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Two microvillar organs, new to Crustacea, in the Mystacocarida
Arthropod Structure & Development 37 (2008) 522–534
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Two microvillar organs, new to Crustacea, in the Mystacocarida Rolf Elofsson a, *, Robert R. Hessler b a b
¨ gen 3, SE-223 62 Lund, Sweden Department of Cell and Organism Biology, Zoology Building, University of Lund, Helgonava Scripps Institution of Oceanography, La Jolla, CA 92093-0202, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 19 February 2008 Accepted 24 May 2008
The mystacocarid crustacean Derocheilocaris typica has two microvillar organs, one new, the other previously unappreciated in crustacean literature. The first is situated on the head-shield and consists of three pairs of cells: one with microvilli and a ballooned nucleus; one smaller and without special features; the third large and investing the other two and extending down to the foregut. We call this new organ the ‘‘cephalic microvillar organ’’ and discuss the value of the concept ‘‘dorsal organ’’, to which it might have been included. The second organ consists of about 21 cells that cover the proximal part of the dorsal surface of the labrum. The cells are alike, being characterized by an apical field of microvilli and a large residual body. This organ is here called the ‘‘labral microvillar organ’’. Both organs are neither sensory nor secretory and do not qualify for membership in any of the other recognized organ systems. We are unable to deduce their Dero-cheilocaris functions. Ó 2008 Elsevier Ltd. All rights reserved.
Keywords: Crustacea Mystacocarida Dorsal organ Labral organ
1. General introduction In our attempt to study the anatomy of the mystacocarid, Derocheilocaris typica down to an ultrastructural level (Elofsson and Hessler, 2005a,b; Hessler and Elofsson, 2007), we have stumbled upon two organs whose purpose we cannot fathom. In spite of this inability, we feel it is essential to make the existence of these organs known to other investigators, in the hope that someone may be able to show what they are for. Both organs are cephalic. Both have microvillar fields that abut the cuticle. One is located middorsally, and thus qualifies for membership among the ‘‘dorsal organs’’. However, as shown below, as a category, ‘‘dorsal organ’’ is not meaningful. The other organ presented here is located on the atrium oral side of the labrum. Because these two organs constitute separate problems, we describe and discuss them separately. 2. Material and methods Abundant specimens were obtained from Stony Beach at Woods Hole, MA, USA. Sand was sieved through an approximately 60 mm filter from pits dug to a depth of 2–4 dm above high-tide line. They were fixed for 1 h in a refrigerator in either 0.3 M cacodylate buffer or seawater with 2.5% glutaraldehyde added. Postfixation was performed, after washing in buffer or seawater, with 1% osmium
* Corresponding author. Tel.: þ46 46 51040. E-mail addresses: [email protected] (R. Elofsson), [email protected] (R.R. Hessler). 1467-8039/$ – see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.asd.2008.05.002
tetroxide in the same buffer for 1 h. The specimens were embedded in Epon. Thin sections were cut on a LKB Ultrotome and poststained in 2% uranyl acetate and lead citrate in an LKB Ultrastainer. A JEOL transmission microscope was used. For scanning electron microscopy, the specimens were fixed as above and left in sucrose overnight. They were then critical point dried and coated with gold/palladium. Observations were made with a JEOL T330 microscope. 3. Cephalic microvillar organ (CMO) 3.1. Introduction A more or less well defined organ, situated dorsally on the head seems to be a widespread feature in crustaceans. It has been described under a variety of names: dorsal organ, Nackenorgan/neck organ/nucal organ, lattice organ, cephalic dorsal hump, Hafptorgan, salt gland, and integumental window to mention some. As the names suggest, the ideas about function are equally diverse. Martin and Laverack (1992) made a survey of the occurrence of dorsal organs in crustaceans, including their presumed function, the long history of their recognition and their appearance in fossil arthropods. Relevant ultrastructural information is further found in Lake et al. (1974) on syncarid crustaceans, Laverack and Sinclair (1994) on the decapod shrimp Macrobrachium intermedium, Laverack et al. (1996) on the syncarid Anaspides tasmaniae and the decapod shrimp Crangon crangon, and Hosfeld and Schminke (1997) and Hosfeld (1999) on harpacticoid crustaceans. Recent publications have widened the scope to include cirripeds (Ho¨eg et al., 1998) and facetotectans (Ho¨eg and Kolbasov, 2002). The discovery of the
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cephalic microvillar organ (CMO) in Derocheilocaris typica further widens the scope of such organs. 3.2. Results
Fig. 1. A. This scanning electron micrograph of the dorsal head-shield illustrates the external position of the cephalic microvillar organ (CMO) (arrow). Anterior is at the top of the photo where also the hinge in the head-shield is seen. Arrowheads point at four anterior sensory pegs of the head-shield forming a rectangle whose diagonals cross each other approximately where the CMO is situated. Scale bar 10 mm.
A small round area defining the position of the CMO is situated approximately 20 mm behind the hinge of the head-shield on Derocheilocaris typica. Externally, it is located at the crossing of diagonals of the imaginary rectangle formed by the two anterior pairs of pegs of the posterior head-shield (Elofsson and Hessler, 2005b). It is hardly seen from the outside since it appears only as a slightly elevated dome in the cuticle about 2 mm in diameter (Fig. 1). The head cuticle is approximately 0.5 mm thick, but over the organ it is only half of that thickness. There are no pores or other structures in its cuticle. Dorsally, the CMO itself is also small. It is squeezed between strong muscles and occupies an area below the cuticle about 6 mm in diameter. Its position, seen in a midsagittal section, is almost directly above the anterior-most margin of the loop formed by the foregut (Fig. 2). Three pairs of cells form the CMO (Fig. 3). One pair, the posterior cells, has apical microvilli, which are tightly apposed and perpendicular to the cuticle (Fig. 4A and inset). The tips of the microvilli are under the cuticular dome. They are
Fig. 2. A midsagittal section of the anterior portion of the head shows the position of both the cephalic microvillar organ (CMO) and the labral microvillar organ (LMO). The relation to part of the brain (B), foregut (F), and labrum (L) is also evident. Scale bar 10 mm.
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Fig. 3. A schematic drawing, from sections, of the CMO viewed from the side. The relation of the three paired cells, anterior (A), posterior (P) and ventral (V) of the CMO, to one another is given. Their position in the body is illustrated by the inclusion of the posterior portion of the brain (B), and the anterior ascending portion of the foregut (F). One large gland cell (G) represents the position of several gland cells, situated on both sides of the ventral cell (V). The lines numbered 1–5 illustrate the approximate levels at which the micrographs 4–6 are taken (1 ¼ Fig. 4A, 2 ¼ Fig. 4B, 3 ¼ Fig. 5A, 4 ¼ Fig. 5B and 5 ¼ Fig. 6). All Figs. 4–6 are horizontal sections from a section series through the organ from one animal.
approximately 50 nm in diameter and a little more than 1 mm in length (Fig. 4, inset). A non-microvillar border of the cell encircles the microvilli except where the two cells meet each other. The microvilli are thus enclosed in a cup formed by the two cells. Anterior to and below the base of the microvilli, much of the cell volume is filled with a vacuole almost 2 mm in diameter with a fine, even, granular content. This vacuole is formed by an outpocketing of the outer membrane of the nucleus (Figs. 4B and 5A). That same granular material forms an almost continuous layer between the inner and outer nuclear membranes all the way around the nucleus. This layer is continuous with the vacuole, but defined from it by a constriction. The ribosomes of the outer nuclear membrane follow the membrane around the vacuole. Since this structure appears in all the animals we studied, we do not judge it to be an artifact. The nucleus, contained in its inner membrane, is situated in a side or ventral portion of the cell (Figs. 4B and 5A). Its size is less than that of the vacuole. It has one large nucleolus and no heterochromatin. The cytoplasm contains cisternae of rough endoplasmic reticulum, a Golgi apparatus, relatively few small mitochondria (0.5 mm in diameter) with few cristae, and occasional lysosome-like vesicles the size of a mitochondrion. The pair of anterior cells is smaller than the preceding. They lack microvilli and the unusual nuclear vacuole and envelope (Figs. 4B and 5A). They meet the cuticle in front of the cuticular dome and follow the anterior outlines of the posterior cells tightly. The nucleus fills most of the cell. Its cytoplasm is similar to that of the posterior cells. These two cell types, the anterior and posterior cell pair, are intimately associated with a pair of large cells with an irregular outline (Fig. 3). Dorsally, these cells adjoin the previous two pairs anteriorly (Fig. 4B). More deeply, they also invest them along their sides and below. They extend between the previous cell pairs and the foregut, a distance of about 4 mm, narrowing ventrally to form together a thin lamella squeezed in between large lateral glands (Fig. 5B). Anteriorly, the cells meet the brain and end against the
circular muscles of the ascending foregut. The main portion of the cells continues posteriorly along the midline for 15–20 mm, whereupon the cells widen to accommodate the nucleus (Fig. 6). The nucleus (2–4 mm) has one large nucleolus and no heterochromatin. This posterior portion of this pair of cells sits broadly on the circular muscles of the foregut in a region where the foregut has turned posteriorly. We term this cell pair the ventral cells because of the position of the main portion of the cell in relation to the anterior and posterior cells. The cells are light due to only sparse rough endoplasmic reticulum and a loose granular cytoplasm. The anterior portion of the ventral cells houses some mitochondria and occasional dark lysosome-like bodies, both of which increase considerably in number in the posterior portion of the cells (Fig. 6). The size (1–2 mm) and shape of the mitochondria varies. They are characterized by having few and short cristae. The dark bodies vary in shape and size (more than 1 mm). Their content can be evenly dark or contain membrane whorls and dark particles. Often these lysosome-like vesicles are associated with mitochondria. There are no specific membrane specializations in any of the cells. A dark fibrous structure stretches out in intercellular spaces to surround the organ. We interpret this structure as an organ-limiting structure similar to a basal lamina.
