INTRODUCTION
1. The “hyaluronic acid” alias “hyaluronan”
In 1934, Karl Meyer and his assistant, John Palmer, described a procedure for isolating a novel glycosaminoglycan from the vitreous of bovine eyes (Meyer and Palmer, 1934). They showed that this substance contained an uronic acid and an aminosugar, but no sulfoesters. In their words: “we propose, for convenience, the name "hyaluronic acid", from hyaloid (vitreous) and uronic acid.” This marked the birth announcement for one of nature's most versatile and fascinating macromolecules. Today, this macromolecule is most frequently referred to as “hyaluronan” (in his abbreviated form: HA), reflecting the fact that it exists in vivo as a polyanion and not in the protonated acid form (Hascall and Laurent, 1997).
1.1. Chemical properties
During the following 20 years it has been determined the precise chemical structure of the basic disaccharide motif that forms the glycosaminoglycan hyaluronan (Weissman and Meyer, 1954). It is a negatively charged linear polymer of N-acetylglucosamine and D-glucuronic acid groups combined with β(1-4) and β(1-3) glycosidic bonding. The result is an energetically very stable configuration, with the larger groups in equatorial position while all of the small hydrogen atoms in the less sterically favourable axial position. Some H-bindings, parallel to the chains axis, rearrange the hyaluronan polymers in a very compact two fold helix secondary structure, leading to the formation of large hydrophobic patches. As important consequence, the two fold helix hyaluronan polymers immobilize an extraordinary quantity of water molecules (100 time more than this weight) organizing a highly hydrated extracellular matrix (Fig. 1).
In normal physiological conditions, hyaluronan is a very long polymer,
consisting in 10,000 or more disaccharides, which corresponds to
polysaccharides with relative molecular masses of at least 4x10
6Da and polymer
lengths of 10μm (Hascall and Laurent, 1997).
1.2. Hyaluronan biosynthesis
Although hyaluronan belongs to the family of glycosaminoglycans, which includes heparan sulphate and chondroitin sulphate, it differs from these in many ways. Other glycosaminoglycans are made as proteoglycans that are synthesized and assembled in the rough endoplasmic reticulum and Golgi apparatus, and are secreted in a similar way to other glycoproteins (Toole, 2004). Hyaluronan, however, is synthesized, as an unmodified polysaccharide, at the plasma membrane by any one of three different hyaluronan synthases (Weigel et al., 1997). Synthases are multipass transmembrane enzymes with the active sites protruding from the inner face of the plasma membrane where they draw off a cytosolic pool of activated UDP-sugar precursors as substrate. While the synthases byosinthesize hyaluronan chain, they also extruded it through the plasma membrane (Philipson and Schwartz, 1984) (Fig. 2).
CDNAs encoding putative hyaluronan synthases, Has1, Has2 and Has3, were
identified in human (Monslow et al., 2003), mouse (Spicer et al., 1996; Spicer et
al., 1997; Semino et al., 1996;), chicken (Spicer and McDonald, 1998), Xenopus
laevis (Rosa et al., 1988; Spicer and McDonald, 1998; Vigetti et al., 2003) and
zebrafish (Semino et al., 1996; Bakkers et al., 2004) showing that the sequences
were highly conserved between the different species. Has1, Has2 and Has3 share
not only some similarity on the aminoacidic sequence but also some molecular
and structural features. Furthermore in vitro experiments showed that expression
of any mammalian hyaluronan synthase led to hyaluronan biosynthesis by
mammalian cell lines, suggesting that each functions independently as a
synthase. Despite this, it has been shown that each synthase differs in the
enzymatic properties (Spicer and McDonald, 1998). In fact Has3 is more
catalytically active than Has2, which is in turn more active than Has1. They also
differ in the relative binding affinities for the two UDP-sugar substrates (Itano et
al., 1999). Moreover Has1 and Has2 proteins polymerize hyaluronan chains of
similar lengths (up to 2 X 10
6Da), whereas Has3 polymerize much shorter
hyaluronan chains, in the range of 2x10
5Da to 3x10
5Da. Hyaluronan chains of
different lengths have different effects on in vitro cell behaviour: very short
hyaluronan chains stimulate cell proliferation and initiate signalling cascades and
may be involved in angiogenesis and inflammatory responses; on the other hand
high-molecular-weight hyaluronan chains have the opposite effect, inhibiting cell
proliferation (Goldberg and Toole, 1987) (Fig. 2). Short hyaluronan chains can also be generated by degradation of extracellular hyaluronan through the action of hyaluronidase or oxidants (Girish and Kemparaju, 2007).
Thus, it is possible that while each hyaluronan synthases catalyzes the biosynthesis of hyaluronan, the three enzymes may not be able to replace each other functionally during all aspects of development or adult life (McDonald and Camenisch 2003).
Figure 1 - A tetrasaccharide from a hyaluronan chain, consisting of two disaccharide repeating units showing the prefer- red configuration in water. G = D- glucuronic acid, N = N-acetylglucosamine. The dotted lines indicate hydrogen bonds. The G1, N1 disaccharide is rotated through 180 degrees about the axis of the chain, compared with the disaccharide G2, N2, thus the chain is a two-fold helix. Modified from Scott (1989).
Figure 2 - The hyaluronan synthases. The hyaluronan synthases are multipass transmembrane proteins that use intracellular UDP-sugar precursors as substrate. While the synthases byosinthesize hyaluronan chain, they also extruded it through the plasma membrane. Each synthase differs in the enzymatic properties:
Has1 produces small amounts of high molecular weight hyaluronan.
Has2 produces significantly higher molecular weight hyaluronan. Has3 is the most active of the hyaluronan synthases, yet produces low molecular weight hyaluronan chains. Hyaluronan chains of different lengths have different effects on in vitro cell behaviour. Modified from Spicer and McDonald (1998).
1.3. Hyaluronan catabolism
It has been estimated that about a third of hyaluronan in the human body is removed and replaced each day (Fraser et al., 1997). Labelled hyaluronan in the epidermal compartments of human skin organ cultures disappears with a half-life of about 1 day (Tammi et al., 1991), in contrast to 20-70 days in the cartilage (Morales and Hascall, 1988) and vitreous body (Fraser et al., 1997) respectively.
