mucochondroid (mucous connective) tissues in the heads of teleosts

Anat Embryol (1988) 178:461 474 Anatomy and Embryology �9 Springer-Verlag 1988

Mucochondroid (mucous connective) tissues in the heads of teleosts Michael Benjamin Department of Anatomy, University College, Cathays Park, Cardiff CF1 1XL, Wales, United Kingdom

Summary. The distribution and structure of mucochondroid tissues in the heads of teleosts has been studied in 56 species from 26 families. The tissues could also be called mucous connective tissue and have previously been known as basophilic gelatinous tissue. They are a heterogeneous group of tissues that contain fibroblasts (type example - the subcutaneous mucochondroid of the Cobitidae) or hyaline cells (type example - the mucochondroid around the medulla oblongata and spinal cord of Rasbora heteromorpha), embedded in a pale-staining matrix in which there are variable numbers of collagen fibres and blood vessels. Mucochondroid tissue is especially common beneath the skin, in labial folds, around olfactory and accessory olfactory sacs, in opercular valves and beneath the sensory epithelia of the stato-acoustic organ. The histochemistry of several mucochondroid tissues has been studied in Misgurnus anguillicaudatus, Acanthopsis choirorhynchus, Gnathonemus petersi and Cyclopterus lumpus. They have widely different amounts and types of glycoproteins, glycosaminoglycans and connective tissue fibres. The ultrastructure of subcutaneous mucochondroid is described in Acanthophthalmus semicinctus. Its cells contain little rough endoplasmic reticulum and a small Golgi apparatus, but numerous plasmalemmal vesicles, especially in the cell processes. The matrix contains 23 35 nm diameter granules, collagen and 11-12 nm diameter microfibrils. The similarities between mucochondroid and hyaline cell chondroid (cartilage) at the ultrastructural level, are more obvious than their differences.

Key words: Mucochondroid - Mucous connective tissue - Cartilage Teleosts

Introduction

In his comprehensive monograph on supporting tissues, Schaffer (1930) distinguished between notochordal tissues, those that resembled the notochord ('chordoid'), cartilage, and tissues that resembled cartilage (' chondroid'). He further described 4 subcategories of chondroid tissues (vesicu- lar, hyaline cell, mucochondroid with branching cells (called fibroblasts in the current manuscript), and mucochondroid with hyaline cells).

Moss and Moss-Salentijn (1983) have proposed that we now view all of Schaffer's (1930) tissues as forms of cartilage. They argue that his tissues are modulations of acom-

mon stem cell and form a continuous spectrum of tissues in lower vertebrates. However, at a different hierarchical level, the need remains for a classification that reflects the diversity of tissues encountered by the comparative histo- logist. A good classification is a convenient shorthand that saves the investigator from describing every new finding of a tissue in detail. It has been my experience in examining fish heads, that the hyaline cell and mucochondroid tissues of Schaffer (1930) are very common in teleosts. Any classification must take them into account. Yet, they have largely been ignored in the last fifty years.

The present paper describes the distribution, structure and histochemistry of Schaffer's (1930) mucochondroid tissues in the heads of teleosts. It is argued that the tissues should also be called mucous connective tissue. A subse- quent paper will deal with Schaffer's (1930) hyaline cell chondroid. The aim is to re-investigate the whole range of supporting tissues in teleosts, in order to provide a basis for a modern classification of these tissues in fish.

Materials and methods

Routine histology. The survey of the structure and distribution of mucochondroid tissues is based on fish selected from the author's large collection of serial sections of the heads of small and/or young teleosts. The collection is available for viewing by arrangement with the author. The species examined are those included in Table 1. All measurements are standard lengths of the fish (i.e. the distance from the tip of the snout to the base of the tail), and the common names quoted in Table 1 are the ones most frequently used in the literature. The details of processing varied slightly, but the following largely applied.

One to three specimens each of the fish named in Ta- ble 1, were fixed in 10% neutral buffered formal saline. The heads were decalcified in 2% nitric acid, dehydrated with graded alcohols, cleared with inhibisol and embedded in 56 ~ C paramat wax or histowax. Serial sections (in transverse, sagittal or coronal planes) were cut at 8 gm on a Leitz rotary microtome. Alternate slides were stained with haematoxylin and eosin (H & E), and Masson's trichrome. Sections from occasional heads were also stained with van Gieson's connective tissue stain and Weigert's elastic stain, and with Mayer's haematoxylin, celestine blue B, alcian blue 8G-X and direct red (Hall 1986). Because of the enormous number of sections available for studying the distribution of the tissues, one section from every other slide was

462

Table 1. Species examined in the present paper

Scientific name Common name Length(s) (in mm)

Family Anguillidae

1. Anguilla anguilla European eel 80, 140

Family Anostomidae 2. Anostomus anostomus Striped anostomus 70

Family Aplocheilidae 3. Aplocheilus panchax Blue panchax 35, 35

Family Atherinidae

4. Telmatherina ladigesi Celebes rainbow fish 33, 38

Family Belontiidae 5. Betta splendens Siamese fighting fish 42 6. Trichogaster trichopterus Opaline gourami 36 7. Trichopsis vittatus Croaking gourami 33, 34

Family Callichthyidae 8. Corydoras metae Masked corydoras 32

Family Characidae 9. Alestes longipinnis Long-finned characin 37, 53 10. Astyanax fasciatus mexicanus Blind cave fish 26, 27, 28 11. Hemigrammus rhodostomus Rummy-nosed tetra 37, 37 12. Cheirodon innesi Neon tetra 27, 28 13. Moenkhausia sanctaefilomenae Red-eyed tetra 31, 33

Family Cichlidae 14. Aequidens maronii Keyhole cichlid 50 15. Cichlasoma nigrofasciatum Convict cichlid 22, 24

Family Clariidae 16. Heteropneustes fossilis Asiatic catfish 106

Family Cobitidae 17. Acanthophthalmus semicinctus Half-banded coolie loach 55, 63 18. Acanthopsis choirorhynchus Horse-faced loach 45, 75, 85 19. Botia horae Hora's loach 28, 28, 29 20. Botia hymenophysa Banded loach 53, 54 21. Botia modesta Orange-finned loach 57 22. Misgurnus anguillicaudatus Weatherfish 105 23. Noemacheilus barbatulus Stone loach 3 species; unknown lengths 24. Noemacheilus botia Mottled loach 39, 40

