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Organization of the cuticle of an aquatic fly larva
TISSUE @
& CELL
1983 Longman
1983 15 (3) Group
477-488
Ltd
PETER F. CREDLAND
ORGANIZATION OF THE CUTICLE AQUATIC FLY LARVA Key
words:
Cuticle,
ultrastructurc,
ABSTRACT.
The
(up to 5 pm),
readily
newly
moulted
using routine From
methods
and
being cndocuticle.
nematocerous information
dipterans
about
cuticle
techniques. It
also
Insect cuticles play an enormous diversity of roles. In many terrestrial insects the cuticle forms a tough, essentially waterproof, exoskeleton but in the aquatic, apneustic larvae of the midge Chironomus riparius Mg., it is a thin barrier against which the hydrostatic skeleton can operate and through which all respiratory gaseous exchange must take place. Nevertheless all cuticles, except perhaps those which constitute arthrodial membranes or those rich in resilin, can be referred to a common plan comprising a thin, superficial epicuticle lacking chitin, and a thicker procuticle, constructed primarily from proteins and chitin, between the epicuticle and epidermis (Neville, 1975; Filshie, 1980). It has become common practice to divide these cuticles into two loose categories. One of these may be termed ‘soft’ and the other ‘solid’ (Andersen, 1977) or alternatively ‘untanned solid’ and ‘tanned solid’ (Neville, 1975). The essential structural difference between the .-~___-__ Dcpartmcnt
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and acetic acid.
that
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unlike
from
has hcen examined
formamide
it is infcrrcd
solid cuticles,
has a very thin
of this cuticle
is the first
cuticle
from
as a source
of
structure.
Introduction
of London,
shed at pupation.
seem that this exceptionally
or sclerotized
such as blowflies
larvae.
ultrastructurc
of proteins
of the thickness
it would
experimental
Chironomus riprim, The
and cxuvia
the extraction
25%
Therefore,
dipteran
cuticle.
and using established
about
sclerotization
larva of the midge,
instar animals,
plan of tanned
of cyclorrhaphous
proteins.
post-cephalic
and also after
represents
using
final
described,
the conventional account,
apncustic
deformable,
and older
the results
present
aquatic,
Chironomus,
OFAN
University U.K.
1982. 1983. 477
two kinds lies in the absence or presence of an exocuticle, although\ most authors allow that a very thin exocuticle may be present in ‘soft’ cuticles, perhaps being indistinguishable from the overlying epicuticle (Locke, 1974). There are some biochemical data which may support this idea (Andersen, 1977) but few structural or ultrastructural studies of a type which could distinguish between the two categories. On secretion, exo- and endocuticle are indistinguishable in any one location although Neville (1965) has identified the exocuticle as that part of the procuticle secreted between apolysis and ecdysis. Locke (1974) stated that ‘the exocuticle may be defined as the region of the endocuticle adjacent to the epicuticle which is so stabilized that it is not attacked by molting fluid and is left behind with the exuvium’. Conversely, ‘the endocuticle is also, by definition, the labile part of the cuticle which is resorbed at molting’. Using these criteria it should be possible to distinguish the exofrom the endocuticle by examining newly moulted insects or their exuvia, circumventing the problems of identifying them in thin sections of integument (Zacharuk, 1976) or the use of histological techniques such as
CREDLAND
478
Mallory’s triple stain which are of limited value in the study of very thin cuticles. However, thin sections of the exuvia of any larval, holometabolous insects do not appear to have been examined previously. Stabilization of the exocuticle by the process known variously as tanning, hardening or sclerotization (Filshie, 1980) renders the vast majority of its proteins insoluble in water (Hackman, 1974), formamide, which will dissolve hydrogen-bonded proteins leaving those covalently bonded to each other or to chitin (Andersen, 1971, 1976; Andersen and Barrett, 1971) or 1 M acetic acid (Andersen, 1973). In contrast to the exocuticle, ‘few covalent bonds are present in endocuticle to reduce the protein solubility’ (Thompson and Hepburn, 1978). The effect of formamide or acetic acid should therefore produce a reflection of the relative solubilities or sclerotization in different parts of the procuticle. The larval integument of C. riparius has been described previously (Credland, 1978) but no attempt was made to identify an exocuticle at that time. Nobody appears to have examined the cuticle of this or any other larval, nematocerous dipteran insect before or since then, except for the pioneering study by Richards and Anderson (1942) of the mosquito, Culex pipiens, leaving a notable void where this important group of insects is concerned. The objective of the present paper is to expand our knowledge of the larval cuticle of C. riparius and, especially. to determine, using the criteria outlined, whether an exocuticle exists in what is apparently the thinnest cuticle yet observed to cover most of the body of any pterygote insect.