Hyaluronan turnover in the body occurs through three separate pathways:
a- A local cellular turnover, which includes binding, internalization and degradation within cells. Binding occurs through the predominant hyaluronan receptors CD44 (Culty et al., 1992) and RHAMM (Evanko et al., 2004).
b- At tissue level hyaluronan is released from tissue matrices, drained into the vasculature and lymphatic, with final steps that include liver, kidney and possibly spleen. This pathway involves unique receptors such as hyaluronan receptors for endocytosis (HARE) (Zhou et al., 2000), lymphatic vessel endothelial HA receptor (LYVE)-1 (Banerji et al., 1999) and layilin (Bono et al., 2005).
c- Scission of hyaluronan can occur by free radicals under oxidative conditions. Free radicals and hyaluronidase may have their activities coordinated under certain pathologic situations (Girish and Kemparaju, 2007).
1.4. Hyaluronan as one of the main extracellular matrix components
The extracellular matrix is a heterogeneous mixture of water, saccharides
and various protein components, i.e., collagens, proteoglycans, noncollagenous
glycoproteins, and elastins, that upon deposition are organized into three-
dimensional tissue-specific meshwork, the structural environment in which cells
are embedded. The extracellular matrix structure is highly dynamic and
undergoes constant remodelling controlled by a delicate balance between
extracellular matrix synthesis and degradation. The extracellular matrix,
however, not only provides a physical framework for cells but also influences a
number of cellular functions via two basic mechanisms (Toole, 2000). First, the
extracellular matrix serves as storage depot for transient components like
growth factors, cytokines, chemokines, and enzymes. Resident proteins bind to
some of these molecules and modulate their activity, bioavailability or presentation to cell surface receptors (Aszodi et al., 2006). Second, most, if not all, matrix proteins directly interact with cellular receptors affecting cell adhesion and migration. The active interplay between cells and the extracellular matrix culminates in intracellular events associated with signal transduction cascades, which in turn regulate the expression of genes necessary for cell differentiation, proliferation, and survival (Aszodi et al., 2006).
The integrity of the pericellular matrix (the extracellular matrix surrounding cells) is dependent on hyaluronan, given that the treatment of cells exhibiting pericellular matrices with hyaluronidase destroys their structure (Lee et al., 1993). Tethering of hyaluronan to different cell types can occur by two independent mechanisms: by transmembrane interaction of "nascent"
hyaluronan with hyaluronan synthase on the cytoplasmic face of the plasma membrane (Fig. 3A) or by binding to specific hyaluronan receptors on the cell surface (Fig. 3B). In many embryonic cells that exhibit pericellular matrices, hyaluronan is most likely tethered by sustained attachment to hyaluronan synthases (Knudson and Knudson, 1993).
Figure 3 - Models of the structure of hyaluronan-dependent pericellular matrices. Coat formation usually requires that hyaluronan is tethered to the cell surface and proteoglycan is bound to the hyaluronan. Tethering of hyaluronan to the cell surface can occur by transmembrane interaction with hyaluronan synthase (A) or by binding to cell surface receptors such as CD44 (B).
HA, hyaluronan; PG, proteoglycan. Modified from Toole (2001).
1.5. Intracellular hyaluronan
The glycosaminoglycan hyaluronan in vertebrate exists as a pool associated with the cell surface, a pool bounded to other matrix components and as a largely mobile pool, but hyaluronan can also been taken up by cells through CD44 (Culty et al., 1992) and RHAMM (Evanko et al., 2004) receptors. It is possible that not all the hyaluronan taken up by cells is immediately degraded.
Intact hyaluronan chains have been detected within cells, in cytoplasm of vascular smooth muscle cells during late prophase/early prometaphase of mitosis, in nucleus and even within the nucleolus (Evanko et al., 2004). It has also been found within lamellae during cell locomotion and following serum stimulation (Collis et al., 1998). Intracellular hyaluronan can be derived from either the extracellular environment or from an as yet unidentified intracellular source (Collis et al., 1998) and may be involved in nuclear function, chromosomal rearrangement, and other events associated with cell proliferation and motility.
1.6. Hyaluronan functions
Despite the presence of the polysaccharide seems to be of a vital importance, since no inherited diseases lacking hyaluronan are known today for several decades it was assumed that its major functions were in the biophysical and homeostatic properties of tissues.
Furthermore, it has been shown that hyaluronan is essential in some dynamic cellular systems such as morphogenesis (Toole, 2001), tissue regeneration (Toole et al., 1989), inflammation (Hascall et al., 2004), wound repair (Chen and Abatangelo, 1999) and tumorigenesis (Knudson et al., 1989) due to his capability to influence cell proliferation, differentiation and migration.
Striking examples of dynamic events during which cells are surrounded by hyaluronan-rich matrices are: mesenchymal cells dividing and migrating during embryonic limb development (Knudson and Toole, 1985; Toole et al., 1989);
neural crest cells travelling from the neural tube to form ganglia of the peripheral
nervous system (Toole, 2001); cushion cells migrating from the endocardium
towards the myocardium during formation of heart valves (Camenisch et al.,
2000); neuronal and glial precursors moving and proliferating during brain
development (Baier et al., 2007); mesenchymal cells invading the primary corneal stroma to form the mature chick cornea (Toole and Trelstad, 1971);
tendon regeneration (Majima et al., 2007); wound repair (Chen and Abatangelo, 1999) and tumor cell growth and invasion (Knudson et al., 1989).
In many cases hyaluronan is removed during final differentiation of cells subsequent to morphogenetic events (Toole 2001). An experimental model for the importance of hyaluronan in embryonic differentiation is the culture of trypsinised chick embryo cardiac cells. These grow, become confluent, fuse, synthesize cardiac actin and myosin and ultimately being to undergo waves of contraction. These same cells, however, cultured on hyaluronan, will grow but remain myoblasts, failing to undergo differentiation, and never undergo contractility (Kujawa et al., 1986).
Thus great and intricate are the mechanisms by which hyaluronan could achieve his different functions on cell migration and differentiation.
One way in which hyaluronan facilitates cell migration is by creating hydrated pathways that allow cellular or fibrous barriers to be penetrated by cells. In fact hyaluronan-rich areas within developing tissues exert internal pressures that can cause the separation of physical structures and create
“highways” for cell migration, as seen during the migration of mesenchymal cells into the cornea following increased hyaluronan deposition, hydration and concomitant swelling of the migratory pathway (Toole and Trelstad, 1971).
Moreover hyaluronan synthases activity has been shown to fluctuate with the cell cycle and to peak at mitosis. Thus, extrusion of hyaluronan onto the cell surface at mitosis would create a hydrated microenvironment that promotes partial detachment and rounding of the proliferating cells. In support of this idea, inhibition of hyaluronan synthesis has been shown to lead to cell cycle arrest at mitosis, just before cell rounding and detachment (Brecht et al., 1986).
