and temporal map of FGF/Erk1/2 activity and response reper-toires in the early chick embryo. Dev Biol 302:536–552

154. Aragon F, Pujades C (2009) FGF signaling controls caudal hind-brain specification through Ras-ERK1/2 pathway. BMC Dev Biol 9:61

155. Roelink H, Nusse R (1991) Expression of two members of the Wnt family during mouse development—restricted temporal and spatial patterns in the developing neural tube. Genes Dev 5:381–388

156. Molven A, Njolstad PR, Fjose A (1991) Genomic structure and restricted neural expression of the zebrafish wnt-1 (int-1) gene. EMBO J 10:799–807

157. McGrew LL, Otte AP, Moon RT (1992) Analysis of Xwnt-4 in embryos of Xenopus laevis: a Wnt family member expressed in the brain and floor plate. Development 115:463–473

158. Hume CR, Dodd J (1993) Cwnt-8C: a novel Wnt gene with a potential role in primitive streak formation and hindbrain organi-zation. Development 119:1147–1160

159. Wolda SL, Moody CJ, Moon RT (1993) Overlapping expres-sion of Xwnt-3A and Xwnt-1 in neural tissue of Xenopus laevis embryos. Dev Biol 155:46–57

160. Kelly GM, Lai CJ, Moon RT (1993) Expression of wnt10a in the central nervous system of developing zebrafish. Dev Biol 158:113–121

161. Hollyday M, McMahon JA, McMahon AP (1995) Wnt expression patterns in chick embryo nervous system. Mech Dev 52:9–25

162. Augustine K, Liu ET, Sadler TW (1993) Antisense attenuation of Wnt-1 and Wnt-3a expression in whole embryo culture reveals roles for these genes in craniofacial, spinal cord, and cardiac morphogenesis. Dev Genet 14:500–520

163. Augustine KA, Liu ET, Sadler TW (1995) Interactions of Wnt-1 and Wnt-3a are essential for neural tube patterning. Teratology 51:107–119

164. McMahon AP, Joyner AL, Bradley A, McMahon JA (1992) The midbrain–hindbrain phenotype of Wnt-1-/Wnt-1- mice results from stepwise deletion of engrailed-expressing cells by 9.5 days postcoitum. Cell 69:581–595

165. Liu P, Wakamiya M, Shea MJ, Albrecht U, Behringer RR, Brad-ley A (1999) Requirement for Wnt3 in vertebrate axis formation. Nat Genet 22:361–365

166. Nordstrom U, Jessell TM, Edlund T (2002) Progressive induction of caudal neural character by graded Wnt signaling. Nat Neurosci 5:525–532

167. Nordstrom U, Maier E, Jessell TM, Edlund T (2006) An early role for WNT signaling in specifying neural patterns of Cdx and Hox gene expression and motor neuron subtype identity. PLoS Biol 4:e252

168. Lekven AC, Thorpe CJ, Waxman JS, Moon RT (2001) Zebrafish wnt8 encodes two wnt8 proteins on a bicistronic transcript and is required for mesoderm and neurectoderm patterning. Dev Cell 1:103–114

169. Erter CE, Wilm TP, Basler N, Wright CV, Solnica Krezel L (2001) Wnt8 is required in lateral mesendodermal precursors for neural posteriorization in vivo. Development 128:3571–3583

170. Shimizu T, Bae YK, Muraoka O, Hibi M (2005) Interaction of Wnt and caudal-related genes in zebrafish posterior body forma-tion. Dev Biol 279:125–141

171. Lekven AC, Buckles GR, Kostakis N, Moon RT (2003) Wnt1 and wnt10b function redundantly at the zebrafish midbrain-hindbrain boundary. Dev Biol 254:172–187

172. Riley BB, Chiang MY, Storch EM, Heck R, Buckles GR, Lekven AC (2004) Rhombomere boundaries are Wnt signaling centers that regulate metameric patterning in the zebrafish hindbrain. Dev Dyn 231:278–291

173. Kiecker C, Niehrs C (2001) A morphogen gradient of Wnt/beta-catenin signaling regulates anteroposterior neural patterning in Xenopus. Development 128:4189–4201