3.3. Discussion The term ‘‘dorsal organ’’ has been used here temporarily as a handy name based only on the position of the structure. The term is, however, unsuitable as a morphologically and functionally unified feature and should be abandoned. Usually the ‘‘dorsal organ’’ is described on the basis of its external morphology, for which there is a rich literature, predominantly used for taxonomic work. However, external appearance will not be dealt with here; an ultrastructural description of internal anatomy is mandatory to appreciate the true nature of these organs.
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Fig. 4. A. The most dorsal horizontal section of the series, just below the small cuticular dome of the CMO (level 1 in Fig. 3), cuts the microvilli (MV) of the posterior cells (P). The tip of one anterior cell is emerging (A). Scale bar 2 mm. Inset: A sagittal section through the dome with the microvilli. This animal is ready to moult. Scale bar 0.5 mm. B. A horizontal section below that of A (see also level 2 in Fig. 3) cuts through all three cells of the CMO. Note the crammed position between large muscle cells (M). The ventral cell pair (V) sends a dorsal extension anterior to the anterior cell pair (A). The two posterior cells (P) are cut through the ballooned portion of the nucleus (asterisk). The right posterior cell shows the entire nucleus. Scale bar 1 mm.
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Fig. 5. A. A horizontal section further down in the series below that of Fig. 4B (see also level 3 in Fig. 3). At this level the anterior (A) and the posterior (P) cells are disappearing and the ventral (V) cells dominates the organ. Scale bar 1 mm. B. Next horizontal section is cut somewhat below that of A (see also level 4 in Fig. 3). Here only the ventral (V) cells of the CMO are left. Large gland (G) cells of different appearance and muscles (M) on both sides of the organ squeeze the ventral cell into a thin lamella. The arrow points in the direction of the caudal portion of the ventral cells. Scale bar 2 mm.
The problem of dorsal organ terminology is aggravated by the use of the term for temporary structures in arthropod embryos (Fioroni, 1980) that are said to be sites of cellular degeneration during development. Fioroni realized that there was no unifying
structure and suggested multiple evolutionary events. Anderson (1973) commented on transitional dorsal organs with degenerative function in malacostracans, and on a pair of transitional dorsolateral organs in peracaridean isopods with a different presumed
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Fig. 6. This last horizontal section of the series is taken just above the foregut (F) and shows the posterior portion of the CMO where only the ventral (V) cells are present (see also level 5 in Fig. 3). It is clearly seen here that the ventral cells dip down on both sides of the roof of the foregut. Their nuclei (N) are present in this region, the left nucleus with its nucleolus. Scale bar 2 mm.
function. All these investigations were light microscopical. An ultrastructural investigation of the transitional dorsal organ of Orchestia cavimana (Meschenmoser, 1989) found evidence to believe it to be an ion-transporting organ like the ones discussed below. The ‘‘dorsal organ’’ is not restricted to the head-shield. It can be paired and occur on body segments under the name of integumental windows (e.g. Hosfeld and Schminke, 1997; Hosfeld, 1999). Scanning electron microscopic and light microscopical observations have noted the presence of a ‘‘dorsal organ’’ in the form of four pits on the head-shield in fossil maxillopod (Mu¨ller and Walossek, 1988) and branchiopod (Walossek, 1993) crustaceans. It has also been found in trinucleid trilobites (Barrientos and Laverack, 1986) as well as in recent chelicerates, chilopods, diplopods, and insects (Fiorini, 1980). The present discussion will be restricted to crustaceans. Here, ultrastructural investigations reveal two major types. ‘‘Dorsal organs’’ of the taxon Thecostraca, comprising Ascothoracida, Cirripedia, and Facetotecta, form a distinct group (Ho¨eg et al., 1998; Ho¨eg and Kolbasov, 2002). They are termed lattice organs and appear in five pairs on the head-shield of cypris larvae. It is a sensory organ whose main components are two primary
receptor cells, each furnished with two (or four) modified cilia. The fate of the axons from the receptor cells has not been reported. Two cells sheath the receptor cells. The modified cilia reach into a chamber in the cuticle whose roof has pores. Within the thecostracans, this generalized morphology differs somewhat in details. One or two gland cells are situated between the pairs of the lattice organ. The gland cells have a microvillous border against the cuticle. The morphology of the lattice organ strongly suggests a chemosensory function. Another sensory ‘‘dorsal organ’’ is found in two malacostracan taxa, decapods (Crangon crangon) and syncarids (Anaspides tasmaniae) (Laverack et al., 1996). Here, one circumscribed median area on the head-shield contains four pits. Associated with the pits are four sensory cells, each bearing a modified cilium, which ends at the cuticle. The receptor cells and their cilia are wrapped in three sheath cells insulating the cilia from one another. Centrally in the area of the dorsal organ, a secretory cell opens into a pore of the cuticle. The secretory cell is microvillous and contains numerous vacuoles. Laverack and Sinclair (1994) found with methylene blue studies, that the dorsal organ of the decapod shrimp
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Fig. 7. This exsagittal section through the labrum is given to show the entire LMO. Anterior is to the left. The arrow to the very left indicates the approximate level of the mouth. The microvilli (MV) and nucleus (N) of LMO cells are clearly seen in this magnification. The cephalo-some relation to the atrium oris (AO), mandible (M) and the dorsal transverse fold of the labrum (DTF) is displayed in the photo. Scale bar 5 mm.
Macrobrachium intermedium was innervated from the tritocerebrum. Although Laverack believed the organ to be mechanosensory, the lack of structures associated with this modality make a chemosensory function more probable. Nishida (1989) studied a structure he called the cephalic dorsal hump in calanoid copepods. It was found to be a male specific feature in many calanoids, situated anterodorsally on the cephalosome. An ultrastructural study on Calanus sinicus and Paracalanus parvus found that the organ consists of two gland cells, each opening in a pore, and a receptor, which ends in a thin porous field on the cuticle. They form a row with the receptor being most posterior. The receptor consists of two cells and two enveloping cells. Two cilia from each receptor cell runs through a cavity enveloped by cells having distal microvilli. The cilia branch and end in an unexplained way within the porous cuticle. Nishida claims the organ to be homologous with the one described by Laverack et al. (1996) (see above) e.g. the sensory dorsal organ. However, the innervation, which was not investigated by Nishida, is crucial to the understanding of the organ. If the organ is innervated from the protocerebrum it could be the ‘‘third unit’’ of copepod crustaceans (Elofsson, 1971), similar to the cavity receptor organ of Artemia salina (Elofsson and Lake, 1971); only if it is innervated from the tritocerebrum could it be similar to the dorsal organ described by Laverack et al. (1996). Another type of ‘‘dorsal organ’’ is not innervated and possesses a transporting epithelium. Hootman and Conte (1975) described the ultrastructure of an ion-transporting epithelium of the neck organ in the nauplii of the anostracan Artemia salina. Fifty to 60 cuboidal cells contained many mitochondria associated with apical
and basal membrane infoldings. They found a great resemblance to salt secreting cells of other taxa. Halcrow (1982) investigated the dorsal (nuchal) organ of juvenile Daphnia magna, a cladoceran crustacean, formed as an expanded portion of a dorsal ridge. The organ comprises two types of cells, one darker due to the presence of abundant mitochondria and the other lighter; both bear microvilli on their apical surfaces. The microvilli differ somewhat on the two types of cells. A large number of mitochondria with prominent cristae are situated at the bases of the microvilli and deeper in the cell. They can also be numerous in the basal portion of the cells where infolding of the membrane is frequent. In a later work Halcrow (1985) investigated Leptodora kindtii. The dorsal (neck) organ occupies a large area of the head. The basal portion of the cells is extensively infolded and interdigitated. Most of the cell is filled with mitochondria with tubular cristae. Microvilli are lacking. Meurice and Goffinet (1983) described a larval dorsal (neck) organ in the marine cladocerans Evadne nordmanni, E. spinifera, E. tergestina and Podon intermedius, where it is situated at the midline on the cephalic shield. Externally, it has a smooth surface surrounded by a cuticular ring. Below this, a dozen cells form a mass where all cells exhibit the same ultrastructural organization. The apical portion of each cell contains a dense bundle of microtubules directed towards the cuticle. Microvilli are lacking. The nucleus, surrounded by a dense cytoplasm containing a few cisternae of rough endoplasmic reticulum, is also apical. The basal portion of the cells is deeply infolded, giving rise to an intricate system of channels separated by numerous mitochondria.
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Fig. 8. This photo gives a frontal view of the dorsoanterior surface of the labrum. Anterior is slightly obliquely toward the top. Two pairs of muscles (M), one pair of gland pores (arrows), and several sensillar pores (arrow heads) are visible in an area lacking LMO cells (center of photo). Surrounding LMO cells form a tight microvillar field with transversely cut microvilli. Scale bar 2 mm.
