The unique nature of mammalian cleavage

With all these difficulties, knowledge of mammalian cleavage was worth waiting for, as mammalian cleavage turned out to be strikingly different from most other patterns of embryonic cell division. The mammalian oocyte is released from the ovary and swept by the fimbriae into the oviduct (Figure 11.20). Fertilization occurs in the ampulla of the oviduct, a region close to the ovary. Meiosis is completed at this time, and first cleavage begins about a day later (see Figure 7.32). Cleavages in mammalian eggs are among the slowest in the animal kingdom—about 12–24 hours apart. Meanwhile, the cilia in the oviduct push the embryo toward the uterus; the first cleavages occur along this journey.

Figure 11.20

Development of a human embryo from fertilization to implantation. Compaction of the human embryo occurs on day 4 when it is at the 10-cell stage. The embryo “hatches” from the zona pellucida upon reaching the uterus. During its migration (more...)

In addition to the slowness of cell division, there are several other features of mammalian cleavage that distinguish it from other cleavage types. The second of these differences is the unique orientation of mammalian blastomeres with relation to one another. The first cleavage is a normal meridional division; however, in the second cleavage, one of the two blastomeres divides meridionally and the other divides equatorially (Figure 11.21). This type of cleavage is called rotational cleavage (Gulyas 1975).

Figure 11.21

Comparison of early cleavage in (A) echinoderms and amphibians (radial cleavage) and (B) mammals (rotational cleavage). Nematodes also have a rotational form of cleavage, but they do not form the blastocyst structure characteristic of mammals. (After (more...)

The third major difference between mammalian cleavage and that of most other embryos is the marked asynchrony of early cell division. Mammalian blastomeres do not all divide at the same time. Thus, mammalian embryos do not increase exponentially from 2- to 4- to 8-cell stages, but frequently contain odd numbers of cells. Fourth, unlike almost all other animal genomes, the mammalian genome is activated during early cleavage, and produces the proteins necessary for cleavage to occur. In the mouse and goat, the switch from maternal to zygotic control occurs at the 2-cell stage (Piko and Clegg 1982; Prather 1989).

Most research on mammalian development has focused on the mouse embryo, since mice are relatively easy to breed throughout the year, have large litters, and can be housed easily. Thus, most of the studies discussed here will concern murine (mouse) development.

Compaction

The fifth, and perhaps the most crucial, difference between mammalian cleavage and all other types involves the phenomenon of compaction. As seen in Figure 11.22, mouse blastomeres through the 8-cell stage form a loose arrangement with plenty of space between them. Following the third cleavage, however, the blastomeres undergo a spectacular change in their behavior. They suddenly huddle together, maximizing their contact with one another and forming a compact ball of cells (Figure 11.22C, Figure 11.22D; Figure 11.23). This tightly packed arrangement is stabilized by tight junctions that form between the outside cells of the ball, sealing off the inside of the sphere. The cells within the sphere form gap junctions, thereby enabling small molecules and ions to pass between them.

Figure 11.22

The cleavage of a single mouse embryo in vitro. (A) 2-cell stage. (B) 4-cell stage. (C) Early 8-cell stage. (D) Compacted 8-cell stage. (E) Morula. (F) Blastocyst. (From Mulnard 1967; photographs courtesy of J. G. Mulnard.)

Figure 11.23

Scanning electron micrographs of (A) uncompacted and (B) compacted 8-cell mouse embryos. (Photographs courtesy of C. Ziomek.)

The cells of the compacted 8-cell embryo divide to produce a 16-cell morula (Figure 11.22E). The morula consists of a small group of internal cells surrounded by a larger group of external cells (Barlow et al. 1972). Most of the descendants of the external cells become the trophoblast (trophectoderm) cells. This group of cells produces no embryonic structures. Rather, it forms the tissue of the chorion, the embryonic portion of the placenta. The chorion enables the fetus to get oxygen and nourishment from the mother. It also secretes hormones that cause the mother's uterus to retain the fetus, and produces regulators of the immune response so that the mother will not reject the embryo as she would an organ graft.