Hyaluronan concentration may decreases in the pericellular matrix of
differentiating cells that have assembly in the correct number and place by
proliferation and migration promoting cell-aggregation, which is essential for the
differentiation process. Cell-aggregation may be achieved as cell surface
hyaluronan could cross-bridge cells via interaction with receptors on adjacent
cells, in fact removal of this hyaluronan would thus block aggregation while the
addition of excess hyaluronan would cause occupation of all receptors by hyaluronan, blocking cross bridging (Toole, 1998) (Fig. 4).
Figure 4 – Hyaluronan-mediated cell aggregation.
Hyaluronan cross-bridges cells bearing hyaluronan receptors such as CD44, causing cell-aggregation (B). If the cells are treated with excess hyaluronan, that occupies all receptors, cross bridging is blocked and aggregation inhibited (C).
HA, hyaluronan. Modified from Toole (1998).
Thus hyaluronan facilitates regional detachment of a cell from its substratum, a necessary step in both mitosis and migration, and creates a hydrated zone around cells that separates them from physical barriers to penetration or shape changes. In the other way, the hyaluronan instructive effect is via interaction with specific cell-surface hyaluronan receptors, such as CD44 or RHAMM that initiates signalling pathways that promote cell movement, proliferation or differentiation.
Several investigators have demonstrated that cell movement in vitro is
promoted in the presence of hyaluronan, that invasion into three-dimensional
collagen gels is dependent on hyaluronan synthesis and that cell movement is
inhibited as a result of degrading the hyaluronan itself or of blocking the binding
of hyaluronan to either of its receptors, e.g., CD44 and RHAMM (Toole, 1998).
Interaction of hyaluronan with RHAMM stimulates tyrosine phosphorylation of several proteins, including a key component of focal adhesions, p125
FAK, resulting in regulation of focal adhesion turnover and promotion of cell motility (Hall et al., 1994). Interaction of hyaluronan with cell surface CD44 also stimulates cell migration in some tumour cell types, such as glioma and melanoma cells (Radotra et al., 2001). Thus it seems that interaction of hyaluronan with either receptor can stimulate cell movement, but their relative importance may depend on the cell type or other physiological factors. Similarly, interaction of hyaluronan with CD44 or RHAMM stimulates signalling pathways involved in cell proliferation in different cell types (Toole, 1998). Nevertheless, the details of these putative pathways are by no means clear for either cell proliferation or locomotion.
2. Hyaluronan synthases
It was demonstrated that temporal expression patterns for the three Has genes differ both in the developing embryo and in adult tissues.
Different studies carried out in the last years have detailed the distribution of the Has genes expression during development of mouse (Tien and Spicer, 2005), zebrafish (Bakkers et al., 2004) and Xenopus (Vigetti et al., 2003;
Nardini et al., 2004), highlighting that the three Has genes are expressed in distinct spatial and temporal patterns. While Has1 is prominently expressed in the early stages of midblastula, gastrulation, and neurulation, Has3 is expressed later in development when sensory organs begin to form and Has2 is expressed throughout all developmental stages.
Has2 is the major hyaluronan synthase during development as it is the most widely expressed, consistently with the phenotype observed in Has2-deficient animals. In fact while the targeted inactivation of mouse Has1 and Has3 produce viable homozygous null animals, Has2 deficiency results in an embryonic lethal phenotype at midgestation (E10.5) due to heart and vascular defects (Camenisch et al., 2000).
In zebrafish embryos, loss of Has2 function (the unique hyaluronan
synthase presents in this specie during development) leads to blockage of
specific gastrulation movements. Zebrafish Has2 is also required for migration of
presumptive slow muscle and sclerotomal cells during morphogenesis of somites (Bakkers et al., 2004).
Recently we have demonstrated that in Xenopus laevis the loss of Has2 function alters somitogenesis and leads to a defective myogenesis. XHas2 activity is also required for the migration of hypaxial muscle cells and trunk neural crest cells (Ori et al., 2006).
Thus, hyaluronan is indispensable for normal vertebrate embryogenesis and the Has2 enzyme is the major source for hyaluronan during development.
3. The hyaladherins
A number of cell-associated and extracellular hyaluronan binding proteins (HABPs or hyaladherins) specifically recognize the hyaluronan structure mediating his biological functions. Among these CD44 (Underhill, 1992), RHAMM (Hardwick et al., 1992), link protein (Binette et al., 1994), aggrecan (Watanabe et al., 1997), versican (Zimmermann et al.,1989), LYVE-1 (Banerji et al., 1999), TSG-6 (Lee et al., 1992), IHABP (Hofmann et al., 1998), cdc37 (Grammatikakis et al., 1995), P-32 (Deb et al., 1996), SPACRCAN (Acharya et al., 2000) and SHAP (Huang et al., 1993).
Some HABPs contain a common domain of about 100 amino acids, termed
“link module”, through which they couple hyaluronan (Kohda et al., 1996). By contrast, some HABPs have a linear 9–11 residue “hyaluronan-binding motif”
containing multiple basic amino acids and bind to hyaluronan via this motif. This hyaluronan-binding motif is termed a B-X
7-B motif, where B is a basic amino acid residue and X is a non-acidic amino acid. For example, link protein, CD44, aggrecan, versican, LYVE-1 and TSG-6 all bind to hyaluronan via a link module, whereas RHAMM, IHABP, cdc37, P-32, SPACR and SPACRCAN bind to HA via a B- X
7-B motif.
This kind of interactions binds hyaluronan with proteoglycans to stabilize the structure of the matrix, and with cell surfaces to modify cell behaviour.
The cDNAs coding for two of such proteins, CD44 and RHAMM, have been
cloned and characterized during vertebrate development. CD44 has been
proposed to bind multiple ligands, not only hyaluronan, including collagens,
fibronectin (Jalkanen et al., 1992), heparin-binding growth factors (Bennett et
al., 1995), osteopontin (Weber et al., 1996) and chondroitin sulfate (Naujokas et al., 1993). Although the receptors bind other ligands than hyaluronan, mutation of a hyaluronan-binding domain in CD44 or RHAMM, ablates the enhanced motility and proliferation associated with tumorigenesis (Yang et al., 1994), underling an involvement of these receptors in cell proliferation and migration.
Both CD44 and RHAMM mediate hyaluronan signalling and participate in growth factor-regulated signalling. However, they likely regulate signalling by different mechanisms because they are not homologous proteins, are compartmentalized differently in the cell and differ in the mechanisms by which they bind to hyaluronan (Day and Prestwich, 2001). The role in cell signalling of the other identified hyaladherins has not yet been reported.