Embryonic and adult dorsal organs are similar in structure. Lanthanum hydroxide experiments made in the same investigation demonstrated specific deposition in the dorsal organ cuticle suggesting its permeable nature. To this category of non-sensory ‘‘dorsal organs’’ belong the integumental windows of harpacticoid copepods (Hosfeld and Schminke, 1997). These occur on the head-shields as well as on the abdominal somites. The integumental windows of the species Parastenocaris vicesima and Chappuisius inopinus have a smooth, naked cuticle. The organ consists of one cell type, which is characterized by apical invaginations and abundant basal glycogen granules and mitochondria. Between these, a granular or striated structure is intercalated. Hosfeld (1999) reported finding similar structures on adults of the harpacticoids Tachidius disciples and Bryocamptus pygmaeus and for T. discipes
also from the first nauplius. In T. discipes the apical portion of the cells has a rich tubular system and in B. pygmaeus the apical portion of the cone-shaped cells have finger-like invaginations, which together with the ramifying mitochondria form a very distinct pattern. A gland cell opens in a pore anterior to this ‘‘dorsal organ’’. Lake et al. (1974) described the ultrastructure of the fenestra dorsalis of syncarid crustaceans Allanaspides helonomus and Allanaspides hickmani. This is a middorsal organ on the cephalothoracic tergite. This organ has all the characteristics of an ion-transporting epithelium and aligns with those described above. The authors give a schematic drawing (their Fig. 1), which shows both the so-called fenestra dorsalis and the four-celled organ, probably similar to the one described by Laverack et al. (1996) for Anaspides tasmaniae. However, they do not mention the four-celled organ in the text.
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Fig. 9. Semidiagrammatic plan view of the LMO field (shaded areas), essentially as it contacts the dorsal labral cuticle. Anterior is toward the top. The mandibular gnathobase is shown on the left side. The level of the mouth is approximately indicated by the line just anterior to the shaft of the mandible. Note that the tip of the mandible extends into the mouth. Muscle insertions on the labral cuticle are dotted. Sensillar openings are shown as small circles. Two pairs of gland pores (one of which is indicated in Fig. 8) in this region are indicated by arrow heads. The dashed line along three of the LMO cells indicates areas not explored in our survey of the specimen. Scale bar 5 mm.
Here, we have an example of two kinds of ‘‘dorsal organs’’ in the same animal (see below). The so-called ‘‘Haftapparat’’ or anchor organ of the cladoceran Sida crystallina, is an anchoring device, and only a cuticular specialization (Gu¨nzl, 1980). The position on the head has invited it into the realm of ‘‘dorsal organs’’, but it is not an organ and has nothing to do with the two types described above. From this review, it seems safe to conclude on the basis of present ultrastructural knowledge that all the crustacean structures called ‘‘dorsal organs’’ fall, in fact, into two different, unrelated categories, one sensory, the other ion-transporting. We suggest that they be called ‘‘dorsal sensory pit organs’’ and ‘‘dorsal iontransporting complexes’’, respectively. The dorsal sensory pit organ should, as Laverack et al. (1996) and Laverack and Sinclair (1994) described it, be a (chemo)sensory organ innervated from the tritocerebrum (see description above). However, innervation is not established for more than Macrobrachium intermedium and remains to be clarified in all other taxa. Only those organs innervated from the tritocerebrum would then classify as dorsal sensory pit organs. An innervation from e.g. protocerebrum should demand a demarcation from other protocerebral sensory organs and be given another term. The dorsal sensory pit organs are, provided tritocerebral innervation, ‘‘lattice’’, ‘‘sensory dorsal’’ and ‘‘cephalic dorsal hump’’ organs. The dorsal ion-transporting complexes consist of transporting epithelia. Their ultrastructure can vary in details, but clearly they share this same role, as comparisons with other ion-transporting tissues reveal (Conte, 1984). The question of different organs in larvae and adults probably only reflects differing needs during development. Conte referred to the organs as a ‘‘crustacean larval salt gland’’ or in adults ‘‘cephalothoracic organ’’. In the present
presentation these, as well as the ‘‘dorsal nuchal’’, ‘‘dorsal neck’’, ‘‘neck’’, ‘‘integumental windows’’ and ‘‘fenestra dorsalis’’ all belong to this category. The term ‘‘dorsal ion-transporting complex’’ encompasses cephalic, thoracic, single and paired types. The morphology of the mystacocarid ‘‘dorsal organ’’ has the impudence of disallowing membership in the two types of ‘‘dorsal organ’’ reviewed here. At first sight, the microvillous cells suggest a light sensory organ of some sort, but we have failed, in our serial sections, to establish a nervous connection of any kind and thus light cannot be mediated through the nervous system. The lack of a nervous connection eliminates the possibility of this organ being any other kind of peripheral nervous structure as well. In this regard, it should be emphasized that none of the cells of this organ displayed any of the ultrastructural features of endocrine cells. Similarly, it is by no means a transporting epithelium. Microvilli can be part of such an organ but the few small, weak mitochondria in their neighborhood and the lack of membrane infoldings and associated mitochondria speak against such a function. The thin cuticle suggests a connection with the external environment, although not an effective one due to the lack of cuticular pores. The only likely connection of the anterior and posterior cells with the interior of the animal is the large ventral cells whose function is unclear. Finally, in all of our experience with ultrastructure of crustacean organ systems, we have never seen anything like the nuclear vacuole of posterior cells. Thus, for the time being, speculation about the function is unwarranted. The mystacocaridean ‘‘dorsal organ’’ forms a ‘‘dorsal organ’’ group of its own with an enigmatic function. For now we label it the cephalic microvillar organ (CMO). 4. Labral microvillar organ (LMO) 4.1. Introduction In their description of the digestive system in Derocheilocaris remanei, Herrera-Alvarez et al. (1996) briefly noted the presence of microvillar cells in the same position as described here in D. typica, but did not comment on them. As noted below, we were also dismissive of similar cells in our description of the digestive system of the cephalocarid Hutchinsoniella macracantha (Elofsson et al., 1992). As shown by D. typica, the organ formed by these cells deserves greater scrutiny. 4.2. Gross anatomy The labrum of mystacocarids is relatively large compared to that of other crustaceans, being a tenth the body length of the entire organism. The resulting atrium oris envelops the tips of the endites of the mandible and both maxillae. The proximal third of the labrum is closely appressed to the ventral surface of the adjacent cephalon in preserved specimens (it has not been studied in living animals in their natural habitat); more distally, it gradually diverges (Fig. 7). The LMO is located on the dorsal surface of the labrum (Fig. 7). Its outline is roughly oval and is about 22 mm long and 16 mm wide. Anteriorly, it begins just inside the mouth, in the region near the tips of the mandibular incisor process, and extends posteriorly to somewhat past a dorsal transverse fold approximately two-fifths back the length of the labrum. Laterally, it its widest at its midpoint, where it is almost as wide as the labrum. It forms a continuous sheet of microvilli just under the cuticle, except where other organs must contact (muscle insertions) or penetrate (sensillae) the cuticle, and where there is a 4.5 3 mm oval patch of the labral surface just posterior to the mouth (Fig. 8). There are 21 LMO cells on ‘‘Horiz. II’’, the one specimen that was studied in sufficient detail to determine this. Three of these are located midsagittally and are
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Fig. 10. A. This is a sagittal view of a whole LMO cell. The labral cuticle (C) at the atrium oris is at the top. The most prominent organelles of the LMO cells, the microvilli (MV), nucleus (N), and residual body (RB) are seen. Scale bar 1 mm. B. A cross-section of the cuticle (C) overlying the LMO, and the dark material (DM) overlying the microvilli (MV) in an intermolt animal. Scale bar 0.5 mm. C. A similar cross-section as in B but of a premolt individual, showing that the layer of dark material (DM) is sandwiched between the old (top of figure) and new cuticles (C). The tips of microvilli (MV) is seen. Scale bar 0.5 mm.
unpaired; the others are paired (Fig. 9). The layer of LMO cells occupies approximately half the depth of the labrum (Fig. 7). 4.3. Ultrastructure Labral microvillar cells are irregularly columnar (Fig. 10A). Approximately the top two to three-fifths of these cells comprise a tightly packed layer of microvilli. The bases of the microvilli join the rest of the cell in a concave depression in the cell body such that the edge of the cell body rises to a conspicuous degree around the periphery of each cell’s microvillar tuft. Some short microvilli arise from the lip of this ring. The cuticle that overlies the LMO does not seem different from cuticle covering other parts of the atrium oris (Fig. 7). It bears no openings related to these cells. The tips of the microvilli do not always contact the overlying cuticle. Instead, they might be separated from the cuticle by a layer of irregular clots of dark, granular particles (Fig. 10B). Typically, this
layer is also seen throughout the mouth region, including the foregut near the mouth and the dorsal surface of the anterior atrium oris. It may be as thick as the cuticle, approximately 0.3 mm. In premolt specimens, the new cuticle forms under this layer of granular material (Fig. 10C), such that upon ecdysis, this layer would be discarded along with the old cuticle. At this point, the microvilli are in direct contact with the cuticle, and will remain so until a new layer of dark material develops. Occasional columns of fine, dark particles that look like the clotted material beneath the cuticle fill the narrow space between microvilli (Fig. 11A). Individual microvilli are as long as 4.5 mm and are approximately 0.1 mm in diameter. The microvilli of some of the cells have a slightly larger diameter than those of the others (Fig. 8). Some of the microvilli would merge with others before contacting the main body of the cell (Fig. 11A). Microvillar interiors are finely granular, with occasional dark particles. No fibrillar components were detected. The microvilli open directly into the main body of the cell, without any apparent terminal web (Fig. 11A).