Family Cyclopteridae 25. Cyclopterus lumpus Lumpsucker 30, 150

Family Cyprinidae 26. Balantiocheilus melanopterus Silver shark 57 27. Barbus conchonius Rosy barb 36, 36 28. Barbus titteya Cherry barb 23, 24 29. Brachydanio rerio Zebra danio 38, 40, 40 30. Carassius auratus Goldfish 40, 47 31. Epalzeorhynchus kalopterus Flying fox 43, 53, 55 32. Garra taeniata Siamese stone-lapping fish 50 33. Labeo bicolor Red-tailed black shark 40, 42, 42 34. Morulius chrysophekadion Black shark 35, 40 35. Rasbora heteromorpha Harlequin fish 20, 28 36. Rasbora trilineata Scissor tail 44, 50 37. Rhodeus sericeus Bitterling 45 38. Tanichthys albonubes White cloud mountain minnow 21 39. Tinca tinca Tench 80

Family Cyprinodontidae 40. Jordanella floridae American flagfish 23, 28

Table 1 (continued)

Scientific name Common name Length(s) (in ram)

463

Family Exocoetidae 41. Dermogenys pusillus Half-beak 50, 50, 55

Family Gasteropelecidae 42. Carnegiella strigata fasciata Marbled hatchet fish 26, 26, 30

Family Gobididae 43. Brachygobius sp. Bumble-bee fish 17, 20, 20 44. Periophthalmus sp. Mudskipper 34, 45

Family Goodeidae 45. Ataeniobius toweri 32, 43

Family Gyrinocheilidae 46. Gyrinocheilus aymonieri Algal eater 35, 38, 47

Family Helostomatidae 47. Helostoma temmincki Kissing gourami 30, 33

Family Loricariidae 48. Farlowella sp. 145 49. Loricaria filamentosa Whiptail catfish 75 50. Otocinclus sp. Sucker catfish 30, 31, 33

Family Mastacembelidae 51. Macrognathus siamensis Spiny eei 130

Family Mormyridae 52. Gnathonemus petersi Elephant-nosed fish 77

Family Muraenidae 53. Gymnothorax sp. Moray eel 180

Family Poecilidae 54. Poecilia sphenops Black molly 32, 34, 40, 43

Family Serrasalmidae 55. Serrasalmus nattereri Red-bellied piranha 23, 23

Family Trachinidae 56. Trachinus vipera Lesser weeverfish 65

systematically examined throughout the head. Neighbour- ing sections were inspected as necessary, when anything interesting was found. The line drawings were made with the aid of a Zeiss Dokumato r and a Gillert and Sibert projection microscope.

Histochemistry. Two specimens of Acanthopsis choirorhynchus (42 and 45 mm standard length) and Misgurnus anguillicaudatus (88 and 93 mm), and one of Gnathonemus petersi (97 mm) were processed as above, but were decalcified in aqueous E D T A (ethylenediaminetetra-acetic acid - a che- lating agent). This decalcifying agent was recommended by Charman and Reid (1972) as the most suitable one for histochemical studies of glycoproteins. The end point for decalcification was determined radiographically. A piece of the dorsal jelly hump (the extensive, gelatinous mass of subcutaneous tissue on the dorsal aspect of the lumpsucker that accounts for about 18% of the fish weight (Davenport and Kjorsvik 1986)) from one specimen (150 mm) of Cyc- lopterus lumpus fixed in unbuffered 10% formalin, was also taken. The histochemical methods employed are listed in

Table 3. Sections of rat tracheal cartilage were included for comparison in all histochemical tests. Additional method controls were used where appropriate.

Electron microscopy. Pieces of subcutaneous, fibroblastic mucochondroid from the dorsal surface of the head in three specimens o f Acanthophthalmus semicinctus (45-57 ram), were fixed for 2 h at 4 ~ C, with 3% glutaraldehyde in 0.1 M cacodylate buffer, washed overnight and post-fixed in 1% osmium tetroxide for 1 h. The material was treated with 3% aqueous uranyl acetate, dehydrated with graded alcohols and embedded in Spurr 's resin. Sections were cut on an LKB III ultratome, stained with lead citrate and uranyl acetate and examined on a Phillips EM 401 electron microscope.

Observat ions

Criteria for the identification of mucochondroid. Mucochon- droid is easier to identify in Masson's trichrome than in H & E sections. It consists of fibroblasts (more usually)

Fig. 1 a-j. Mucochondroid in the heads of teleosts. In a-d and f-j, the cells are fibroblasts. In e they are hyaline and in i they are both. a Subcutaneous mucochondroid in Acanthopsis choirorhynchus. Note the widely-spaced fibroblasts (F) and the wavy bundles of collagen f/bres (C) that lie at right angles to the skin (S). b Mucochondroid in the labial folds of Betta splendens stands out clearly (arrows) in low power micrographs by virtue of its pale-staining matrix. D, dentary; 0 V, oral valve. Sagittal LS. e Large cells (arrows) with prominent cytoplasmic granules in the mucochondroid at the base of the maxillary barbel in M~sgurnus anguillieaudatus, d In the mucochondroid on the snout of Acanthopsis choirorhynchus, the stellate fibroblasts have elongate, dark-staining nuclei and paler cytoplasmic processes that are difficult to see. Some of the cells lie in lacunae (arrows). M, Melanophores. e In the mucochondroid between the spinal cord (SC) and the vertebral body (V) of Rasbora heterornorpha, there are hyaline cells arranged in ch0ndrones (arrows). The tissue is partly surrounded by a ~ (P) and contains bundles of collagen fibres (C) and occasional blood vessels (B). f Thick bundles of collagen fibres (C) and blood vessels (B) in the jelly lump of Cyclopterus Iumpus. g In Balantiocheilus rnelanopterus, there are many blood vessels (B) in the mueochondroid that supports the statoacoustic organ (E). h The lower jaw proboscis of Gnathonernus petersi has a supporting rod of mucochondroid that is surrounded by a thick 'perichondrium' (P) into which are inserted skeletal muscle fibres (SM) that move the organ, i At the base of the nasal skin flaps (NS) in Acanthopsis choirorhynchus, the mucochondroid has fibroblasts to the left (F) and hyaline cells to the right (H). ] An intimate association between the fibres of the extrinsic ocular muscles (SM) in Periophthalmus sp., and mucochondroid (asterisk). All sections stained with Masson's trichrome