174. Monsoro-Burq AH, Wang E, Harland RM (2005) Msx1 and Pax3 cooperate to mediate FGF8 and WNT signals during Xenopus neural crest induction. Dev Cell 8:167–178

175. Wu J, Yang J, Klein PS (2005) Neural crest induction by the canonical Wnt pathway can be dissociated from anterior–poste-rior neural patterning in Xenopus. Dev Biol 279:220–232

176. Li B, Kuriyama S, Moreno M, Mayor R (2009) The posterioriz-ing gene Gbx2 is a direct target of Wnt signaling and the earliest factor in neural crest induction. Development 136:3267–3278

177. Kim CH, Oda T, Itoh M, Jiang D, Artinger KB, Chan-drasekharappa SC, Driever W, Chitnis AB (2000) Repressor activity of Headless/Tcf3 is essential for vertebrate head forma-tion. Nature 407:913–916

178. MacDonald BT, Adamska M, Meisler MH (2004) Hypomor-phic expression of Dkk1 in the doubleridge mouse: dose depend-ence and compensatory interactions with Lrp6. Development 131:2543–2552

179. Gavalas A, Krumlauf R (2000) Retinoid signalling and hindbrain patterning. Curr Opin Genet Dev 10:380–386

180. Schilling TF, Nie Q, Lander AD (2012) Dynamics and preci-sion in retinoic acid morphogen gradients. Curr Opin Genet Dev 22:562–569

181. van der Wees J, Schilthuis JG, Koster CH, Diesveld-Schipper H, Folkers GE, van der Saag PT, Dawson MI, Shudo K, van der Burg B, Durston AJ (1998) Inhibition of retinoic acid receptor-mediated signalling alters positional identity in the developing hindbrain. Development 125:545–556

182. Berggren K, McCaffery P, Dräger U, Forehand CJ (1999) Dif-ferential distribution of retinoic acid synthesis in the chicken embryo as determined by immunolocalization of the retinoic acid synthetic enzyme, RALDH-2. Dev Biol 210:288–304


1 3

183. Maden M, Gale E, Kostetskii I, Zile M (1996) Vitamin A-defi-cient quail embryos have half a hindbrain and other neural defects. Curr Biol 6:417–426

184. Gale E, Zile M, Maden M (1999) Hindbrain respecification in the retinoid-deficient quail. Mech Dev 89:43–54

185. Dupé V, Ghyselinck NB, Wendling O, Chambon P, Mark M (1999) Key roles of retinoic acid receptors alpha and beta in the patterning of the caudal hindbrain, pharyngeal arches and otocyst in the mouse. Development 126:5051–5059

186. Niederreither K, Vermot J, Schuhbaur B, Chambon P, Dollé P (2000) Retinoic acid synthesis and hindbrain patterning in the mouse embryo. Development 127:75–85

187. Dickman ED, Thaller C, Smith SM (1997) Temporally-regu-lated retinoic acid depletion produces specific neural crest, ocu-lar and nervous system defects. Development 124:3111–3121

188. White JC, Highland M, Kaiser M, Clagett-Dame M (2000) Vitamin A deficiency results in the dose-dependent acquisition of anterior character and shortening of the caudal hindbrain of the rat embryo. Dev Biol 220:263–284

189. Hill J, Clarke JD, Vargesson N, Jowett T, Holder N (1995) Exogenous retinoic acid causes specific alterations in the development of the midbrain and hindbrain of the zebrafish embryo including positional respecification of the Mauthner neuron. Mech Dev 50:3–16

190. Maves L, Kimmel CB (2005) Dynamic and sequential pattern-ing of the zebrafish posterior hindbrain by retinoic acid. Dev Biol 285:593–605

191. Cai AQ, Radtke K, Linville A, Lander AD, Nie Q, Schilling TF (2012) Cellular retinoic acid-binding proteins are essential for hindbrain patterning and signal robustness in zebrafish. Development 139:2150–2155

192. Samarut E, Fraher D, Laudet V, Gibert Y (2015) ZebRA: an overview of retinoic acid signaling during zebrafish develop-ment. Biochim Biophys Acta 1849:73–83

193. Sirbu IO, Gresh L, Barra J, Duester G (2005) Shifting bounda-ries of retinoic acid activity control hindbrain segmental gene expression. Development 132:2611–2622