The mouse embryo proper is derived from the descendants of the inner cells of the 16-cell stage, supplemented by cells dividing from the trophoblast during the transition to the 32-cell stage (Pedersen et al. 1986; Fleming 1987). These cells generate the inner cell mass (ICM), which will give rise to the embryo and its associated yolk sac, allantois, and amnion. By the 64-cell stage, the inner cell mass (approximately 13 cells) and the trophoblast cells have become separate cell layers, neither contributing cells to the other group (Dyce et al. 1987; Fleming 1987). Thus, the distinction between trophoblast and inner cell mass blastomeres represents the first differentiation event in mammalian development. This differentiation is required for the early mammalian embryo to adhere to the uterus (Figure 11.24). The development of the embryo proper can wait until after that attachment occurs. The inner cell mass actively supports the trophoblast, secreting proteins (such as FGF4) that cause the trophoblast cells to divide (Tanaka et al. 1998).

Figure 11.24

Implantation of the mammalian blastocyst into the uterus. (A) Mouse blastocysts entering the uterus. (B) Initial implantation of the blastocyst in a rhesus monkey. (A from Rugh 1967; B courtesy of the Carnegie Institution of Washington, Chester Reather, (more...)

Initially, the morula does not have an internal cavity. However, during a process called cavitation, the trophoblast cells secrete fluid into the morula to create a blastocoel. The inner cell mass is positioned on one side of the ring of trophoblast cells (see Figures 11.23 and 11.25). The resulting structure, called the blastocyst, is another hallmark of mammalian cleavage.

Figure 11.25

Mouse blastocyst hatching from the zona pellucida. (Photograph from Mark et al. 1985, courtesy of E. Lacy.)

Modifications for development within another organism

The mammalian embryo obtains nutrients directly from its mother and does not rely on stored yolk. This adaptation has entailed a dramatic restructuring of the maternal anatomy (such as expansion of the oviduct to form the uterus) as well as the development of a fetal organ capable of absorbing maternal nutrients. This fetal organ—the chorion—is derived primarily from embryonic trophoblast cells, supplemented with mesodermal cells derived from the inner cell mass. The chorion forms the fetal portion of the placenta. It will induce the uterine cells to form the maternal portion of the placenta, the decidua. The decidua becomes rich in the blood vessels that will provide oxygen and nutrients to the embryo.

The origins of early mammalian tissues are summarized in Figure 11.26. The first segregation of cells within the inner cell mass results in the formation of the hypoblast (sometimes called the primitive endoderm) layer (Figure 11.27A). The hypoblast cells delaminate from the inner cell mass to line the blastocoel cavity, where they give rise to the extraembryonic endoderm, which forms the yolk sac. As in avian embryos, these cells do not produce any part of the newborn organism. The remaining inner cell mass tissue above the hypoblast is now referred to as the epiblast. The epiblast cell layer is split by small clefts that eventually coalesce to separate the embryonic epiblast from the other epiblast cells, which form the amnionic cavity (Figures 11.27 B , C). Once the lining of the amnion is completed, it fills with a secretion called amnionic (amniotic) fluid, which serves as a shock absorber for the developing embryo while preventing its desiccation. The embryonic epiblast is believed to contain all the cells that will generate the actual embryo, and it is similar in many ways to the avian epiblast.

Figure 11.26

Schematic diagram showing the derivation of tissues in human and rhesus monkey embryos. (After Luckett 1978; Bianchi et al. 1993.)

Figure 11.27

Tissue formation in the human embryo between days 7 and 11. (A, B) Human blastocyst immediately prior to gastrulation. The inner cell mass delaminates hypoblast cells that line the blastocoel, forming the extraembryonic endoderm of the primitive yolk (more...)