465

or hyaline cells (less often), embedded in a pale-staining matrix that is not completely extracted in routine processing, as in other loose connective tissues (Fig. 1 a, e). It contains a variable number of collagen fibres and its matrix stains much less strongly after neutral fixatives than does that of hyaline cartilage. The tissue stands out clearly at low power (Fig. I b). There is more matrix and fewer cells than in hyaline cell chondroid (cartilage) (Benjamin 1986, 1988). The fibroblasts have dark-staining, elongate nuclei and delicate cytoplasmic processes (Fig. 1 d). Hyaline cells are larger, rounded or oval cells, with round or oval nuclei and a pale-staining cytoplasm (Fig. 1 e). They have no secretory granules or lipid droplets and are similar to the cells of hyaline cell chondroid (cartilage) (Benjamin 1986, 1988).

Type examples. A good ' type ' example of fibroblastic mucochondroid is the subcutaneous tissue of loaches belonging to the family Cobitidae (Figs. 1 a, d; 2b, c). Hyaline cell mucochondroid is well represented around the medulla oblongata and adjacent spinal cord of the harlequin fish, Ras- bora heteromorpha (Figs. I e, 3). This hyaline cell mucochondroid contains numerous blood vessels, and is surrounded by a thin per ichondrium or periosteum where it is next to bone (Fig. 1 e). Elsewhere, notably at its boundary with nervous tissue, there is no perichondrium.

Structural variations and distribution of mucochondroid. Ex- cept where otherwise stated, the mucochondroid tissues have fibroblastic cells.

The distribution of mucochondroid in the heads of teleosts is summarized in Table 2 and selected examples of the tissue are illustrated in Figs. 2-6. There is considerable structural variat ion within the limits defined above, especially in the amount of amorphous matrix, and the number of collagen fibres within it. Cell density also varies greatly, even in the same fish. There are more ceils per unit area in the labial folds ( ' l ips ' ) of Periophthalmus sp., than there are around the eyes, or in the ' ros t ra l fold ' (a term used in this paper to describe any crest-like fold of tissue that partly or completely covers the upper lip) (Fig. 2d). The cells may lie in lacunae (as in A. ehoirorhyn- chus (Fig. 1 d)). Irregularly-shaped melanophores are found within the mucochondroid of A. choirorhynchus (Fig. 1 d). In M. anguillicaudatus, there are some round cells, packed with melanin granules in the mucochondroid of the maxillary barbels (Figs. 1 c, 2 b).

The matrix of both types of mucochondroid contains occasional blood vessels, nerves, and bundles of collagen fibres (Fig. 1 a, c, e). The collagen bundles are thick and prominent in the jelly hump of C. lumpus (Fig. 1 f). In some fish with subcutaneous mncochondroid (e.g.A. choirorhynchus; Fig. 2c), wavy bundles of collagen fibres regularly traverse the tissue f rom its superficial to its deep aspects (Fig. 1 a). Vessels and nerves are especially conspicuous where mucochondroid supports the sensory epithelia of the statoacoustic organ (Figs. 1 g, 4b). However, collagen fibres are here more difficult to see in H & E or Masson 's trichrome sections, though the tissue does stain pale red with van Gieson's connective tissue stain. For these reasons, the mucochondroid of the stato-acoustic organ is particularly distinctive and very similar in different species.

There is a thick per ichondrium around the mucochondroid tissue that supports the lower jaw proboscis or barbel of G. petersi (Figs. lh , 6c) and the maxillary barbels of

Table 2. A summary of the distribution of mucochondroid tissues in the heads of teleosts. The numbers refer to the species numbers in Table 1. An asterisk signifies hyaline cell mucochondroid. Where there is no asterisk, the tissue is fibroblastic mucochondroid

Atrium: 12 Barbels (in core or at base): 18, 19, 22, 22* Basioccipital chewing pad (between bone and pad) : 26, 26*, 36* Beak (in its delicate membrane): 41 Blood vessels (surrounding large vessels): 16 Branchial arches: 16" Buccal mucosa (excluding basihyal covering): 3, 6, 9, 16, 18, 18",

45, 52 Cranial nerves (surrounding them): 16, 32 Epidermal neuroblasts (forming a sheath for them) : 47 Eyes (around them ol- between extrinsic ocular muscles): 9, 22,

36, 44 Heart valves (within the core): 5, 12, 24, 34, 39, 52 Lateral line canals (surrounding them) : 4, 23, 45, 52 Labial folds (including oral suckers): 1, 3, 5, 9, 14, 15, 16, 22,

23, 27, 29, 38, 39, 42, 43, 44, 46, 47, 49, 54 Medulla oblongata/spinal cord (around it): 19, 22, 24, 26, 30, 33,

34", 35", 36, 37 Nares: 8, 17, 18", 19, 33, 34, 47, 48, 49, 50 Olfactory and accessory olfactory sacs (around them): 1, 2, 9, 11,

15, 17, 18, 19, 19", 22, 29, 44, 48, 54 Opercular valves: 9, 9", 10, 11, 13, 16, 17, 18, 19, 19", 20, 22,

26, 26", 29, 30, 31, 33, 34, 36, 39, 39", 46, 52, 54 Oral valves (within the core) : 3, 14, 49, 49* Pharyngeal mucosa: 5, 6, 6", 14, 15, 18, 18", 37, 45, 46, 54* Pharyngeal bones (in region of teeth) : 39 Proboscis (within the core): 52 Rostral fold: 15, 27, 33, 44, 45 Skin (beneath it): 1, 4, 9, 15, 16, 17, 19, 20, 21, 22, 23, 24, 28,

31, 35, 37, 38, 39, 40, 44, 51, 53, 54", 56 Stato-acoustic organ (supporting the sensory epithelia): 2, 3, 4,

7, 8, 9, t0, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 24, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 39, 42, 46, 47, 48, 49, 50, 52

Scales: 15, 45 Tongue (tissue covering basihyal): 1, 9, 13, 14, 45, 52, 54, 55,

56

M. anguillicaudatus (Fig. 2b). It is clear f rom watching live specimens of G. petersi in captivity that the tissue provides the nose of this fish with enough rigidity for it to be probed into a soft substrate, yet enough flexibility for it to be bent backwards by muscle action.

