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The Search for the Magic Bullet
Chapter
The Search for the Magic Bullet
The message is ‘speed up, not start up.’
Although neither notochord nor spinal cord provides information to sclerotomal mesenchyme that is essential for chondrogenesis to commence in vitro, both tissues provide important and necessary information in vivo. While the inducer can be ‘bypassed’ or ‘mimicked’ in vitro by conditioned medium, embryos cannot compensate for the absence of spinal cord or notochord. Rather, vertebral differentiation and/or morphogenesis will be disrupted. Consequently, I think we are justified in continuing to refer to spinal cord and notochord as inducers of sclerotomal chondrogenesis and to their interaction with sclerotome as inductive. In this Chapter I explore what we know of the inducers and of their mechanisms of action on vertebral chondrogenesis and morphogenesis.
INTEGRITY OF NOTOCHORD/SPINAL CORD AND VERTEBRAL MORPHOGENESIS As discussed at the beginning of Chapter 41, intact spinal cord, notochord and spinal ganglia influence vertebral morphogenesis. But must notochord and spinal cord be intact to act? What type of registration, if any, exists between the axial tissues and vertebral segmentation? Normal neural arches develop in the presence of disorganized neural tissue in the killifish, Fundulus heteroclitus. In urodele amphibians and birds, on the other hand, abnormalities of the spinal cord lead to vertebral anomalies; administering nicotine sulphide to twoday-old chick embryos results in twisting of the notochord in the neck within hours and vertebral deformities within days. An interesting difference in the integrity of cartilage appears to depend on whether induction is by spinal cord
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or notochord. Cartilage induced by spinal cord dissipates between the 9th and 16th days in culture. Notochordinduced cartilage persists until at least the 25th day. Lash extrapolated this result to suggest that the spinal cord helps to remodel the vertebral column as it grows in vivo, thereby preventing compression of the spinal cord, an interesting idea that should be explored further, especially in relation to vertebral anomalies in fish raised under aquaculture conditions.1 Fish skeletal defects Laboratory rearing, like domestication (Box 7.2), influences variation; there is greater variation in all parts of the vertebral column in laboratory-reared than in wild-caught Red Sea bream, Pagrus major (Matsuoka, 1982). Defects represent variation outside that expected for the species. Skeletal defects in hatchery-reared fish are sufficiently common to be an accepted cost of aquaculture and a major concern for aquaculturalists. Depending on species and hatchery conditions, as many as 50 per cent of the fish will have vertebral defects. In one study, 44 per cent of hatchery-reared Senegal sole, Solea senegalensis, showed vertebral abnormalities, 28 per cent of which affected the tail and vertebral column. Although clues to the aetiology of some defects can be obtained from teratological studies – wavy notochords and bent gill arches result when retinoic acid or collagen maturation is blocked in flounder embryos (Suzuki et al., 2001) – determining the origins of vertebral defects has been an intractable problem. From studies on Atlantic salmon, Salmo salar, cultured in Tasmania, Australia, we know that gender has no influence but that chromosome number (ploidy) significantly affects the incidence of deformities. For example, only
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1 per cent of diploid freshwater smolts have lower jaw deformities. By contrast, the incidence of deformities in triploid fry, freshwater and seawater smolts is 2, 7 and 14 per cent, respectively. Similarly, primary gill filaments fail to form in up to 60 per cent of triploid individuals but only in 4 per cent of diploids. Triploidy provides the clue. The underlying mechanism remains elusive. Triploid rainbow trout, Oncorhynchus mykiss, were found to develop an additional caudal vertebra than found in diploid individuals, No other skeletal changes were found.2 Development of ‘short-tails’ is a common problem in farmed Atlantic salmon. These are fish in which the vertebral column is compressed along the A-P axis to such an extent that body length is reduced and body depth so enlarged that the body form hardly resembles that of a salmon (see Figure 1 in Witten et al., 2005). The defect at the cellular level involves the metaplasia of intervertebral tissues to cartilage, essentially replacing the notochord with cartilage (Witten et al., 2005).
Table 42.1 Inductive ability of notochord and spinal cord from chick and mice on somites of 9-day-old mice and H.H. 15, 17 or 18 chick embryosa H.H. stage or age in days of notochord/ spinal cord donor
Stage 18 Stages 27–37.5
Cartilage induced in transfilter recombinations (%)
Cartilage induced in direct recombination (%)
Chick–chick chimaeras – notochord 100 – 100 100
Stages 21–24 Stages 12–15
Chick–mouse chimaeras – notochord 100 100 83 –
5.5 daysb 7.5 daysb 5.5–6.5 daysc
Chick–mouse chimaeras – spinal cord 100 – 100 – 0 –
9 days
Mouse–mouse chimaeras – notochord 83 –
9 daysb
Mouse–mouse chimaeras – spinal cord 87.5 100
a
FOR HOW LONG ARE NOTOCHORD AND SPINAL CORD ACTIVE? In many inductive interactions the inducing cells lose their ability to induce at the same time as the responding cells lose their ability to respond, a mutually convenient temporal association that assures that induction will not be prolonged. As the spinal cord and notochord appear to play specific roles in inducing somitic cartilage, it is of interest to know whether they lose their inductive ability. I can summarize by stating that, as late as has been tested, notochords retain inductive activity but spinal cords do not. The evidence comes from combining notochord or spinal cord from older embryos or neonates with somites from younger embryos to determine whether cartilage forms. Some typical studies are summarized below and in Tables 36.1–36.3. Cooper (1965) used chick–chick and chick–mouse chimaeras to show that notochord from embryos as old as H.H. 37 (11.5 days) can induce chick or mouse somites to form cartilage (Fig. 37.1 and Table 42.1). Notochords from older chick embryos were not tested but notochord from 19-day-old mice remains inductively active (Fig. 37.1). The spinal cord does not retain inductive ability for the same period, which may reflect, in part, earlier maturation of spinal cord in comparison with notochord. Spinal cords from mouse embryos older than nine days of gestation no longer induce (Tables 42.1 and 42.2). We found that spinal cord from chick embryos as old as H.H. 44 (18 days of incubation) allow H.H. 17 somites to chondrify in vitro under conditions wherein somites alone fail to chondrify (Tremaine and Hall, 1979). Ability or inability to induce correlates with state of differentiation. Notochord (and cartilage; Chapter 22) only induces when the cells are undergoing hypertrophy or vacuolation (Table 42.3).3
Based on data from Cooper (1965). Ventral half of spinal cord. c Dorsal half of spinal cord. b
Table 42.2 Relative amount of cartilage in somites cultured with spinal cord from mice of different agesa Age of spinal cord (days)
Relative % of cartilage/culture
9 11 12 and 13
100 80 25
a
Based on data from Grobstein and Holtzer (1955).
CAN DERMOMYOTOME OR LATERAL-PLATE MESODERM CHONDRIFY? During embryonic development the sclerotome chondrifies, the dermotome produces connective tissue, and the myotome forms muscle (Fig. 16.1). Can dermotome or myotome form cartilage if suitably challenged? An unstated assumption underlying such a question is that distance from the normal inducers is all that prevents dermotome and myotome from chondrifying. Three major types of experiments have been designed to answer this question. Although muscle development is delayed, cartilage differentiation proceeds normally after 75 per cent of the medial aspect of the somites is removed from tail budstage urodele embryos. Apparently, areas of the somites normally destined for connective tissue and muscle regulate (Box 35.1) and chondrify. Implanting spinal cord or notochord into the dermomyotome of embryonic chicks elicits a secondary (ectopic) neural arch as chondrogenic differentiation and morphogenesis are initiated in non-sclerotomal mesenchyme.
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The Search for the Magic Bullet Table 42.3 Cartilage-inducing activity of embryonic chick and mouse notochord at four developmental stagesa Stage of developing notochord
H.H. stage/age of embryo
Number of cultures tested in notochord stage
Cartilageinducing activity
1 Aggregating chordamesoblast cells
Chick H.H. 12–15 Mice 9 days Chick H.H. 18 Chick H.H. 21–24 Chick H.H. 27–37.5 Chick H.H. 37.5 vertebral canal
2 3 20 18 77 2
None None Cartilage induced Cartilage induced Cartilage induced None
2 Intercellular vacuolation 3 Cellular hypertrophy 4 Non-hypertrophied state
a
Based on data from Cooper (1965). Notochord and somites were either recombined directly or transfilter and maintained in vitro.
Table 42.4 Chondrification in lateral-plate mesoderm from chick embryos of H.H. 9–12 grafted to the CAMa Grafted tissue
Chondrification (%)
Lateral-plate mesoderm Lateral-plate mesoderm ectoderm and endoderm Lateral-plate mesoderm spinal cord
4 37
a
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Based on data from O’Hare (1972c).
Cartilage forms in CAM-grafted lateral-plate mesoderm adjacent to the last four pairs of somites from embryos of H.H. 9–12, provided that ectoderm/endoderm or spinal cord is included in the grafts (Table 42.4). This result removes an interpretation applied to some earlier experiments – that transplanted spinal cord or notochord did not induce lateral mesoderm, but attracted sclerotomal cells into the lateral mesoderm where they formed the cartilage.4
THE SEARCH FOR THE MAGIC BULLET The search for a pure inducer that pervaded so many areas of embryology and developmental biology throughout the 20th century also directed, channeled, perhaps even hamstrung, students of somite chondrogenesis. For a brief period, beginning with a study by George Strudel in 1953, it looked as if progress was being made, but no significant advances were made in the two or three decades that followed.5 Strudel found that a saline extract of spinal cords or notochords from H.H. 18–21 chick embryos could induce somitic chondrogenesis in vitro.6 A decade later, he extended this finding back to earlier stages, concluding: …even very young somite cells are genetically determined cells and all that they need to undergo or accomplish their phenotypic differentiation is a microenvironment favouring chondrogenesis. This does not mean that the inducing action of the spinal cord and the notochord is dispensable. It may be that the spinal cord and/or the notochord exercise their inducing effect very early. (pers. comm.)