Materials and Methods Larvae of Chironomus riparius were reared in permanent laboratory cultures maintained at about 24°C with a photoperiod of 14 hr each day (Credland. 1973). Third instar larvae were isolated in vials until they moulted. Animals described as ‘young’ were sacrificed within 3 hr of their ecdysis to the fourth instar. Older larvae, about 10 mm in length, were removed from the culture vessels. Only those which exhibited no signs of maturation in their thoracic imaginal discs
were used. These animals had not undergone apolysis but were of somewhat variable age. Exuvia were collected by isolating pharate pupae and removing their exuvia within about 4 hr of ecdysis. Since the exuvium is incompletely shed, but merely displaced posteriorly, the precise time of pupal ecdysis is difficult to assess. Young larvae were immersed directly in 5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.2), the primary fixative used throughout for electron microscopy. The head capsule was removed and the section of the body comprising the seventh to eleventh post-cephalic segments was isolated from each larva. This piece of the animal was then split laterally and the larger organs removed. leaving the integument for further preparative treatment. The seventh to tenth post-cephalic segments of older larvae were isolated initially as a single piece but they were not exposed to a fixative. Instead, they were immersed in 1% potassium tetraborate which has been found to facilitate removal of the epidermis from its overlying cuticle (Andersen. 1974). After 30 min the epidermis was peeled and scraped away, taking great care to avoid excessive damage to the cuticle. The eighth, ninth and tenth segment were then isolated from each other and subjected to one of three procedures. Each was placed in either formamide or 1 M acetic acid or the primary fixative. Segments in the former two reagents were left at room temperature for 24 hr before being washed for at least 4 hr in several changes of distilled water and finally transferred to the primary fixative. Exuvia were rinsed in distilled water. The individual segments that were required were isolated and then subjected to any one of the three procedures employed for segments of older larvae. Regardless of prior treatment, all primary fixation lasted for 3-4 hr at about 4°C. Integuments or cuticles were then washed in several changes of fresh 0.1 M phosphate buffer and postfixed in 1% osmium tetroxide in the same buffer for 1 hr. Dehydration was completed in graded ethanols, the material was rinsed briefly in 1: 2 epoxypropane, and then embedded in TAAB or Spurr’s (1969) resin. This sections. 60-90 nm, were cut. double-stained with uranyl acetate and lead citrate (Reynolds, 1963), and examined in a
LARVAL
CUTICLE
OF CHIRONOMUS
Zeiss 109 or an AEI 6B electron microscope. It should be appreciated that chitin is not stained in this procedure and appears electron-lucent; proteins are stained by heavy metals and therefore appear relatively electron-dense (Rudall, 1967; Hackman, 1974). Larvae of various ages were cut into small pieces, fixed in Bouin’s fluid and routinely processed for histological examination. Paraffin or ester wax sections were stained with Mallory’s triple stain in the usual manner (Pantin, 1946). Results Preliminary observations
For the first few hours following ecdysis the larvae are orange-red in colour throughout their length due to the haemoglobin in their ha;molymph. The head capsule is unpigmented and, like the rest of the cuticle, stains blue with Mallory’s stain. However, the head capsule of older larvae is pigmented, bearing characteristic deep brown patterning on a tan ground colour. The remainder of the cuticle, with the exception of the hooks on the prolegs, is virtually colourless and the body consequently appears red. In these larvae the head capsule and proleg hooks appear yellowish after staining with Mallory’s, but that cuticle closest to the epidermis is frequently red or orange. The cuticle of the post-cephalic segments stains blue except for a thin distal band which is red or refractory. Electron microscopical examination of the cuticle of the eighth, ninth and tenth postcephalic segments reveals that they are all alike and of similar thickness in any individual larva. Therefore, these isolated segments of single larvae, subjected to different experimental treatments, have been used as the units of comparison in the work which is described below. Young larvae
The post-cephalic integument of newly ecdysed, fourth instar larvae is extensively folded both in relation to the underlying tissues and its appearance in older larvae. The cuticle itself is only about 1 pm thick at this stage (Fig. 1). The epicuticle, at least its outer layers, is folded in relation to the underlying procuti-
479
cle throughout the stadium. There is no evidence that this folding is more extensive in young larvae. Although clearly identifiable in these larvae, the epicuticle is less distinctively organized than in older individuals. The inner epicuticle (Weis-Fogh, 1970), sometimes also known as the ‘dense layer’ or protein epicuticle (Locke, 1974; Filshie, 1980), varies in thickness such as that an even face is presented to the procuticle but its outer margin is irregular. Outside the inner epicuticle is a narrow zone within which one can distinguish a denser proximal layer and more electron-lucent distal layer. This disparity in electron density is more pronounced in older larvae where the identity of the layers is more readily resolved (see below). The extensive and irregular folding of the epicuticle results in it appearing to vary greatly in thickness when seen in sectional view. Taking the narrowest view to represent the most perpendicular section, the epicuticle is about 150 nm in total thickness at its widest, of which the inner epicuticle comprises more than 100 nm. It should be noted that in addition to the layers already described, there is occasional evidence of an extremely thin superficial layer. The procuticle is lamellate. Usually two lamellae are visible at the time of ecdysis. Pore canals are absent from post-cephalic cuticle but some small inclusions are present. The precise orientation of the chitin microfibres has not been established but the arcuate pattern of protein assumed to be associated with the chitin suggests an organization of the conventional type consisting of parallel microfibres in helicoidally arranged laminae (Bouligand, 1965; Neville, 1975). More detail of the cuticular components is visible in the cuticles of older larvae. Schmidt’s layer is present but the cuticular material is well ordered within a short distance of the apical plasma membranes of the epidermal cells. Chitin microfibres appear to be organized at or very quickly after secretion. The epidermis is very thin (Credland, 1978) but the cytoplasm of the constituent cells is packed with ribosomes, numerous microtubules and mitochondria. Although the density of organelles, inclusions and secreted cuticle prevents their easy recognition, there are some plaques on the apical plasma membrane as seen in Culpodes
CREDLAND
Fig.
I.
procuticle.
Post-cephalic X5
Fig. 2. Cuticlc (srrowcd) procuticlc;
cuticlc
of a ncwlq
moultcd
final
inatar
larva.
i.c
inner
epicuticle:
P,
I .oOO.
clearly
of the heed capwlc diffcrcntiatc
E. epidermis.
xSI
of :I similar
this cuticle
from
larva.
The wry
smooth
that
covering
the post-cephalic
surface
and pore canals segments.
P.