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The nucleus tends to reside in the lower part of the cell (Fig. 7). It is irregularly oval and about 2 mm long (Fig. 10A). The nucleolus is peripheral. The nuclear contents are nearly homogeneously granular, some of the granules being very dark. Occasional pores penetrate the nuclear membrane. A continuous layer of rough endoplasmic reticulum (RER) closely surrounds the nucleus (Figs. 10A and 11C). Concentrations of RER occur in many places in the body of the cell. These do not form orderly layers, but rather surround irregular cisternae of varying size. Clusters of free ribosomes are also abundant. Golgi structures are also present (Fig. 11B). Seven mitochondria reside in one cell where we were able to count them. These vary in size, from oval ones that were somewhat smaller than half the diameter of the nucleus to one large, irregularly elongate mitochondrion that wanders through much of the cell. Small vesicles abundantly occupy the cytoplasm of the main body of the cell, particularly in the region near the base of the microvilli (Fig. 11A). They vary in size from 50–80 nm, but the smaller ones are particularly abundant. These have granular contents, and the larger ones may contain smaller vesicles. The smaller vesicles match approximately the size of what might be pinocytotic vesicles at the proximal tips of the microvillar interstices (Fig. 11A). Each cell possesses one very large residual body (Figs. 10A and 11D). In one cell, it measures 2.5 mm – longer than the nucleus. They are oval, but their surfaces are roughly angular. Their contents are darker than the rest of the cell and are heterogeneous. This includes a granular groundmass, and vesicles of varying size, some of which include other vesicles themselves. There are also irregular dark patches and, most conspicuously, large, stacked membrane whorls. There may also be much smaller residual bodies with similar contents. 4.4. Discussion Nervous tissue occurs in the region of the LMO (Elofsson and Hessler, 2005), but no nervous system structures impinge on LMO cells in a meaningful way. This shows the LMO is not a sensory organ. The lack of secretory vesicles indicates that the LMO is not a secretory organ. There are clues, however, that suggest what LMO cells are doing. Abundant small vesicles in the vicinity of the bases of the microvilli suggest active uptake. The presence of the large residual body is clear testimony that byproducts of LMO metabolism must be stored. The size of the residual body shows that this is a major activity. We favor the possibility that the endocytotic mechanism of the LMO cells has a hepatic function, but what material is it processing? Its position and orientation suggest that it works on material found in the proximal atrium oris, i.e. material that is about to be ingested. If so, that material must cross an unbroken cuticular layer, which limits the kind of substance available to it. Masses of dark granules are often seen in microvillar interstitial spaces. Their appearance matches that of the clots of dark, subcutaneous granules that form a layer of varying thickness in the walls of the atrium oris and foregut. This layer is discharged at ecdysis. Therefore, it is unlikely that this material is being taken up by the LMO. Rather, the presence of these dark granules in the microvillar interstices might indicate production by the LMO and
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transport to the subcuticular layer. If so, LMO cells possess both exo- and endocytotic functions. The widespread distribution in the oral-esophageal region of the dark material may be the result of the need for a larger temporary storage area during intermolt periods. Mystacocarids may not be alone in having an LMO. In a description of the digestive system in the cephalocarid Hutchinsoniella macracantha, Elofsson et al. (1992) mention a patch of tissue that is reminiscent of D. typica’s LMO. Fig. 4 of that publication shows a sagittal section of the labral epithelium of H. macracantha that is remarkably similar, but not identical to that of D. macracantha (Fig. 10A). The columnar epithelial cells’ dorsal surface is a dense layer of microvilli. At their distal ends is a layer of dark bodies, which lies just below the amorphous cuticle. There are indications that the dark bodies may be shed during ecdysis. Unfortunately, our old TEM micrographs and notes are insufficient to describe the cephalocarid LMO cells adequately. One cell that could be measured was 15 mm high. A microvillus measured 0.1 mm, distinctly smaller than that of D. typica. The nucleus measures 4 mm. Mitochondria are abundant. No large residual body was seen, but there may be numerous smaller ones. Undoubtedly, some aspects of size and abundance are scaling issues; cephalocarids are an order of magnitude larger than mystacocarids. Elofsson et al. (1992) states that what we here are calling LMO cells are best developed in approximately the same labral area as in D. typica. But they also claim the same kind of epithelial cell is found on the atrial oral surface of the cephalon, in the esophagus, and even in the tip of the mandible. These are the regions where the dark subcuticular bodies are found. From the perspective of our new findings in D. typica, it seems possible that the dark bodies in H. macracantha were not in contact with the shafts of microvilli over all of their anatomical distribution. Figs. 9 and 10 in the cephalocarid publication show portions of the esophagus where the layer of dark bodies is well developed, but where microvillar shafts appear to be absent. This issue can only be resolved by a reinspection of cephalocarid material. Better knowledge of other taxa might allow the LMO to be used in phylogenetic speculation. Assuming the LMOs of the mystacocarids and cephalocarids are homologous, this organ is very old, since the separation of the two clades are thought to date back to the Cambrian (Walossek, 1999). 5. General conclusion Lacking any convincing understanding of what the cephalic and labral microvillar organs are doing, little can be said by way of a general conclusion. The remark in Section 1 that these two organs share some common features refers to the microvilli and the importance of residual bodies. However, surely this does not imply that they have the same function. We hope that future work will cast light on the meaning of these organs. Acknowledgements This work is dependent on the invaluable and skilled technical work of Rita Walle´n. It is also made possible by the generous support of Professor Dan Nilsson. When the animals were collected, Dr. Lauren Mullineaux (Woods Hole Oceanographic Institution)
Fig. 11. This figure shows details from sagittal sections of the organelles of the LMO cells. A is a view from the base of microvilli, showing concentrations of dark granular material (DM) in their interstices and numerous small vesicles (arrow heads) just below the base of the microvilli. Scale bar 0.5 mm. B. Just below the base of the microvilli (MV), Golgi structures (GS) and numerous free ribosomes (black dots) are found. A pinocytotic figure (arrow head) stemming from the base of a microvillus is visible. Scale bar 0.5 mm. C. A slice through the nucleus (N) and adjacent cytoplasm shows irregular RER cisternae (RER), mitochondrion (M), and cell membranes (CM). Scale bar 0.5 mm. D. A cut through the main residual body shows its varied inclusions. Scale bar 0.5 mm.
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kindly offered us her facilities. This work was funded by the U.M. Egenficka Foundation.
References Anderson, D.T., 1973. Embryology and phylogeny in annelids and arthropods. In: Kerkut, G.A. (Ed.), International Series of Monographs in Pure and Applied Biology, Zoology Division, vol. 50. Pergamon Press, Oxford. Barrientos, Y., Laverack, M.S., 1986. The larval crustacean dorsal organ and its relationship to the trilobite median tubercle. Lethaia 19, 309–313. Conte, F.P., 1984. Structure and function of the crustacean larval salt gland. In: Bourne, G.H., Danielli, J.F. (Eds.), International Review of Cytology. Membranes (ed Danielli, J.F.), 91. Academic Press Inc., Orlando, pp. 45–106. Elofsson, R., 1971. The ultrastructure of a chemoreceptor organ in the head of copepod crustaceans. Acta Zoologica 52, 299–315. Elofsson, R., Hessler, R.R., 2005a. The tegumental glands of Derocheilocaris typica (Crustacea, Mystacocarida). Arthropod Structure & Development 34, 125–138. Elofsson, R., Hessler, R.R., 2005b. The tegumental sensory organs and nervous system of Derocheilocaris typica (Crustacea, Mystacocarida). Arthropod Structure & Development 34, 139–152. Elofsson, R., Lake, P.S., 1971. On the cavity receptor organ (X-organ or organ of Bellonci) of Artemia salina (Crustacea: Anostraca). Zeitschrift fu¨r Zellforschung und mikroskopische Anatomie 121, 319–326. Elofsson, R., Hessler, R.R., Hessler, A.Y., 1992. Digestive system of the cephalocarid Hutchinsoniella macracantha. Journal of Crustacean Biology 12, 571–591. Fioroni, P.,1980. Das Dorsalorgan der Arthropoden unter besonderer Berucksichtigung der malacostraken Krebse. Zoologische Jahrbu¨cher. Abteilung Anatomie 104, 425–465. Gu¨nzl, H., 1980. Der Ankerapparat von Sida crystallina (Crustacea, Cladocera) II. Feinbau und Neubildung. Zoomorphologie 95, 149–157. Halcrow, K., 1982. Some ultrastructural features of the nuchal organ of Daphnia magna Straus (Crustacea: Branchiopoda). Canadian Journal of Zoology 60, 1257– 1264. Halcrow, K., 1985. A note on the significance of the neck organ of Leptodora kindtii (Focke) (Crustacea, Cladocera). Canadian Journal of Zoology 63, 738–740. Herrera-Alvarez, L., Ferna´ndez, I., Benito, J., Pardos, F., 1996. Ultrastructure of the labrum and foregut of Derocheilocaris remanei (Crustacea, Mystacocarida). Journal of Morphology 230, 199–217. Hessler, R.R., Elofsson, R., 2007. An excretory organ in the Mystacocarida. Arthropod Structure & Development 36 1711–1181.