In 1957, Jay Lash and his colleagues introduced an ingenious experimental procedure adopted by a number of us. Embryonic spinal cords were placed transfilter to somites in vitro for 10 hours, during which time the spinal cord deposited extracellular material onto and into the filter. The spinal cords were removed and somites then placed onto the matrix products deposited into the filter and allowed to develop. Three days later, cartilage differentiated. ‘Control’ somites cultured on filters without ECM products formed no cartilage. Lash and colleagues concluded that only a short period of contact between somite and inducing spinal cord is needed to transfer a chemical signal between spinal cord and somites (Lash et al., 1957). A role for ectoderm? In studies carried out in the 1920s to 1940s ectoderm and possibly also subjacent endoderm were often included with grafted somitic mesoderm, either deliberately or inadvertently (see Chapter 41). Potential problems associated with ectodermal contamination were taken up as researchers asked whether ectoderm adjacent to the somites plays any role in cartilage induction. The results are contradictory and the final story is not in even now. We can say that epithelia exert no direct inductive action in vivo; chondrogenesis is not initiated when notochord or spinal cord are extirpated but dorsal epidermis is left intact. If epithelia play a role, it is secondary to and dependent on notochord and/or spinal cord. A sample of the results and conclusions include the following. ●
●
In somites from embryos of H.H. 10–13 grafted with or without ectoderm or endoderm, cartilage forms only when an epithelium is present. Seventy-five per cent form cartilage when ecto- and endoderm are present, 57 per cent when only ectoderm, and 14 per cent when only endoderm is present. Seno and Büyüközer (1958) concluded that epithelia, especially ectoderm, provide mechanical support for sclerotomal mesenchyme, thus preventing cells from dispersing. They ruled out a more physiological role. Stockdale et al. (1961) wrapped ectoderm around pellets of somitic cells equivalent in size to two to four somites and cultured them. Cartilage failed to develop, a result
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Bones and Cartilage: Developmental and Evolutionary Skeletal Biology
that does not necessarily rule out a role for ectoderm acting on intact somites in ovo. Similarly, no cartilage formed in any of 90 cultures of strips or clusters of somite wrapped in ectoderm or endoderm and either cultured or grafted into the coelomic cavities of host embryos (Lash, 1963b). Among the tissues Holtzer (1964b) tested for inductive ability, epidermis (source and age not specified) would not induce H.H. 18 somites to chondrify in vitro.
The British came to the rescue with an extensive (and definitive?) series of experiments by M. J. O’Hare, published in three parts in 1972. O’Hare grafted the last four somites from embryos of H.H. 9–12 after associating them with one of a variety of epithelia. As you can see from Table 42.5, somites grafted alone fail to chondrify, but a variety of epithelia permit chondrogenesis in anywhere from 10 to 30 per cent of the grafts. O’Hare also showed that the ability to augment chondrogenesis was not restricted to epithelia adjacent to somitic mesoderm but also was present in limb bud and trunk epithelia from older embryos (Table 42.5). He explained these temporal differences as based on a correlation with presence of a basement membrane and associated extracellular matrix (ECM) in ‘active’ epithelia. Such temporal differences argue against a strict mechanical effect, as does the ability of lateral-plate mesoderm to chondrify when associated with ectoderm (Table 42.5). The tentative conclusion from these studies is that epithelia allow somites to chondrify in the absence of notochord or spinal cord, if somites are CAM-grafted but not if cultured. Conclusions relating to any role for epithelia in vivo must be confined to the statement that epithelia cannot act in the absence of spinal cord or notochord. Cartilage cells as cartilage inducers
the container – brain and notochord induce skull, notochord and spinal cord induce vertebral column. The inducer is a very different tissue than the tissue induced. It may come as a surprise then, if I tell you that the container can induce more container. Chondroblasts and chondrocytes from a variety of skeletal sites – including ribs, long bones, vertebrae and trachea – can induce sclerotomal mesenchyme to chondrify. But the cartilage cells have to be at a particular stage in their differentiation. Cooper (1965) established this relationship by culturing mouse or chick somites transfilter to cartilage (Table 42.6). Only chondrocytes that were enlarging or becoming hypertrophic elicited cartilage. Thus, with mouse tissues, induced cartilage was found adjacent to flattened or enlarging cells, but not adjacent to fully hypertrophic chondrocytes (Table 42.6). Small-celled chondroblasts fail to become flattened chondroblasts in vitro and fail to induce. Murine cartilage remains terminally hypertrophic in vitro and fails to induce, while chick chondrocytes regress from their hypertrophic state, cytolyse and fail to induce. This association of onset of inductive ability with onset of chondrocyte hypertrophy and concomitant loss of inductive ability with attainment of hypertrophy is analogous to the attainment and loss of inductive ability by the notochord as its cells hypertrophy, discussed in Chapter 41. Are similar factors involved?7 It is curious indeed that the tissue to be induced (cartilage) has the ability to induce cartilage. It was therefore exciting to find that the products deposited by chondroblasts into their ECM are strikingly similar to those produced and deposited by inductively active cartilage, spinal cord and notochord. Do contents induce container (vertebral column) using the same molecules that will come to characterize the cartilaginous container? To begin to answer this question, I review briefly the nature of the ECMs produced by chondroblasts/cytes before
When considering skull and vertebral column development we can draw the generalization that the contents induce
Table 42.5 Chondrogenesis in the last four somites from chick embryos of H.H. 9–12 grafted in contact with different epitheliaa Treatment
Chondrogenesis % (N)
Isolated somites Somites ectoderm and endoderm Somites ectoderm and endoderm after trypsinization Somites trunk epithelium from two-day-old embryos Somites trunk epithelium from three-day-old embryos Somites trunk epithelium from four-day-old embryos Somites limb-bud epithelium from four-day-old embryos
0 (0/89) 21 (17/81) 10 (5/48)
a
Based on data from O’Hare (1972b).
0 (0/21) 30 (10/33) 23 (7/30) 24 (7/29)
Table 42.6 Incidence of cartilage in somites from mouse and chick embryos exposed to chondroblasts of different differentiation stages a Stage of cartilage differentiationb Mouse Small-celled chondroblasts Flattened and enlarging chondrocytes Enlarging or terminal chondrocytes Chick Small-celled chondroblasts Flattened chondrocytes Enlarging chondrocytes Terminal chondrocytes
Incidence of cartilage % (N)
0 (0/51) 91 (51/56) 0 (0/12) 0 (0/17) 93 (27/28 mouse somites) 88 (15/17 chick somites) 21 (3/14) 95 (20/21)
a Based on data from Hall (1977a). Mouse embryos were nine days of gestation, chick embryos were H.H. 12–15 and 17/18. Somites were cultured transfilter to chondroblasts. b Murine terminal chondrocytes are hypertrophic; chick terminal chondrocytes have lost their hypertrophic state.
The Search for the Magic Bullet discussing the matrices produced by notochord and spinal cord.
CHONDROCYTE EXTRACELLULAR MATRIX The production of glycosaminoglycans (GAGs), especially chondroitin sulphate (CS), is characteristic of functioning chondroblasts and chondrocytes. Onset of such synthesis, as detected by 35S autoradiography, was used for a long time to detect chondrogenic cells. 35S is taken up by prechondrogenic somitic mesoderm of chick and mammalian embryos, with maximal uptake in mesenchyme immediately adjacent to the embryonic axis. Lash and colleagues sought to determine whether GAG accumulation preceded histological differentiation of cartilage, but could not detect incorporation of 35S chondroitin sulphate earlier than they could detect metachromatic ECM.8 Somites explanted in vitro without spinal cord or notochord neither chondrify nor do they produce collagen or CS. Somites from embryos of H.H. 16 and 17 can be cultured for eight days without observing any activity of the enzymes ATP-sulphurylase or APS-kinase – both of which are involved in sulphation of chondroitin – unless spinal cord or notochord is included. With refinements in technique it became possible to detect CS before ECM could be detected histochemically. For example, H.H. 11 somites (Fig. 16.2) cultured with notochord or spinal cord produce what were then called chondroitin sulphates A and C (now chondroitin-4-sulphate and chondroitin-6-sulphate).9 With the development of thin-layer chromatographic techniques to isolate precursors of CS, it became apparent that many early embryonic tissues – somites, epidermis, spinal cord, notochord, extraembryonic membranes and mesonephros (see Box 38.2 for the latter) – metabolize glucosamine to form UDP-N-acetylgalactosamine (UDPNaGal), but in much smaller amounts than cartilage (Fig. 26.1). Two conclusions were reached: (i) many tissues
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can synthesize CS but not all tissues can accumulate it, and (ii) isolated somites can produce CS but need a stimulus to augment and stabilize the rate of deposition. Exposing somites to spinal cords or notochord does not confer upon them the ability to synthesize CS but rather enhances the rate at which already functioning synthetic pathways operate. The ‘instruction’ from the ‘inducer’ is speed-up, not start-up. Which extracellular molecules are being deposited by notochord and spinal cord when these two tissues interact with sclerotomal mesenchyme?
NOTOCHORD AND SPINAL CORD EXTRACELLULAR MATRICES The evident role played by the notochord and spinal cord in evoking chondrogenesis in vivo, and the accumulating evidence that normal constituents of cartilage ECM such as CS, collagen and hyaluronan can influence the rate of synthesis of cartilage ECM through feedback inhibition and stimulation (Hall, 1973a; and see Chapter 26), led to a search for ECM around the notochord and spinal cord, a search that demonstrated that notochord and the ventral portion of the spinal cord in chick embryos as young as H.H. 10 synthesize and accumulate GAGs into an ECM. Table 42.7 presents a summary of the major events in the formation of notochordal and spinal cordal ECMs and the changes occurring in sclerotomal mesenchyme at the same stages.
GLYCOSAMINOGLYCANS The first studies utilized autoradiographic techniques to show that GAGs can be visualized around the notochord when chondrogenesis is commencing within adjacent sclerotome. The impetus came when Johnson and Comar
Table 42.7 Chronology of vertebral chondrogenesis in the embryonic chicka H.H. stage
Age
Notochord, spinal cord
3 10 11 12 13 17
12 hours 36 hours 42 hours 47 hours 50 hours 60 hours
35
18
3 days
23
4 days
27–30 31–36 36–39
5–7 days 7–10 days 10–13 days
a
Based on data from Hall (1977a).
S in presumptive notochord Collagen, GAGs as ECM Basement membrane present
Youngest age shown to produce collagen when isolated in vitro Considerable hyaluronan around notochord. Treatment with collagenase or hyaluronidase prevents induction Notochord vacuolated, now induces 100% of cultured somites to chondrify
More CS than hyaluronan Ventral spinal cord loses inductive ability Notochord loses inductive ability (H.H. 37.5)
Sclerotome
Intact mass of ovoid cells Nest of cells breaking up Mesenchymal cells beginning to migrate Sclerotomites starting to form
Sclerotomal cells finished migration ECM around cells closes to notochord – now prechondroblasts ECM spreading into peripheral cells Chondroblasts and chondrocytes Cell death in area nearest notochord
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(1957) injected 35S into the albumen of eggs and detected uptake into notochord and primitive streak in embryos as young as H.H. 3, long before somite chondrogenesis. Notochord and spinal cord from embryos of H.H. 11 and older are surrounded by sulphated GAGs. 35S was localized over the spinal cord, notochord and immediately adjacent ECM when these regions were cultured, but Lash and his colleagues could not judge where this material was synthesized.10 Application of electron microscopy and histochemistry demonstrated sulphated GAGs (hyaluronan and CS) adjacent to the notochord at H.H. 16 and 17, respectively. By H.H. 17, there was 2.5 times more hyaluronate than CS. By H.H. 28 the ratio had reversed. Therefore, it was suggested that hyaluronan plays a role in sclerotome aggregation. In 1972, my fellow Australian Bryan Toole began what became a career-long study of hyaluronan and its relationship with CS (see Chapters 14 and 20). The ratio of hyaluronan to CS is high at H.H. 23 but low thereafter, the decline correlating with increasing levels of hyaluronidase (Table 42.8) and with formation of metachromatic ECM in perinotochordal mesenchyme; removal of hyaluronan accompanies formation of ECM.11 Further application of electron microscopy laid the basis for our understanding of the organization of the ECM, demonstrating: ●
●
●
amorphous material (GAGs) on microfibrils around the notochords of two- to four-day-old embryonic chicks; the importance of the metachromatic ECM between sclerotome and spinal cord and around the notochord, with the interpretation that these matrix products progressively become the ECM of perinotochordal cartilage (Box 4.4); and the accumulation of 200–400-Å GAG granules along with 150-Å unbanded collagen fibrils around the notochord and the ventral portion of the spinal cord at H.H. 10; recall that ventral but not dorsal spinal cord is inductively active. By H.H. 17, this ECM was prominent and a basement membrane surrounded notochord and spinal cord, a matrix maturation preceding the arrival of migrating sclerotomal mesenchyme around the notochord at H.H. 18.12
Both ECM and basement membrane are lost following trypsinization to isolate somites. The finding that products Table 42.8 Ratios of hyaluronan to CS in somites isolated from chick embryos and maintained in vitroa H.H. stage
Ratio of hyaluronan to CS
23 26 28 30
1.92 0.60 0.38 0.16
a Based on data from Toole (1972). Somite numbers 20–32 were isolated from embryos of H.H. 23–30.