By labeling individual cells of the epiblast with horseradish peroxidase, Kirstie Lawson and her colleagues (1991) were able to construct a detailed fate map of the mouse epiblast (Figure 1.6). Gastrulation begins at the posterior end of the embryo, and this is where the node forms^* (Figure 11.28). Like the chick epiblast cells, the mammalian mesoderm and endoderm migrate through a primitive streak, and like their avian counterparts, the migrating cells of the mammalian epiblast lose E-cadherin, detach from their neighbors, and migrate through the streak as individual cells (Burdsall et al. 1993). Those cells migrating through the node give rise to the notochord. However, in contrast to notochord formation in the chick, the cells that form the mouse notochord are thought to become integrated into the endoderm of the primitive gut (Jurand 1974; Sulik et al. 1994). These cells can be seen as a band of small, ciliated cells extending rostrally from the node (Figure 11.29). They form the notochord by converging medially and folding off in a dorsal direction from the roof of the gut.

Figure 11.28

Amnion structure and cell movements during human gastrulation. (A) Human embryo and uterine connections at day 15 of gestation. In the upper view, the embryo is cut sagittally through the midline; the lower view looks down upon the dorsal surface of the (more...)

Figure 11.29

Formation of the notochord in the mouse. (A) The ventral surface a the 7.5-day mouse embryo, seen by scanning electron microscopy. The presumptive notochord cells are the small, ciliated cells in the midline that are flanked by the larger endodermal cells (more...)

The ectodermal precursors end up anterior to the fully extended primitive streak, as in the chick epiblast; but whereas the mesoderm of the chick forms from cells posterior to the farthest extent of the streak, the mouse mesoderm forms from cells anterior to the primitive streak. In some instances, a single cell gives rise to descendants in more than one germ layer, or to both embryonic and extraembryonic derivatives. Thus, at the epiblast stage, these lineages have not become separate from one another. As in avian embryos, the cells migrating in between the hypoblast and epiblast layers are coated with hyaluronic acid, which they synthesize as they leave the primitive streak. This acts to keep them separate while they migrate (Solursh and Morriss 1977). It is thought (Larsen 1993) that the replacement of human hypoblast cells by endoderm precursors occurs on days 14–15 of gestation, while the migration of cells forming the mesoderm does not start until day 16 (Figure 11.28B).

Formation of extraembryonic membranes

While the embryonic epiblast is undergoing cell movements reminiscent of those seen in reptilian or avian gastrulation, the extraembryonic cells are making the distinctly mammalian tissues that enable the fetus to survive within the maternal uterus. Although the initial trophoblast cells of mice and humans divide like most other cells of the body, they give rise to a population of cells wherein nuclear division occurs in the absence of cytokinesis. The original type of trophoblast cells constitute a layer called the cytotrophoblast, whereas the multinucleated type of cell forms the syncytiotrophoblast. The cytotrophoblast initially adheres to the endometrium through a series of adhesion molecules. Moreover, these cells also contain proteolytic enzymes that enable them to enter the uterine wall and remodel the uterine blood vessels so that the maternal blood bathes fetal blood vessels. The syncytiotrophoblast tissue is thought to further the progression of the embryo into the uterine wall by digesting uterine tissue (Fisher et al. 1989). The uterus, in turn, sends blood vessels into this area, where they eventually contact the syncytiotrophoblast. Shortly thereafter, mesodermal tissue extends outward from the gastrulating embryo (see Figure 11.27D). Studies of human and rhesus monkey embryos have suggested that the yolk sac (and hence the hypoblast) is the source of this extraembryonic mesoderm (Bianchi et al. 1993). The extraembryonic mesoderm joins the trophoblastic extensions and gives rise to the blood vessels that carry nutrients from the mother to the embryo. The narrow connecting stalk of extraembryonic mesoderm that links the embryo to the trophoblast eventually forms the vessels of the umbilical cord. The fully developed extraembryonic organ, consisting of trophoblast tissue and blood vessel-containing mesoderm, is called the chorion, and it fuses with the uterine wall to create the placenta. Thus, the placenta has both a maternal portion (the uterine endometrium, which is modified during pregnancy) and a fetal component (the chorion). The chorion may be very closely apposed to maternal tissues while still being readily separable from them (as in the contact placenta of the pig), or it may be so intimately integrated with maternal tissues that the two cannot be separated without damage to both the mother and the developing fetus (as in the deciduous placenta of most mammals, including humans).^†