3.1. The hyaluronan-receptor CD44
Probably the most widely expressed and extensively studied hyaluronan- receptor is CD44 (Fig. 5). It is a widely distributed cell surface glycoprotein that is encoded by a single gene but it is expressed as numerous isoforms as a result of alternative splicing. The simplest and most widespread is termed the
“standard isoform” and denoted CD44s (often alternatively termed the
hematopoietic isoform or CD44H). The extracellular region of all CD44 isoforms
includes an amino-terminal domain (link-module) responsible for the binding to
hyaluronan. However, hyaluronan binding to CD44 is subject to numerous
additional positive and negative influences from other regions of the molecule,
e.g., glycosylation, alternative splicing, dimerization, clustering in the plasma
membrane and integrity of the highly conserved cytoplasmic domain (Lesley et
al., 2000; Skelton et al., 1998; Kincade et al., 1997). In response to hyaluronan
binding, and depending on the cellular context, the cytoplasmic tail of CD44
interacts with many regulatory and adaptor molecules, such as SRC kinases,
RHO GTPases, VAV2, GAB1, ankyrin and ezrin (Ponta et al., 2003; Thorne et al.,
2004). In culture, it has been shown that hyaluronan-CD44 interaction results in
activation of various components of intracellular signalling pathways, e.g. Rac1
(Oliferenko et al., 2005), PI3K (Kamikura et al., 2000), erbB2 (Bourguignon et
al., 2007) and NF-kB (McKee et al., 1997) and in rearrangement of cytoskeletal
elements, e.g., ankyrin (Zhu and Bourguignon, 2000) and ezrin/radixin/moesin
(Yonemura et al., 1998). In some cases, these effects cause a quite dramatic change in cell proliferation (Bourguignon et al., 2007), motility (Oliferenko et al., 2000; Zhu and Bourguignon, 2000) and invasion (Svee et al., 1996). CD44 also mediates the cellular uptake and degradation of hyaluronan, which in turn affects growth regulation and tissue integrity (Kaya et al., 1997).
During critical stages of morphogenesis, CD44 is expressed at high levels in developing heart, somites and condensing limb-bud mesenchyme at critical stages of morphogenesis. These sites correlate with regions where hyaluronan has been demonstrated to regulate morphogenetic events (Wheatley et al., 1993; Sretavan et al., 1994; Fenderson et al., 1993) suggesting that CD44 might play a critical role during embryogenesis.
Figure 5 - Model of the structure of the standard isoform of CD44s. The products of various combinations of ~10 alternatively spliced variant exons are inserted at the position indicated by the brace, giving rise to numerous variant isoforms of CD44. The extracellular domains of CD44 are highly, but variably, glycosylated, and several serine residues in the cytoplasmic domain can be phosphorylated.
Modified from Ponta et al., (1998).
3.2. The Receptor for Hyaluronan-Mediated Motility (RHAMM)
Another hyaluronan-binding protein implicated in cell behaviour, especially motility, is the “receptor for hyaluronan mediated motility” (RHAMM) (Hardwick et al., 1992). As CD44, these hyaladherin lacking a link module, is subject to alternative splicing generating several isoforms (Lynn et al., 2001a). RHAMM is transiently expressed on the cell surface, where it is now designated CD168, and in the cytoplasm, as well as in the cytoskeleton and in the nucleus (Turley and Harrison, 1999) (Fig. 6).
After its interaction with hyaluronan, but also with other ligands such as the fibronectin, RHAMM delivers signals for cell migration and proliferation in normal and malignant cells (Akiyama et al., 2001; Turley et al., 2002; Mohapatra et al., 1996; Cheung et al., 1999). In particular RHAMM is involved in the Ras and ERK signalling pathways (Hall et al. 1995; Zhang et al., 1998) and it is associated with the cytoskeleton (Assmann et al., 1999). Interaction of hyaluronan with RHAMM stimulates tyrosine phosphorylation of several proteins, including a key component of focal adhesions, p125
FAK, resulting in promotion of cell motility (Hall et al., 1994). Moreover RHAMM has been identified as a microtubule- associated protein that also interacts with the actin (Assmann et al., 1999):
probably, RHAMM influences the cellular motility owing to the polymeralization of actin and regulates the cell mitoses through spindle poles (Groen et al., 2004).
Figure 6 - Cell surface and intracellular RHAMM.
Cell surface RHAMM (or CD168) is proposed to modify erk1 activation via an association with growth factor receptors such as the PDGF receptor. Intracellular RHAMM forms are erk1 binding proteins and act downstream of both src and ras at the level of MEK1 to control signalling through ras. Modified from Turley and Harrison (1999).
A complete study of RHAMM protein expression in different adult or embryonic tissues has not been published, but unreported accounts utilizing RT- PCR analyses indicate that the largest intracellular RHAMM is present in most tissues (Turley and Harrison, 1999). In particular, it has been demonstrated RHAMM mRNA synthesis during development at morula and blastocyst stages of in vitro-produced bovine embryos (Stojkovic et al., 2003), in neurons of cerebral cortex and most subcortical and brainstem structures at postnatal day 1 rat (Lynn et al., 2001b) and in association with Xenopus laevis micro-tubules during mitotic spindle assembly (Groen et al., 2004). A few reports of immunocytochemical analyses from a limited number of adult tissues suggest that RHAMM expression may vary with specific cell types within tissues (Nagy et al., 1998; Pilarksi et al., 1994).
Although it is clear that CD44 and RHAMM can participate independently in proliferative and migratory phenomena, their relative contributions to any given event have not been fully resolved in most cases and it is likely that they have redundant or overlapping functions in some situations (Nedvetzki et al., 2004).
3.3. Hyaluronan-receptors loss of function studies
Unlike genetic deletion of HAS2, deletion of either CD44 (Schmits et al., 1997; Protin et al., 1999) or RHAMM (Tolg and Turley, unpublished data in Tammi et al., 2002) genes in mice does not result in embryonic lethality.