Mucochondroid is particularly evident beneath the skin on the dorsal and ventral surfaces of the head (Figs. 1 a, 2). Here, it is sometimes possible to appreciate the gelatinous properties of the tissue in gross dissections. Subcutane- ous mucochondroid is most characteristic of bottom-living fish, including the mudskipper, Periophthalmus sp. (Fig. 2 d) and numerous members of the Cobitidae (as stated above

see also Table 2). By far the largest quanti ty of a subcutaneous gelatinous tissue is in the jelly hump along the dorsal aspect of C. lumpus (see Fig. 1 of Davenpor t and Kjorsvik 1986). It is also present beneath the skin of larval lumpfish before a definitive lump develops. Subcutaneous mucochondroid commonly lies between the integument and any underlying bone (Fig. 2a, c). It is less prominent between the skin and the head musculature, though it is found in this position on the ventral surface of the head in M. anguillicaudatus (Fig. 2b).

In other areas too, mucochondroid tissues underlie and support epithelial membranes - in the labial folds (Figs. 1 b;

466

a

,7/

\

o,-~ ~ r , - : ' i ' C

O O" "

r

LF

S 4~.

q t b

SM

Fig. 2a-d. Mucochondroid (*). a Superficial to the lacrimal bone (L) of Rasbora heterornorpha. Transverse section, b Subcutaneously, and in the maxillary barbels (MB) of Misgurnus anguillicaudatus. BC, buccal cavity; SM, skeletal muscle. Sagittal LS. e Around the olfactory sac (O), in the nares (N) and beneath the skin (S) of Acanthopsis choirorhynchus. Sagittal LS. d Beneath the skin (S), around the eyes (E) and extrinsic ocular muscles (SM), and in the labial (LF) and rostral folds (R) of Periophthalrnus sp. Sagittal LS

467

~O

a

Fig. 3. Hyaline cell mucochondroid (*) around the medulla oblongata (MO) of Rasbora heteromorpha. B, bone; N, notochord

6 a, b, d), rostra1 folds (Figs. 2 d; 6 b, d), buccal and pharyngeal mucosae (Fig. 6c), the ' t ongue ' (the free end of the basihyal and its overlying tissues, that is separated from the rest of the floor of the buccal cavity by a groove (Fig. 4c)), nares (Figs. li, 2c, 5c), lateral line canals (Fig. 4a), in the stato-acoustic organ (Figs. lg , 4b) and in oral and opercular valves (Fig. 5 a, b). Hyaline cell mucochondroid is particularly common in the opercular valves of several teleosts (Table 2).

Transitions to other tissues. In the opercular valves of Bar- bus conchonius, Noemacheilus botia, Brachydanio rerio and Tinca tinca, one or both types of mucochondroid merge gradually with hyaline cell chondroid (cartilage). The latter tissue is always nearer to the attached margin of the valve than is either mucochondroid tissue. In Brachydanio rerio, the transition is directly from fibroblastic mucochondroid to hyaline cell chondroid (cartilage), but in the other teleosts, hyaline cell mucochondroid is an intermediate tissue. The intimate association of mucochondroid with hyaline cell chondroid (cartilage) is also found elsewhere e.g. lateral to the olfactory sac in Poeeilia sphenops, at the base of the nasal skin flaps in Acanthopsis choirorhynchus (Figs. 1 c, 2c) and on the free margin of the premaxillae in Misgurnus anguillicaudatus.

In many of the opercular valves listed in Table 2 (e.g. those of Tinca tinca and Botia hymenophysa), the mucochondroid tissues are intimately associated with skeletal muscle fibres. This admixture also occurs in the periorbital tissue o f Periophthalmus barbarus (Figs. l j, 2d) and in the pharyngeal mucosae of Aequidens maroni and Cichlasoma nigrofasciatum. In C. nigrofasciatum, there are occasional fat cells in the mucochondroid tissue, as there are in the tongue of Alestes longipinnis (Fig. 4c).

The tissue merges gradually with dense fibrous connective tissue on the lateral side of the lacrimal bone in Barbus

b SE

BP

C

kJ

Fig. 4a-e. Mucochondroid (*). a Around a lateral line canal (LC) in Telmatherina ladigesi. E, Eye. Transverse section, b Supporting the sensory epithelia (SE) of the stato-acoustic organ of MoruIius chrysophekadion. BP, basioccipital chewing pad; PC, pharyngeal cavity. Sagittal LS. e In the 'tongue' (7) of Moenkhausia sanctaefilomenae. FC, fat cells; L J, lower jaw. Sagittal LS

conchonius, and is in direct contact with hyaline cartilage (e.g. on the dorsum of the head of Poecilia sphenops (Fig. 6b) and with bone (e.g. in Morulius chrysophekadion, Acanthopsis choirorhynchus and Rasbora heteromorpha

a

468

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t

i t

\

OP

d A

~ BV BA

V Fig. 5a-d. Mucochondroid (*). a In the oral valves (OV) and in the lateral skin folds (F) on the beak (B) of Dermogenys pusillus. Sagittal LS. h In the opercuIar valve (OP) of Atestes longipinnis. Transverse section, e In the nasal skin flaps (N) of Corydoras metae. Transverse section of olfactory region (O). d In the bulbo-ventricular (BV) and atrioventricular (A V) valves of Betta splendens. A, atrium; BA, bulbus arteriosus; V, ventricle. Sagittal LS

Figs. 2a, c; 6d). However, no transitional tissues have yet been seen where mucochondroid meets hyaline cartilage or bone. Certain of the sites in which mucochondroid is found are occupied in closely related fish by hyaline cell chondroid (cartilage), or by fat. Thus, the considerable quantity of subcutaneous mucochondroid in many members of the Co- bitidae is replaced by adipose tissue in Botia horae, and the rostral cap of Labeo bicolor and Garra laeniata is sup- ported by hyaline cell chondroid (cartilage).