of the ECM reform when notochord or spinal cord is maintained in vitro provided evidence for the production of the ECM by these tissues. Analysis of enzymatic digests of products from cultures of isolated notochords and neural tubes reveals mostly chondroitin and heparan sulphates, including shared CS–protein complexes.13 In a series of insightful papers, Nagaswamisri Vasan showed that perinotochordal ECM from chick embryos contains small proteoglycans and cartilage-type proteoglycans, and that only the large aggregated forms are required for notochord to induce somitic cartilage. Unstimulated somites – i.e. somites not exposed to notochord – do not contain any link protein for proteoglycan. Adding notochord to somite cultures activates the synthesis of link protein and stabilizes the ECM. On the other hand neither hyaluronan nor hyaluronidase were affected by adding notochord to somite cultures. Proteoglycan monomers from sternal cartilage also evoke chondrogenesis from somitic mesoderm, further implicating ECM components in the inductive interaction.14 Collagen was also being studied. Indeed, these studies were among the first to demonstrate a family of collagen molecules and tissue-specific collagen types. Collagens Until the late 1950s, and in large part as a consequence of the methodologies available, collagen had been demonstrated only in mesenchymal tissues. Consequently, collagen was regarded as a mesenchymal protein. From the early 1960s onwards, there was a suggestion that epithelial structures such as notochord and spinal cord might have the ability to synthesize and export collagen. The initial findings came from ultrastructural studies, as TEM became an important tool revealing cellular organization. The ECM of the notochord in embryonic chicks and mice was described as containing perinotochordal fibrils, some of which were thought to be derivatives of sclerotomal cells, some from the notochord sheath. Then came a series of studies by Low and colleagues describing microfilaments on the notochord and between notochord and spinal cord at H.H. 11. Amorphous material – interpreted as GAG – is attached to these microfibrils, which are of two types: 150–200-Å diameter, unbanded and beaded fibrils close to the notochord and sensitive to removal by collagenase, and 100-Å tubular banded fibrils some distance from the notochord, sensitive to removal by hyaluronidase and amylase but not by collagenase.15 Georges Strudel’s group described collagen fibrils in the ECMs of spinal cord in chick embryos. Because these fibrils were sensitive to collagenase and coincided in position with localized uptake of 3H-proline as measured autoradiographically, Strudel suggested that they are incorporated into the ECM of differentiating vertebral cartilage.16 Direct evidence that collagen around the spinal cord is produced by the spinal cord was obtained by Cohen and Hay (1971), who showed that: (i) ventral
The Search for the Magic Bullet spinal cord from two-day-old chick embryos synthesizes collagen when cultured alone; (ii) the collagen is deposited as fibrils into the ECM; and (iii) the deposited collagen is associated with the basement membrane. Similarly, direct evidence that the collagen around the notochord is produced by the notochord was obtained by isolating notochords from two-day-old chick embryos using trypsinization to remove ECM and basement lamina and culturing the ‘naked’ notochords. After two or three days ECM microfibrils accumulated next to a reconstituted basal lamina. Culture for longer times in the presence of 100 g/ml ascorbic acid (a co-factor for collagen synthesis) demonstrated that the fibrils – some as wide as 1500 Å and laid down in sheets – were cross-striated with an axial periodicity of 510 Å, within the range for collagen. With advances in knowledge of collagen types, these perinotochordal and perispinal cordal fibrils were shown to be type II (cartilage-type) collagen as is the collagen synthesized in vitro by spinal cords from two-day-old chick embryos. Subsequently, antibodies against type II collagen were shown to bind to embryonic notochord. A decade later, notochords from chick embryos were shown to secrete type IX collagen just prior to vertebral chondrogenesis.17 The developmental significance of the discovery that notochord and spinal cord, which promote chondrogenesis in sclerotomal cells, produce the same collagen type (type II) as the tissue they induce is the possibility that type II collagen may play an inductive role in chondrogenesis. The evolutionary significance is that notochord is phylogenetically older than cartilage; notochord may have acquired the ability to synthesize and deposit type II collagen before cartilage did. Rather than saying that notochord possesses cartilage-type collagen, we should be saying that cartilage possesses notochord-type collagen.
FUNCTION OF NOTOCHORD AND SPINAL CORD MATRIX PRODUCTS An obvious question that follows from these studies is whether collagen or GAGs produced by notochord and/or spinal cord play any role in initiating chondrogenesis from sclerotomal mesenchyme; the desire to answer this question motivated the search for matricial materials in the first place. Familiar names are associated with these studies – Strudel, O’Hare, Lash and Kosher – as two experimental approaches were initiated in the early 1970s: ●
●
removal of the ECM from notochord or spinal cord following digestion with enzymes, and testing for retention of inductive activities; and adding matrix products, individually or in combinations, to the media in which otherwise unstimulated somites are cultured, and looking for chondrogenesis.
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Differentiation of sclerotomal cartilage was inhibited when Strudel cultured axial rudiments from chick embryos in the presence of 20 g collagenase to remove collagen, or 10 g testicular hyaluronidase to remove GAGs. Transferring these rudiments to a control medium allowed chondrogenesis to commence. Of course, enzymatic treatment of entire axial rudiments removes collagen and GAG from somites as well as from the other tissues. Inhibiting chondrogenesis could result from the inability of the sclerotomal cells to secrete ECM, rather than an inability of the notochord or spinal cord to induce. This difficulty in interpretation applies to other studies in which the proline analogue L-azetidinecarboxylic acid (LACA) was injected in ovo or added to culture media containing somites and axial organs, and in which secretion of ECM was retarded and myotomal differentiation favoured at the expense of chondrogenesis.18 O’Hare (1972c) surmounted such difficulties by impregnating Millipore filters with collagenase and hyaluronidase, placing embryonic somites and spinal cord onto the filters, and CAM-grafting them. Without enzyme pretreatment of the filters, 52 per cent of the grafts produced cartilage. Only 14 per cent produced cartilage on filters impregnated with enzymes. O’Hare attributed the lack of chondrogenesis to loss of the spinal-cord basement membrane and/or ECM rather than to a direct effect of the enzymes on the somites. Another approach was to treat isolated notochord with chondroitinase or testicular hyaluronidase, combine the treated notochords with intact somites, and maintain them in vitro. Cartilage did not form unless some ECM remained on the notochord. Trypsinizing notochords prevents them from inducing, but after a time in vitro, perinotochordal materials reform – including a new basement membrane, reappearance of microfibrils and uptake of 3 H-proline – and the notochords regain their ability to induce.19 Experiments in which matrix products were added to somites in Robert Kosher’s laboratory did not produce results allowing such clear-cut interpretations. Chondroitin-4- and chondroitin-6-sulphates extracted from vertebrae were added to the media in which somites were cultured. Although the percentage of cultures forming cartilage was the same in treated and control media, accumulation of GAG dropped off after two days in control medium but was maintained at the high initial rate in treated media. Similar results for maintenance of collagen synthesis were obtained when procollagen or collagen was added to the medium used to maintain somites; synthesis of type II collagen is maintained at a high level, indicating positive feedback from product to synthesis of product.20
KEY ROLES FOR Pax-1 AND Pax-9 Pax-1, a member of the paired box of homeobox genes, is expressed in all somites early in development, but is
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down-regulated in the five most caudal somites. Expression also is seen in the perichordal zone (Table 1.1) of the vertebral column of embryonic but not adult mice. It appears that Pax-1 plays two roles. Transient expression in condensing mesenchyme before the onset of chondrogenesis suggests an early role in sclerotomal differentiation. The second role is in the differentiation of intervertebral discs.21 Pax-9 plays an even more ancient role, a role that was uncovered from studies on the Japanese lamprey, Entosphenus (Lampetra) japonicus, in which Pax-9 is expressed in the endodermal pharyngeal pouches, mesenchyme of the velum (but not the velar muscle), hyoid arch and nasohypophyseal plate, but not in somitic mesoderm; involvement of Pax-9 in somitic chondrogenesis is a feature of jawed not jawless vertebrates. Ogasawara et al. (2000) used these expression patterns to argue that Pax-9 tracks neural crest-derived mesenchyme in the velum (which first arises from a premandibular segment) and that, as a consequence, the agnathan velum is homologous with the gnathostome jaw. If correct, this homology has important consequences for our views on the origin of gnathostome jaws. Reinforcing this possibility, Pax-9-deficient mice lack the angular and coronoid processes of the dentary, have supernumerary preaxial digits, are missing all teeth, which are arrested at the bud stage, and fail to develop derivatives of the pharyngeal arches such as the thymus, pituitary and ultimobranchial glands (Peters et al., 1998). Pax-1 and Pax-9 – a paired box-containing gene closely related to Pax-1 – act synergistically in vertebral development. Pax-9 is expressed in the vertebral column, pharyngeal pouches, tail, head and limbs; the mutant Danforth’s short tail involves loss of caudal expression of Pax-9 and therefore loss of the notochord inducer. Pax-1 is involved in ventralizing the sclerotome; loss of Pax-1 is associated with sclerotomal and vertebral anomalies. Similarly, Hox group 3 paralogous genes (Hoxa-3, Hoxb-3 and Hoxd-3) act synergistically to alter neural tissues, and neural-crest and somitic mesenchyme. Surprisingly, the identity of specific Hox genes is less critical than the number of genes expressed in a particular region or tissue. An especially nice example is that redundancy among Hox-10 and Hox-11 paralogues is so great that transformatioin of vertebral identity from sacral → lumbar or lumbar → thoracic does not take place in mice that are mutant for five of the six alleles, all six having to be knocked out to effect the transformation.22 Defects seen in Pax-1 mutant mice are not seen in Pax-9 mutants. Double mutants (Pax-1 /Pax-9 ) lack vertebral bodies, intervertebral discs and the proximal portion of the ribs (see Box 16.2 for genes that influence rib development). Neural arches are normal. The chondrocyte lineage is induced to produce a loose mesenchyme rather than segmental elements. Sox-9 and collagen type II are down-regulated in this loose mesenchyme, which shows a low rate of proliferation but is not maintained,
undergoing apoptosis. Misexpressing Sox-9 in the dermomyotome leads to production of type II collagen and Pax-1 by dermomyotomal cells, which switch to a chondrogenic fate.23 In a substantive study of mouse sclerotome development, Furumoto et al. (1999) demonstrated that Pax-1 and Mfh-1 – both of which depend on Shh from the notochord for their expression – act synergistically to control vertebral column development. Double mutants (Pax-1 / Mfh-1 ) lack the dorso-medial elements of the vertebrae and consequently fail to form either vertebral bodies or intervertebral discs. Both genes regulate somitogenesis at the proliferation (condensation) stage.24 Shh-null mice have normal molecular markers in the three portions of the somites. Marcelle et al. (1999) therefore separated the paraxial mesoderm from the axial structures and examined the role of Shh in expression of Pax-1, Myo-D and Pax-3. Pax-1 is rescued by Shh, Myo-D is maintained but not induced by Shh, while Pax-3 is expressed independently of Shh. Thus, Shh is a potent mitogen for somitic cells. Murtaugh et al. (1999) showed that Shh has several actions on somitic precursor cells in chick embryos. Presomitic mesodermal cells respond to Shh by initiating chondrogenesis, the later stages of their response depending on Bmp signaling. Shh enhances the response of somitic cells to Bmps, suggesting at least two phases. Finally, Shh enhances precursor cells competent to respond to Bmp by initiating chondrogenesis. Msx-2 and notochordal influences are differentially expressed in dorsal and neural-arch development in chick embryos. Bmps are also involved; there is parallel expression of Bmp-4 and Msx-1 and Msx-2 in the lateral neural plate and then in the dorsal neural tube and midline ectoderm in chick embryos. These are locations from which signaling could be transferred to the sclerotome. Grafting Bmp-4 or Bmp-2 into the neural tube up-regulates Msx-1 and Msx-2 in the adjacent mesenchyme, resulting in production of ectopic cartilage in the pectoral girdle that develops. Grafting Bmp-4 or Bmp-2 lateral to the neural tube down-regulates Pax-1 and Pax-3. Subsequently, Shh was shown to antagonize Bmp-4 and Msx during vertebral development.25 The paired type homeodomain transcription factor Uncx4.1, which acts upstream of Pax-9, is required before neural arches can form from lateral sclerotomal cells. Mice homozygous for a targeted mutation in Uncx4.1 die perinatally with severe malformations of the axial skeleton – all lateral sclerotomal derivatives (neural arches) fail to form because chondrogenesis is inhibited. The defect is early – anlagen form but fail to condense – further reinforcing the roles of these genes at the condensation stage of vertebral chondrogenesis.26 By using morpholino antisense oligonucleotides against the transcription factor twist, Yasutake et al. (2004) showed that twist is required for neural arches to form in embryos of the Japanese medaka, Oryzias latipes, because of involvement of twist after sclerotomal-cell migration.