Figure 11.30 shows the relationships between the embryonic and extraembryonic tissues of a 6-week human embryo. The embryo is seen encased in the amnion and is further shielded by the chorion. The blood vessels extending to and from the chorion are readily observable, as are the villi that project from the outer surface of the chorion. These villi contain the blood vessels and allow the chorion to have a large area exposed to the maternal blood. Although fetal and maternal circulatory systems normally never merge, diffusion of soluble substances can occur through the villi (Figure 11.31). In this manner, the mother provides the fetus with nutrients and oxygen, and the fetus sends its waste products (mainly carbon dioxide and urea) into the maternal circulation. The maternal and fetal blood cells, however, usually do not mix.

Figure 11.30

Human embryo and placenta after 40 days of gestation. The embryo lies within the amnion, and its blood vessels can be seen extending into the chorionic villi. The small sphere to the right of the embryo is the yolk sac. (The Carnegie Institution of Washington, (more...)

Figure 11.31

Relationship of the chorionic villi to the maternal blood in the uterus.

Two signaling centers

The mammalian embryo appears to have two signaling centers: one in the node (“the organizer”) and one in the anterior visceral endoderm (Figure 11.34A; Beddington et al. 1994). The node appears to be responsible for the creation of all of the body, and these two signaling centers work together to form the forebrain (Bachiller et al. 2000). Both the mouse node and the anterior visceral endoderm express many of the genes known to be expressed in the chick and frog organizer tissue. The node produces Chordin and Noggin (which the anterior visceral endoderm does not), while the anterior visceral endoderm expresses several genes that are necessary for head formation. These include the genes for transcription factors Hesx-1, Lim-1, and Otx-2, as well as the gene for the paracrine factor Cerberus. The anterior visceral endoderm is established before the node, and the primitive streak always forms on the side of the epiblast opposite this anterior site. Homozygosity for mutant alleles of any of the above-mentioned head-specific organizing genes produces mice lacking forebrains (Figure 11.35; Thomas and Beddington 1996; Beddington and Robertson 1999). While knockouts of either chordin or noggin do not affect development, mice missing both sets of genes develop a body lacking forebrain, nose, and facial structures.

Figure 11.34

The two signaling centers of the mammalian embryo. (A) In the day 7 mouse embryo, the dorsal surface of the epiblast (embryonic ectoderm) is in contact with the amnionic cavity. The ventral surface of the epiblast contacts the newly formed mesoderm. In (more...)

Figure 11.35

Headless phenotype of the Lim-1 knockout mouse. A Lim-1-deficient mouse is shown next to a wild-type littermate. The ear pinnae (arrows) are the most anterior structures seen in these mutants. (From Shawlot and Behringer 1995, photograph courtesy of the (more...)

Patterning the anterior-posterior axis: the hox code hypothesis

Once gastrulation begins, anterior-posterior polarity in all vertebrates becomes specified by the expression of Hox genes. These genes are homologous to the homeotic gene complex (Hom-C) of the fruit fly (see Chapter 9). The Drosophila homeotic gene complex on chromosome 3 contains the Antennapedia and bithorax clusters of homeotic genes (see Figure 9.28), and can be seen as a single functional unit. (Indeed, in other insects, such as the flour beetle Tribolium, it is a single unit.) The Hom-C genes are arranged in the same general order as their expression pattern along the anterior-posterior axis, the most 3´ gene (labial) being required for producing the most anterior structures, and the most 5´ gene (AbdB) specifying the development of the posterior abdomen. Mouse and human genomes contain four copies of the Hox complex per haploid set, located on four different chromosomes (Hoxa through Hoxd in the mouse, HOXA through HOXD in humans: Boncinelli et al. 1988; McGinnis and Krumlauf 1992; Scott 1992). Not only are the same general types of homeotic genes found in both flies and mammals, but the order of these genes on their respective chromosomes is remarkably similar. In addition, the expression of these genes follows the same pattern: those mammalian genes homologous to the Drosophila labial, proboscipedia, and deformed genes are expressed anteriorly, while those genes that are homologous to the Drosophila Abdominal-B gene are expressed posteriorly. Another set of genes that controls the formation of the fly head (orthodenticle and empty spiracles) has homologues in the mouse that show expression in the midbrain and forebrain.