Despite CD44 expression pattern during mouse development, the loss of this
gene has no overt effect on mouse development, viability and differentiation of
tissues and organs (Protin et al., 1999), probably due to a redundancy effect of
other hyaluronan receptors. On the other side, adult mutant mice shown a
complex phenotype, exhibiting defective trafficking of leukocytes, altered
responses to tissue injury and transformation by oncogenic viruses such as SV40
(Schmits et al., 1997; Protin et al., 1999). It has been shown that in inflamed
CD44 knock-out mice, RHAMM compensates for the loss of CD44 in binding to
hyaluronan, supporting cell migration, up-regulating genes involved in the
inflammation and exacerbating collagen-induced arthritis. Interestingly, the
compensation for loss of the CD44 gene does not occur because of enhanced expression of the redundant gene (RHAMM), but rather because the lack of CD44 allows increased accumulation of the hyaluronan substrate, with which both CD44 and RHAMM engage, thus enabling augmented signalling through RHAMM (Nedvetzki et al., 2004).
In Xenopus laevis the down-regulation of XCD44 gene does not perturb the somitogenesis process that is instead altered in Has2 down-regulated embryos, but strongly impair hypaxial muscle precursor cell migration and the subsequent formation of the ventral body wall musculature (Ori et al., 2006). Moreover In contrast to XHas2, loss of function of XCD44 does not seem to be essential for trunk neural crest cells migration, suggesting that the HA dependence of trunk neural crest cells movement was rather associated with an altered macromolecular composition of the extracellular matrix structuring the cells migratory pathways or a redundancy effect of other hyaluronan receptors (Ori et al., 2006).
4. The hyaluronan binding proteoglycan versican
Versican is a chondroitin sulfate proteoglycan (PG) belonging to the hyalectan subgroup of PGs including aggrecan, neurocan and brevican, having in common an N-terminal G1 domain embodying the binding site for hyaluronan. In addition to the hyaluronan-binding domain, hyalectans show common structural features such as a set of epidermal growth factor (EGF), lectin and complement regulatory protein (CRP) in the carboxy-terminal portion (G3 domain) of the molecule (Zimmermann and Ruoslahti, 1989; Shinomura et al., 1993). The glycosaminoglycan (GAG) attachment regions are located in the middle portion of the protein and are different in the four ECM PGs of the hyalectan subgroup.
In versican, two different GAG attachment domains have thus far been identified
in higher vertebrates and denoted GAGα and GAGβ. Both domains (exons) are
present within the parental V0 isoform while they are differentially spliced out in
isoforms V1 and V2 (Zimmermann and Ruoslahti, 1989; Shinomura et al.,
1993). In the nervous system, versican seems to mainly act as barrier molecule
affecting axon growth, cell migration and plasticity and these functions appear to
be performed through its GAG chains (Schmalfeldt et al., 2000; Bradbury et al.,
2002). Recently, the V1 versican isoform has been shown to be involved in
mesenchymal-epithelial conversion (Sheng et al., 2006) and much of the above biological effects of the PG may be contributed by the interactions of the PG with several matrix molecules (Wu et al., 2005; Cattaruzza and Perris, 2006).
Versican has been found to be colocalized with hyaluronan, CD44, and tenascin in the pericellular matrix of cultured fibroblasts (Yamagata et al., 1993), and in epidermal keratinocyte tumours, levels of hyaluronan, CD44 and versican correlated with the aggressiveness of the disease (Karvinen et al., 2003).
Moreover, it has been shown that versican may interact directly with CD44 through its GAG chains, independently from hyaluronan, inducing CD44 activation and signals transduction (Kawashima et al., 2000).
A striking example that hyalectan proteoglycans are essential in forming and stabilizing hyaluronan-rich matrices derives from the study of the heart defect (hdf) mouse, arising from a transgene insertional mutation in the versican gene causing its almost complete abrogation (Mjaatvedt et al., 1998). In the hyaluronan-rich matrices in the developing mouse heart are present many other hyaluronan binding molecules, in particular the hyaluronan binding proteoglycan versican (McDonald and Camenisch, 2003). Has2 and versican display similar domains of expression in E9.5–10.5 mouse heart and the cardiac jelly is rich in HA and versican (Camenisch et al., 2000). Moreover, Has2 and versican null mice (Mjaatvedt et al., 1998; Yamamura et al., 1997) exhibit a common abnormal cardiac morphogenesis, demonstrating that both matrix molecules are essential for formation of cardiac jelly and endocardial cushions (Camenisch et al., 2000). It has also been underscored a crucial role of isoforms V0 and V1 during trunk neural crest cell migration suggesting a chemorepulsive function (Landolt et al., 1995; Dutt et al., 2006). Other studies propose instead that versican is a component of the extracellular matrix of neural crest pathways controlling directionality of cell movement through a mechanism of “inverse haptotaxis” (Perris and Perissionotto, 2002), consistent with previous findings in the white mutant axolotl embryo harbouring a genetic deficiency in versican expression (Perris et al., 1990).
However, the complex structure of versican provides the basis for its
multiple and sometime contrasting effects on cellular functions, such cell
adhesion, cell migration, cell proliferation and apoptosis, as well as on
morphogenesis.
5. The amphibian Xenopus laevis as model system
The African claw-toad frog Xenopus laevis is one of the most popular systems used in the field of developmental biology. Its advantages come from the easy accessibility and large size (1 mm in diameter) of the egg, which can be easily fertilized in vitro. Moreover, the embryos can be obtained in high number (up to 1500 per female), develop rapidly (see Fig. 7 for life cycle), are easy to handle in culture, and survive well to microsurgical manipulations (microinjections and transplantations). The egg has an initial radial (animal- vegetal) axis of asymmetry. Sperm entry induces the microtubule-driven rotation of the egg cortex by 30° relative to the inner cytoplasm (Gerhart et al., 1989). As a result, maternal determinants are displaced to the equatorial region opposite the sperm entry point; this defines the future dorso-ventral axis of the embryo (Harland and Gerhart, 1997; De Robertis et al., 2000).
Under the control of maternal regulators (cyclins and cyclin-dependent kinases), the fertilized egg undergoes synchronised divisions, which lead successively to the morula (mass of cell without an internal cavity), and the blastula, which has an internal cavity (blastocoel). The morphology and radial symmetry of the embryo are maintained until late blastula stage. Following the onset of zygotic transcription at midblastula, a signalling center (the Spemann organizer) becomes established in the future dorsal side of the embryo. In a process known as gastrulation, cells migrate and extend from this so-called blastopore lip, to form the three germ layers of the gastrula embryo. The ectoderm is at the outside of the embryo, the endoderm is at the inside and the mesoderm is in between the two. Cells of different germ layers interact with each other to initiate differentiation; these interactions are known as primary inductions. At the completion of gastrulation/early neurulation, a second signalling centre called notochord is form by a rod of mesodermal cells in the most dorsal portion of the embryo (Scott, 2003; Johnson and Raven, 2002).