The Search for the Magic Bullet
CONCLUSIONS The major events occurring in sclerotome, ventral spinal cord and notochord during chick development discussed in Chapters 41 and 42 are summarized in Table 42.7 and below. Sclerotomal mesenchyme has an inherent potential for chondrogenesis. Nevertheless, expression of chondrogenic potential is exquisitely sensitive to the environment. Depending on the nature of supplements added in vitro, sclerotomal mesenchyme from embryonic chicks as young as H.H. 10 (36 hours of incubation) can chondrify. Grafting somites to a vascularized environment such as the CAM permits somites from even younger embryos to chondrify, presumably not because of vascularization per se, but because of exposure to molecules in the circulation. The presence of the synthetic machinery for, and the synthesis of CS – attributes previously thought to apply only to induced mesoderm – are shared with many nonchondrogenic tissues. What is special about sclerotomal mesenchyme and its interaction with notochord and spinal cord is the ability to augment the rate of synthesis of CS and other GAGs, and to deposit these products into an ECM. Control is not at the level of presence or absence of the synthetic pathways, but in regulating those pathways. Notochord and spinal cord provide the major in-vivo environmental factors that augment somitic chondrogenesis. Notochord and spinal cord are also the most potent agents active in vitro. Whilst other factors can substitute in vitro, their absence is not compensated for in vivo. Collagen and CS enhance and maintain chondrogenesis by positive feedback to matrix products, chondrogenesis having been initiated at the condensation stage through the action of such regulatory genes as Pax-1 and Pax-9. Normal morphogenesis of the neural arches and centra also depends on influences from spinal cord, notochord and spinal ganglia. Ablating these tissues provides the evidence for this conclusion. The underlying mechanisms await resolution, although Msx and Bmps are possible players. If epidermal ectoderm plays a role in vivo, it is subordinate to and dependent upon notochord and/or spinal cord.
NOTES 1. See Watterson (1952) for the studies on Fundulus, Detwiler and Holtzer (1956) and Strudel and Gateau (1971) for abnormalities in urodele spinal cord and chick notochord, Lash (1968b) for maintenance of cartilage by notochordal influences, and Roy et al. (2004) and Witten et al. (2005) for vertebral anomalies in aquacultured fish. 2. See Sadler et al. (2001) for triploid Atlantic salmon, and Kacem et al. (2004) for triploid rainbow trout. 3. See Cooper (1965) for the correlation with hypertrophy, and Bancroft and Bellairs (1976) for associated ultrastructural features.
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4. See Holtzer and Detwiler (1953) for the urodele study, Strudel (1953b) and Watterson et al. (1954) for secondary neural arches, and O’Hare (1972c) for the CAM-grafts. Culturing myotome plus notochord and spinal cord from chick embryos of H.H. 18 did not result in chondrogenesis (Cooper, 1965). It may be, however, that by H.H. 18, the myotome is fixed with respect to ability to form muscle and cannot switch. 5. See Lash (1963a) for a review of the first decade of the search (1953–1963), Hall (1977a) for the research to 1977, and Monsoro-Burq and Le Douarin (2000) and Stockdale et al. (2000) for more recent approaches. 6. See Strudel (1953b, 1962, 1962). 7. Epithelial otic vesicles from mouse and chick embryos are amongst the other tissues that have been tested for an ability to induce sclerotomal mesenchyme and found not to induce (Grobstein and Holtzer, 1955; Strudel, 1955, 1962; Benoit, 1960b; O’Hare, 1972b). See Benoit (1955, 1960a,c, 1964) and Hall (1991b) for the ability of the otic capsule to form in ectopic sites. 8. See Amprino (1955a) and Johnson and Comar (1957) for 35S uptake into somatic mesoderm, and Lash et al. (1960) for uptake and CS. Lash (1963a) showed that CS is sensitive to RNA’ase digestion. 9. See Glick et al. (1964) for production of ATP-sulphurylase and APS-kinase, Franco-Browder et al. (1963) for production of chondroitin-4-sulphate and chondroitin-6-sulphate, and Okayama et al. (1976) for a discussion of early determination for chondrogenesis. 10. See Franco-Browder et al. (1963) for the studies with H.H. 11 embryos, and Lash (1963) and Lash et al. (1964) for 35S uptake. 11. See O’Connell and Low (1970) for ultrastructure, Kvist and Finnegan (1970a,b) for histochemistry, Toole (1972) for HA:CS, and Goldberg and Toole (1984) for hyaluronan as part of the pericellular coat. 12. See Frederickson and Low (1971), Strudel (1971) and Corsin (1974) for the amorphous material and importance of the ECM, and Minor (1973) and Bancroft and Bellairs (1976) for the GAG granules and collagen fibrils. 13. See Hay and Meier (1974) for separation of chondroitin and heparin sulphates, and Mathews (1971, 1975) for the similarity of CS–protein complexes. 14. See Vasan (1981, 1983, 1987) for proteoglycan accumulation, Vasan et al. (1986a,b) for core protein, hyaluronan and hyaluronidase, and Vasan and Miller (1985) for sternal cartilage monomers. 15. See Jurand (1962, 1974), O’Connell and Low (1970), Frederickson and Low (1971), Minor (1973) and Bancroft and Bellairs (1976) for perinotochordal fibrils. 16. See Bazin and Strudel (1972, 1973) and Ruggeri (1972) for independent autoradiographic studies. 17. See Carlson et al. (1974) and Carlson and Upson (1974). Hydroxylation of proline and lysine is inhibited in ascorbic acid (vitamin C) deficiency. Consequently, biosynthesis of collagen and elastin is impaired (Barnes et al., 1969; Wu et al., 1989). See Linsenmayer et al. (1973a) and Trelstad et al. (1973) for notochord and spinal-cord type II collagen, von der Mark et al. (1976) and von der Mark (1980) for the work with type II antibodies, and Hayashi et al. (1992) for notochordal secretion of type X collagen. 18. See Strudel (1972, 1973a–c) for the enzyme studies and Strudel (1975a–c) for the studies with LACA (which inhibits
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19.
20.
21.
22.
Bones and Cartilage: Developmental and Evolutionary Skeletal Biology
excretion of collagen and consequently inhibits growth and delays chondrogenesis, for which see Aydelotte and Kochhar, 1972 and Hall, 1978c). See Kosher and Lash (1975) and Lauscher and Carlson (1975) for studies with chondroitinase and hyaluronidase. Mice immunized with chondroitinase ABC, proteoglycan and adjuvant develop polyarthritis, an ankylosing spondylitis (Glant et al., 1987). See Kosher et al. (1973) for the studies with CSs, and Kosher and Church (1975) and Kosher and Savage (1979) for the studies with procollagen and collagen. See Deutsch et al. (1988) for Pax-1 expression, Wallin et al. (1994) for early and late roles in chondrogenesis, and Wilting et al. (1995) for caudal expression. Wilting and colleagues also used the boundary of expression to determine the head/trunk (cervical/occipital) boundary in Japanese quail and mice. See Manley and Capecchi (1997) for the concept of redundancy among Hox genes and see Wellik and Capecchi (2003) for the Hox-10/Hox-11 example.
23. See Peters et al. (1998, 1999) for Pax-1 and Pax-9, Neubüser et al. (1995) for the distribution of Pax-9 and Danforth’s short tail, and Balling et al. (1996) for Pax-1 knockout. Pax-1 and Pax-9 are also expressed in complex, non-overlapping patterns in chick limb mesenchyme, especially at the condensation stage, when expression is strongest (Le Clair et al., 1999). See Healy et al. (1999) for expression of Pax-1 in the dermomyotome. 24. The winged helix transcription factor, MFH-1, is expressed in mouse cranial neural crest and head mesoderm and then in cartilage. MFH-1 mice die as embryos or at birth (Iida et al., 1997). 25. See Monsoro-Burq et al. (1994, 1996), Liem et al. (1995), Pourquié et al. (1996), Watanabe and Le Douarin (1996) and Watanabe et al. (1998a). Tgf-1 enhances chondrogenesis from chick sclerotomal micromass cultures (Sanders et al., 1993), probably because of a generic proliferative effect. 26. See Leitges et al. (2000) and Mansouri et al. (2000) for the studies with Uncx4.1.