Ultrastructure. The cells are widely dispersed in the abun- dant matrix and have numerous attenuated cytoplasmic processes by which they contact adjacent cells. The stellate shape of the cells, and the absence of any distinction between territorial and interterritorial matrix, make the tissue resemble loose connective tissue in low power electron micrographs (Fig. 7 a).

The nuclei have a moderate amount of heterochromatin that is particularly prominent at their periphery (Fig. 7 a). Nucleoli are also visible. There is little rough endoplasmic reticulum and that present is scattered throughout the cells as short cisternae. There are small Golgi bodies, many free ribosomes, a moderate number of mitochondria and a few intermediate filaments (Figs. 7a, e). Centrioles are quite common.

The most conspicuous feature o f the cells is the large number of plasmalemmal vesicles, many of which are fused with the cell membrane or each other. They may contain a pale, flocculent material that is continuous with a similar substance in the extracellular space (Fig. 7 b). Vesicles dominate the cell processes (Fig. 7c), except for the thinnest processes, which lacked any membranous organelles. There are also occasional coated vesicles and thickenings on the cytoplasmic aspect of the cell membrane (Fig. 7d).

469

b

BC

C ~ J- """

B M '

~ p ' ~ : N ~ c < ; U " ' \

PC P "~-~.~-~- Kt"

Fig. 6a-~l. Mucochondroid (*). a In the labial folds (LF) of Betta splendens. BC, buccal cavity. Sagittal LS. b In the upper labial (LF) and rostral folds (RF) of Poecilia sphenops. BC, buccal cavity. Sagittal LS. e In the lower jaw proboscis (P) and buccal mucosa (BM) of Gnathonemus petersi. BC, buccal cavity; P, 'perichondrium'. Sagittal LS. d In the rostral fold (RF) of Morulius chrysophekadion. BC, buccal cavity. Sagittal LS

The matrix contains two types of filaments, and a small number of matrix granules (23-35 nm in diameter) and in- tergranular material (Fig. 7e). The filaments include cross- banded collagen fibres (30-40 nm in diameter) that are most evident at the periphery of the tissue and around blood vessels and nerves. There are also a much larger number of thinner (11-12 nm), unbanded and often wavy, connective tissue microfibrils (Fig. 7e). These can be intimately related to the cell membrane and its associated vesicles. Sometimes they appear to lead directly from the cell cytoplasm to the matrix, through the cell membrane (Fig. 7f).

Histochemistry. The histochemical reactions of selected mucochondroid tissues are summarized in Table 3. All the subcutaneous tissues are PAS-negative (periodic acid Schiff- negative). The positive reaction of the tissue in the nose of G. petersi is associated with the glycogen content of the cells (the cells are no longer positive after diastase treat- ment). However, the PAS-positivity of the tissue supporting the sensory epithelia of the stato-acoustic organ is undim- ished in diastase-treated sections.

The subcutaneous mucochondroid tissues and the rod of tissue in the proboscis of G. petersi react strongly with

Fig. 7a -L The ultrastructure of subcutaneous fibroblastic mucochondroid in Acanthophthalmus semicinctus, a The tissue resembles loose connective tissue. Note the branching processes of its cells (arrows) and the absence of any distinction between territorial and interterritorial matrix. Microfibrils, but not collagen fibres are present in this micrograph. M, mitochondria, b Plasmalemmal vesicles (P) fused with the cell membrane. The vesicles contain a pale-staining material that is continuous with a coating of similar material outside the cell (arrows). e Plasmalemmal vesicles (P) completely encircle the periphery of this cytoplasmic process, d In areas devoid of attached plasmalemmal vesicles, there are thickenings on the cytoplasmic aspect of the cell membrane (arrows). e Matrix granules (MG), microfibrils (MF) and collagen fibres (C) in the extracellular space next to the elongated process of a fibroblast. Note the granules attached to the cell membrane (arrows) and the many ribosomes (R) within the cell process, f Microfibrils (MF) intimately related to the cell membrane (CM) and its associated vesicles (P)

471

Table 3. The histochemistry of fibroblastic mucochondroid tissue in A canthopsis choirorhynchus (fish number 18 in Table 1), Misgurnus anguillicaudatus (fish number 22 in Table 1), Gnathonemus petersi (fish number 52 in Table 1) and Cyclopterus lumpus (fish number 25 in Table ~). A, subcutaneous tissue; B, tissue supporting the sensory epithelia of the state-acoustic organ; C, tissue forming the core of the proboscis in G. petersi. A highly cellular ' t rue ' cartilage (Z, cartilage in gill arches) of Schaffer (1930) is also included for comparison. + + strongly positive, + positive, ( + ) weakly positive, - negative

A. choirorhynchus M. anguillicaudatus G. petersi C. lumpus

A B Z A B Z C A

Elastic stains Verhoeff's . . . . . . + + Weigert's + + - - + + - + + - Orcein + + - - + + - - + + - Aldehydefuchsin + + - + + + - + + + + - Schiff's reaction SchifFs reagent for demonstrating . . . . . ( + ) - + free aldehyde groups PAS for adjacent glycol groups - ( + ) + + - - + + + + Diastase-PAS for glycogen - ( + ) + + - - + - + Alcian blue pH 2.5 (acidic mucosubstances) ( + ) ( + ) + + + + + + + + pH 1.0 (sulphated mucosubstances) ( + ) + + - + + + + + + Alcian blue critical electrolyte concentrations of magnesium chloride 0.9 M for keratan sulphate . . . . . . . . 0.7 M for heparin/heparan sulphate & keratan sulphate -- + ( + ) - + - - - 0.5 M for strongly sulphated GAGs - + + - + + + + + - 0.3 M for weakly & strongly sulphated GAGs - + + + - + + + + + ( + ) 0.06 M for all GAGs + + + + + + + + + + + Toluidine blue for metachromasia pH 5.0 - ( + ) + + - ( + ) + + + - pU 4.0 - ( + ) + + -- ( + ) + + + -- pH 3.0 - - + -- -- ( + ) + -- pH 2.0 - - ( + ) -- -- ( + ) + -- pH 1.0 - - + -- -- + + --

orcein, a ldehyde fuchsin and Weiger t ' s elast ic stains. H o w - ever, it is only in G. petersi tha t m u c o c h o n d r o i d stains wi th Verhoe fFs haematoxy l in . This stain reveals a n e t w o r k o f deIicate, yet in tense ly-s ta in ing fibres t h r o u g h o u t the tissue and c i rcumferen t ia l a r rays o f fibres in the pe r i chondr ium. It is clear f r o m c o m p a r i n g ad jacent sect ions in G. petersi that mate r ia l in add i t ion to these fibres is s ta ining wi th the o ther elastic dyes.