While Hox genes appear to specify the anterior-posterior axis throughout the vertebrates, we shall discuss mammals here, since the experimental evidence is particularly strong for this class. The mammalian Hox/HOX genes are numbered from 1 to 13, starting from that end of each complex that is expressed most anteriorly. Figure 11.36 shows the relationships between the Drosophila and mouse homeotic gene sets. The equivalent genes in each mouse complex (such as Hoxa-1, Hoxb-1, and Hoxd-1) are called a paralogous group. It is thought that the four mammalian Hox complexes were formed from chromosome duplications. Because there is not a one-to-one correspondence between the Drosophila Hom-C genes and the mouse Hox genes, it is likely that independent gene duplications have occurred since these two animal branches diverged (Hunt and Krumlauf 1992; see Chapter 22).

Figure 11.36

Evolutionary conservation of homeotic gene organization and transcriptional expression in fruit flies and mice. (A) Similarity between the Hom-C cluster on Drosophila chromosome 3 and the four Hox gene clusters in the mouse genome. The shaded regions (more...)

Expression of hox genes along the dorsal axis

Hox gene expression can be seen along the dorsal axis (in the neural tube, neural crest, paraxial mesoderm, and surface ectoderm) from the anterior boundary of the hindbrain through the tail. The different regions of the body from the midbrain through the tail are characterized by different constellations of Hox gene expression, and the pattern of Hox gene expression is thought to specify the different regions. In general, the genes of paralogous group 1 are expressed from the tip of the tail to the most anterior border of the hindbrain. Paralogue 2 genes are expressed throughout the spinal cord, but the anterior limit of expression stops two segments more caudally than that of the paralogue 1 genes (see Figure 11.36; Wilkinson et al. 1989; Keynes and Lumsden 1990). The higher-numbered Hox paralogues are expressed solely in the posterior regions of the neural tube, where they also form a “nested” set.

Experimental analysis of the hox code

The expression patterns of the murine Hox genes suggest a code whereby certain combinations of Hox genes specify a particular region of the anterior-posterior axis (Hunt and Krumlauf 1991). Particular sets of paralogous genes provide segmental identity along the anterior-posterior axis of the body. Evidence for such a code comes from three sources:

•  Gene targeting or “knockout” experiments (see Chapter 4), in which mice are constructed that lack both copies of one or more Hox genes
•  Retinoic acid teratogenesis, in which mouse embryos exposed to retinoic acid show a different pattern of Hox gene expression along the anterior-posterior axis and abnormal differentiation of their axial structures
•  Comparative anatomy, in which the types of vertebrae in different vertebrate species are correlated with the constellation of Hox gene expression

Gene targeting

When Chisaka and Capecchi (1991) knocked out the Hoxa-3 gene from inbred mice, these homozygous mutants died soon after birth. Autopsies of these mice revealed that their neck cartilage was abnormally short and thick and that they had severely deficient or absent thymuses, thyroids, and parathyroid glands (Figure 11.37). The heart and major blood vessels were also malformed. Further analysis showed that the number and migration of the neural crest cells that normally form these structures were not affected. Rather, it appears that the Hoxa-3 genes are responsible for specifying cranial neural crest cell fate and for enabling these cells to differentiate and proliferate into neck cartilage and the glands that form the fourth and sixth pharyngeal pouches (Manley and Capecchi 1995).

Figure 11.37

Deficient development of neural crest-derived pharyngeal arch and pouch structures in Hoxa-3-deficient mice. The arches are numbered. (Right) A 10.5-day embryo of a heterozygous Hoxa-3 mouse (wild-type), showing normal development of pouch 3 (thymus), (more...)