The Search for the Magic Bullet
The message is ‘speed up, not start up.’
Although neither notochord nor spinal cord provides information to sclerotomal mesenchyme that is essential for chondrogenesis to commence in vitro, both tissues provide important and necessary information in vivo. While the inducer can be ‘bypassed’ or ‘mimicked’ in vitro by conditioned medium, embryos cannot compensate for the absence of spinal cord or notochord. Rather, vertebral differentiation and/or morphogenesis will be disrupted. Consequently, I think we are justified in continuing to refer to spinal cord and notochord as inducers of sclerotomal chondrogenesis and to their interaction with sclerotome as inductive. In this Chapter I explore what we know of the inducers and of their mechanisms of action on vertebral chondrogenesis and morphogenesis.
INTEGRITY OF NOTOCHORD/SPINAL CORD AND VERTEBRAL MORPHOGENESIS As discussed at the beginning of Chapter 41, intact spinal cord, notochord and spinal ganglia influence vertebral morphogenesis. But must notochord and spinal cord be intact to act? What type of registration, if any, exists between the axial tissues and vertebral segmentation? Normal neural arches develop in the presence of disorganized neural tissue in the killifish, Fundulus heteroclitus. In urodele amphibians and birds, on the other hand, abnormalities of the spinal cord lead to vertebral anomalies; administering nicotine sulphide to twoday-old chick embryos results in twisting of the notochord in the neck within hours and vertebral deformities within days. An interesting difference in the integrity of cartilage appears to depend on whether induction is by spinal cord
42
or notochord. Cartilage induced by spinal cord dissipates between the 9th and 16th days in culture. Notochordinduced cartilage persists until at least the 25th day. Lash extrapolated this result to suggest that the spinal cord helps to remodel the vertebral column as it grows in vivo, thereby preventing compression of the spinal cord, an interesting idea that should be explored further, especially in relation to vertebral anomalies in fish raised under aquaculture conditions.1 Fish skeletal defects Laboratory rearing, like domestication (Box 7.2), influences variation; there is greater variation in all parts of the vertebral column in laboratory-reared than in wild-caught Red Sea bream, Pagrus major (Matsuoka, 1982). Defects represent variation outside that expected for the species. Skeletal defects in hatchery-reared fish are sufficiently common to be an accepted cost of aquaculture and a major concern for aquaculturalists. Depending on species and hatchery conditions, as many as 50 per cent of the fish will have vertebral defects. In one study, 44 per cent of hatchery-reared Senegal sole, Solea senegalensis, showed vertebral abnormalities, 28 per cent of which affected the tail and vertebral column. Although clues to the aetiology of some defects can be obtained from teratological studies – wavy notochords and bent gill arches result when retinoic acid or collagen maturation is blocked in flounder embryos (Suzuki et al., 2001) – determining the origins of vertebral defects has been an intractable problem. From studies on Atlantic salmon, Salmo salar, cultured in Tasmania, Australia, we know that gender has no influence but that chromosome number (ploidy) significantly affects the incidence of deformities. For example, only
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1 per cent of diploid freshwater smolts have lower jaw deformities. By contrast, the incidence of deformities in triploid fry, freshwater and seawater smolts is 2, 7 and 14 per cent, respectively. Similarly, primary gill filaments fail to form in up to 60 per cent of triploid individuals but only in 4 per cent of diploids. Triploidy provides the clue. The underlying mechanism remains elusive. Triploid rainbow trout, Oncorhynchus mykiss, were found to develop an additional caudal vertebra than found in diploid individuals, No other skeletal changes were found.2 Development of ‘short-tails’ is a common problem in farmed Atlantic salmon. These are fish in which the vertebral column is compressed along the A-P axis to such an extent that body length is reduced and body depth so enlarged that the body form hardly resembles that of a salmon (see Figure 1 in Witten et al., 2005). The defect at the cellular level involves the metaplasia of intervertebral tissues to cartilage, essentially replacing the notochord with cartilage (Witten et al., 2005).
Table 42.1 Inductive ability of notochord and spinal cord from chick and mice on somites of 9-day-old mice and H.H. 15, 17 or 18 chick embryosa H.H. stage or age in days of notochord/ spinal cord donor
Stage 18 Stages 27–37.5
Cartilage induced in transfilter recombinations (%)
Cartilage induced in direct recombination (%)
Chick–chick chimaeras – notochord 100 – 100 100
Stages 21–24 Stages 12–15
Chick–mouse chimaeras – notochord 100 100 83 –
5.5 daysb 7.5 daysb 5.5–6.5 daysc
Chick–mouse chimaeras – spinal cord 100 – 100 – 0 –
9 days
Mouse–mouse chimaeras – notochord 83 –
9 daysb
Mouse–mouse chimaeras – spinal cord 87.5 100
a
FOR HOW LONG ARE NOTOCHORD AND SPINAL CORD ACTIVE? In many inductive interactions the inducing cells lose their ability to induce at the same time as the responding cells lose their ability to respond, a mutually convenient temporal association that assures that induction will not be prolonged. As the spinal cord and notochord appear to play specific roles in inducing somitic cartilage, it is of interest to know whether they lose their inductive ability. I can summarize by stating that, as late as has been tested, notochords retain inductive activity but spinal cords do not. The evidence comes from combining notochord or spinal cord from older embryos or neonates with somites from younger embryos to determine whether cartilage forms. Some typical studies are summarized below and in Tables 36.1–36.3. Cooper (1965) used chick–chick and chick–mouse chimaeras to show that notochord from embryos as old as H.H. 37 (11.5 days) can induce chick or mouse somites to form cartilage (Fig. 37.1 and Table 42.1). Notochords from older chick embryos were not tested but notochord from 19-day-old mice remains inductively active (Fig. 37.1). The spinal cord does not retain inductive ability for the same period, which may reflect, in part, earlier maturation of spinal cord in comparison with notochord. Spinal cords from mouse embryos older than nine days of gestation no longer induce (Tables 42.1 and 42.2). We found that spinal cord from chick embryos as old as H.H. 44 (18 days of incubation) allow H.H. 17 somites to chondrify in vitro under conditions wherein somites alone fail to chondrify (Tremaine and Hall, 1979). Ability or inability to induce correlates with state of differentiation. Notochord (and cartilage; Chapter 22) only induces when the cells are undergoing hypertrophy or vacuolation (Table 42.3).3
Based on data from Cooper (1965). Ventral half of spinal cord. c Dorsal half of spinal cord. b
Table 42.2 Relative amount of cartilage in somites cultured with spinal cord from mice of different agesa Age of spinal cord (days)
Relative % of cartilage/culture
9 11 12 and 13
100 80 25
a
Based on data from Grobstein and Holtzer (1955).
CAN DERMOMYOTOME OR LATERAL-PLATE MESODERM CHONDRIFY? During embryonic development the sclerotome chondrifies, the dermotome produces connective tissue, and the myotome forms muscle (Fig. 16.1). Can dermotome or myotome form cartilage if suitably challenged? An unstated assumption underlying such a question is that distance from the normal inducers is all that prevents dermotome and myotome from chondrifying. Three major types of experiments have been designed to answer this question. Although muscle development is delayed, cartilage differentiation proceeds normally after 75 per cent of the medial aspect of the somites is removed from tail budstage urodele embryos. Apparently, areas of the somites normally destined for connective tissue and muscle regulate (Box 35.1) and chondrify. Implanting spinal cord or notochord into the dermomyotome of embryonic chicks elicits a secondary (ectopic) neural arch as chondrogenic differentiation and morphogenesis are initiated in non-sclerotomal mesenchyme.
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The Search for the Magic Bullet Table 42.3 Cartilage-inducing activity of embryonic chick and mouse notochord at four developmental stagesa Stage of developing notochord
H.H. stage/age of embryo
Number of cultures tested in notochord stage
Cartilageinducing activity
1 Aggregating chordamesoblast cells
Chick H.H. 12–15 Mice 9 days Chick H.H. 18 Chick H.H. 21–24 Chick H.H. 27–37.5 Chick H.H. 37.5 vertebral canal
2 3 20 18 77 2
None None Cartilage induced Cartilage induced Cartilage induced None
2 Intercellular vacuolation 3 Cellular hypertrophy 4 Non-hypertrophied state
a
Based on data from Cooper (1965). Notochord and somites were either recombined directly or transfilter and maintained in vitro.
Table 42.4 Chondrification in lateral-plate mesoderm from chick embryos of H.H. 9–12 grafted to the CAMa Grafted tissue
Chondrification (%)
Lateral-plate mesoderm Lateral-plate mesoderm ectoderm and endoderm Lateral-plate mesoderm spinal cord
4 37
a
42
Based on data from O’Hare (1972c).
Cartilage forms in CAM-grafted lateral-plate mesoderm adjacent to the last four pairs of somites from embryos of H.H. 9–12, provided that ectoderm/endoderm or spinal cord is included in the grafts (Table 42.4). This result removes an interpretation applied to some earlier experiments – that transplanted spinal cord or notochord did not induce lateral mesoderm, but attracted sclerotomal cells into the lateral mesoderm where they formed the cartilage.4
THE SEARCH FOR THE MAGIC BULLET The search for a pure inducer that pervaded so many areas of embryology and developmental biology throughout the 20th century also directed, channeled, perhaps even hamstrung, students of somite chondrogenesis. For a brief period, beginning with a study by George Strudel in 1953, it looked as if progress was being made, but no significant advances were made in the two or three decades that followed.5 Strudel found that a saline extract of spinal cords or notochords from H.H. 18–21 chick embryos could induce somitic chondrogenesis in vitro.6 A decade later, he extended this finding back to earlier stages, concluding: …even very young somite cells are genetically determined cells and all that they need to undergo or accomplish their phenotypic differentiation is a microenvironment favouring chondrogenesis. This does not mean that the inducing action of the spinal cord and the notochord is dispensable. It may be that the spinal cord and/or the notochord exercise their inducing effect very early. (pers. comm.)