Organogenesis starts when the notochord signals to the dorsal ectoderm
and induces it to form the neural tube (neurula stage). Once the neural tube has
formed, it induces changes in the neighbouring cells. During organogenesis,
already differentiated cells (or tissues) interact with each other, in processes
known as secondary inductions. At the end of organogenesis, the neurons have
made connections between themselves, and with muscles. The larva (tadpole) is
then ready to hatch (Scott, 2003; Johnson and Raven, 2002).
Figure 7 - Xenopus laevis life cycle. From Wolpert et al., (1998).
6. The neural crest cells
In 1868, the Swiss embryologist Wilhelm His discovered a band of cells sandwiched between the developing neural tube and the future epidermal ectoderm in neurula stage chick embryos. He identified this group of cells as the source of spinal and cranial ganglia and named them “Zwischenstrang-the intermediate cord”.
In a 1878 paper on the development of the cranial nerves in chick embryos, Arthur Milnes Marshall used the term “neural ridge” for the cells that give rise to cranial and spinal nerve ganglia. Realizing that this term was less descriptive than was desirable, a year later he replaced neural ridge with “neural crest”.
6.1. Neural crest cells migration
The vertebrate neural crest is a migratory embryonic cell population that forms at the border between the neural plate and the future epidermis. Neural crest cells (NCCs) delaminate from the neuroepithelium in a rostro-caudal wave and migrate throughout the embryo to form a wide range of derivatives. The NCCs migration occurs in three stages: initiation, dispersion and cessation of migration.
6.1.1. Initiation of migration
Although derived from the ectoderm, the neural crest has sometimes been called “the fourth germ layer” because of its importance. Neural crest is a transitory structure that originates during the process of neurulation. The neural tube closes first in the head and then the closure progresses tailward;
consequently NCCs delamination and migration occur in a wave along the rostro-
caudal axis of the embryo following the progression of neural tube closure. While
in mammals the cranial neural crest begins to migrate when the neural tube is
still open (Morriss-Kay and Tucket, 1991), in amphibians, as well as in the chick,
the onset of crest cell migration coincides with the closure of the neural tube
(Tosney, 1982). In particular, the segregation of Xenopus laevis neural crest
material from the neural plate and surrounding epidermis begins at stage 15,
indicated by a loosening of the cells. At stage 16 the neural crest has already
been delimitated in the anterior and middle regions of the neural anlagen, while at stage 17 this process is just in progress in the caudal region. At these stages the neural crest material has already been displaced to the lateral side of the future neural tube in the anterior half of the embryo. The anterior border of the massive neural crest, developing from the lateral margins of the neural anlagen, lies at the level of the posterior half of the eye anlagen at stage 18, at which stage also a first segregation takes place of mes- and rhombencephalic neural crest (Nieuwkoop and Faber, 1956)
NCCs are specified as the result of an inductive action by the non-neural ectoderm (lateral cells of the neural plate), mediated by the bone morphogenetic proteins BMP-4 and BMP-7 (Liem et al., 1995). BMP-4 and BMP-7 induce the expression of the Slug and the RhoB proteins in the cells destined to become neural crest (Nieto et al., 1994; Mancilla and Mayor 1996; Liu and Jessell 1998).
Slug is a transcription factor of the zinc finger family that induce NCCs to break away from the neural tube by changing their shape and properties from those of typical neuroepithelial cells to those of mesenchymal cells (Epithelial- Mesenchymal Transition). The RhoB protein establishes the cytoskeletal conditions that promote migration (Carlson, 1999). However, the cells cannot leave the neural tube as long as they are tightly connected to one another and one of the functions of the Slug protein is to activate the factors that dissociate the tight junctions between the cells (Savagner et al., 1997). In fact, another important factor in the initiation of NCCs migration is the downregulation of the N-cadherin and N-CAM, transmembrane glycoproteins responsible for Ca
2+- dependent cell-cell adhesion. As final effect of all these and other changes, that may be achieved by a genetic control or a response to environmental triggers, the NCCs become more motile (Monier-Gavelle and Duband, 1995).
6.1.2. Dispersion
The NCCs migrate from the dorsal side of the neural tube to various parts of the embryo along defined pathways that can be divided into two main functional domains (Fig 8):
(1) the cranial (or cephalic) NCCs, that migrate dorso-laterally to originate
cephalic ganglia, sclera, choroid and cornea of the eye, but also to some
mesenchymal cells that differentiate into the cartilages and bones of the skull and the head skeleton. Cranial NCCs are the only crest population able to give rise to skeletal elements (Nakamura and Ayer-Le Lievre, 1982). Sometimes the neural crest-derived mesenchyme is know as ‘‘mesectoderm’’ to reflect its ectodermal origin (Hall, 2005). The mesectoderm also provides head connective- tissue cells and the vascular smooth-muscle cells associated with the vessels derived from the aortic arches and with the vessels irrigating the forebrain and face (Etchevers et al., 2001).
(2) the trunk NCCs, that migrate ventrally between the somites and neural tube along various pathways to give rise melanocytes, dorsal root ganglia, the adrenal medulla and the nerve clusters surrounding the aorta. A portion of the anterior trunk neural crest enters the heart and forms the separation between the pulmonary artery and aorta (Le Lièvre and Le Douarin, 1975). The most posterior trunk NCCs, the vagal and sacral NCCs, generate the parasympathetic (enteric) ganglia of the gut (Le Douarin and Teillet, 1973; Pomeranz et al., 1991).
Figure 8 – Schematic representation of the principal cranial and trunk neural crest cells derivatives
.
In Xenopus laevis, the migration of the cranial neural crest is quite distinct
from that of the trunk. In the trunk, the NCCs are subjected to the alternating
permissive and inhibitory halves of somites, which result in their segmental
organisation (Rickmann et al., 1985). In the head, the NCCs pour out from the
developing brain into a periphery that is devoid of somites. Yet, the cranial NCCs do not migrate individually but are organised into separate streams, three of which can be identified in the developing head of all vertebrate embryos:
mandibular, hyoid, and branchials streams (Nieuwkoop and Faber, 1956) (Fig.9).
Figure 9 – Schematic representation of Xenopus laevis skeletogenic cranial neural crest cells development. I, mandibular crest stream; II, hyoid crest stream; III-IV, branchial crest streams; ET, ethmoidal-trabecular plate cartilage; Q, palatoquadrate cartilage; So, subocular cartilage; M, Meckel’s cartilage; C, ceratohyal cartilage; G, gills cartilages. Modified from Sadaghiani and Thiebaud (1987).