194. White RJ, Nie Q, Lander AD, Schilling TF (2007) Complex regulation of cyp26a1 creates a robust retinoic acid gradient in the zebrafish embryo. PLoS Biol 5:e304

195. Hernandez RE, Putzke AP, Myers JP, Margaretha L, Moens CB (2007) Cyp26 enzymes generate the retinoic acid response pattern necessary for hindbrain development. Development 134:177–187

196. McCaffery P, Dräger UC (1993) Retinoic acid synthesis in the developing retina. Adv Exp Med Biol 328:181–190

197. Fujii H, Sato T, Kaneko S, Gotoh O, Fujii-Kuriyama Y, Osawa K, Kato S, Hamada H (1997) Metabolic inactivation of retinoic acid by a novel P450 differentially expressed in developing mouse embryos. EMBO J 16:4163–4173

198. Moss JB, Xavier-Neto J, Shapiro MD, Nayeem SM, McCaffery P, Dräger UC, Rosenthal N (1998) Dynamic patterns of reti-noic acid synthesis and response in the developing mammalian heart. Dev Biol 199:55–71

199. Swindell EC, Thaller C, Sockanathan S, Petkovich M, Jessell TM, Eichele G (1999) Complementary domains of retinoic acid production and degradation in the early chick embryo. Dev Biol 216:282–296

200. Begemann G, Schilling TF, Rauch GJ, Geisler R, Ingham PW (2001) The zebrafish neckless mutation reveals a requirement for raldh2 in mesodermal signals that pattern the hindbrain. Development 128:3081–3094

201. Dobbs-McAuliffe B, Zhao Q, Linney E (2004) Feedback mech-anisms regulate retinoic acid production and degradation in the zebrafish embryo. Mech Dev 121:339–350

202. Emoto Y, Wada H, Okamoto H, Kudo A, Imai Y (2005) Retinoic acid-metabolizing enzyme Cyp26a1 is essential for determining territories of hindbrain and spinal cord in zebrafish. Dev Biol 278:415–427

203. Linville A, Radtke K, Waxman JS, Yelon D, Schilling TF (2009) Combinatorial roles for zebrafish retinoic acid receptors in the hindbrain, limbs and pharyngeal arches. Dev Biol 325:60–70

204. McCaffery PJ, Adams J, Maden M, Rosa-Molinar E (2003) Too much of a good thing: retinoic acid as an endogenous regulator of neural differentiation and exogenous teratogen. Eur J Neurosci 18:457–472

205. Reijntjes S, Blentic A, Gale E, Maden M (2005) The control of morphogen signalling: regulation of the synthesis and catabolism of retinoic acid in the developing embryo. Dev Biol 285:224–237

206. Giguere V, Ong ES, Segui P, Evans RM (1987) Identification of a receptor for the morphogen retinoic acid. Nature 330:624–629

207. Petkovich M, Brand NJ, Krust A, Chambon P (1987) A human retinoic acid receptor which belongs to the family of nuclear receptors. Nature 330:444–450

208. Mangelsdorf DJ, Evans RM (1995) The RXR heterodimers and orphan receptors. Cell 83:841–850

209. Tang XH, Gudas LJ (2011) Retinoids, retinoic acid receptors, and cancer. Annu Rev Pathol 6:345–364

210. Rhinn M (2012) Dollé P Retinoic acid signalling during develop-ment. Development 139:843–858

211. Popperl H, Featherstone MS (1993) Identification of a retinoic acid response element upstream of the murine Hox-4.2 gene. Mol Cell Biol 13:257–265

212. Marin F, Puelles L (1994) Patterning of the embryonic avian midbrain after experimental inversions: a polarizing activity from the isthmus. Dev Biol 163:19–37

213. Dupé V, Davenne M, Brocard J, Dollé P, Mark M, Dierich A, Chambon P, Rijli FM (1997) In vivo functional analysis of the Hoxa-1 3′ retinoic acid response element (3′RARE). Develop-ment 124:399–410

214. Bel-Vialar S, Itasaki N, Krumlauf R (2002) Initiating Hox gene expression: in the early chick neural tube differential sensitivity to FGF and RA signaling subdivides the HoxB genes in two distinct groups. Development 129:5103–5115