,tH)O
(Locke and Huie, 1979). The chironomid epidermal cells do not, however, have such distinctive apical microvilli. For purposes of comparison, it should be noted that the cuticle of the head capsule in the same animals is similar to that already described but has a smooth, unfolded epicuticle and pore canals (Fig. 2). Older lurvae The cuticle of older larvae (as defined in Materials and Methods) is always thicker
than that of the comparable segment in young animals in the same instar. The maximum thickness reached in the eighth, ninth or tenth post-cephalic segment is about 5 pm (Fig. 3). Although the epicuticle is still folded in relation to the subjacent procuticle, the entire integument is less folded than that of younger individuals. The epicuticle is clearly differentiated into a number of layers. The nomenclature of each layer of insect epicuticle is not standardized (Filshie, 1980) and it is therefore
LARVAL
CUTICLE
OF CHIRONOMUS
difficult to draw simple parallels between those found in C. riparius and those of other insects. There is an outer epicuticle, using Weis-Fogh’s terminology (1970) which was followed by Neville (1975), which is probably homologous with the cuticulin layer of Lucilia (Filshie, 1970). Distally there is a layer of greater electron lucency which may represent a lipoprotein layer equivalent to the ‘outerepicuticle’of Lucilia(Filshie, 1970). The thin superficial layer which is only occasionally seen in young larvae is usually visible but is fragmentary. It could be evidence of a layer which is substantially removed in the preparation of cuticles for electron microscopy or perhaps represent the incipient dispersal of compounds from another layer. A ‘dense layer’ (Locke, 1961, 1974; Filshie, 1970) or ‘inner epicuticle’ (Weis-Fogh, 1970; Neville, 1975) cannot always be differentiated from the procuticle in these older larvae, but there is invariably a thin electron-dense layer internal to the ‘outer epicuticle’ (sensu Weis-Fogh, 1970). A conventional ‘dense layer’ is most apparent after cuticles have been exposed to formamide or acetic acid. Its thickness varies, for the same reason as that explained in connection with young larvae, from about 50 lo 200 nm. The procuticle is entirely lamellate. Usually all the lamellae are of similar thickness. There is no indication that those nearer the epicuticle and deposited earliest are either thicker or thinner than those secreted later in the stadium. There is, however, a tendency for the more distal lamellae to accept more heavy metal stain than the more proximal ones. A marked discontinuity in the electron density of the lamellae commonly occurs about 0.8-1.0 pm below the epicuticle. The lamellae are composed of microfibres and associated material which appear to follow the arcuate or ‘C’ pattern which is widespread in insect cuticles (Neville, 1975; Locke, 1974). The internal margin of the procuticle, after treatment with potassium tetraborate and removal of the epidermis, is fragmentary or diffuse. It sometimes has a fibrillar appearance. Only in larvae which may be approaching apolysis and in which cuticle deposition may have ceased, does the internal surface of the procuticle appear smooth. In intact integuments a Schmidt’s layer is
usually visible although, as mentioned previously, it is thin, suggesting that the cuticle is rapidly organized into its normal lamellate form. No evidence of pore canals has been found in the cuticle of post-cephalic segments, nor do wax canals appear in association with the epicuticle. Larval exuvia
The exuvium of the segments examined following larval-pupal ecdysis is about 0.8-l pm thick. It consists of the epicuticle, which appears virtually unchanged from its pre-moult condition, and a portion of the procuticle (Fig. 7). The procuticular component is divisible into three zones. Beneath the epicuticle is the broadest zone, 0.6-0.8 pm, which appears homogeneous at low magnification (below about ~50,000) but is actually composed of electron-lucent microfibres, probably chitin, and much flocculent, electron-dense material presumed to be proteinaceous. Internal to this zone is a band which appears fragmentary. It is composed of strands or sheets of material aligned parallel with the cuticle surface and separated by spaces which get progressively wider as the distance from the epicuticle increases. The internal boundary of this band is delimited by the third zone which is a continuous layer of material about 35 nm thick, broken at very few points. This innermost part of the exuvium may represent the ecdysial membrane. The distinct lamellae visible in the procuticle prior to apolysis are not as obvious in the exuvium. Incubation of cuticles in formamide acid
or acetic
Formamide and 1 M acetic acid produce similar changes in the appearance of cuticles of final instar larvae (Figs. 4, 5). The effect of incubating cuticles in either reagent is to produce a significant loss of electron-dense material, protein, from the proximal regions of the procuticle but a minor or negligible loss from the distal procuticle. The consequence of this action is to produce an enhanced contrast between the proximal and distal regions, the discontinuity occurring approximately 0.6-0.7 pm below the epicuticle. It is also noticeable that whole cuticles are thinner after incubation in formamide or
LARVAL
CUTICLE
CHIRONOMUS
OF
483
appear to have an electron-dense core. The majority of the microfibres run parallel with the cuticle surface and, when seen in transverse section, they appear to be associated into sheets by interposed proteins. There is also some evidence of vertical linking of the fibres in different planes. Each ‘C’ in the gross lamellate pattern appears to be a composite structure made up of sections through a number of microfibres when seen at high magnifications (Fig. 12).
acetic acid because of shrinkage, especially in the proximal procuticle. On a few occasions sections have been cut through sites of muscle attachment to the cuticle. In such regions the tonofibrillae or muscle attachment fibres follow an apparently helicoidal path through the cuticle. The proximal procuticle in these areas is much less affected by incubation in formamide or acetic acid and relatively little material is lost (Fig. 6). The distinction between the distal and proximal procuticle seen elsewhere is not apparent at these sites. The two reagents have rather different effects on exuvial cuticle. Both result in some indication of lamellae in the most distal zone of the procuticle although they are more distinct after incubation in 1 M acetic acid which does seem to cause the loss of a little electron-dense material from the layer (Figs. 8, 9). However, the central, fragmentary zone of the procuticle is little affected by acetic acid except for some swelling whereas the structure of this band is totally lost after incubation in formamide. Formamide has little effect on the most distal zone. Treatment with either reagent causes the almost total disruption of the innermost zone of the exuvium. Following immersion in formamide or acetic acid, the size and deployment of chitin microfibres are much more clearly visible (Figs. 10-13). Each microfibre has a diameter of about 3 nm when seen in the negative contrast resulting from its association with proteins (Fig. 13). Occasionally, some of these electron-lucent microfibres
Fig.
3. Cuticle
of an older
layers.
exocuticlc
(exe),
epidermis Fig. contrast
was removed
4. Cuticle between
Fig. 5. Cuticle before marked.
before
of the
fixztion.
same
laarva (exe)
The
contrast
larva
without
exposure
to protein
than proximal
solvents.
ones cndocuticle
The (endo).
distal The
x46,0(H). but
incubated
in I M
and endocuticle
to that shown
between
the
exe
acetic
(endo)
acid before
is greatly
fixation.
enhanced.
The
x46,ooO.
in Figs. 4 and 5, but incubated
in formamide
(NO)
is again
and
endocuticle
(endo)
very
x46.000.
Fig.
6. Cuticle
of a larva,
region
of muscle
attachment.
adjacent
prepared
electron-dense
the cxocuticle of a similar
fixation.
larva
are more
Discussion
The fundamental difference between exocuticle and endocuticle is that the proteins of the former have undergone the process known as sclerotization, which has conferred unique properties upon them and therefore the exocuticle itself. As there is no universally acceptable understanding of the chemical basis of sclerotization, the presence of particular chemical groups in cuticles, or extracts made from them, cannot be regarded as incontrovertible proof that sclerotization has occurred (Andersen, 1979; Hillerton and Vincent, 1979). Furthermore, such demonstations as the presence of ketocatechols in cuticle extracts do not indicate the location of these compounds or their precursors in the original cuticle (Andersen and Barrett, 1971). Analysis of amino acid extracts have also been used to try and provide a biochemical distinction between ‘solid’ and ‘soft’ cuticles (Andersen, 1977). It is argued that cuticles containing relatively more amino acids with polar side chains are
area.