Ho¨eg, J.T., Kolbasov, G.A., 2002. Lattice organs in y-cyprids of the Facetotecta and their significance in the phylogeny of the Crustacea Thecostraca. Acta Zoologica 83, 67–79. Ho¨eg, J.T., Hosfeld, B., Jensen, P.G., 1998. TEM studies on the lattice organ of cirriped cypris larvae (Crustacea, Thecostraca, Cirripedia). Zoomorphology 118, 195–205. Hootman, S.R., Conte, F.P., 1975. Functional morphology of the neck organ in Artemia salina nauplii. Journal of Morphology 145, 371–385. Hosfeld, B., 1999. Ultrastructure of ionocytes from osmoregulatory integumental windows of Tachidius discipes and Bryocamptus pygmaeus (Crustacea, Copepoda, Harpacticoida) with remarks on the homology of nonsensory dorsal organs of crustaceans. Acta Zoologica 80, 61–74. Hosfeld, B., Schminke, H.K., 1997. The ultrastructure of ionocytes from osmoregulatory integumental windows of Parastenocaris vicesima (Crustacea, Copepoda, Harpacticoida). Archives fu¨r Hydrobiologie 139, 389–400. Lake, P.S., Swain, R., Ong, J.E., 1974. The ultrastructure of the fenestra dorsalis of the syncarid crustaceans Allanaspides helonomus and Allanaspides hickmani. Zeitschrift fu¨r Zellforschung und Mikroskopische Anatomie 147, 335–351. Laverack, M.S., Sinclair, A., 1994. Innervation of the dorsal organ of the shrimp Macrobrachium intermedium (Decapoda: Natantia). Journal of Crustacean Biology 14, 1–5. Laverack, M.S., Macmillan, D.L., Ritchie, G., Sandow, S.L., 1996. The ultrastructure of the sensory dorsal organ of Crustacea. Crustaceana 69, 636–651. Martin, J.W., Laverack, M.S., 1992. On the distribution of the crustacean dorsal organ. Acta Zoologica 73, 357–368. Meschenmoser, M., 1989. Ultrastucture of the embryonic dorsal organ of Orchestia cavimana (Crustacea, Amphipoda); with a note on localization of chloride and on the change in calcium-deposition before the embryonic moult. Tissue & Cell 21, 431–442. Meurice, J.Cl., Goffinet, G., 1983. Ultrastructural evidence of the ion-transporting role of the adult and larval neck organ of the marine gymnomeran Cladocera (Crustacea, Branchipoda). Cell and Tissue Research 234, 351–363. Mu¨ller, K.J., Walossek, D., 1988. External morphology and larval development of the upper Cambrian maxillipod Bredocaris admirabilis. Fossils and Strata 23, 1–70. Nishida, S., 1989. Distribution, structure and importance of the cephalic dorsal hump, a new sensory organ in calanoid copepods. Marine Biology 101, 173–185. Walossek, D., 1993. The upper Cambrian Rehbachiella and the phylogeny of Brachiopoda and Crustacea. Fossil and Strata 32, 1–202. Walossek, D., 1999. On the Cambrian diversity of Crustacea. In: Schram, F.R., von Vaupel Klein (Eds.), Crustaceans and the Biodiversity Crisis, vol. 1. Brill, Leiden, pp. 3–27.
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Arthropod Structure & Development journal homepage: www.elsevier.com/locate/asd
Two microvillar organs, new to Crustacea, in the Mystacocarida Rolf Elofsson a, *, Robert R. Hessler b a b
¨ gen 3, SE-223 62 Lund, Sweden Department of Cell and Organism Biology, Zoology Building, University of Lund, Helgonava Scripps Institution of Oceanography, La Jolla, CA 92093-0202, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 19 February 2008 Accepted 24 May 2008
The mystacocarid crustacean Derocheilocaris typica has two microvillar organs, one new, the other previously unappreciated in crustacean literature. The first is situated on the head-shield and consists of three pairs of cells: one with microvilli and a ballooned nucleus; one smaller and without special features; the third large and investing the other two and extending down to the foregut. We call this new organ the ‘‘cephalic microvillar organ’’ and discuss the value of the concept ‘‘dorsal organ’’, to which it might have been included. The second organ consists of about 21 cells that cover the proximal part of the dorsal surface of the labrum. The cells are alike, being characterized by an apical field of microvilli and a large residual body. This organ is here called the ‘‘labral microvillar organ’’. Both organs are neither sensory nor secretory and do not qualify for membership in any of the other recognized organ systems. We are unable to deduce their Dero-cheilocaris functions. Ó 2008 Elsevier Ltd. All rights reserved.
Keywords: Crustacea Mystacocarida Dorsal organ Labral organ
1. General introduction In our attempt to study the anatomy of the mystacocarid, Derocheilocaris typica down to an ultrastructural level (Elofsson and Hessler, 2005a,b; Hessler and Elofsson, 2007), we have stumbled upon two organs whose purpose we cannot fathom. In spite of this inability, we feel it is essential to make the existence of these organs known to other investigators, in the hope that someone may be able to show what they are for. Both organs are cephalic. Both have microvillar fields that abut the cuticle. One is located middorsally, and thus qualifies for membership among the ‘‘dorsal organs’’. However, as shown below, as a category, ‘‘dorsal organ’’ is not meaningful. The other organ presented here is located on the atrium oral side of the labrum. Because these two organs constitute separate problems, we describe and discuss them separately. 2. Material and methods Abundant specimens were obtained from Stony Beach at Woods Hole, MA, USA. Sand was sieved through an approximately 60 mm filter from pits dug to a depth of 2–4 dm above high-tide line. They were fixed for 1 h in a refrigerator in either 0.3 M cacodylate buffer or seawater with 2.5% glutaraldehyde added. Postfixation was performed, after washing in buffer or seawater, with 1% osmium
* Corresponding author. Tel.: þ46 46 51040. E-mail addresses: [email protected] (R. Elofsson), [email protected] (R.R. Hessler). 1467-8039/$ – see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.asd.2008.05.002
tetroxide in the same buffer for 1 h. The specimens were embedded in Epon. Thin sections were cut on a LKB Ultrotome and poststained in 2% uranyl acetate and lead citrate in an LKB Ultrastainer. A JEOL transmission microscope was used. For scanning electron microscopy, the specimens were fixed as above and left in sucrose overnight. They were then critical point dried and coated with gold/palladium. Observations were made with a JEOL T330 microscope. 3. Cephalic microvillar organ (CMO) 3.1. Introduction A more or less well defined organ, situated dorsally on the head seems to be a widespread feature in crustaceans. It has been described under a variety of names: dorsal organ, Nackenorgan/neck organ/nucal organ, lattice organ, cephalic dorsal hump, Hafptorgan, salt gland, and integumental window to mention some. As the names suggest, the ideas about function are equally diverse. Martin and Laverack (1992) made a survey of the occurrence of dorsal organs in crustaceans, including their presumed function, the long history of their recognition and their appearance in fossil arthropods. Relevant ultrastructural information is further found in Lake et al. (1974) on syncarid crustaceans, Laverack and Sinclair (1994) on the decapod shrimp Macrobrachium intermedium, Laverack et al. (1996) on the syncarid Anaspides tasmaniae and the decapod shrimp Crangon crangon, and Hosfeld and Schminke (1997) and Hosfeld (1999) on harpacticoid crustaceans. Recent publications have widened the scope to include cirripeds (Ho¨eg et al., 1998) and facetotectans (Ho¨eg and Kolbasov, 2002). The discovery of the
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cephalic microvillar organ (CMO) in Derocheilocaris typica further widens the scope of such organs. 3.2. Results
Fig. 1. A. This scanning electron micrograph of the dorsal head-shield illustrates the external position of the cephalic microvillar organ (CMO) (arrow). Anterior is at the top of the photo where also the hinge in the head-shield is seen. Arrowheads point at four anterior sensory pegs of the head-shield forming a rectangle whose diagonals cross each other approximately where the CMO is situated. Scale bar 10 mm.
A small round area defining the position of the CMO is situated approximately 20 mm behind the hinge of the head-shield on Derocheilocaris typica. Externally, it is located at the crossing of diagonals of the imaginary rectangle formed by the two anterior pairs of pegs of the posterior head-shield (Elofsson and Hessler, 2005b). It is hardly seen from the outside since it appears only as a slightly elevated dome in the cuticle about 2 mm in diameter (Fig. 1). The head cuticle is approximately 0.5 mm thick, but over the organ it is only half of that thickness. There are no pores or other structures in its cuticle. Dorsally, the CMO itself is also small. It is squeezed between strong muscles and occupies an area below the cuticle about 6 mm in diameter. Its position, seen in a midsagittal section, is almost directly above the anterior-most margin of the loop formed by the foregut (Fig. 2). Three pairs of cells form the CMO (Fig. 3). One pair, the posterior cells, has apical microvilli, which are tightly apposed and perpendicular to the cuticle (Fig. 4A and inset). The tips of the microvilli are under the cuticular dome. They are
Fig. 2. A midsagittal section of the anterior portion of the head shows the position of both the cephalic microvillar organ (CMO) and the labral microvillar organ (LMO). The relation to part of the brain (B), foregut (F), and labrum (L) is also evident. Scale bar 10 mm.