The mandibular crest stream
At stage 20 the first crest segment (mandibular stream), originated from the
future midbrain and the first two rhombomers of the future hindbrain, has
already grown beyond the optic vesicle. By stage 21-22 it moves and curves
ventrally to the optic vesicle; some cells from its dorsal part migrate rostrally
over the eye. By stage 23 the caudo-ventral portion of the mesencephalic crest
has reached the most anterior part of the eye vesicle and its dorsal portion has
located dorso-rostrally to the eye, but at this stage both portions have not yet
rejoined. On the anterior part of the brain almost all crest material has by now
migrated to a lateral position and the brain is free of crest cells on its dorsal
side, anterior to the otic vesicle. By stage 24 the dorsal and the caudo-ventral
portions of the mandibular crest stream have already rejoined each other,
surrounding the optic vesicle. At stage 25, the cells of the first crest segment
penetrated in the maxillary mesoderm of the first pharyngeal arch (mandibular
arch). Here the caudo-ventral portion of the mandibular stream will take part in the formation of the palatoquadrate (Q) and Meckel’s (M) cartilages, while the dorsal portion the ethmoidal-trabecular plate (Et) cartilage. The primordia of these cartilages are already laid down at stage 40, become pro-cartilaginous at stage 41 and cartilaginous at stage 43. In the most rostral part of the embryo head the Meckel’s cartilages of both sides are connected through the infrarostral cartilage (IR) that is not a neural crest derivative (Nieuwkoop and Faber, 1956).
The hyoid crest stream
The second crest stream, the hyoid stream, arises from the fourth rhombomere of the future hindbrain. It begins to migrate at stage 20 descending downward over the mesoderm of the second pharyngeal arch (hyoid arch) from the rostral part of the otic vesicle and by stage 23 it has descended just halfway toward its final position. At stage 24 the hyoid crest has moved to the ventral part of the hyoid arch mesoderm and has already penetrated into it. The massive stream of the hyoid stream cells, which takes part in the formation of the ceratohyal cartilage, migrates ventrally to the bottom of the pharynx and become located in the ventral midline in the second arch connective tissue at stage 29/30. It should be mentioned that there is a small contribution of the crest cells in the formation of the muscles connected to the ceratohyal cartilage.
Later in development the ceratohyal articulates with the ventral surface of the first branchial stream derived palatoquadrate. The ceratohyal is present as a mesenchymatous condensation at stage 40, it is pro-cartilaginous at stage 41, begins to chondrify at stage 42, while chondrification is completed at stage 43 (Nieuwkoop and Faber 1956).
The branchial crest streams
Finally, the third segment originates from the most posterior rhombomers and contributes to the formation of the third and fourth pharyngeal arch (branchial arches): the gills. The deep groove between the second and the third crest segments is occupied by the otic placode, thus the branchial streams begin to migrate, at stage 21, over the branchial region, posterior to the otic vesicle.
By stage 23, for the first time, a split can be seen that subdivides the branchial
crest segments into two portions (anterior and posterior) in such a way that the anterior portion moves in front of the posterior portion during the following stages. The dorso-posterior border of the posterior branchial stream represents a continuation of the trunk neural crest material that has not yet started to migrate and is still located on the middle of the neural tube. At stage 25 two contacting cell masses of the anterior and the posterior branchial stream are located on the swelling branchial region, and their ventral borders cannot be seen in the lateral view. Primordial gills have already been laid down at stage 40 in the form of mesechymatous condensation and the chondrification has started at stage 43 to completed at stage 46 (Nieuwkoop and Faber 1956).
6.1.3. Cessation of migration
After a phase of migration, the NCCs stop in defined embryonic areas where they differentiate into the specific cell types. Among the possibilities that can be put forward to explain when and why NCCs terminate their migration and initiate cytodifferentiation at their destination site, are that some kind of obstacle forces the cells to stop and aggregate (Le Douarin, 1982). Obstacles or barriers can include basal laminae, blood vessels, somites or other cell clusters. The increasing of the production of chondroitin sulphate also represents a barrier to crest cell migration. Actually cessation of migration is not as simple as the cells just hitting a barrier and stopping; it is more complex than that.
Furthermore the cessation of migration is achieved in part by a reversal of the mechanisms used to initiate migration. The adhesion properties change:
certain integrins are down-regulated and, dependent upon the fate of the cell, N- cadherin, N-CAM and E-cadherin can be re-expressed.
7. The extracellular matrix and neural crest cells migration
The extracellular matrix components undergo structural alterations and
biochemical changes when NCCs migrate in or on it. There is some extracellular
matrix molecules that promotes migration, since they are present throughout the
matrix encountered by the trunk NCCs, such as hyaluronan (Pintar, 1978),
fibronectin (Newgreen and Thiery, 1980), laminin (Krotoski et al., 1986),
tenascin (Tan, 1987) and various collagens (Perris et al., 1991). Other proteins are instead involved in the prevention of NCCs migration, such as the chondroitin sulfate proteoglycans aggrecan and versican (Tan, 1987; Perris et al., 1991) that are generally present in regions from which NCCs are absent. In fact these proteoglycans tend to inhibit NCCs migration in vitro, consistent with the idea that they may restrict or inhibit migration in the embryo. In chick embryos hyaluronan was found widely distributed along neural crest migratory pathways, while as the migration ceases and gangliogenesis commences, its levels appear reduced in the immediate vicinity of NCCs (Perris et al., 1991). To ensure successful neural crest migration, each matrix molecule appears to be required in specific quantities, at particular locations, according to a tightly regulated temporal programme of development. Moreover, expression of the various molecules must be precisely coordinated with each other. It seems most likely that a balance between permissive and non-permissive molecules is the crucial factor. Permissive molecules allow cell attachment and spreading, whereas non- permissive molecules are necessary to ensure that attachment to the substrate is not so tight as to preclude forward migration (Henderson and Copp, 1997). It is worth to underlie that cranial and trunk cells use different mechanisms of attachment to extracellular matrices. In fact, while trunk NCCs attach to fibronectin, laminin and collagens, cranial NCCs do not attach to collagens although they do attach to basal laminae.