215. Houle M, Prinos P, Iulianella A, Bouchard N, Lohnes D (2003) Retinoic acid regulation of Cdx1: an indirect mechanism for reti-noids and vertebral specification. Mol Cell Biol 20:6579–6586

216. Nolte C, Amores A, Kovacs EN, Postlethwait J, Featherstone M (2003) The role of a retinoic acid response element in establish-ing the anterior neural expression border of Hoxd4 transgenes. Mech Dev 120:325–335

217. Bertrand S, Thisse B, Tavares R, Sachs L, Chaumot A, Bardet PL, Escrivà H, Duffraisse M, Marchand O, Safi R, Thisse C, Laudet V (2007) Unexpected novel relational links uncovered by extensive developmental profiling of nuclear receptor expression. PLoS Genet 3:e188

218. Pouilhe M, Gilardi-Hebenstreit P, Desmarquet-Trin Dinh C, Charnay P (2007) Direct regulation of vHnf1 by retinoic acid signaling and MAF-related factors in the neural tube. Dev Biol 309:344–357

219. Ahn Y, Mullan HE, Krumlauf R (2014) Long-range regulation by shared retinoic acid response elements modulates dynamic expression of posterior Hoxb genes in CNS development. Dev Biol 388:134–144

220. Studer M, Pöpperl H, Marshall H, Kuroiwa A, Krumlauf R (1994) Role of a conserved retinoic acid response element in rhombomere restriction of Hoxb-1. Science 265:1728–1732

221. Marshall H, Studer M, Pöpperl H, Aparicio S, Kuroiwa A, Bren-ner S, Krumlauf R (1994) A conserved retinoic acid response element required for early expression of the homeobox gene Hoxb-1. Nature 370:567–571


1 3

222. Niederreither K, McCaffery P, Dräger UC, Chambon P, Dollé P (1997) Restricted expression and retinoic acid-induced downregulation of the retinaldehyde dehydrogenase type 2 (RALDH-2) gene during mouse development. Mech Dev 62:67–78

223. Laue K, Jänicke M, Plaster N, Sonntag C, Hammerschmidt M (2008) Restriction of retinoic acid activity by Cyp26b1 is required for proper timing and patterning of osteogenesis during zebrafish development. Development 135:3775–3787

224. Strate I, Min TH, Iliev D, Pera EM (2009) Retinol dehydrogenase 10 is a feedback regulator of retinoic acid signalling during axis formation and patterning of the central nervous system. Develop-ment 136:461–472

225. Feng L, Hernandez RE, Waxman JS, Yelon D, Moens CB (2010) Dhrs3a regulates retinoic acid biosynthesis through a feedback inhibition mechanism. Dev Biol 338:1–14

226. Vitobello A, Ferretti E, Lampe X, Vilain N, Ducret S, Ori M, Spetz JF, Selleri L, Rijli FM (2011) Hox and Pbx factors control retinoic acid synthesis during hindbrain segmentation. Dev Cell 20:469–482

227. Kim YK, Wassef L, Hamberger L, Piantedosi R, Palczewski K, Blaner WS, Quadro L (2008) Retinyl ester formation by lecithin: retinol acyltransferase is a key regulator of retinoid homeostasis in mouse embryogenesis. J Biol Chem 283:5611–5621

228. Zhang YR, Zhao YQ, Huang JF (2012) Retinoid-binding pro-teins: similar protein architectures bind similar ligands via com-pletely different ways. PLoS One 7:e36772

229. Addison M, Xu Q, Cayuso J, Wilkinson DG (2018) Cell identity switching regulated by retinoic acid signaling maintains homo-geneous segments in the hindbrain. Dev Cell 45:606–620

230. Wilkinson DG (2018) Establishing sharp and homogeneous seg-ments in the hindbrain. F1000 Res 7. https ://doi.org/10.12688 /f1000 resea rch.15391 .1

231. Schilling TF, Sosnik J, Nie Q (2016) Visualizing retinoic acid morphogen gradients. Methods Cell Biol 133:139–163

232. Rossant J, Zirngibl R, Cado D, Shago M, Giguère V (1991) Expression of a retinoic acid response element-hsplacZ transgene defines specific domains of transcriptional activity during mouse embryogenesis. Genes Dev 5:1333–1344