W h e n s ta ined wi th alcian blue in the presence o f 0.9 M m a g n e s i u m chlor ide, only the ma t r ix o f ra t t racheal cart i- lage cont inues to stain. Indeed, in the crit ical e lec t ro ly te concen t r a t i on me thod , the subcu taneous m u c o c h o n d r o i d tissues only stain wi th alcian blue in the presence o f the lowest c o n c e n t r a t i o n ( 0 . 0 6 M ) o f m a g n e s i u m chlor ide. H o w e v e r , the m u c o c h o n d r o i d in the s ta to-acous t ic o rgan o f A. choirorhynehus and M. anguillicaudatus stains in the presence o f 0.7 M m a g n e s i u m chlor ide. In con t ras t to the cel lular car t i lage o f the gill f i laments and m a m m a l i a n hyaline cart i lage, the mat r ix o f fish m u c o c h o n d r o i d does n o t stain wi th to lu id ine blue.

In the je l ly h u m p of C. lumpus, m o s t o f the ma t r ix reacts wi th ne i ther a lc ian b lue n o r PAS. H o w e v e r , there are dis- crete, ovo id s tructures , the shape and size o f nuclei , that react posi t ively wi th bo th o f these stains.

Discussion

Schaffer (1930) dis t inguishes be tween ' t r u e ' car t i lages and c h o n d r o i d (tissues wi th proper t ies in t e rmed ia t e be tween those o f the n o t o c h o r d and carti lage). His n a r r o w concep t

o f ' t r u e ' car t i lage as a tissue tha t largely acts as a p recu r so r o f the ver tebra te axial skeleton, and which differs funda- menta l ly f rom chondro id , has been widely cri t icized by G a b e (1967), Person and Phi Ipot t (1969) and Person (1983). Moss and Moss-Sa len t i jn (1983) fur ther a rgue that exces- sive a t t en t ion to t e rmino logy obscures the con t inuous inter- g rad ing of tissues a long a spec t rum o f car t i l ag inous types. The i r a d o p t i o n o f the un i ta ry t e rm ' ca r t i l age ' signifies their bel ief tha t Schaffer ' s (1930) tissues can all be usefully in- c luded under this single rubric. A uni ta ry t e rm cer ta inly emphas izes deve lopmen ta l re la t ions be tween the tissues, yet in o the r ways, it re turns us to the state o f affairs in the early pa r t o f the C19th. Schaffer (1930) reminds us (p 210 o f his m o n o g r a p h ) tha t the t e rm ' c a r t i l a g e ' was or iginal ly no t a h is to logica l one, bu t was appl ied to a wide range o f noncalc i f ied tissues tha t were mechan ica l ly robust , resil- ient, pressure-res is tant and elastic. I t was agains t this back- g round tha t he subdiv ided ' c a r t i l a g e ' tissues in to n o t o c h o r - dal, chordo ida l , c h o n d r o i d a l and ' t r u l y ' car t i lag inous tissues. Schaffer ' s (1930) classif icat ion was mos t careful ly con- s idered and based on cri ter ia tha t were reasonab le at the time. It should no t be a b a n d o n e d wi thou t g o o d reason, bu t buil t u p o n by the app l ica t ion o f m o d e r n techniques .

His to logis ts cou ld regard Schaffer ' s (1930) hyal ine cell c h o n d r o i d as m u c h a f o r m o f car t i lage as Schaffer ' s Zellk- norpel (the cartilage gt stroma capsulaire of G a b e 1967 and Bert in 1958). But the ' c a r t i l a g i n o u s ' s tatus o f Schaffer ' s (1930) m u c o c h o n d r o i d needs fur ther c o m m e n t . As Schaffer (1930) h imse l f recognized, m u c o c h o n d r o i d is clearly different f r o m the centra l n o t i o n o f car t i lage ep i tomized by mare-

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malian hyaline cartilage. The cells are not shrunken in lacunae and the staining properties of the collagen fibres are not obscured by those of the matrix. To call the tissues cartilage and not mucous connective tissue, obscures a more obvious similarity to the mucous connective tissues of mammals. Indeed Schaffer (1930) himself (p. 203 of his monograph) pointed out that mucochondroid is only a peculiar form of connective tissue. Modern texts of mammalian histology always give Wharton's jelly in the umbilical cord as an example of mucous connective tissue (e.g. Fawcett 1986). However, older texts also refer to tooth pulp as such a tissue. This was included by Schaffer (1930 p 202) in his category of mucochondroid tissue with branching cells.

Wright and Youson (1982) thought that the mucocartilages of ammocoetes (which Schaffer (1930) included as mucochondroid tissues) should be viewed as special kinds of loose connective tissue - yet they still call the tissue 'mucocartilage' (Armstrong et al. 1987)! Tretjakoff (1928) referred to what Schaffer (1930) later called mucochondroid, as 'basophilic gelatinous tissues', though he had also referred to them earlier in his career as chondroid tissues. He regarded mucochondroid as differing from embryonic mucous connective tissue in its content of fibrous material.

I have not adopted Tretjakoff's (1928) term as it fails to convey any relation of the tissue either to ordinary connective tissue or to cartilage. Furthermore, the basophilia, which he says is as intense as that of hyaline cartilage, is fickle and depends (as he was aware) on the choice of fixative. The present results confirm his observation that mucochondroid is not basophilic with neutral fixatives. The variations in basophilia with fixation conditions make it likely that the histochemical reactions of the mucoid ground substance vary according to the preparation procedures. Further studies will be necessary therefore before confi- dence can be expressed in the present finding that glycoproteins are inconspicuous in subcutaneous mucochondroid.