Fragments
incubated
in I M acetic
Far less protein of muscle
(M)
acid before
has been removed
remain
internal
fixation, from
sectioned
this cuticle
to the cuticle.
~13,500.
(CM)
through
a
than the
.’
,-
LARVAL
CUTICLE
OF CHIRONOMUS
‘soft’ and those with fewer are ‘solid’. Unfortunately, it is not always clear whether the designation ‘soft’ or ‘solid’ is made before or after the analysis is undertaken. Therefore, biochemical analyses of cuticles cannot yet provide conclusive evidence of the extent of cuticle sclerotization or a ready distinction between solid and soft cuticles. Examination of the cuticle’s appearance would therefore appear to be the most appropriate means of determining the extent of its stabilization. The use of Mallory’s triple stain suggests that the bulk of the post-cephalic procuticle of C. riparius is not sclerotized. The red staining or refractory nature of the distal margin is, however, suggestive of its impregnation or sclerotization (Andersen, 1977). The extreme narrowness of this zone inhibits its clear resolution with the light microscope and resort to the electron microscope is required for further information. Nevertheless, the contrast between the chemical
nature of the head capsule and post-cephalic cuticle is abundantly clear and there is no doubt that the former is subject to far more extensive stabilization; it is shed with minimal resorption of endocuticle at the moult (Credland, unpublished observations). Ultrastructural examination of the postcephalic cuticle has revealed that a similar thickness of procuticle (about 04-1.0 pm) is secreted pre-ecdysially, is apparently slightly more electron-dense than postecdysial procuticle, and remains in the exuvium. Using the conventional criteria, such cuticle can appropriately be termed exocuticle. Whilst such observations reveal nothing about the chemical nature of the exocuticle, they are strongly suggestive of the fact that the layer is far more stable than the underlying endocuticle. The only explanations for the presence of procuticle in the exuvium depend either on the different chemical natures of the exo- and
Fig. 7. Exuvium of a post-cephalic segment shed at pupation. The procuticular element consists of a distal layer(h) which appears homogeneous at low magnifications, an intermediate fragmentary zone (f) and an internal continuous layer (arrowed). ~65,000. Fig. 8. Similar piece of exuvium but incubated in formamide before fixation. The fragmentary zone (f) and inner continuous layer (arrowed) show signs of further disruption. ~65,000. Fig. 9. An adjacent piece of exuvium to that shown in Fig. 8 but incubated in 1 M acetic acid and not formamide before fixation. Some swelling has occurred, especially in the fragmentary aone (f). rendering the exuvium thicker. ~65,ooO. Figs. 10-13. before fixation,
Areas of post-cephalic cuticlc from larvae, all incubated in protein to show the organization of chitin microfibres in the procuticle.
Fig. 10. Incubated in 1 M acetic acid and snowing the conventional cuticle with the ‘C’ pattern of microfibres. ~44,000. Fig. 11. Incubated in formamide and showing the characteristic of resolving individual microfibres. ~40,000.
lamellate
solvents
appearance
of
pattern but also the difficulty
Fig. 12. A small area of Fig. 11 much enlarged to show the discontinuities within each element of the pattern. Each apparently single microtibre is seen at this magnification to consist of a number of components whose plane varies, probably in accord with a model of sequential laminae laid down in a helicoidal manner. ~132,Otx). Fig. 13. Incubated in 1 M acetic acid and showing microfibres in different planes, each surrounded by a thin protein sheath. Unlike the protein, chitin is electron-htcent as it does not accept a heavy metal stain. When seen in transverse section, the microfibre appears as a lucent disc surrounded by a dense ring (arrowed). ~255,000.
32
486
endocuticles or on the failure of enzymes in moulting fluid to digest the cuticle. The latter could occur because the supply of enzymes is exhausted, they are inactivated or because they simply do not have time to complete the dissolution of the entire procuticle. A simple and rather crude way of assessing which of these explanations may be correct is provided by the use of formamide or 1 M acetic acid. These two reagents are believed to remove from cuticles those proteins which are not sclerotized by the formation of covalent links between themselves or with chitin (Andersen and Barrett, 1971; Andersen, 1973, 1976; Thompson and Hepburn, 1978). Therefore one might anticipate that if a cuticle or exuvium contains many more loosely bound proteins which are electron-dense after heavy metal staining, it will appear significantly more electronlucent if treated with either of the reagents prior to fixation and preparation for electron microscopy. Chitin, because it does not stain with heavy metals, will not markedly affect the images seen although it is normally removed by enzymes in the moulting fluid (Jeuniaux, 1971). Actually, some proteins are inextricably bound to the chitin (Hillerton, 1980; Brine and Austin, 1981) and can be expecLed to remain even after incubation in formamide or acetic acid. Therefore, the essential difference between an unsclerotized cuticle from which certain proteins have been removed with these reagents and an exuvium will lie in the presence of chitin and its protein coat in the former. This is precisely what is seen in the endocuticle of the post-cephalic segments of C. ripari~~ which was rendered substantially electronlucent, in contrast to the exocuticle, after incubation in either formamide or 1 M acetic acid. Thus, the results substantiate the assertion that the majority of the proteins of the endocuticle are not covalently bound to each other, but those in the exocuticle, or at least a large proportion of them, are indeed sclerotized, being bound to each other and/ or the chitin. It is in cuticles from which loosely bound proteins have been extracted that the chitin microfibres can most easily be resolved. Seen effectively in negative contrast, surrounded by their protein coat, their diameter of about 3 nm is typical of the value observed in other insects (Filshie, 1980).