233. Perz-Edwards A, Hardison NL, Linney E (2001) Retinoic acid-mediated gene expression in transgenic reporter zebrafish. Dev Biol 229:89–101

234. Waxman JS, Yelon D (2011) Zebrafish retinoic acid receptors function as context-dependent transcriptional activators. Dev Biol 352:128–140

235. Li J, Hu P, Li K, Zhao Q (2012) Identification and characteri-zation of a novel retinoic acid response element in zebrafish cyp26a1 promoter. Anat Rec 295:268–277

236. Mandal A, Rydeen A, Anderson J, Sorrell MR, Zygmunt T, Torres-Vázquez J, Waxman JS (2013) Transgenic retinoic acid sensor lines in zebrafish indicate regions of available embryonic retinoic acid. Dev Dyn 242:989–1000

237. Shimozono S, Iimura T, Kitaguchi T, Higashijima S, Miyawaki A (2013) Visualization of an endogenous retinoic acid gradient across embryonic development. Nature 496:363–366

238. Sosnik J, Zheng L, Rackauckas CV, Digman M, Gratton E, Nie Q, Schilling TF (2016) Noise modulation in retinoic acid signal-ing sharpens segmental boundaries of gene expression in the embryonic zebrafish hindbrain. Elife 5:e14034

239. Hollemann T, Chen Y, Grunz H, Pieler T (1998) Regionalized metabolic activity establishes boundaries of retinoic acid signal-ling. EMBO J 17:7361–7372

240. Grandel H, Lun K, Rauch GJ, Rhinn M, Piotrowski T, Houart C, Sordino P, Küchler AM, Schulte-Merker S, Geisler R, Holder N, Wilson SW, Brand M (2002) Retinoic acid signalling in the zebrafish embryo is necessary during pre-segmentation stages

to pattern the anterior-posterior axis of the CNS and to induce a pectoral fin bud. Development 129:2851–2865

241. Kastner P, Messaddeq N, Mark M, Wendling O, Grondona JM, Ward S, Ghyselinck N, Chambon P (1997) Vitamin A deficiency and mutations of RXRalpha, RXRbeta and RARalpha lead to early differentiation of embryonic ventricular cardiomyocytes. Development 124:4749–4758

242. Dupé V, Lumsden A (2001) Hindbrain patterning involves graded responses to retinoic acid signalling. Development 128:2199–2208

243. Escriva H, Bertrand S, Germain P, Robinson-Rechavi M, Umb-hauer M, Cartry J, Duffraisse M, Holland L, Gronemeyer H, Lau-det V (2006) Neofunctionalization in vertebrates: the example of retinoic acid receptors. PLoS Genet 2:e102

244. Bouillet P, Oulad-Abdelghani M, Ward SJ, Bronner S, Chambon P, Dolle P (1996) A new mouse member of the Wnt gene family, mWnt-8, is expressed during early embryogenesis and is ectopi-cally induced by retinoic acid. Mech Dev 58:141–152

245. Hans S, Westerfield M (2007) Changes in retinoic acid signaling alter otic patterning. Development 134:2449–2458

246. Niederreither K, Subbarayan V, Dollé P, Chambon P (1999) Embryonic retinoic acid synthesis is essential for early mouse post-implantation development. Nat Genet 21:444–448

247. Urness LD, Paxton CN, Wang X, Schoenwolf GC, Mansour SL (2010) FGF signaling regulates otic placode induction and refine-ment by controlling both ectodermal target genes and hindbrain Wnt8a. Dev Biol 340:595–604

248. Mahoney Rogers AA, Zhang J, Shim K (2011) Sprouty1 and Sprouty2 limit both the size of the otic placode and hindbrain Wnt8a by antagonizing FGF signaling. Dev Biol 353:94–104

249. In der Rieden PM, Vilaspasa FL, Durston AJ (2010) Xwnt8 directly initiates expression of labial Hox genes. Dev Dyn 239:126–139

250. Janssens S, Denayer T, Deroo T, van Roy F, Vleminckx K (2010) Direct control of Hoxd1 and Irx3 expression by Wnt/beta catenin signaling during anteroposterior patterning of the neural axis in Xenopus. Int J Dev Biol 54:1435–1442