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Fig. 3. A schematic drawing, from sections, of the CMO viewed from the side. The relation of the three paired cells, anterior (A), posterior (P) and ventral (V) of the CMO, to one another is given. Their position in the body is illustrated by the inclusion of the posterior portion of the brain (B), and the anterior ascending portion of the foregut (F). One large gland cell (G) represents the position of several gland cells, situated on both sides of the ventral cell (V). The lines numbered 1–5 illustrate the approximate levels at which the micrographs 4–6 are taken (1 ¼ Fig. 4A, 2 ¼ Fig. 4B, 3 ¼ Fig. 5A, 4 ¼ Fig. 5B and 5 ¼ Fig. 6). All Figs. 4–6 are horizontal sections from a section series through the organ from one animal.
approximately 50 nm in diameter and a little more than 1 mm in length (Fig. 4, inset). A non-microvillar border of the cell encircles the microvilli except where the two cells meet each other. The microvilli are thus enclosed in a cup formed by the two cells. Anterior to and below the base of the microvilli, much of the cell volume is filled with a vacuole almost 2 mm in diameter with a fine, even, granular content. This vacuole is formed by an outpocketing of the outer membrane of the nucleus (Figs. 4B and 5A). That same granular material forms an almost continuous layer between the inner and outer nuclear membranes all the way around the nucleus. This layer is continuous with the vacuole, but defined from it by a constriction. The ribosomes of the outer nuclear membrane follow the membrane around the vacuole. Since this structure appears in all the animals we studied, we do not judge it to be an artifact. The nucleus, contained in its inner membrane, is situated in a side or ventral portion of the cell (Figs. 4B and 5A). Its size is less than that of the vacuole. It has one large nucleolus and no heterochromatin. The cytoplasm contains cisternae of rough endoplasmic reticulum, a Golgi apparatus, relatively few small mitochondria (0.5 mm in diameter) with few cristae, and occasional lysosome-like vesicles the size of a mitochondrion. The pair of anterior cells is smaller than the preceding. They lack microvilli and the unusual nuclear vacuole and envelope (Figs. 4B and 5A). They meet the cuticle in front of the cuticular dome and follow the anterior outlines of the posterior cells tightly. The nucleus fills most of the cell. Its cytoplasm is similar to that of the posterior cells. These two cell types, the anterior and posterior cell pair, are intimately associated with a pair of large cells with an irregular outline (Fig. 3). Dorsally, these cells adjoin the previous two pairs anteriorly (Fig. 4B). More deeply, they also invest them along their sides and below. They extend between the previous cell pairs and the foregut, a distance of about 4 mm, narrowing ventrally to form together a thin lamella squeezed in between large lateral glands (Fig. 5B). Anteriorly, the cells meet the brain and end against the
circular muscles of the ascending foregut. The main portion of the cells continues posteriorly along the midline for 15–20 mm, whereupon the cells widen to accommodate the nucleus (Fig. 6). The nucleus (2–4 mm) has one large nucleolus and no heterochromatin. This posterior portion of this pair of cells sits broadly on the circular muscles of the foregut in a region where the foregut has turned posteriorly. We term this cell pair the ventral cells because of the position of the main portion of the cell in relation to the anterior and posterior cells. The cells are light due to only sparse rough endoplasmic reticulum and a loose granular cytoplasm. The anterior portion of the ventral cells houses some mitochondria and occasional dark lysosome-like bodies, both of which increase considerably in number in the posterior portion of the cells (Fig. 6). The size (1–2 mm) and shape of the mitochondria varies. They are characterized by having few and short cristae. The dark bodies vary in shape and size (more than 1 mm). Their content can be evenly dark or contain membrane whorls and dark particles. Often these lysosome-like vesicles are associated with mitochondria. There are no specific membrane specializations in any of the cells. A dark fibrous structure stretches out in intercellular spaces to surround the organ. We interpret this structure as an organ-limiting structure similar to a basal lamina.
3.3. Discussion The term ‘‘dorsal organ’’ has been used here temporarily as a handy name based only on the position of the structure. The term is, however, unsuitable as a morphologically and functionally unified feature and should be abandoned. Usually the ‘‘dorsal organ’’ is described on the basis of its external morphology, for which there is a rich literature, predominantly used for taxonomic work. However, external appearance will not be dealt with here; an ultrastructural description of internal anatomy is mandatory to appreciate the true nature of these organs.
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Fig. 4. A. The most dorsal horizontal section of the series, just below the small cuticular dome of the CMO (level 1 in Fig. 3), cuts the microvilli (MV) of the posterior cells (P). The tip of one anterior cell is emerging (A). Scale bar 2 mm. Inset: A sagittal section through the dome with the microvilli. This animal is ready to moult. Scale bar 0.5 mm. B. A horizontal section below that of A (see also level 2 in Fig. 3) cuts through all three cells of the CMO. Note the crammed position between large muscle cells (M). The ventral cell pair (V) sends a dorsal extension anterior to the anterior cell pair (A). The two posterior cells (P) are cut through the ballooned portion of the nucleus (asterisk). The right posterior cell shows the entire nucleus. Scale bar 1 mm.
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Fig. 5. A. A horizontal section further down in the series below that of Fig. 4B (see also level 3 in Fig. 3). At this level the anterior (A) and the posterior (P) cells are disappearing and the ventral (V) cells dominates the organ. Scale bar 1 mm. B. Next horizontal section is cut somewhat below that of A (see also level 4 in Fig. 3). Here only the ventral (V) cells of the CMO are left. Large gland (G) cells of different appearance and muscles (M) on both sides of the organ squeeze the ventral cell into a thin lamella. The arrow points in the direction of the caudal portion of the ventral cells. Scale bar 2 mm.
The problem of dorsal organ terminology is aggravated by the use of the term for temporary structures in arthropod embryos (Fioroni, 1980) that are said to be sites of cellular degeneration during development. Fioroni realized that there was no unifying
structure and suggested multiple evolutionary events. Anderson (1973) commented on transitional dorsal organs with degenerative function in malacostracans, and on a pair of transitional dorsolateral organs in peracaridean isopods with a different presumed
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Fig. 6. This last horizontal section of the series is taken just above the foregut (F) and shows the posterior portion of the CMO where only the ventral (V) cells are present (see also level 5 in Fig. 3). It is clearly seen here that the ventral cells dip down on both sides of the roof of the foregut. Their nuclei (N) are present in this region, the left nucleus with its nucleolus. Scale bar 2 mm.
function. All these investigations were light microscopical. An ultrastructural investigation of the transitional dorsal organ of Orchestia cavimana (Meschenmoser, 1989) found evidence to believe it to be an ion-transporting organ like the ones discussed below. The ‘‘dorsal organ’’ is not restricted to the head-shield. It can be paired and occur on body segments under the name of integumental windows (e.g. Hosfeld and Schminke, 1997; Hosfeld, 1999). Scanning electron microscopic and light microscopical observations have noted the presence of a ‘‘dorsal organ’’ in the form of four pits on the head-shield in fossil maxillopod (Mu¨ller and Walossek, 1988) and branchiopod (Walossek, 1993) crustaceans. It has also been found in trinucleid trilobites (Barrientos and Laverack, 1986) as well as in recent chelicerates, chilopods, diplopods, and insects (Fiorini, 1980). The present discussion will be restricted to crustaceans. Here, ultrastructural investigations reveal two major types. ‘‘Dorsal organs’’ of the taxon Thecostraca, comprising Ascothoracida, Cirripedia, and Facetotecta, form a distinct group (Ho¨eg et al., 1998; Ho¨eg and Kolbasov, 2002). They are termed lattice organs and appear in five pairs on the head-shield of cypris larvae. It is a sensory organ whose main components are two primary
receptor cells, each furnished with two (or four) modified cilia. The fate of the axons from the receptor cells has not been reported. Two cells sheath the receptor cells. The modified cilia reach into a chamber in the cuticle whose roof has pores. Within the thecostracans, this generalized morphology differs somewhat in details. One or two gland cells are situated between the pairs of the lattice organ. The gland cells have a microvillous border against the cuticle. The morphology of the lattice organ strongly suggests a chemosensory function. Another sensory ‘‘dorsal organ’’ is found in two malacostracan taxa, decapods (Crangon crangon) and syncarids (Anaspides tasmaniae) (Laverack et al., 1996). Here, one circumscribed median area on the head-shield contains four pits. Associated with the pits are four sensory cells, each bearing a modified cilium, which ends at the cuticle. The receptor cells and their cilia are wrapped in three sheath cells insulating the cilia from one another. Centrally in the area of the dorsal organ, a secretory cell opens into a pore of the cuticle. The secretory cell is microvillous and contains numerous vacuoles. Laverack and Sinclair (1994) found with methylene blue studies, that the dorsal organ of the decapod shrimp
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Fig. 7. This exsagittal section through the labrum is given to show the entire LMO. Anterior is to the left. The arrow to the very left indicates the approximate level of the mouth. The microvilli (MV) and nucleus (N) of LMO cells are clearly seen in this magnification. The cephalo-some relation to the atrium oris (AO), mandible (M) and the dorsal transverse fold of the labrum (DTF) is displayed in the photo. Scale bar 5 mm.