CREDLAND
Their arrangement, particularly in the ‘C’ pattern, conforms with the model proposed by Bouligand (1965) and indicates a structure of helicoidally arranged laminae (Figs. 11, 12). Their size and spacing is such that artefacts due to wetting or preparative techniques pointed out by Rudall and Kenchington (1973) appear to have been avoided although some shrinkage in cuticle thickness (Figs. 4, 5) after protein removal does seem to have occurred as expected (Neville, 1965). The possible argument that could be advanced that the exocuticle is relatively unaffected by formamide or acetic acid because it is masked or protected by the epicuticle and endocuticle is refuted by the exposure of exuvia to the same reagents. Similarly, the questions about the timing and availability of enzymes for dissolution of the exocuticle under normal moulting conditions are also answered. As has been demonstrated, very little electron-dense material is removed by either reagent from exuvia and one may conclude that the protein present in the exocuticle is therefore firmly bound or sclerotized. The correspondence between the thickness of the pre-ecdysial cuticle, the thickness of the procuticle left in the exuvium, and the extractability of the proteins in this layer, as compared with those in the more proximal part of the procuticle, would appear to provide ample justification for the assertion that a sclerotized exocuticle is present in the post-cephalic larval sclerites of Chironomus riparius. Furthermore, it represents about 25% of the maximum thickness of the procuticle except at sites of muscle attachment, which were previously reported to avoid digestion by moulting fluid in larvae of Cuffiphoru (Wolfe, 1954) on the basis of light microscope observations. At such sites, the exocuticle may constitute almost the entire thickness of the cuticle (Fig. 6). It should be noted, however, that no inference can be drawn either about the chemical nature of the sclerotization or the timing of its occurrence from the results presented. One feature which emerges clearly from these results is that the cuticle of C. riparius is not fundamentally different from the solid, tanned cuticle (Neville, 1975) despite it being very thin and ‘soft’ in the sense that it is deformed by slight forces, as used by
LARVAL
CUTICLE
487
OF CHIRONOMUS
Andersen (1977). The thicker cuticles of larval Luciliu (Filshie, 1970), Sarcophaga (Dennell, 1946), and pupal Tenebrio (Locke, 1961) are each reported to lack an exocuticle. However, in at least the first of these cases, Fig. 29 of Filshie’s paper (1970) appears to show a part of the procuticle of the second instar about to be shed in the exuvium. Similarly, illustrations of the Drosophila larva1 cuticle (Mitchell et al., 1971) suggest that about one half of the procuticle of the second instar is lost in the exuvium, a similar proportion to that secreted preecdysially. In itself, such evidence is not enough to prove the existence of an exocuticle but, conversely, if Locke’s (1974) definitions of exo- and endocuticles are accepted and it is sclerotization which prevents dissolution of endocuticle, then it is sufficient to question a simple statement that the cuticle of each larval instar is ‘soft and untanned’ as Filshie (1970) said of Luciliu. Whilst it would be most unwise to argue that all cuticles may have an exocuticle because ‘there is an inherent danger in attempting to formulate generalizations based largely on circumstantial evidence’ (Filshie, 1980), one might question whether any cuticle has ever been irrefutably shown to lack one. It should be appreciated that all the
conclusions reached as a result of this work are based on the defnitions and descriptive statements concerning cuticles cited previously. Evidence for them may therefore be regarded as circumstantial. However, until the chemical basis of sclerotization has been explained satisfactorily and a means of identifying the sclerotized proteins in situ has been produced, any distinction between exo- and endocuticle can only be made by deduction of the properties which sclerotization is presumed to confer. On a functional basis (Neville, 1975), there can be no doubt that the extremely thin cuticle of C. ripurius larvae does have an exocuticle whose behaviour precisely corresponds with that in other, larger insects with thicker cuticles. In itself, such a fact exposes the real problems of considering ‘soft’ cuticles as uniquely different from ‘solid’ ones, and highlights the dangers of thinking of thin larval cuticles as being distinct from thicker ones without a critical experimental approach being allied with straightforward observations. Acknowledgements I am grateful to Graham Lawes, Tony King
and Kevin Jennings for their technical assistance.
References Andersen, S. 0. 1971. R&in. Comprehensive Biochemkfry (eds. M. Florkin and E. H. Stotz), Vol. 26C, pp. 633-657. Elsevier, Amsterdam. Andersen. S. 0. 1973. Comparison between the sclerotization of adult and larval cuticle in Schtirocerca gregaria. J. Insect fhysiol., 19, 1603- 1614. Andersen, S. 0. 1974. Cuticular sclerotization in larval and adult locusts, Schistocerca gregaria. 1. Insect Physiol., 20, 1537-1552. Andersen. S. 0. 1976. Cuticular enzymes and sclerotization in insects. TheInsect Inregumenr (ed. H. R. Hepburn), pp. l21- 144. Elscvicr, Amsterdam. Andersen, S. 0. 1977. Arthropod cuticles: their composition, properties and functions. Symp. zooi. Sot. Land., 39, 7-32. Andersen, S. 0. 1979. Biochemistry of insect cuticle. A. Rev. Em., 24, 29-61. Andersen, S. 0. and Barrett, F. M. 1971. The isolation of ketocatechols from insect cuticle and their possible role in sclerotization. J. Insect fhysiol., 17, 69-83. Bouligand, Y. 1965. Sur une architecture torsadee rCpandue dans de nombreuses cuticles d’arthropodes. C. r. hebd. SPanc. Acud. Sci., Paris, 261, 3665-3668. Brine, C. J. and Austin, P. R. 1981. Chitin isolates: species variation in residual amino acids. Camp. Biochem. PhysioL, 708, 173-178. Credland, P. F. 1973. A new method for establishing a permanent laboratory culture of Chironomur riparius Meigen (Diptera: Chironomidae). Freshwat. Biol., 3, 45-51.