251. Diez del Corral R, Storey KG (2004) Opposing FGF and retinoid pathways: a signalling switch that controls differentiation and patterning onset in the extending vertebrate body axis. BioEssays 26:857–869

252. Shimizu T, Bae YK, Hibi M (2006) Cdx-Hox code controls competence for responding to Fgfs and retinoic acid in zebrafish neural tissue. Development 133:4709–4719

253. Zhao X, Duester G (2009) Effect of retinoic acid signaling on Wnt/beta-catenin and FGF signaling during body axis extension. Gene Expr Patterns 9:430–435

254. Martínez S, Marín F, Nieto MA, Puelles L (1995) Induction of ectopic engrailed expression and fate change in avian rhom-bomeres: intersegmental boundaries as barriers. Mech Dev 51:289–303

255. Crossley PH, Martinez S, Martin GR (1996) Midbrain develop-ment induced by FGF8 in the chick embryo. Nature 380:66–68

256. Irving C, Mason I (2000) Signalling by FGF8 from the isthmus patterns anterior hindbrain and establishes the anterior limit of Hox gene expression. Development 127:177–186

257. Wendling O, Ghyselinck NB, Chambon P, Mark M (2001) Roles of retinoic acid receptors in early embryonic morphogenesis and hindbrain patterning. Development 128:2031–2038

258. Linville A, Gumusaneli E, Chandraratna RA, Schilling TF (2004) Independent roles for retinoic acid in segmentation and neuronal differentiation in the zebrafish hindbrain. Dev Biol 270:186–199

259. Nittenberg R, Patel K, Joshi Y, Krumlauf R, Wilkinson DG, Brickell PM, Tickle C, Clarke JD (1997) Cell movements, neu-ronal organisation and gene expression in hindbrains lacking morphological boundaries. Development 124:2297–2306

https://doi.org/10.12688/f1000research.15391.1

https://doi.org/10.12688/f1000research.15391.1


1 3

260. Sato T, Joyner AL (2009) The duration of Fgf8 isthmic organizer expression is key to patterning different tectal-isthmo-cerebellum structures. Development 136:3617–3626

261. Diez del Corral R, Olivera-Martinez I, Goriely A, Gale E, Maden M, Storey K (2003) Opposing FGF and retinoid pathways control ventral neural pattern, neuronal differentiation, and segmentation during body axis extension. Neuron 40:65–79

262. Duester G (2013) Retinoid signaling in control of progenitor cell differentiation during mouse development. Semin Cell Dev Biol 24:694–700

263. Parker HJ, Bronner ME, Krumlauf R (2016) The vertebrate Hox gene regulatory network for hindbrain segmentation: evolution and diversification: Coupling of a Hox gene regulatory network to hindbrain segmentation is an ancient trait originating at the base of vertebrates. BioEssays 38:526–538

264. Skromne I, Thorsen D, Hale M, Prince VE, Ho RK (2007) Repression of the hindbrain developmental program by Cdx

factors is required for the specification of the vertebrate spinal cord. Development 134:2147–2158

265. Chang J, Skromne I, Ho RK (2016) CDX4 and retinoic acid interact to position the hindbrain-spinal cord transition. Dev Biol 410:178–189

266. Lee K, Skromne I (2014) Retinoic acid regulates size, pattern and alignment of tissues at the head-trunk transition. Development 141:4375–4384

267. Isaacs HV, Pownall ME, Slack JM (1998) Regulation of Hox gene expression and posterior development by the Xenopus cau-dal homologue Xcad3. EMBO J 17:3413–3427

268. Gaunt SJ, Drage D, Cockley A (2003) Vertebrate caudal gene expression gradients investigated by use of chick cdx-A/lacZ and mouse cdx-1/lacZ reporters in transgenic mouse embryos: evidence for an intron enhancer. Mech Dev 120:573–586

Documents

Patterning the Vertebrate Neuraxismateriais.dbio.uevora.pt/BD/Morfogenese/segment_encefalo.pdf · 2020. 5. 19. · plane of the ectoderm (7, 8). Whereas the molecular identity of