Knockout of the Hoxa-2 gene also produces mice whose neural crest cells have been respecified, but the defects in these mice are anterior to those in the Hoxa-3 knockouts. Cranial elements normally formed by the neural crest cells of the second pharyngeal arch (stapes, styloid bones) are missing and are replaced by duplicates of the structures of the first pharyngeal arch (incus, malleus, etc.) (Gendron-Maguire et al. 1993; Rijli et al. 1993). Thus, without certain Hox genes, some regionally specific organs along the anterior-posterior axis fail to form, or become respecified as other regions. Similarly, when the Hoxc-8 gene is knocked out (Le Mouellic et al. 1992), several axial skeletal segments resemble more anterior segments, much like what is seen in Drosophila loss-of-function homeotic mutations. As can be seen in Figure 11.38, the first lumbar vertebra of the mouse has formed a rib—something characteristic of the thoracic vertebrae anterior to it.

Figure 11.38

Partial transformation of the first lumbar vertebra into a thoracic vertebra by the knockout of the Hoxc-8 gene. The first lumbar vertebra of this mouse has formed a rib—a structure normally formed only by the thoracic vertebrae anterior to it. (more...)

One can get severe axial transformations by knocking out two or more genes of a paralogous group. Mice homozygous for the Hoxd-3 deletion have mild abnormalities of the first cervical vertebra (the atlas), while mice homozygous for the Hoxa-3 deletion have no abnormality of this bone, though they have other malformations (see the discussion of this mutant above). When both sets of mutations a bred into the same mouse, both sets of problems become more severe. Mice with neither Hoxa-3 nor Hoxd-3 have no atlas bone at all, and the hyoid and thyroid cartilage is so reduced in size that there are holes in the skeleton (Condie and Capecchi 1994; Greer et al. 2000). It appears that there are interactions occurring between the products of the Hox genes, and that in some functions, one of the paralogues can replace the other.

Thus, the evidence from gene knockouts supports the hypotheses that (1) different sets of Hox genes are necessary for the specification of any region of the anterior-posterior axis, (2) that the members of a paralogous group of Hox genes may be responsible for different subsets of organs within these regions, and (3) that the defects caused by knocking out particular Hox genes occur in the most anterior region of that gene's expression.

Retinoic acid teratogenesis

Homeotic changes are also seen when mouse embryos are exposed to teratogenic doses of retinoic acid (RA). Exogenous retinoic acid given to mouse embryos in utero can cause certain Hox genes to become expressed in groups of cells that usually do not express them (Conlon and Rossant 1992; Kessel 1992). Moreover, the craniofacial abnormalities of mouse embryos exposed to teratogenic doses of RA (Figure 11.39) can be mimicked by causing the expression of Hoxa-7 throughout the embryo (Balling et al. 1989). If high doses of RA can activate Hox genes in inappropriate locations along the anterior-posterior axis, and if the constellation of active Hox genes specifies the region of the anterior-posterior axis, then mice given RA in utero should show homeotic transformations manifested as rostralizing malformations occurring along that axis.

Figure 11.39

Mouse embryos cultured at day 8 in control medium (A, C) or in medium containing retinoic acid (B, D). At day 10 (A, B), the first pharyngeal arch of the treated embryos has a shortened and flattened appearance and has apparently fused with the second (more...)

Kessel and Gruss (1991) found this to be the case. Wild-type mice have 7 cervical (neck) vertebrae, 13 thoracic (ribbed) vertebrae, and 6 lumbar (abdominal) vertebrae, in addition to the sacral and caudal (tail) vertebrae. In embryos exposed to RA on day 8 of gestation (during gastrulation), the first one or two lumbar vertebrae were transformed into thoracic (ribbed) vertebrae, while the first sacral vertebra often became a lumbar vertebra. In some cases, the entire posterior region of the mouse embryo failed to form (Figure 11.39E). These changes in structure were correlated with changes in the constellation of Hox genes expressed in these tissues. For example, when RA was given to embryos on day 8, Hoxa-10 expression was shifted posteriorly, and an additional set of ribs formed on what had been the first lumbar vertebra. When posterior Hox genes were not expressed at all, the caudal part of the embryo failed to form.^‡