CREDLAND
488
Credland, P. F. 1978. An ultrastructural study of the larval integument of the midge, Chironomus ripariur Meigen (Diptera: Chironomidae). Cell Tiss. Res., 186, 327-335. Dennell, R. 1946. The study of an insect cuticle: the larval cuticle of Sarcophaga folculato Pand. (Diptera). Proc. R. SOL, 133B, 348-373. Filshie, B. K. 1970. The fine structure and deposition of the larval cuticle of the sheep blowfly (Lucilia cuprina). Tissue & Cell, 2, 478-498. Filshie, B. K. 1980. Insect cuticle through the electron microscope-distinguishing fact from artifact. Inrec! Biology in the Future, ‘VBW 80’ (eds. M. Locke and D. S. Smith), pp. 59-77. Academic Press. New York. Hackman, R. H. 1974. Chemistry of the insect cuticle. The Physio/ogy of fnrecta (ed. M. Rockstein), 2nd edn, Vol6, pp. 215-270. Academic Press, New York. Hillerton, J. E. 1980. Electron microscopy of fibril-matrix interactions in a natural composite, insect cuticle. J. mater. Sci., l&3109-3112. Hillerton, J. E. and Vincent, .I. F. V. 1979. The stabilization of insect cuticles. J. fnrect Physiol., 25, 957-963. Jeuniaux, C. 1971. Chitinous structures. Comprehemive Biochemistry (eds. M. FIorkin and E. H. Stotz). Vol. 26C, pp. 595-632. Elsevier. Amsterdam. Locke, M. 1961. Pore canals and related structures in insect cuticle. .I. biophys. biochem. Cytol.. 10, 589-618. Locke, M. 1974. The structure and formation of the integument in insects. The Physiology of Insecta (ed. M. Rockstein), 2nd edn, Vol. 6, pp. 123-213. Academic Press. New York. Locke, M. and Huie, P. 1979. Apolysis and the turnover of plasma membrane plaques during cuticle formation m an insect. Tissue & Cell, 11, 277-291. Mitchell, H. K., Weber-Tracy, U. M. and Schaar, G. 1971. Aspects of cuticle formation in Drosophila melanogaster. J. exp. Zool., 116, 429-444.
Neville, A. C. 1965. Chitin lamellogenesis in locust cuticle. Q. .I1Microsc. Sci., 106.269-286. Neville, A. C. 1975. Biology of the Arthropod Cuticle. Springer-Verlag, New York. Pantin, C. F. A. 1946. Notes on Microscopical Technique for Zoologists. University Press, Cambridge. Reynolds, E. S. 1963. The use of lead citrate at high pH as an electron opaque stain in electron microscopy. J. Cell Biol., 17, 208-212. Richards. A. G. and Anderson, T. F. 1942. Electron microscope studies of insect cuticle, with a discussion of the application of electron optics to this problem. .I. Morph., 71, 135-183. Rudall, K. M. 1967. Conformation in chitin-protein complexes. Conformation in BiopoIymers (ed. G. N. Ramachandran), Vol. 2, pp. 751-765. Academic Press, New York. Rudall, K. M. and Kenchington, W. 1973. The chitin system. Rio/. Rev., 49, 597-636. Spurr, A. R. 1969. A low-viscosity epoxy resin embedding medium for electron microscopy. 1. U[trastruct. Res.. 26, 31-48.
P. R. and Hepburn, H. R. 1978. Changes in chemical and mechanical properties of honeybee (Apu mellifera adansonii) cuticle during development. J. camp. Physiol., 126, 257-262. Weis-Fogh, T. 1970. Structure and formation of insect cuticle. Insect (Ilrrastructure (ed. A. C. Ncville), Symp. R. em
Thompson,
Sot. Land., No. 5, pp. 165-185. Blackwell, Oxford. Wolfe, L. S. 1954. The deposition of the third instar larval cuticle of Calliphora eryrhrocephala. Q. JI microsc. Sci., 9.5, 49-66. Zacharuk, R. Y. 1976. Structural changes of the cuticle associated with moulting. The Insect fnregument (ed. H. R. Hepburn), pp. 299-321. Elsevier, Amsterdam.
& CELL
1983 Longman
1983 15 (3) Group
477-488
Ltd
PETER F. CREDLAND
ORGANIZATION OF THE CUTICLE AQUATIC FLY LARVA Key
words:
Cuticle,
ultrastructurc,
ABSTRACT.
The
(up to 5 pm),
readily
newly
moulted
using routine From
methods
and
being cndocuticle.
nematocerous information
dipterans
about
cuticle
techniques. It
also
Insect cuticles play an enormous diversity of roles. In many terrestrial insects the cuticle forms a tough, essentially waterproof, exoskeleton but in the aquatic, apneustic larvae of the midge Chironomus riparius Mg., it is a thin barrier against which the hydrostatic skeleton can operate and through which all respiratory gaseous exchange must take place. Nevertheless all cuticles, except perhaps those which constitute arthrodial membranes or those rich in resilin, can be referred to a common plan comprising a thin, superficial epicuticle lacking chitin, and a thicker procuticle, constructed primarily from proteins and chitin, between the epicuticle and epidermis (Neville, 1975; Filshie, 1980). It has become common practice to divide these cuticles into two loose categories. One of these may be termed ‘soft’ and the other ‘solid’ (Andersen, 1977) or alternatively ‘untanned solid’ and ‘tanned solid’ (Neville, 1975). The essential structural difference between the .-~___-__ Dcpartmcnt
Rcccivcd Rcviscd
of Zoology, Regent’s
Park.
23 September Ih February
Bedford London
College, NW1
4NS.
criteria,
with
of the mature
of
or other the
indicates
fine the
value
of
an cxocuticle
cuticle
insects. of
the
conforms
to
in the larvae
This
sectioned cxuvium
is
the remainder
those rcportcd
soft-bodied structure
and acetic acid.
that
procuticle. dclicatc
unlike
from
has hcen examined
formamide
it is infcrrcd
solid cuticles,
has a very thin
of this cuticle
is the first
cuticle
from
as a source
of
structure.
Introduction
of London,
shed at pupation.
seem that this exceptionally
or sclerotized
such as blowflies
larvae.
ultrastructurc
of proteins
of the thickness
it would
experimental
Chironomus riprim, The
and cxuvia
the extraction
25%
Therefore,
dipteran
cuticle.
and using established
about
sclerotization
larva of the midge,
instar animals,
plan of tanned
of cyclorrhaphous
proteins.
post-cephalic
and also after
represents
using
final
described,
the conventional account,
apncustic
deformable,
and older
the results
present
aquatic,
Chironomus,
OFAN
University U.K.