There is no simple solution to the terminological and classification problems that are here discussed. I currently favour regarding the tissues that I have described, as both mucous connective and mucochondroid tissue, but for the moment at least, of maintaining Schaffer's (1930) distinction between such tissues and cartilage. I f we accept that teleosts have a continuous spectrum of connective and supporting tissues, it follows that it is impossible to devise any scheme of classification that is not subjective. The boundaries drawn between the different tissues are artificial and there will inevitably be occasions when they are inter- preted differently by different researchers. But this has always been so, and the choices open to the classifier can lie anywhere between forming many subcategories or none. In my opinion, a classification of teleostean supporting tissues that recognizes many subcategories of tissue is still both necessary and possible. The absence of a workable scheme, but with the bewildering variety of cartilages and related tissues, causes much confusion to ichthyologists. It has also contributed to the neglect of fish cartilage and related tissues by those biologists who are interested in fun- damental aspects of the generalities and specifics of tissues. There are few people who are prepared to study tissues they cannot easily name.

One cannot impose on mucochondroid any concept of a ' typical' cartilage cell. Indeed, it has long been recognized that the morphology of cartilage cells does not help in defin- ing cartilage (see Schaffer (1930) p 211 for a review of the

early literature). If the tissues are viewed as mucous connective tissue, their cells can hardly be regarded as chondrocytes at all! Certainly, in the subcutaneous mucochondroid of A. semicinctus, the ultrastructure of the cells differs mark- edly from that of typical mammalian chondrocytes (Kosher 1983). There is no extensive zone of RER and no prominent Golgi apparatus or secretory vacuoles. As in Schaffer's (1930) hyaline cell chondroid, there is a general paucity of organelles (Benjamin 1986). The cytoplasm of the cells in both tissues is dominated by free ribosomes and large numbers of plasmalemmal vesicles. Indeed the similarity of fibroblastic and hyaline cells is more striking ultrastruc- rurally than it is at the light microscopy level.

Plasmalemmal vesicles also characterize the fibrocytes of dermal connective tissue in the gular skin of the teleost, Agonus cataphractus (Whitear and Mittal 1986). They have been reported in lamprey cartilage (Wright and Youson 1982) and in mammalian chondrocytes (Cox and Peacock 1977; Wilsman et al. 1981) and are also typical of smooth muscle cells, including those in the teleostean bulbus arteriosus (Benjamin etal. 1983). However, the vesicles in smooth muscle cells may have more to do with ion seques- tration and stimulus-contraction coupling than do those in connective tissue cells. In connective tissues, they could indicate a high rate of matrix turnover.

The most obvious difference between mucochondroid fibroblasts and the cells of hyaline cell chondroid/cartilage (Benjamin 1986, 1988) is one of size and shape. The fibroblasts lack the voluminous cytoplasm of the hyaline cells, but have more numerous and attenuated cell processes. It is perhaps because of these differences that intermediate filaments are less conspicuous in them. It was suggested previously that the filaments gave structural support to the turgid-looking cells of hyaline cell chondroid (Benjamin 1986). Smaller cells may require less support. As discussed earlier, teleostean mucochondroid is similar to the mucous connective tissue of mammals - both have fibroblastic cells embedded in a gelatinous matrix. It is therefore worth not- ing the interesting comparison of Reynolds (1952) between the appearance of the cells of Wharton's jelly in a collapsed umbilical cord and the form of the cells in cords clamped with artery forceps before fixation. In the former, the cells are stellate and the tissue looks like a cushion for the con- tained blood vessels. In the latter, the cells are bipolar. The impression now is of a tissue that forms a resistant sheath, which with the pressure developed in the umbilical vessels, makes the cord a semi-rigid structure, somewhat like erectile tissue. We must therefore bear in mind that the stellate shape of mucochondroid cells in fish may be artefactual.

The 11 12 nm microfibrils that are prominent in the mucochondroid matrix of A. semicinctus, are common con- stituents of connective tissue in general (Inou6 and LeBlond 1986). They have also been called oxtalan fibres, one of a group of 'elastic system fibres' that are frequently reported in mammals (e.g. Cotta-Pereira et al. 1977). They are a source of much confusion - especially in lower vertebrates. According to Gawlik (1965), oxytalan and elaunin fibres (in contrast to true elastic fibres) do not stain with Verhoeff's haematoxylin, but do stain with orcein and aldehyde fnchsin. This rationale could explain the staining pattern of subcutaneous mucochondroid in the Cobitidae in the absence of amorphous deposits of elastin. However, Wright and Youson (1983) found that cartilage in the sea

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lamprey, Petromyzon marinus, does not contain biochemi- cally-detectable elastin, even though it is Verhoeff-positive. Kostovid-Knezevic et al. (1986) claim that elaunin and oxytalan fibres can be present in tissues devoid of elastic ele- ments. It is worth remembering that over half a century ago, Tretjakoff (1928) mooted the idea of a third class of fibre (fibril) other than elastic and collagenous/procollagen- ous (reticular) fibres. Inou6 and LeBlond (1986) distinguished between typical microfibrils that have a hollow core surrounded by a surface band, and atypical microfibrils that lack the hollow core. On this basis, the microfibrils in A. semicinctus are atypical.

The apparent continuity of cytoplasmic and extracellular microfibrils (Fig. 71) is difficult to interpret. Similar findings have been described previously in fibroblasts from the lamina propria of the mammalian gut (Pitha 1968) and from the mucous connective tissue of the umbilical cord (Parry 1976). It is tempting to suggest that the microfibrils are formed within the cell and gain exit to the matrix through deficiencies in the cell membrane. However, one cannot exclude the possibility that the appearance results from tangential sectioning.