In 1957, Jay Lash and his colleagues introduced an ingenious experimental procedure adopted by a number of us. Embryonic spinal cords were placed transfilter to somites in vitro for 10 hours, during which time the spinal cord deposited extracellular material onto and into the filter. The spinal cords were removed and somites then placed onto the matrix products deposited into the filter and allowed to develop. Three days later, cartilage differentiated. ‘Control’ somites cultured on filters without ECM products formed no cartilage. Lash and colleagues concluded that only a short period of contact between somite and inducing spinal cord is needed to transfer a chemical signal between spinal cord and somites (Lash et al., 1957). A role for ectoderm? In studies carried out in the 1920s to 1940s ectoderm and possibly also subjacent endoderm were often included with grafted somitic mesoderm, either deliberately or inadvertently (see Chapter 41). Potential problems associated with ectodermal contamination were taken up as researchers asked whether ectoderm adjacent to the somites plays any role in cartilage induction. The results are contradictory and the final story is not in even now. We can say that epithelia exert no direct inductive action in vivo; chondrogenesis is not initiated when notochord or spinal cord are extirpated but dorsal epidermis is left intact. If epithelia play a role, it is secondary to and dependent on notochord and/or spinal cord. A sample of the results and conclusions include the following. ●
●
In somites from embryos of H.H. 10–13 grafted with or without ectoderm or endoderm, cartilage forms only when an epithelium is present. Seventy-five per cent form cartilage when ecto- and endoderm are present, 57 per cent when only ectoderm, and 14 per cent when only endoderm is present. Seno and Büyüközer (1958) concluded that epithelia, especially ectoderm, provide mechanical support for sclerotomal mesenchyme, thus preventing cells from dispersing. They ruled out a more physiological role. Stockdale et al. (1961) wrapped ectoderm around pellets of somitic cells equivalent in size to two to four somites and cultured them. Cartilage failed to develop, a result
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Bones and Cartilage: Developmental and Evolutionary Skeletal Biology
that does not necessarily rule out a role for ectoderm acting on intact somites in ovo. Similarly, no cartilage formed in any of 90 cultures of strips or clusters of somite wrapped in ectoderm or endoderm and either cultured or grafted into the coelomic cavities of host embryos (Lash, 1963b). Among the tissues Holtzer (1964b) tested for inductive ability, epidermis (source and age not specified) would not induce H.H. 18 somites to chondrify in vitro.
The British came to the rescue with an extensive (and definitive?) series of experiments by M. J. O’Hare, published in three parts in 1972. O’Hare grafted the last four somites from embryos of H.H. 9–12 after associating them with one of a variety of epithelia. As you can see from Table 42.5, somites grafted alone fail to chondrify, but a variety of epithelia permit chondrogenesis in anywhere from 10 to 30 per cent of the grafts. O’Hare also showed that the ability to augment chondrogenesis was not restricted to epithelia adjacent to somitic mesoderm but also was present in limb bud and trunk epithelia from older embryos (Table 42.5). He explained these temporal differences as based on a correlation with presence of a basement membrane and associated extracellular matrix (ECM) in ‘active’ epithelia. Such temporal differences argue against a strict mechanical effect, as does the ability of lateral-plate mesoderm to chondrify when associated with ectoderm (Table 42.5). The tentative conclusion from these studies is that epithelia allow somites to chondrify in the absence of notochord or spinal cord, if somites are CAM-grafted but not if cultured. Conclusions relating to any role for epithelia in vivo must be confined to the statement that epithelia cannot act in the absence of spinal cord or notochord. Cartilage cells as cartilage inducers
the container – brain and notochord induce skull, notochord and spinal cord induce vertebral column. The inducer is a very different tissue than the tissue induced. It may come as a surprise then, if I tell you that the container can induce more container. Chondroblasts and chondrocytes from a variety of skeletal sites – including ribs, long bones, vertebrae and trachea – can induce sclerotomal mesenchyme to chondrify. But the cartilage cells have to be at a particular stage in their differentiation. Cooper (1965) established this relationship by culturing mouse or chick somites transfilter to cartilage (Table 42.6). Only chondrocytes that were enlarging or becoming hypertrophic elicited cartilage. Thus, with mouse tissues, induced cartilage was found adjacent to flattened or enlarging cells, but not adjacent to fully hypertrophic chondrocytes (Table 42.6). Small-celled chondroblasts fail to become flattened chondroblasts in vitro and fail to induce. Murine cartilage remains terminally hypertrophic in vitro and fails to induce, while chick chondrocytes regress from their hypertrophic state, cytolyse and fail to induce. This association of onset of inductive ability with onset of chondrocyte hypertrophy and concomitant loss of inductive ability with attainment of hypertrophy is analogous to the attainment and loss of inductive ability by the notochord as its cells hypertrophy, discussed in Chapter 41. Are similar factors involved?7 It is curious indeed that the tissue to be induced (cartilage) has the ability to induce cartilage. It was therefore exciting to find that the products deposited by chondroblasts into their ECM are strikingly similar to those produced and deposited by inductively active cartilage, spinal cord and notochord. Do contents induce container (vertebral column) using the same molecules that will come to characterize the cartilaginous container? To begin to answer this question, I review briefly the nature of the ECMs produced by chondroblasts/cytes before
When considering skull and vertebral column development we can draw the generalization that the contents induce
Table 42.5 Chondrogenesis in the last four somites from chick embryos of H.H. 9–12 grafted in contact with different epitheliaa Treatment
Chondrogenesis % (N)
Isolated somites Somites ectoderm and endoderm Somites ectoderm and endoderm after trypsinization Somites trunk epithelium from two-day-old embryos Somites trunk epithelium from three-day-old embryos Somites trunk epithelium from four-day-old embryos Somites limb-bud epithelium from four-day-old embryos
0 (0/89) 21 (17/81) 10 (5/48)
a
Based on data from O’Hare (1972b).
0 (0/21) 30 (10/33) 23 (7/30) 24 (7/29)
Table 42.6 Incidence of cartilage in somites from mouse and chick embryos exposed to chondroblasts of different differentiation stages a Stage of cartilage differentiationb Mouse Small-celled chondroblasts Flattened and enlarging chondrocytes Enlarging or terminal chondrocytes Chick Small-celled chondroblasts Flattened chondrocytes Enlarging chondrocytes Terminal chondrocytes
Incidence of cartilage % (N)
0 (0/51) 91 (51/56) 0 (0/12) 0 (0/17) 93 (27/28 mouse somites) 88 (15/17 chick somites) 21 (3/14) 95 (20/21)
a Based on data from Hall (1977a). Mouse embryos were nine days of gestation, chick embryos were H.H. 12–15 and 17/18. Somites were cultured transfilter to chondroblasts. b Murine terminal chondrocytes are hypertrophic; chick terminal chondrocytes have lost their hypertrophic state.
The Search for the Magic Bullet discussing the matrices produced by notochord and spinal cord.
CHONDROCYTE EXTRACELLULAR MATRIX The production of glycosaminoglycans (GAGs), especially chondroitin sulphate (CS), is characteristic of functioning chondroblasts and chondrocytes. Onset of such synthesis, as detected by 35S autoradiography, was used for a long time to detect chondrogenic cells. 35S is taken up by prechondrogenic somitic mesoderm of chick and mammalian embryos, with maximal uptake in mesenchyme immediately adjacent to the embryonic axis. Lash and colleagues sought to determine whether GAG accumulation preceded histological differentiation of cartilage, but could not detect incorporation of 35S chondroitin sulphate earlier than they could detect metachromatic ECM.8 Somites explanted in vitro without spinal cord or notochord neither chondrify nor do they produce collagen or CS. Somites from embryos of H.H. 16 and 17 can be cultured for eight days without observing any activity of the enzymes ATP-sulphurylase or APS-kinase – both of which are involved in sulphation of chondroitin – unless spinal cord or notochord is included. With refinements in technique it became possible to detect CS before ECM could be detected histochemically. For example, H.H. 11 somites (Fig. 16.2) cultured with notochord or spinal cord produce what were then called chondroitin sulphates A and C (now chondroitin-4-sulphate and chondroitin-6-sulphate).9 With the development of thin-layer chromatographic techniques to isolate precursors of CS, it became apparent that many early embryonic tissues – somites, epidermis, spinal cord, notochord, extraembryonic membranes and mesonephros (see Box 38.2 for the latter) – metabolize glucosamine to form UDP-N-acetylgalactosamine (UDPNaGal), but in much smaller amounts than cartilage (Fig. 26.1). Two conclusions were reached: (i) many tissues
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can synthesize CS but not all tissues can accumulate it, and (ii) isolated somites can produce CS but need a stimulus to augment and stabilize the rate of deposition. Exposing somites to spinal cords or notochord does not confer upon them the ability to synthesize CS but rather enhances the rate at which already functioning synthetic pathways operate. The ‘instruction’ from the ‘inducer’ is speed-up, not start-up. Which extracellular molecules are being deposited by notochord and spinal cord when these two tissues interact with sclerotomal mesenchyme?
NOTOCHORD AND SPINAL CORD EXTRACELLULAR MATRICES The evident role played by the notochord and spinal cord in evoking chondrogenesis in vivo, and the accumulating evidence that normal constituents of cartilage ECM such as CS, collagen and hyaluronan can influence the rate of synthesis of cartilage ECM through feedback inhibition and stimulation (Hall, 1973a; and see Chapter 26), led to a search for ECM around the notochord and spinal cord, a search that demonstrated that notochord and the ventral portion of the spinal cord in chick embryos as young as H.H. 10 synthesize and accumulate GAGs into an ECM. Table 42.7 presents a summary of the major events in the formation of notochordal and spinal cordal ECMs and the changes occurring in sclerotomal mesenchyme at the same stages.
GLYCOSAMINOGLYCANS The first studies utilized autoradiographic techniques to show that GAGs can be visualized around the notochord when chondrogenesis is commencing within adjacent sclerotome. The impetus came when Johnson and Comar
Table 42.7 Chronology of vertebral chondrogenesis in the embryonic chicka H.H. stage
Age
Notochord, spinal cord
3 10 11 12 13 17
12 hours 36 hours 42 hours 47 hours 50 hours 60 hours
35
18
3 days
23
4 days
27–30 31–36 36–39
5–7 days 7–10 days 10–13 days
a
Based on data from Hall (1977a).
S in presumptive notochord Collagen, GAGs as ECM Basement membrane present
Youngest age shown to produce collagen when isolated in vitro Considerable hyaluronan around notochord. Treatment with collagenase or hyaluronidase prevents induction Notochord vacuolated, now induces 100% of cultured somites to chondrify
More CS than hyaluronan Ventral spinal cord loses inductive ability Notochord loses inductive ability (H.H. 37.5)
Sclerotome
Intact mass of ovoid cells Nest of cells breaking up Mesenchymal cells beginning to migrate Sclerotomites starting to form
Sclerotomal cells finished migration ECM around cells closes to notochord – now prechondroblasts ECM spreading into peripheral cells Chondroblasts and chondrocytes Cell death in area nearest notochord
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(1957) injected 35S into the albumen of eggs and detected uptake into notochord and primitive streak in embryos as young as H.H. 3, long before somite chondrogenesis. Notochord and spinal cord from embryos of H.H. 11 and older are surrounded by sulphated GAGs. 35S was localized over the spinal cord, notochord and immediately adjacent ECM when these regions were cultured, but Lash and his colleagues could not judge where this material was synthesized.10 Application of electron microscopy and histochemistry demonstrated sulphated GAGs (hyaluronan and CS) adjacent to the notochord at H.H. 16 and 17, respectively. By H.H. 17, there was 2.5 times more hyaluronate than CS. By H.H. 28 the ratio had reversed. Therefore, it was suggested that hyaluronan plays a role in sclerotome aggregation. In 1972, my fellow Australian Bryan Toole began what became a career-long study of hyaluronan and its relationship with CS (see Chapters 14 and 20). The ratio of hyaluronan to CS is high at H.H. 23 but low thereafter, the decline correlating with increasing levels of hyaluronidase (Table 42.8) and with formation of metachromatic ECM in perinotochordal mesenchyme; removal of hyaluronan accompanies formation of ECM.11 Further application of electron microscopy laid the basis for our understanding of the organization of the ECM, demonstrating: ●
●
●
amorphous material (GAGs) on microfibrils around the notochords of two- to four-day-old embryonic chicks; the importance of the metachromatic ECM between sclerotome and spinal cord and around the notochord, with the interpretation that these matrix products progressively become the ECM of perinotochordal cartilage (Box 4.4); and the accumulation of 200–400-Å GAG granules along with 150-Å unbanded collagen fibrils around the notochord and the ventral portion of the spinal cord at H.H. 10; recall that ventral but not dorsal spinal cord is inductively active. By H.H. 17, this ECM was prominent and a basement membrane surrounded notochord and spinal cord, a matrix maturation preceding the arrival of migrating sclerotomal mesenchyme around the notochord at H.H. 18.12
Both ECM and basement membrane are lost following trypsinization to isolate somites. The finding that products Table 42.8 Ratios of hyaluronan to CS in somites isolated from chick embryos and maintained in vitroa H.H. stage
Ratio of hyaluronan to CS
23 26 28 30
1.92 0.60 0.38 0.16
a Based on data from Toole (1972). Somite numbers 20–32 were isolated from embryos of H.H. 23–30.