Retinoic acid probably plays a role in axis specification during normal development, and the source of retinoic acid is probably the node (Hogan et al. 1992; Maden et al. 1996). It is possible that the specification of mesoderm cells depends on the amount of time spent within the high retinoic acid concentrations of the node: the more time spent in the node, the more posterior the specification. This pattern can be demonstrated in culture, as embryonal carcinoma cells express more “posterior” Hox genes the longer they are exposed to retinoic acid (Simeone et al. 1990). Giving RA exogenously would mimic the RA concentrations normally encountered only by the posterior cells. The evidence points to a Hox code wherein different constellations of Hox genes, activated by different retinoic acid concentrations, specify the regional characteristics along the anterior-posterior axis.

Comparative anatomy

A new type of comparative embryology is now emerging, and it is based on the comparison of gene expression patterns. Gaunt (1994) and Burke and her collaborators (1995) have compared the vertebrae of the mouse and the chick. Although the mouse and the chick have a similar number of vertebrae, they apportion them differently. Mice (like all mammals, be they giraffes or whales) have only 7 cervical vertebrae. These are followed by 13 thoracic vertebrae, 6 lumbar vertebrae, 4 sacral vertebrae, and a variable (20+) number of caudal vertebrae (Figure 11.40). The chick, on the other hand, has 14 cervical vertebrae, 7 thoracic vertebrae, 12 or 13 (depending on the strain) lumbosacral vertebrae, and 5 coccygeal (fused tail) vertebrae. The researchers asked, Does the constellation of Hox gene expression correlate with the type of vertebra formed (e.g., cervical or thoracic) or with the relative position of the vertebrae (e.g., number 8 or 9)?

Figure 11.40

Schematic representation of the mouse and chick vertebral pattern along the anterior-posterior axis. The boundaries of expression of certain Hox gene paralogous groups have been mapped onto these domains. (After Burke et al. 1995.)

The answer is that the constellation of Hox gene expression predicts the type of vertebra formed. In the mouse, the transition between cervical and thoracic vertebrae is between vertebrae 7 and 8; in the chick it is between vertebrae 14 and 15. In both cases, the Hox-5 paralogues are expressed in the last cervical vertebrae, while the anterior boundary of the Hox-6 paralogues extends to the first thoracic vertebra. Similarly, in both animals, the thoracic-lumbar transition is seen at the boundary between the Hox-9 and Hox-10 paralogous groups. It appears there is a code of differing Hox gene expression along the anterior-posterior axis, and this code determines the type of vertebra formed.

WEBSITE

11.11 Why do mammals have only seven cervical vertebrae? Recent speculation predicts that the mammalian Hox genes function simultaneously in several processes. To alter a Hox gene's expression so as to change vertebral type might lead to lethal changes in the other processes. http://www.devbio.com/chap11/link1111.shtml

The dorsal-ventral axis

Very little is known about the mechanisms of dorsal-ventral axis formation in mammals. In mice and humans, the hypoblast forms on the side of the inner cell mass that is exposed to the blastocyst fluid, while the dorsal axis forms from those ICM cells that are in contact with the trophoblast. Thus, the dorsal-ventral axis of the embryo is, in part, defined by the embryonic-abembryonic axis of the blastocyst. This axis (wherein the embryonic region contains the ICM while the abembryonic region is that part of the blastocyst opposite the ICM) may be determined within the oocyte, as it develops perpendicularly to the animal-vegetal axis of the newly fertilized egg (Figure 11.41; Gardner 1997). As development proceeds, the notochord maintains dorsal-ventral polarity by inducing specific dorsal-ventral patterns of gene expression in the overlying ectoderm (Goulding et al. 1993).

Figure 11.41

Relationship between the animal-vegetal axis of the egg and the embryonic-abembryonic axis of the blastocyst. The polar body marks the animal pole of the embryo. The dorsal-ventral axis of the embryo appears to form at right angles to the animal-vegetal (more...)