1982. 1983. 477
two kinds lies in the absence or presence of an exocuticle, although\ most authors allow that a very thin exocuticle may be present in ‘soft’ cuticles, perhaps being indistinguishable from the overlying epicuticle (Locke, 1974). There are some biochemical data which may support this idea (Andersen, 1977) but few structural or ultrastructural studies of a type which could distinguish between the two categories. On secretion, exo- and endocuticle are indistinguishable in any one location although Neville (1965) has identified the exocuticle as that part of the procuticle secreted between apolysis and ecdysis. Locke (1974) stated that ‘the exocuticle may be defined as the region of the endocuticle adjacent to the epicuticle which is so stabilized that it is not attacked by molting fluid and is left behind with the exuvium’. Conversely, ‘the endocuticle is also, by definition, the labile part of the cuticle which is resorbed at molting’. Using these criteria it should be possible to distinguish the exofrom the endocuticle by examining newly moulted insects or their exuvia, circumventing the problems of identifying them in thin sections of integument (Zacharuk, 1976) or the use of histological techniques such as
CREDLAND
478
Mallory’s triple stain which are of limited value in the study of very thin cuticles. However, thin sections of the exuvia of any larval, holometabolous insects do not appear to have been examined previously. Stabilization of the exocuticle by the process known variously as tanning, hardening or sclerotization (Filshie, 1980) renders the vast majority of its proteins insoluble in water (Hackman, 1974), formamide, which will dissolve hydrogen-bonded proteins leaving those covalently bonded to each other or to chitin (Andersen, 1971, 1976; Andersen and Barrett, 1971) or 1 M acetic acid (Andersen, 1973). In contrast to the exocuticle, ‘few covalent bonds are present in endocuticle to reduce the protein solubility’ (Thompson and Hepburn, 1978). The effect of formamide or acetic acid should therefore produce a reflection of the relative solubilities or sclerotization in different parts of the procuticle. The larval integument of C. riparius has been described previously (Credland, 1978) but no attempt was made to identify an exocuticle at that time. Nobody appears to have examined the cuticle of this or any other larval, nematocerous dipteran insect before or since then, except for the pioneering study by Richards and Anderson (1942) of the mosquito, Culex pipiens, leaving a notable void where this important group of insects is concerned. The objective of the present paper is to expand our knowledge of the larval cuticle of C. riparius and, especially. to determine, using the criteria outlined, whether an exocuticle exists in what is apparently the thinnest cuticle yet observed to cover most of the body of any pterygote insect.
Materials and Methods Larvae of Chironomus riparius were reared in permanent laboratory cultures maintained at about 24°C with a photoperiod of 14 hr each day (Credland. 1973). Third instar larvae were isolated in vials until they moulted. Animals described as ‘young’ were sacrificed within 3 hr of their ecdysis to the fourth instar. Older larvae, about 10 mm in length, were removed from the culture vessels. Only those which exhibited no signs of maturation in their thoracic imaginal discs
were used. These animals had not undergone apolysis but were of somewhat variable age. Exuvia were collected by isolating pharate pupae and removing their exuvia within about 4 hr of ecdysis. Since the exuvium is incompletely shed, but merely displaced posteriorly, the precise time of pupal ecdysis is difficult to assess. Young larvae were immersed directly in 5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.2), the primary fixative used throughout for electron microscopy. The head capsule was removed and the section of the body comprising the seventh to eleventh post-cephalic segments was isolated from each larva. This piece of the animal was then split laterally and the larger organs removed. leaving the integument for further preparative treatment. The seventh to tenth post-cephalic segments of older larvae were isolated initially as a single piece but they were not exposed to a fixative. Instead, they were immersed in 1% potassium tetraborate which has been found to facilitate removal of the epidermis from its overlying cuticle (Andersen. 1974). After 30 min the epidermis was peeled and scraped away, taking great care to avoid excessive damage to the cuticle. The eighth, ninth and tenth segment were then isolated from each other and subjected to one of three procedures. Each was placed in either formamide or 1 M acetic acid or the primary fixative. Segments in the former two reagents were left at room temperature for 24 hr before being washed for at least 4 hr in several changes of distilled water and finally transferred to the primary fixative. Exuvia were rinsed in distilled water. The individual segments that were required were isolated and then subjected to any one of the three procedures employed for segments of older larvae. Regardless of prior treatment, all primary fixation lasted for 3-4 hr at about 4°C. Integuments or cuticles were then washed in several changes of fresh 0.1 M phosphate buffer and postfixed in 1% osmium tetroxide in the same buffer for 1 hr. Dehydration was completed in graded ethanols, the material was rinsed briefly in 1: 2 epoxypropane, and then embedded in TAAB or Spurr’s (1969) resin. This sections. 60-90 nm, were cut. double-stained with uranyl acetate and lead citrate (Reynolds, 1963), and examined in a
LARVAL
CUTICLE
OF CHIRONOMUS
Zeiss 109 or an AEI 6B electron microscope. It should be appreciated that chitin is not stained in this procedure and appears electron-lucent; proteins are stained by heavy metals and therefore appear relatively electron-dense (Rudall, 1967; Hackman, 1974). Larvae of various ages were cut into small pieces, fixed in Bouin’s fluid and routinely processed for histological examination. Paraffin or ester wax sections were stained with Mallory’s triple stain in the usual manner (Pantin, 1946). Results Preliminary observations
For the first few hours following ecdysis the larvae are orange-red in colour throughout their length due to the haemoglobin in their ha;molymph. The head capsule is unpigmented and, like the rest of the cuticle, stains blue with Mallory’s stain. However, the head capsule of older larvae is pigmented, bearing characteristic deep brown patterning on a tan ground colour. The remainder of the cuticle, with the exception of the hooks on the prolegs, is virtually colourless and the body consequently appears red. In these larvae the head capsule and proleg hooks appear yellowish after staining with Mallory’s, but that cuticle closest to the epidermis is frequently red or orange. The cuticle of the post-cephalic segments stains blue except for a thin distal band which is red or refractory. Electron microscopical examination of the cuticle of the eighth, ninth and tenth postcephalic segments reveals that they are all alike and of similar thickness in any individual larva. Therefore, these isolated segments of single larvae, subjected to different experimental treatments, have been used as the units of comparison in the work which is described below. Young larvae
The post-cephalic integument of newly ecdysed, fourth instar larvae is extensively folded both in relation to the underlying tissues and its appearance in older larvae. The cuticle itself is only about 1 pm thick at this stage (Fig. 1). The epicuticle, at least its outer layers, is folded in relation to the underlying procuti-
479
cle throughout the stadium. There is no evidence that this folding is more extensive in young larvae. Although clearly identifiable in these larvae, the epicuticle is less distinctively organized than in older individuals. The inner epicuticle (Weis-Fogh, 1970), sometimes also known as the ‘dense layer’ or protein epicuticle (Locke, 1974; Filshie, 1980), varies in thickness such as that an even face is presented to the procuticle but its outer margin is irregular. Outside the inner epicuticle is a narrow zone within which one can distinguish a denser proximal layer and more electron-lucent distal layer. This disparity in electron density is more pronounced in older larvae where the identity of the layers is more readily resolved (see below). The extensive and irregular folding of the epicuticle results in it appearing to vary greatly in thickness when seen in sectional view. Taking the narrowest view to represent the most perpendicular section, the epicuticle is about 150 nm in total thickness at its widest, of which the inner epicuticle comprises more than 100 nm. It should be noted that in addition to the layers already described, there is occasional evidence of an extremely thin superficial layer. The procuticle is lamellate. Usually two lamellae are visible at the time of ecdysis. Pore canals are absent from post-cephalic cuticle but some small inclusions are present. The precise orientation of the chitin microfibres has not been established but the arcuate pattern of protein assumed to be associated with the chitin suggests an organization of the conventional type consisting of parallel microfibres in helicoidally arranged laminae (Bouligand, 1965; Neville, 1975). More detail of the cuticular components is visible in the cuticles of older larvae. Schmidt’s layer is present but the cuticular material is well ordered within a short distance of the apical plasma membranes of the epidermal cells. Chitin microfibres appear to be organized at or very quickly after secretion. The epidermis is very thin (Credland, 1978) but the cytoplasm of the constituent cells is packed with ribosomes, numerous microtubules and mitochondria. Although the density of organelles, inclusions and secreted cuticle prevents their easy recognition, there are some plaques on the apical plasma membrane as seen in Culpodes
CREDLAND
Fig.