of the ECM reform when notochord or spinal cord is maintained in vitro provided evidence for the production of the ECM by these tissues. Analysis of enzymatic digests of products from cultures of isolated notochords and neural tubes reveals mostly chondroitin and heparan sulphates, including shared CS–protein complexes.13 In a series of insightful papers, Nagaswamisri Vasan showed that perinotochordal ECM from chick embryos contains small proteoglycans and cartilage-type proteoglycans, and that only the large aggregated forms are required for notochord to induce somitic cartilage. Unstimulated somites – i.e. somites not exposed to notochord – do not contain any link protein for proteoglycan. Adding notochord to somite cultures activates the synthesis of link protein and stabilizes the ECM. On the other hand neither hyaluronan nor hyaluronidase were affected by adding notochord to somite cultures. Proteoglycan monomers from sternal cartilage also evoke chondrogenesis from somitic mesoderm, further implicating ECM components in the inductive interaction.14 Collagen was also being studied. Indeed, these studies were among the first to demonstrate a family of collagen molecules and tissue-specific collagen types. Collagens Until the late 1950s, and in large part as a consequence of the methodologies available, collagen had been demonstrated only in mesenchymal tissues. Consequently, collagen was regarded as a mesenchymal protein. From the early 1960s onwards, there was a suggestion that epithelial structures such as notochord and spinal cord might have the ability to synthesize and export collagen. The initial findings came from ultrastructural studies, as TEM became an important tool revealing cellular organization. The ECM of the notochord in embryonic chicks and mice was described as containing perinotochordal fibrils, some of which were thought to be derivatives of sclerotomal cells, some from the notochord sheath. Then came a series of studies by Low and colleagues describing microfilaments on the notochord and between notochord and spinal cord at H.H. 11. Amorphous material – interpreted as GAG – is attached to these microfibrils, which are of two types: 150–200-Å diameter, unbanded and beaded fibrils close to the notochord and sensitive to removal by collagenase, and 100-Å tubular banded fibrils some distance from the notochord, sensitive to removal by hyaluronidase and amylase but not by collagenase.15 Georges Strudel’s group described collagen fibrils in the ECMs of spinal cord in chick embryos. Because these fibrils were sensitive to collagenase and coincided in position with localized uptake of 3H-proline as measured autoradiographically, Strudel suggested that they are incorporated into the ECM of differentiating vertebral cartilage.16 Direct evidence that collagen around the spinal cord is produced by the spinal cord was obtained by Cohen and Hay (1971), who showed that: (i) ventral
The Search for the Magic Bullet spinal cord from two-day-old chick embryos synthesizes collagen when cultured alone; (ii) the collagen is deposited as fibrils into the ECM; and (iii) the deposited collagen is associated with the basement membrane. Similarly, direct evidence that the collagen around the notochord is produced by the notochord was obtained by isolating notochords from two-day-old chick embryos using trypsinization to remove ECM and basement lamina and culturing the ‘naked’ notochords. After two or three days ECM microfibrils accumulated next to a reconstituted basal lamina. Culture for longer times in the presence of 100 g/ml ascorbic acid (a co-factor for collagen synthesis) demonstrated that the fibrils – some as wide as 1500 Å and laid down in sheets – were cross-striated with an axial periodicity of 510 Å, within the range for collagen. With advances in knowledge of collagen types, these perinotochordal and perispinal cordal fibrils were shown to be type II (cartilage-type) collagen as is the collagen synthesized in vitro by spinal cords from two-day-old chick embryos. Subsequently, antibodies against type II collagen were shown to bind to embryonic notochord. A decade later, notochords from chick embryos were shown to secrete type IX collagen just prior to vertebral chondrogenesis.17 The developmental significance of the discovery that notochord and spinal cord, which promote chondrogenesis in sclerotomal cells, produce the same collagen type (type II) as the tissue they induce is the possibility that type II collagen may play an inductive role in chondrogenesis. The evolutionary significance is that notochord is phylogenetically older than cartilage; notochord may have acquired the ability to synthesize and deposit type II collagen before cartilage did. Rather than saying that notochord possesses cartilage-type collagen, we should be saying that cartilage possesses notochord-type collagen.
FUNCTION OF NOTOCHORD AND SPINAL CORD MATRIX PRODUCTS An obvious question that follows from these studies is whether collagen or GAGs produced by notochord and/or spinal cord play any role in initiating chondrogenesis from sclerotomal mesenchyme; the desire to answer this question motivated the search for matricial materials in the first place. Familiar names are associated with these studies – Strudel, O’Hare, Lash and Kosher – as two experimental approaches were initiated in the early 1970s: ●
●
removal of the ECM from notochord or spinal cord following digestion with enzymes, and testing for retention of inductive activities; and adding matrix products, individually or in combinations, to the media in which otherwise unstimulated somites are cultured, and looking for chondrogenesis.
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Differentiation of sclerotomal cartilage was inhibited when Strudel cultured axial rudiments from chick embryos in the presence of 20 g collagenase to remove collagen, or 10 g testicular hyaluronidase to remove GAGs. Transferring these rudiments to a control medium allowed chondrogenesis to commence. Of course, enzymatic treatment of entire axial rudiments removes collagen and GAG from somites as well as from the other tissues. Inhibiting chondrogenesis could result from the inability of the sclerotomal cells to secrete ECM, rather than an inability of the notochord or spinal cord to induce. This difficulty in interpretation applies to other studies in which the proline analogue L-azetidinecarboxylic acid (LACA) was injected in ovo or added to culture media containing somites and axial organs, and in which secretion of ECM was retarded and myotomal differentiation favoured at the expense of chondrogenesis.18 O’Hare (1972c) surmounted such difficulties by impregnating Millipore filters with collagenase and hyaluronidase, placing embryonic somites and spinal cord onto the filters, and CAM-grafting them. Without enzyme pretreatment of the filters, 52 per cent of the grafts produced cartilage. Only 14 per cent produced cartilage on filters impregnated with enzymes. O’Hare attributed the lack of chondrogenesis to loss of the spinal-cord basement membrane and/or ECM rather than to a direct effect of the enzymes on the somites. Another approach was to treat isolated notochord with chondroitinase or testicular hyaluronidase, combine the treated notochords with intact somites, and maintain them in vitro. Cartilage did not form unless some ECM remained on the notochord. Trypsinizing notochords prevents them from inducing, but after a time in vitro, perinotochordal materials reform – including a new basement membrane, reappearance of microfibrils and uptake of 3 H-proline – and the notochords regain their ability to induce.19 Experiments in which matrix products were added to somites in Robert Kosher’s laboratory did not produce results allowing such clear-cut interpretations. Chondroitin-4- and chondroitin-6-sulphates extracted from vertebrae were added to the media in which somites were cultured. Although the percentage of cultures forming cartilage was the same in treated and control media, accumulation of GAG dropped off after two days in control medium but was maintained at the high initial rate in treated media. Similar results for maintenance of collagen synthesis were obtained when procollagen or collagen was added to the medium used to maintain somites; synthesis of type II collagen is maintained at a high level, indicating positive feedback from product to synthesis of product.20
KEY ROLES FOR Pax-1 AND Pax-9 Pax-1, a member of the paired box of homeobox genes, is expressed in all somites early in development, but is
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down-regulated in the five most caudal somites. Expression also is seen in the perichordal zone (Table 1.1) of the vertebral column of embryonic but not adult mice. It appears that Pax-1 plays two roles. Transient expression in condensing mesenchyme before the onset of chondrogenesis suggests an early role in sclerotomal differentiation. The second role is in the differentiation of intervertebral discs.21 Pax-9 plays an even more ancient role, a role that was uncovered from studies on the Japanese lamprey, Entosphenus (Lampetra) japonicus, in which Pax-9 is expressed in the endodermal pharyngeal pouches, mesenchyme of the velum (but not the velar muscle), hyoid arch and nasohypophyseal plate, but not in somitic mesoderm; involvement of Pax-9 in somitic chondrogenesis is a feature of jawed not jawless vertebrates. Ogasawara et al. (2000) used these expression patterns to argue that Pax-9 tracks neural crest-derived mesenchyme in the velum (which first arises from a premandibular segment) and that, as a consequence, the agnathan velum is homologous with the gnathostome jaw. If correct, this homology has important consequences for our views on the origin of gnathostome jaws. Reinforcing this possibility, Pax-9-deficient mice lack the angular and coronoid processes of the dentary, have supernumerary preaxial digits, are missing all teeth, which are arrested at the bud stage, and fail to develop derivatives of the pharyngeal arches such as the thymus, pituitary and ultimobranchial glands (Peters et al., 1998). Pax-1 and Pax-9 – a paired box-containing gene closely related to Pax-1 – act synergistically in vertebral development. Pax-9 is expressed in the vertebral column, pharyngeal pouches, tail, head and limbs; the mutant Danforth’s short tail involves loss of caudal expression of Pax-9 and therefore loss of the notochord inducer. Pax-1 is involved in ventralizing the sclerotome; loss of Pax-1 is associated with sclerotomal and vertebral anomalies. Similarly, Hox group 3 paralogous genes (Hoxa-3, Hoxb-3 and Hoxd-3) act synergistically to alter neural tissues, and neural-crest and somitic mesenchyme. Surprisingly, the identity of specific Hox genes is less critical than the number of genes expressed in a particular region or tissue. An especially nice example is that redundancy among Hox-10 and Hox-11 paralogues is so great that transformatioin of vertebral identity from sacral → lumbar or lumbar → thoracic does not take place in mice that are mutant for five of the six alleles, all six having to be knocked out to effect the transformation.22 Defects seen in Pax-1 mutant mice are not seen in Pax-9 mutants. Double mutants (Pax-1 /Pax-9 ) lack vertebral bodies, intervertebral discs and the proximal portion of the ribs (see Box 16.2 for genes that influence rib development). Neural arches are normal. The chondrocyte lineage is induced to produce a loose mesenchyme rather than segmental elements. Sox-9 and collagen type II are down-regulated in this loose mesenchyme, which shows a low rate of proliferation but is not maintained,
undergoing apoptosis. Misexpressing Sox-9 in the dermomyotome leads to production of type II collagen and Pax-1 by dermomyotomal cells, which switch to a chondrogenic fate.23 In a substantive study of mouse sclerotome development, Furumoto et al. (1999) demonstrated that Pax-1 and Mfh-1 – both of which depend on Shh from the notochord for their expression – act synergistically to control vertebral column development. Double mutants (Pax-1 / Mfh-1 ) lack the dorso-medial elements of the vertebrae and consequently fail to form either vertebral bodies or intervertebral discs. Both genes regulate somitogenesis at the proliferation (condensation) stage.24 Shh-null mice have normal molecular markers in the three portions of the somites. Marcelle et al. (1999) therefore separated the paraxial mesoderm from the axial structures and examined the role of Shh in expression of Pax-1, Myo-D and Pax-3. Pax-1 is rescued by Shh, Myo-D is maintained but not induced by Shh, while Pax-3 is expressed independently of Shh. Thus, Shh is a potent mitogen for somitic cells. Murtaugh et al. (1999) showed that Shh has several actions on somitic precursor cells in chick embryos. Presomitic mesodermal cells respond to Shh by initiating chondrogenesis, the later stages of their response depending on Bmp signaling. Shh enhances the response of somitic cells to Bmps, suggesting at least two phases. Finally, Shh enhances precursor cells competent to respond to Bmp by initiating chondrogenesis. Msx-2 and notochordal influences are differentially expressed in dorsal and neural-arch development in chick embryos. Bmps are also involved; there is parallel expression of Bmp-4 and Msx-1 and Msx-2 in the lateral neural plate and then in the dorsal neural tube and midline ectoderm in chick embryos. These are locations from which signaling could be transferred to the sclerotome. Grafting Bmp-4 or Bmp-2 into the neural tube up-regulates Msx-1 and Msx-2 in the adjacent mesenchyme, resulting in production of ectopic cartilage in the pectoral girdle that develops. Grafting Bmp-4 or Bmp-2 lateral to the neural tube down-regulates Pax-1 and Pax-3. Subsequently, Shh was shown to antagonize Bmp-4 and Msx during vertebral development.25 The paired type homeodomain transcription factor Uncx4.1, which acts upstream of Pax-9, is required before neural arches can form from lateral sclerotomal cells. Mice homozygous for a targeted mutation in Uncx4.1 die perinatally with severe malformations of the axial skeleton – all lateral sclerotomal derivatives (neural arches) fail to form because chondrogenesis is inhibited. The defect is early – anlagen form but fail to condense – further reinforcing the roles of these genes at the condensation stage of vertebral chondrogenesis.26 By using morpholino antisense oligonucleotides against the transcription factor twist, Yasutake et al. (2004) showed that twist is required for neural arches to form in embryos of the Japanese medaka, Oryzias latipes, because of involvement of twist after sclerotomal-cell migration.