According to Schaffer (1930), fibroblastic mucochondroid is present in various locations in a wide range of vertebrates and invertebrates. These include the subcutaneous tissue and peribranchial folds of Amphioxus, a large part of the head skeleton of ammocoetes, the lingual cartilage of Valvata piscinalis, the lingual papillae of Limnaea, beneath the skin musculature of Pterotrachea, the mantel of tunicates, the collar of sabellid and terebellid worms, around the semicircular canals of lower vertebrates, in the heart valves of man and Tropidonotus, the tooth pulp of the pig embryo and the epiglottis of the cat. He makes little mention of teleosts, though on page 201 of his monograph, he refers to mucochondroid on the premaxilla of Cobitis taenia and unspecified cranial bones in Cottus gobio. His account also confirms (p. 200) the subcutaneous deposits of the tissue in the Cobitidae. It seems likely that Schaffer's (1930) ideas on mucochondroid were given shape by his earlier work on lampreys and hagfish, and that he was trying to fit teleost tissues into a framework that he had already established in his mind. Tretjakoff (1928) dealt with fish in more detail and indeed Schaffer's (1930) review relies heavily on TretjakoflTs findings. Tretjakoff (1930) described mucochondroid in many locations - in the orbit of A cipenser, the papillae and cushions of the skin of Botkus maeoticus, the scale pockets of Solea, the marrow cavities of Pleuronectidae and Plectognathi ( = Tetraodontiformes), the heart and arteries of all fish and the electric organs of rays.

Whereas the functional characteristics of hyaline cell chondroid (cartilage) must reflect the properties of the cells that dominate the tissue (Benjamin 1986), the functions of any mucochondroid tissue probably depend more on the character of the extracellular materials. Tretjakoff (1928) thought that gelatinous tissues frequently served as packing material e.g. in the orbit and other skull foraminae, and in the labial folds and fin rays. His opinion is endorsed by the present author. It is my experience that areolar connective tissue less frequently performs this function. It gen- erally forms thin sheets and provides a plane of movement between two structures. Where a greater thickness of tissue is required for support, but without sacrificing mobility, mucochondroid tissue or adipose tissues may provide an

answer. What determines whether mucochondroid, rather than adipose tissue is present is unclear. It could depend on the buoyancy requirement of the tissues (Davenport and Kjorsvik 1986) or on the need for mobilizing stored lipid. It is interesting to note that the two tissues can intermix (as in the basihyal covering of Alestes longipinnis), and that the subcutaneous mucochondroid in many loaches is replaced by adipose tissue in Botia korae. Schaffer (1930) discussed this idea of 'substitution-abili ty ' (especially in relation to adipose tissue) on several occasions and even used it as a criterion for a supporting tissue. He too, refers to subcutaneous adipose tissue in both scaleless and scaly fish (Schaffer (1930, p. 94).

Schaffer (1930) paid little attention to the significance of mucochondroid. He described it as having a supporting role by virtue of its thickness and its content of connective tissue fibres, and as being a stage in the development of hyaline cell chondroid and ' t rue ' cartilage. His idea of a replacement (including a de-differentiation) of mucochondroid (mucocartilage) into adult cartilage in the lamprey, Petromyzon marinus (see Hardisty 1981 for an English review of lamprey mucocartilages), has recently been con- firmed by Armstrong et al. (1987).

Johnels (1948) who conducted a most careful investiga- tion into ammocoete mucocartilages, views them as special- ized larval tissues that are adapted to the needs of a burrowing and microphagous animal. His ideas also find support from the distribution pattern of the tissue here described in fish. The conspicuous presence of mucochondroid next to the neurocranium of the Cobitidae and Anguilla anguilIa may be related to the bottom-dwelling habit of these fish. The viscous, lubricating properties of the tissue could pro- tect the skin and/or the underlying bone from damage when the fish burrow. The tissue is also prominent in the mud- probing proboscis or lower jaw barbel of G. petersi.

We know little of the physical properties of the tissue, though its gelatinous nature is often evident where gross dissections are possible. Greer Walker et al. (1985) described a jelly-like substance (which I regard as mucochondroid) around the heart of certain deep-sea teleosts. It may prevent the atrium from collapsing during systole.

One of the most strikingly gelatinous of teleostean tissues is the enormous jelly mass of the lumpfish, C. lumpus. According to Davenport and Kjorsvik (1986), its physical characteristics contributes to the unusually low density of the fish. This low density allows the animal to live in mid- water without a swim bladder and without the needless expenditure of energy. They describe it a s ' stiff' and suggest that it has an exoskeletal function. They also refer to it as elastic. I confirm that it springs back to shape after defor- mation (at least in fixed material), but I have failed to dem- onstrate elastic fibres within it. Its 'elasticity' is thus pre- sumably a property of its amorphous matrix.

Mikuriya (1972) has described the ' tongue ' of Gnathon- emus petersi as a deflated tube of fluid-containing tissue that is surrounded by a thick fibrous envelope. She regards the tissue as more fluid than solid on the basis of the dispersion of indian ink particles injected into the tongue of fresh specimens. I also have examined this tissue micro- scopically (Fig. 1 n) and regard it as mucochondroid. It is similar to that in the lower jaw proboscis, though less cellular and with an even paler-staining matrix. Observations on living specimens of G. petersi suggest that the proboscis is a highly malleable organ that is suited for probing into

474

soft mud. It can be bent double by the action of muscle fibres inserted into the fibrous envelope. What rigidity it has could well depend on its restraining fibrous sheath or 'per ichondr ium' . However, it is not known whether the perichondrium illustrated in Fig. I h can also provide for appositional growth of the tissue. A fibrous envelope also partly or completely surrounds some other mucochondroid tissues including the subcutaneous tissue of the Cobitidae. If their subcutaneous mucochondroid is an adaptat ion to a burrowing habit, then strategically placed fibrous tissue is a necessary complement to the more compression-resistant properties of other matrix components.

Acknowledgements. I wish to thank Dr. G.J. Howes for checking the taxonomic details in Table 1 and Dr. Howes and Professor W.A. Beresford for their invaluable comments on the manuscript. My thanks are also due to Professor M.L. Moss for his encourage- ment and interest in my work. Mr. D. Scarborough helped with the histochemistry and both he and Miss B. Edwards cut the routine sections. Mr. J. Sandhu prepared the material for electron microscopy. The specimens of C. lumpus, T. vipera and Gymnoth- orax sp. were kindly provided by the Ministry of Agriculture, Fisheries and Food, Lowestoft, Suffolk.

References

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Accepted March 1, 1988

mucochondroid (mucous connective) tissues in the heads of teleosts

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