The left-right axis

The mammalian body is not symmetrical. Although the heart begins its formation at the midline of the embryo, it moves to the left side of the chest cavity and loops to the right (Figure 11.42). The spleen is found solely on the left side of the abdomen, the major lobe of the liver forms on the right side of the abdomen, the large intestine loops right to left as it traverses the abdominal cavity, and the right lung has one more lobe than the left lung.

Figure 11.42

Left-right asymmetry in the developing human. (A) Abdominal cross sections show that the originally symmetrical organ rudiments acquire asymmetric positions by week 11. The liver moves to the right and the spleen moves to the left. (B) Not only does the (more...)

Mutations in mice have shown that there are two levels of regulating the left-right axis: a global level and an organ-specific level. Some genes, such as situs inversus viscerum (iv), randomizes the left-right axis for each asymmetrical organ (Hummel and Chapman 1959; Layton 1976). This means that the heart may loop to the left in one homozygous animal, but loop to the right in another (Figure 11.43). Moreover, the direction of heart looping is not coordinated with the placement of the spleen or the stomach. This can cause serious problems, even death. The second gene, inversion of embryonic turning (inv), causes a more global phenotype. Mice homozygous for an insertion mutation at this locus were found to have all their asymmetrical organs on the wrong side of the body (Yokoyama et al. 1993).^§ Since all the organs are reversed, this asymmetry does not have dire consequences for the mice.

Figure 11.43

Asymmetry of gene expression in the mouse embryo. (A) In situ hybridization for nodal mRNA in a wild-type 5-somite mouse embryo. The nodal gene expression is confined to the lateral plate mesoderm on the left side of the embryo. (B) Cross section through (more...)

Recently, several additional asymmetrically expressed genes have been discovered, and their influence on one another has enabled scientists to put them into a possible pathway. The end of this pathway—the activation of Nodal proteins and the Pitx2 transcription factor on the left side of the lateral plate mesoderm—appears to be the same as in frog and chick embryos, although the path leading to this point differs between the species (Figure 11.44; see Figure 11.17; Collignon et al. 1996; Lowe et al. 1996; Meno et al. 1996).

Figure 11.44

Situs formation in mammals. (A) Proposed pathway for left-right axis formation in the mouse. The leftward movement of cilia in the node activates some as yet unidentified factor (possibly the product of the inv gene). This product activates the nodal (more...)

In frogs, the pathway begins with the placement of Vg1; in chicks it begins with the suppression of sonic hedgehog expression. In mammals, the distinction between left and right sides begins in the ciliary cells of the node (Figure 11.44B). The cilia cause the flow of fluid in the yolk sac cavity from right to left. When Nonaka and colleagues (1998) knocked out a mouse gene encoding the ciliary motor protein dynein (see Chapter 7), the nodal cilia did not move, and the situs (lateral position) of each asymmetrical organ was randomized. This finding correlated extremely well with other data. First, it had long been known that humans having a dynein deficiency had immotile cilia and a random chance of having their hearts on the left or right side of the body (Afzelius 1976). Second, when the gene for the iv mutant mice described above was cloned, it was found to encode the ciliary dynein protein (Supp et al. 1997).

In some way (perhaps through the product of the inv gene), this leftward motion of the cilia activates the genes for two paracrine factors, Nodal and Lefty-2, in the lateral plate mesoderm on the left side of the embryo (Figure 11.44A). The proteins produced by these factors spread throughout the left side of the embryo, and appear to be restrained to that side by the Lefty-1 protein, which is secreted by the bottom left side of the neural tube (Meno et al. 1998). Lefty-2 appears to be able to block the Snail protein (which becomes specific for the right side of the body), while Nodal activates pitx2 gene expression (Pedra et al. 1998).

The development of mammals has enormous importance for understanding the bases of numerous human diseases. In the next chapters, we will discuss later aspects of vertebrate development and the relationship between genetics and development during organ formation.