The Search for the Magic Bullet
CONCLUSIONS The major events occurring in sclerotome, ventral spinal cord and notochord during chick development discussed in Chapters 41 and 42 are summarized in Table 42.7 and below. Sclerotomal mesenchyme has an inherent potential for chondrogenesis. Nevertheless, expression of chondrogenic potential is exquisitely sensitive to the environment. Depending on the nature of supplements added in vitro, sclerotomal mesenchyme from embryonic chicks as young as H.H. 10 (36 hours of incubation) can chondrify. Grafting somites to a vascularized environment such as the CAM permits somites from even younger embryos to chondrify, presumably not because of vascularization per se, but because of exposure to molecules in the circulation. The presence of the synthetic machinery for, and the synthesis of CS – attributes previously thought to apply only to induced mesoderm – are shared with many nonchondrogenic tissues. What is special about sclerotomal mesenchyme and its interaction with notochord and spinal cord is the ability to augment the rate of synthesis of CS and other GAGs, and to deposit these products into an ECM. Control is not at the level of presence or absence of the synthetic pathways, but in regulating those pathways. Notochord and spinal cord provide the major in-vivo environmental factors that augment somitic chondrogenesis. Notochord and spinal cord are also the most potent agents active in vitro. Whilst other factors can substitute in vitro, their absence is not compensated for in vivo. Collagen and CS enhance and maintain chondrogenesis by positive feedback to matrix products, chondrogenesis having been initiated at the condensation stage through the action of such regulatory genes as Pax-1 and Pax-9. Normal morphogenesis of the neural arches and centra also depends on influences from spinal cord, notochord and spinal ganglia. Ablating these tissues provides the evidence for this conclusion. The underlying mechanisms await resolution, although Msx and Bmps are possible players. If epidermal ectoderm plays a role in vivo, it is subordinate to and dependent upon notochord and/or spinal cord.
NOTES 1. See Watterson (1952) for the studies on Fundulus, Detwiler and Holtzer (1956) and Strudel and Gateau (1971) for abnormalities in urodele spinal cord and chick notochord, Lash (1968b) for maintenance of cartilage by notochordal influences, and Roy et al. (2004) and Witten et al. (2005) for vertebral anomalies in aquacultured fish. 2. See Sadler et al. (2001) for triploid Atlantic salmon, and Kacem et al. (2004) for triploid rainbow trout. 3. See Cooper (1965) for the correlation with hypertrophy, and Bancroft and Bellairs (1976) for associated ultrastructural features.
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4. See Holtzer and Detwiler (1953) for the urodele study, Strudel (1953b) and Watterson et al. (1954) for secondary neural arches, and O’Hare (1972c) for the CAM-grafts. Culturing myotome plus notochord and spinal cord from chick embryos of H.H. 18 did not result in chondrogenesis (Cooper, 1965). It may be, however, that by H.H. 18, the myotome is fixed with respect to ability to form muscle and cannot switch. 5. See Lash (1963a) for a review of the first decade of the search (1953–1963), Hall (1977a) for the research to 1977, and Monsoro-Burq and Le Douarin (2000) and Stockdale et al. (2000) for more recent approaches. 6. See Strudel (1953b, 1962, 1962). 7. Epithelial otic vesicles from mouse and chick embryos are amongst the other tissues that have been tested for an ability to induce sclerotomal mesenchyme and found not to induce (Grobstein and Holtzer, 1955; Strudel, 1955, 1962; Benoit, 1960b; O’Hare, 1972b). See Benoit (1955, 1960a,c, 1964) and Hall (1991b) for the ability of the otic capsule to form in ectopic sites. 8. See Amprino (1955a) and Johnson and Comar (1957) for 35S uptake into somatic mesoderm, and Lash et al. (1960) for uptake and CS. Lash (1963a) showed that CS is sensitive to RNA’ase digestion. 9. See Glick et al. (1964) for production of ATP-sulphurylase and APS-kinase, Franco-Browder et al. (1963) for production of chondroitin-4-sulphate and chondroitin-6-sulphate, and Okayama et al. (1976) for a discussion of early determination for chondrogenesis. 10. See Franco-Browder et al. (1963) for the studies with H.H. 11 embryos, and Lash (1963) and Lash et al. (1964) for 35S uptake. 11. See O’Connell and Low (1970) for ultrastructure, Kvist and Finnegan (1970a,b) for histochemistry, Toole (1972) for HA:CS, and Goldberg and Toole (1984) for hyaluronan as part of the pericellular coat. 12. See Frederickson and Low (1971), Strudel (1971) and Corsin (1974) for the amorphous material and importance of the ECM, and Minor (1973) and Bancroft and Bellairs (1976) for the GAG granules and collagen fibrils. 13. See Hay and Meier (1974) for separation of chondroitin and heparin sulphates, and Mathews (1971, 1975) for the similarity of CS–protein complexes. 14. See Vasan (1981, 1983, 1987) for proteoglycan accumulation, Vasan et al. (1986a,b) for core protein, hyaluronan and hyaluronidase, and Vasan and Miller (1985) for sternal cartilage monomers. 15. See Jurand (1962, 1974), O’Connell and Low (1970), Frederickson and Low (1971), Minor (1973) and Bancroft and Bellairs (1976) for perinotochordal fibrils. 16. See Bazin and Strudel (1972, 1973) and Ruggeri (1972) for independent autoradiographic studies. 17. See Carlson et al. (1974) and Carlson and Upson (1974). Hydroxylation of proline and lysine is inhibited in ascorbic acid (vitamin C) deficiency. Consequently, biosynthesis of collagen and elastin is impaired (Barnes et al., 1969; Wu et al., 1989). See Linsenmayer et al. (1973a) and Trelstad et al. (1973) for notochord and spinal-cord type II collagen, von der Mark et al. (1976) and von der Mark (1980) for the work with type II antibodies, and Hayashi et al. (1992) for notochordal secretion of type X collagen. 18. See Strudel (1972, 1973a–c) for the enzyme studies and Strudel (1975a–c) for the studies with LACA (which inhibits
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19.
20.
21.
22.
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excretion of collagen and consequently inhibits growth and delays chondrogenesis, for which see Aydelotte and Kochhar, 1972 and Hall, 1978c). See Kosher and Lash (1975) and Lauscher and Carlson (1975) for studies with chondroitinase and hyaluronidase. Mice immunized with chondroitinase ABC, proteoglycan and adjuvant develop polyarthritis, an ankylosing spondylitis (Glant et al., 1987). See Kosher et al. (1973) for the studies with CSs, and Kosher and Church (1975) and Kosher and Savage (1979) for the studies with procollagen and collagen. See Deutsch et al. (1988) for Pax-1 expression, Wallin et al. (1994) for early and late roles in chondrogenesis, and Wilting et al. (1995) for caudal expression. Wilting and colleagues also used the boundary of expression to determine the head/trunk (cervical/occipital) boundary in Japanese quail and mice. See Manley and Capecchi (1997) for the concept of redundancy among Hox genes and see Wellik and Capecchi (2003) for the Hox-10/Hox-11 example.
23. See Peters et al. (1998, 1999) for Pax-1 and Pax-9, Neubüser et al. (1995) for the distribution of Pax-9 and Danforth’s short tail, and Balling et al. (1996) for Pax-1 knockout. Pax-1 and Pax-9 are also expressed in complex, non-overlapping patterns in chick limb mesenchyme, especially at the condensation stage, when expression is strongest (Le Clair et al., 1999). See Healy et al. (1999) for expression of Pax-1 in the dermomyotome. 24. The winged helix transcription factor, MFH-1, is expressed in mouse cranial neural crest and head mesoderm and then in cartilage. MFH-1 mice die as embryos or at birth (Iida et al., 1997). 25. See Monsoro-Burq et al. (1994, 1996), Liem et al. (1995), Pourquié et al. (1996), Watanabe and Le Douarin (1996) and Watanabe et al. (1998a). Tgf-1 enhances chondrogenesis from chick sclerotomal micromass cultures (Sanders et al., 1993), probably because of a generic proliferative effect. 26. See Leitges et al. (2000) and Mansouri et al. (2000) for the studies with Uncx4.1.