Macrobrachium intermedium was innervated from the tritocerebrum. Although Laverack believed the organ to be mechanosensory, the lack of structures associated with this modality make a chemosensory function more probable. Nishida (1989) studied a structure he called the cephalic dorsal hump in calanoid copepods. It was found to be a male specific feature in many calanoids, situated anterodorsally on the cephalosome. An ultrastructural study on Calanus sinicus and Paracalanus parvus found that the organ consists of two gland cells, each opening in a pore, and a receptor, which ends in a thin porous field on the cuticle. They form a row with the receptor being most posterior. The receptor consists of two cells and two enveloping cells. Two cilia from each receptor cell runs through a cavity enveloped by cells having distal microvilli. The cilia branch and end in an unexplained way within the porous cuticle. Nishida claims the organ to be homologous with the one described by Laverack et al. (1996) (see above) e.g. the sensory dorsal organ. However, the innervation, which was not investigated by Nishida, is crucial to the understanding of the organ. If the organ is innervated from the protocerebrum it could be the ‘‘third unit’’ of copepod crustaceans (Elofsson, 1971), similar to the cavity receptor organ of Artemia salina (Elofsson and Lake, 1971); only if it is innervated from the tritocerebrum could it be similar to the dorsal organ described by Laverack et al. (1996). Another type of ‘‘dorsal organ’’ is not innervated and possesses a transporting epithelium. Hootman and Conte (1975) described the ultrastructure of an ion-transporting epithelium of the neck organ in the nauplii of the anostracan Artemia salina. Fifty to 60 cuboidal cells contained many mitochondria associated with apical
and basal membrane infoldings. They found a great resemblance to salt secreting cells of other taxa. Halcrow (1982) investigated the dorsal (nuchal) organ of juvenile Daphnia magna, a cladoceran crustacean, formed as an expanded portion of a dorsal ridge. The organ comprises two types of cells, one darker due to the presence of abundant mitochondria and the other lighter; both bear microvilli on their apical surfaces. The microvilli differ somewhat on the two types of cells. A large number of mitochondria with prominent cristae are situated at the bases of the microvilli and deeper in the cell. They can also be numerous in the basal portion of the cells where infolding of the membrane is frequent. In a later work Halcrow (1985) investigated Leptodora kindtii. The dorsal (neck) organ occupies a large area of the head. The basal portion of the cells is extensively infolded and interdigitated. Most of the cell is filled with mitochondria with tubular cristae. Microvilli are lacking. Meurice and Goffinet (1983) described a larval dorsal (neck) organ in the marine cladocerans Evadne nordmanni, E. spinifera, E. tergestina and Podon intermedius, where it is situated at the midline on the cephalic shield. Externally, it has a smooth surface surrounded by a cuticular ring. Below this, a dozen cells form a mass where all cells exhibit the same ultrastructural organization. The apical portion of each cell contains a dense bundle of microtubules directed towards the cuticle. Microvilli are lacking. The nucleus, surrounded by a dense cytoplasm containing a few cisternae of rough endoplasmic reticulum, is also apical. The basal portion of the cells is deeply infolded, giving rise to an intricate system of channels separated by numerous mitochondria.
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Fig. 8. This photo gives a frontal view of the dorsoanterior surface of the labrum. Anterior is slightly obliquely toward the top. Two pairs of muscles (M), one pair of gland pores (arrows), and several sensillar pores (arrow heads) are visible in an area lacking LMO cells (center of photo). Surrounding LMO cells form a tight microvillar field with transversely cut microvilli. Scale bar 2 mm.
Embryonic and adult dorsal organs are similar in structure. Lanthanum hydroxide experiments made in the same investigation demonstrated specific deposition in the dorsal organ cuticle suggesting its permeable nature. To this category of non-sensory ‘‘dorsal organs’’ belong the integumental windows of harpacticoid copepods (Hosfeld and Schminke, 1997). These occur on the head-shields as well as on the abdominal somites. The integumental windows of the species Parastenocaris vicesima and Chappuisius inopinus have a smooth, naked cuticle. The organ consists of one cell type, which is characterized by apical invaginations and abundant basal glycogen granules and mitochondria. Between these, a granular or striated structure is intercalated. Hosfeld (1999) reported finding similar structures on adults of the harpacticoids Tachidius disciples and Bryocamptus pygmaeus and for T. discipes
also from the first nauplius. In T. discipes the apical portion of the cells has a rich tubular system and in B. pygmaeus the apical portion of the cone-shaped cells have finger-like invaginations, which together with the ramifying mitochondria form a very distinct pattern. A gland cell opens in a pore anterior to this ‘‘dorsal organ’’. Lake et al. (1974) described the ultrastructure of the fenestra dorsalis of syncarid crustaceans Allanaspides helonomus and Allanaspides hickmani. This is a middorsal organ on the cephalothoracic tergite. This organ has all the characteristics of an ion-transporting epithelium and aligns with those described above. The authors give a schematic drawing (their Fig. 1), which shows both the so-called fenestra dorsalis and the four-celled organ, probably similar to the one described by Laverack et al. (1996) for Anaspides tasmaniae. However, they do not mention the four-celled organ in the text.
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Fig. 9. Semidiagrammatic plan view of the LMO field (shaded areas), essentially as it contacts the dorsal labral cuticle. Anterior is toward the top. The mandibular gnathobase is shown on the left side. The level of the mouth is approximately indicated by the line just anterior to the shaft of the mandible. Note that the tip of the mandible extends into the mouth. Muscle insertions on the labral cuticle are dotted. Sensillar openings are shown as small circles. Two pairs of gland pores (one of which is indicated in Fig. 8) in this region are indicated by arrow heads. The dashed line along three of the LMO cells indicates areas not explored in our survey of the specimen. Scale bar 5 mm.
Here, we have an example of two kinds of ‘‘dorsal organs’’ in the same animal (see below). The so-called ‘‘Haftapparat’’ or anchor organ of the cladoceran Sida crystallina, is an anchoring device, and only a cuticular specialization (Gu¨nzl, 1980). The position on the head has invited it into the realm of ‘‘dorsal organs’’, but it is not an organ and has nothing to do with the two types described above. From this review, it seems safe to conclude on the basis of present ultrastructural knowledge that all the crustacean structures called ‘‘dorsal organs’’ fall, in fact, into two different, unrelated categories, one sensory, the other ion-transporting. We suggest that they be called ‘‘dorsal sensory pit organs’’ and ‘‘dorsal iontransporting complexes’’, respectively. The dorsal sensory pit organ should, as Laverack et al. (1996) and Laverack and Sinclair (1994) described it, be a (chemo)sensory organ innervated from the tritocerebrum (see description above). However, innervation is not established for more than Macrobrachium intermedium and remains to be clarified in all other taxa. Only those organs innervated from the tritocerebrum would then classify as dorsal sensory pit organs. An innervation from e.g. protocerebrum should demand a demarcation from other protocerebral sensory organs and be given another term. The dorsal sensory pit organs are, provided tritocerebral innervation, ‘‘lattice’’, ‘‘sensory dorsal’’ and ‘‘cephalic dorsal hump’’ organs. The dorsal ion-transporting complexes consist of transporting epithelia. Their ultrastructure can vary in details, but clearly they share this same role, as comparisons with other ion-transporting tissues reveal (Conte, 1984). The question of different organs in larvae and adults probably only reflects differing needs during development. Conte referred to the organs as a ‘‘crustacean larval salt gland’’ or in adults ‘‘cephalothoracic organ’’. In the present
presentation these, as well as the ‘‘dorsal nuchal’’, ‘‘dorsal neck’’, ‘‘neck’’, ‘‘integumental windows’’ and ‘‘fenestra dorsalis’’ all belong to this category. The term ‘‘dorsal ion-transporting complex’’ encompasses cephalic, thoracic, single and paired types. The morphology of the mystacocarid ‘‘dorsal organ’’ has the impudence of disallowing membership in the two types of ‘‘dorsal organ’’ reviewed here. At first sight, the microvillous cells suggest a light sensory organ of some sort, but we have failed, in our serial sections, to establish a nervous connection of any kind and thus light cannot be mediated through the nervous system. The lack of a nervous connection eliminates the possibility of this organ being any other kind of peripheral nervous structure as well. In this regard, it should be emphasized that none of the cells of this organ displayed any of the ultrastructural features of endocrine cells. Similarly, it is by no means a transporting epithelium. Microvilli can be part of such an organ but the few small, weak mitochondria in their neighborhood and the lack of membrane infoldings and associated mitochondria speak against such a function. The thin cuticle suggests a connection with the external environment, although not an effective one due to the lack of cuticular pores. The only likely connection of the anterior and posterior cells with the interior of the animal is the large ventral cells whose function is unclear. Finally, in all of our experience with ultrastructure of crustacean organ systems, we have never seen anything like the nuclear vacuole of posterior cells. Thus, for the time being, speculation about the function is unwarranted. The mystacocaridean ‘‘dorsal organ’’ forms a ‘‘dorsal organ’’ group of its own with an enigmatic function. For now we label it the cephalic microvillar organ (CMO). 4. Labral microvillar organ (LMO) 4.1. Introduction In their description of the digestive system in Derocheilocaris remanei, Herrera-Alvarez et al. (1996) briefly noted the presence of microvillar cells in the same position as described here in D. typica, but did not comment on them. As noted below, we were also dismissive of similar cells in our description of the digestive system of the cephalocarid Hutchinsoniella macracantha (Elofsson et al., 1992). As shown by D. typica, the organ formed by these cells deserves greater scrutiny. 4.2. Gross anatomy The labrum of mystacocarids is relatively large compared to that of other crustaceans, being a tenth the body length of the entire organism. The resulting atrium oris envelops the tips of the endites of the mandible and both maxillae. The proximal third of the labrum is closely appressed to the ventral surface of the adjacent cephalon in preserved specimens (it has not been studied in living animals in their natural habitat); more distally, it gradually diverges (Fig. 7). The LMO is located on the dorsal surface of the labrum (Fig. 7). Its outline is roughly oval and is about 22 mm long and 16 mm wide. Anteriorly, it begins just inside the mouth, in the region near the tips of the mandibular incisor process, and extends posteriorly to somewhat past a dorsal transverse fold approximately two-fifths back the length of the labrum. Laterally, it its widest at its midpoint, where it is almost as wide as the labrum. It forms a continuous sheet of microvilli just under the cuticle, except where other organs must contact (muscle insertions) or penetrate (sensillae) the cuticle, and where there is a 4.5 3 mm oval patch of the labral surface just posterior to the mouth (Fig. 8). There are 21 LMO cells on ‘‘Horiz. II’’, the one specimen that was studied in sufficient detail to determine this. Three of these are located midsagittally and are
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Fig. 10. A. This is a sagittal view of a whole LMO cell. The labral cuticle (C) at the atrium oris is at the top. The most prominent organelles of the LMO cells, the microvilli (MV), nucleus (N), and residual body (RB) are seen. Scale bar 1 mm. B. A cross-section of the cuticle (C) overlying the LMO, and the dark material (DM) overlying the microvilli (MV) in an intermolt animal. Scale bar 0.5 mm. C. A similar cross-section as in B but of a premolt individual, showing that the layer of dark material (DM) is sandwiched between the old (top of figure) and new cuticles (C). The tips of microvilli (MV) is seen. Scale bar 0.5 mm.