NCCs possess several integrin receptors that recognize a variety of extracellular matrix molecules as demonstrate in vitro, where antibodies against integrins block NCCs attachment to fibronectin, laminin, and collagens, suggesting that these are the primary mediators of NCCs attachment (Lallier and Bronner-Fraser, 1991). These receptors integrate extracellular and intracellular scaffolds, enabling cells to move by contracting the actin microfilaments against the fixed extracellular matrix (Horwitz et al., 1986; Tamkun et al., 1986;
Ruoslahti and Pierschbacher, 1987). It has also been shown that the presence of
bound integrin may prevents the activation of genes that specify apoptosis
(Montgomery et al., 1994), in particular, chondrocytes of chick sternum can
survive and differentiate only if they interact with extracellular matrix through
their integrins (Hirsch et al., 1997). However, the mechanisms by which bound
integrins inhibit apoptosis remain controversial.
Another set of proteins involved in NCCs are the ephrin. These proteins are expressed in the posterior section of each sclerotome, and wherever they are, NCCs do not go (Krull et al., 1997; Wang and Anderson, 1997). The NCCs recognize the ephrin proteins through their cell surface Eph receptors. Binding to the ephrins activates the tyrosine kinase domains of the Eph receptors in the NCCs and these kinases probably phosphorylate proteins that interfere with the actin cytoskeleton that is critical for cell migration.
It is also important for NCCs, especially those in the trunk, to have sufficient cell-free extracellular space to migrate normally. In fact, NCCs exert relatively weak traction forces on the substrate, an attribute that allows them to migrate on or through relatively weak extracellular matrices (Hall, 1999). Among all matrix components, the glycosaminoglycan hyaluronan is the mainly involved in the initial separation of NCCs from the neural tube: due to highly hydrophilic feature it could opens up of spaces through which NCCs migrate (Hall, 1999). On the other hand, NCCs themselves can produce proteases e.g., plasminogen activator, hat help create the spaces through which they migrate and also synthesize some matrix material such as hyaluronan and versican Menoud et al., 1989).
The extracellular matrix is also involved in the cessation of NCCs migration.
Some regions of the embryo undergo a decline in hyaluronan levels, causing a
decrease in the amount of extracellular space. Moreover the extracellular matrix
molecules, along which the cells migrate, such as fibronectin and collagen, can
be decreased in number, reducing the availability of a migratory substrate. NCCs
also use “stop signals”, in fact, as these cells migrate into particular destination
sites, they encounter new extracellular matrix components that serve as stop
signal cues. It has been proposed that NCCs migration is inhibited when motility-
promoting matrix adhesion molecules become in lesser proportion to
hyaluronan-aggregating proteoglycans; in other words, hyaluronan-proteoglycan
complexes inhibit migration (Perris and Johansson, 1990). These investigators
found that in in vitro cartilage tissues aggrecan inhibited NCCs migration on
fibronectin substrata, and that this inhibition was abrogated when hyaluronan
was eliminated from the culture system by Streptomyces hyaluronidase
treatment. Other researchers have observed similar effects on invading tumour
cells. Mammary tumours and gliomas of higher invasive grade have a lower ratio
of hyaluronectin to hyaluronan (Dauce et al., 1992). Thus, hyaluronan- hyaluronectin complexes may also serve as stop signals.
Experiments performed by Knudson and Knudson on NCCs obtained from chick embryo neural tube explants, leads to hypothesize a model of hyaluronan interaction on NCCs stopping migration (Fig. 10). Hyaluronan may provide a hydrated milieu conducive to cellular migration or directly facilitate cell migration via interaction with cell-surface hyaluronan receptors. The interaction of hyaluronan with his receptors results in a chemokinetic response. Moreover the expression of unoccupied CD44 hyaluronan receptors may allow cells to adhere to and be translocated through hyaluronan-enriched extracellular matrices (FIG.
10A). When migrating cells encounter new matrices enriched in hyaluronan as well as matrix hyaladherins, the assembly of new pericellular matrices results.
Such matrices may result in a steric hindrance of the interaction with migration signals. Individual cells may remain isolated in matrix cocoons within connective tissues or may initiate new patterns of cytodifferentiation within this new location (FIG. 10B). NCCs may thus use these same hyaluronan receptors both for migration through hyaluronan-enriched extracellular matrices and in the assembly of hyaluroan/hyaluronectin containing pericellular matrices, blocking their interaction with other matrix adhesion proteins, to terminate migration (Knudson and Knudson, 1993).
Figure 10- Model of hyaluronan interaction on NCCs stopping migration. (A) Hyaluronan provides a hydrated milieu conducive to cellular migration. (B) Matrices enriched in hyaluronan and hyaladherins isolates individual cells that stop migration to initiates new patterns of cytodifferentiation within this new location. Modified from Knudson and Knudson (1993).
8. Cranial neural crest cells differentiation
A portion of NCCs migrate toward their final destinations into the pharyngeal
pouches where they proliferate and differentiate. The most characteristic feature
in the development of the Xenopus laevis foregut is the formation of the visceral
pouches and the oral evagination. The first and second pharyngeal or visceral
pouches are indicated as slight depression in the lateral wall at stage 20. They
deepen gradually at stage 21 and 22. The endodermal wall of the first visceral
pouch evaginates and protrudes towards the simultaneously developing local
thickening of the sensorial layer of the ectoderm. Both layers establish a first
contact at stage 23 and become intimate at stage 24. In this way the first
pharyngeal arch (mandibular arch) is formed. The development of the second
visceral pouch, separating the second visceral arch (hyoid arch) from the
following pharyngeal arches (branchial arches), begins somewhat later. Here
both layers approach each other at stage 24, make a first contact at stage 26
and establish an intimate contact at stage 27. The third visceral pouch is
indicated as a slight depression of the latero-caudal wall of the foregut cavity at
stage 23. The endodermal layer and the only slightly thickened sensorial layer of
the epidermis approach each other at stage 26 and make first contact at stage
27, so that now third visceral arch (the first branchial arch) is separated from
the rest of the branchial mesoderm at stage 27. The other two branchial arches
develop in a similar way and are not well defined before stage 35/36. At stage
29/30 the cell material of the first two, and at stage 31 that of the first four
pharyngeal arches, already shows a clear distinction between future skeletogenic
and myogenic elements, of which the latter are now located more superficially
and are very rich in yolk material. The skeletogenic elements are of neural crest
origin (Fig.11). They were originally located superficially, but have already
migrated into deeper layers (Nieuwkoop and Faber, 1956).
Figure 11 – Xenopus laevis pharyngeal arches. (A) Hoechst staining to visualize nuclear organization of branchial pouches different layers in horizontal section of tailbud stage Xenopus laevis embryo. (A’) schematization of branchial pouches layering indicated by the white square in A showing the distinction between endoderm (ent) and ectoderm (ect) of the branchial pouches and the future skeletogenic (nc) and myogenic (mes) elements.