unpaired; the others are paired (Fig. 9). The layer of LMO cells occupies approximately half the depth of the labrum (Fig. 7). 4.3. Ultrastructure Labral microvillar cells are irregularly columnar (Fig. 10A). Approximately the top two to three-fifths of these cells comprise a tightly packed layer of microvilli. The bases of the microvilli join the rest of the cell in a concave depression in the cell body such that the edge of the cell body rises to a conspicuous degree around the periphery of each cell’s microvillar tuft. Some short microvilli arise from the lip of this ring. The cuticle that overlies the LMO does not seem different from cuticle covering other parts of the atrium oris (Fig. 7). It bears no openings related to these cells. The tips of the microvilli do not always contact the overlying cuticle. Instead, they might be separated from the cuticle by a layer of irregular clots of dark, granular particles (Fig. 10B). Typically, this
layer is also seen throughout the mouth region, including the foregut near the mouth and the dorsal surface of the anterior atrium oris. It may be as thick as the cuticle, approximately 0.3 mm. In premolt specimens, the new cuticle forms under this layer of granular material (Fig. 10C), such that upon ecdysis, this layer would be discarded along with the old cuticle. At this point, the microvilli are in direct contact with the cuticle, and will remain so until a new layer of dark material develops. Occasional columns of fine, dark particles that look like the clotted material beneath the cuticle fill the narrow space between microvilli (Fig. 11A). Individual microvilli are as long as 4.5 mm and are approximately 0.1 mm in diameter. The microvilli of some of the cells have a slightly larger diameter than those of the others (Fig. 8). Some of the microvilli would merge with others before contacting the main body of the cell (Fig. 11A). Microvillar interiors are finely granular, with occasional dark particles. No fibrillar components were detected. The microvilli open directly into the main body of the cell, without any apparent terminal web (Fig. 11A).
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The nucleus tends to reside in the lower part of the cell (Fig. 7). It is irregularly oval and about 2 mm long (Fig. 10A). The nucleolus is peripheral. The nuclear contents are nearly homogeneously granular, some of the granules being very dark. Occasional pores penetrate the nuclear membrane. A continuous layer of rough endoplasmic reticulum (RER) closely surrounds the nucleus (Figs. 10A and 11C). Concentrations of RER occur in many places in the body of the cell. These do not form orderly layers, but rather surround irregular cisternae of varying size. Clusters of free ribosomes are also abundant. Golgi structures are also present (Fig. 11B). Seven mitochondria reside in one cell where we were able to count them. These vary in size, from oval ones that were somewhat smaller than half the diameter of the nucleus to one large, irregularly elongate mitochondrion that wanders through much of the cell. Small vesicles abundantly occupy the cytoplasm of the main body of the cell, particularly in the region near the base of the microvilli (Fig. 11A). They vary in size from 50–80 nm, but the smaller ones are particularly abundant. These have granular contents, and the larger ones may contain smaller vesicles. The smaller vesicles match approximately the size of what might be pinocytotic vesicles at the proximal tips of the microvillar interstices (Fig. 11A). Each cell possesses one very large residual body (Figs. 10A and 11D). In one cell, it measures 2.5 mm – longer than the nucleus. They are oval, but their surfaces are roughly angular. Their contents are darker than the rest of the cell and are heterogeneous. This includes a granular groundmass, and vesicles of varying size, some of which include other vesicles themselves. There are also irregular dark patches and, most conspicuously, large, stacked membrane whorls. There may also be much smaller residual bodies with similar contents. 4.4. Discussion Nervous tissue occurs in the region of the LMO (Elofsson and Hessler, 2005), but no nervous system structures impinge on LMO cells in a meaningful way. This shows the LMO is not a sensory organ. The lack of secretory vesicles indicates that the LMO is not a secretory organ. There are clues, however, that suggest what LMO cells are doing. Abundant small vesicles in the vicinity of the bases of the microvilli suggest active uptake. The presence of the large residual body is clear testimony that byproducts of LMO metabolism must be stored. The size of the residual body shows that this is a major activity. We favor the possibility that the endocytotic mechanism of the LMO cells has a hepatic function, but what material is it processing? Its position and orientation suggest that it works on material found in the proximal atrium oris, i.e. material that is about to be ingested. If so, that material must cross an unbroken cuticular layer, which limits the kind of substance available to it. Masses of dark granules are often seen in microvillar interstitial spaces. Their appearance matches that of the clots of dark, subcutaneous granules that form a layer of varying thickness in the walls of the atrium oris and foregut. This layer is discharged at ecdysis. Therefore, it is unlikely that this material is being taken up by the LMO. Rather, the presence of these dark granules in the microvillar interstices might indicate production by the LMO and
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transport to the subcuticular layer. If so, LMO cells possess both exo- and endocytotic functions. The widespread distribution in the oral-esophageal region of the dark material may be the result of the need for a larger temporary storage area during intermolt periods. Mystacocarids may not be alone in having an LMO. In a description of the digestive system in the cephalocarid Hutchinsoniella macracantha, Elofsson et al. (1992) mention a patch of tissue that is reminiscent of D. typica’s LMO. Fig. 4 of that publication shows a sagittal section of the labral epithelium of H. macracantha that is remarkably similar, but not identical to that of D. macracantha (Fig. 10A). The columnar epithelial cells’ dorsal surface is a dense layer of microvilli. At their distal ends is a layer of dark bodies, which lies just below the amorphous cuticle. There are indications that the dark bodies may be shed during ecdysis. Unfortunately, our old TEM micrographs and notes are insufficient to describe the cephalocarid LMO cells adequately. One cell that could be measured was 15 mm high. A microvillus measured 0.1 mm, distinctly smaller than that of D. typica. The nucleus measures 4 mm. Mitochondria are abundant. No large residual body was seen, but there may be numerous smaller ones. Undoubtedly, some aspects of size and abundance are scaling issues; cephalocarids are an order of magnitude larger than mystacocarids. Elofsson et al. (1992) states that what we here are calling LMO cells are best developed in approximately the same labral area as in D. typica. But they also claim the same kind of epithelial cell is found on the atrial oral surface of the cephalon, in the esophagus, and even in the tip of the mandible. These are the regions where the dark subcuticular bodies are found. From the perspective of our new findings in D. typica, it seems possible that the dark bodies in H. macracantha were not in contact with the shafts of microvilli over all of their anatomical distribution. Figs. 9 and 10 in the cephalocarid publication show portions of the esophagus where the layer of dark bodies is well developed, but where microvillar shafts appear to be absent. This issue can only be resolved by a reinspection of cephalocarid material. Better knowledge of other taxa might allow the LMO to be used in phylogenetic speculation. Assuming the LMOs of the mystacocarids and cephalocarids are homologous, this organ is very old, since the separation of the two clades are thought to date back to the Cambrian (Walossek, 1999). 5. General conclusion Lacking any convincing understanding of what the cephalic and labral microvillar organs are doing, little can be said by way of a general conclusion. The remark in Section 1 that these two organs share some common features refers to the microvilli and the importance of residual bodies. However, surely this does not imply that they have the same function. We hope that future work will cast light on the meaning of these organs. Acknowledgements This work is dependent on the invaluable and skilled technical work of Rita Walle´n. It is also made possible by the generous support of Professor Dan Nilsson. When the animals were collected, Dr. Lauren Mullineaux (Woods Hole Oceanographic Institution)
Fig. 11. This figure shows details from sagittal sections of the organelles of the LMO cells. A is a view from the base of microvilli, showing concentrations of dark granular material (DM) in their interstices and numerous small vesicles (arrow heads) just below the base of the microvilli. Scale bar 0.5 mm. B. Just below the base of the microvilli (MV), Golgi structures (GS) and numerous free ribosomes (black dots) are found. A pinocytotic figure (arrow head) stemming from the base of a microvillus is visible. Scale bar 0.5 mm. C. A slice through the nucleus (N) and adjacent cytoplasm shows irregular RER cisternae (RER), mitochondrion (M), and cell membranes (CM). Scale bar 0.5 mm. D. A cut through the main residual body shows its varied inclusions. Scale bar 0.5 mm.
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kindly offered us her facilities. This work was funded by the U.M. Egenficka Foundation.
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