I.
procuticle.
Post-cephalic X5
Fig. 2. Cuticlc (srrowcd) procuticlc;
cuticlc
of a ncwlq
moultcd
final
inatar
larva.
i.c
inner
epicuticle:
P,
I .oOO.
clearly
of the heed capwlc diffcrcntiatc
E. epidermis.
xSI
of :I similar
this cuticle
from
larva.
The wry
smooth
that
covering
the post-cephalic
surface
and pore canals segments.
P.
,tH)O
(Locke and Huie, 1979). The chironomid epidermal cells do not, however, have such distinctive apical microvilli. For purposes of comparison, it should be noted that the cuticle of the head capsule in the same animals is similar to that already described but has a smooth, unfolded epicuticle and pore canals (Fig. 2). Older lurvae The cuticle of older larvae (as defined in Materials and Methods) is always thicker
than that of the comparable segment in young animals in the same instar. The maximum thickness reached in the eighth, ninth or tenth post-cephalic segment is about 5 pm (Fig. 3). Although the epicuticle is still folded in relation to the subjacent procuticle, the entire integument is less folded than that of younger individuals. The epicuticle is clearly differentiated into a number of layers. The nomenclature of each layer of insect epicuticle is not standardized (Filshie, 1980) and it is therefore
LARVAL
CUTICLE
OF CHIRONOMUS
difficult to draw simple parallels between those found in C. riparius and those of other insects. There is an outer epicuticle, using Weis-Fogh’s terminology (1970) which was followed by Neville (1975), which is probably homologous with the cuticulin layer of Lucilia (Filshie, 1970). Distally there is a layer of greater electron lucency which may represent a lipoprotein layer equivalent to the ‘outerepicuticle’of Lucilia(Filshie, 1970). The thin superficial layer which is only occasionally seen in young larvae is usually visible but is fragmentary. It could be evidence of a layer which is substantially removed in the preparation of cuticles for electron microscopy or perhaps represent the incipient dispersal of compounds from another layer. A ‘dense layer’ (Locke, 1961, 1974; Filshie, 1970) or ‘inner epicuticle’ (Weis-Fogh, 1970; Neville, 1975) cannot always be differentiated from the procuticle in these older larvae, but there is invariably a thin electron-dense layer internal to the ‘outer epicuticle’ (sensu Weis-Fogh, 1970). A conventional ‘dense layer’ is most apparent after cuticles have been exposed to formamide or acetic acid. Its thickness varies, for the same reason as that explained in connection with young larvae, from about 50 lo 200 nm. The procuticle is entirely lamellate. Usually all the lamellae are of similar thickness. There is no indication that those nearer the epicuticle and deposited earliest are either thicker or thinner than those secreted later in the stadium. There is, however, a tendency for the more distal lamellae to accept more heavy metal stain than the more proximal ones. A marked discontinuity in the electron density of the lamellae commonly occurs about 0.8-1.0 pm below the epicuticle. The lamellae are composed of microfibres and associated material which appear to follow the arcuate or ‘C’ pattern which is widespread in insect cuticles (Neville, 1975; Locke, 1974). The internal margin of the procuticle, after treatment with potassium tetraborate and removal of the epidermis, is fragmentary or diffuse. It sometimes has a fibrillar appearance. Only in larvae which may be approaching apolysis and in which cuticle deposition may have ceased, does the internal surface of the procuticle appear smooth. In intact integuments a Schmidt’s layer is
usually visible although, as mentioned previously, it is thin, suggesting that the cuticle is rapidly organized into its normal lamellate form. No evidence of pore canals has been found in the cuticle of post-cephalic segments, nor do wax canals appear in association with the epicuticle. Larval exuvia
The exuvium of the segments examined following larval-pupal ecdysis is about 0.8-l pm thick. It consists of the epicuticle, which appears virtually unchanged from its pre-moult condition, and a portion of the procuticle (Fig. 7). The procuticular component is divisible into three zones. Beneath the epicuticle is the broadest zone, 0.6-0.8 pm, which appears homogeneous at low magnification (below about ~50,000) but is actually composed of electron-lucent microfibres, probably chitin, and much flocculent, electron-dense material presumed to be proteinaceous. Internal to this zone is a band which appears fragmentary. It is composed of strands or sheets of material aligned parallel with the cuticle surface and separated by spaces which get progressively wider as the distance from the epicuticle increases. The internal boundary of this band is delimited by the third zone which is a continuous layer of material about 35 nm thick, broken at very few points. This innermost part of the exuvium may represent the ecdysial membrane. The distinct lamellae visible in the procuticle prior to apolysis are not as obvious in the exuvium. Incubation of cuticles in formamide acid
or acetic
Formamide and 1 M acetic acid produce similar changes in the appearance of cuticles of final instar larvae (Figs. 4, 5). The effect of incubating cuticles in either reagent is to produce a significant loss of electron-dense material, protein, from the proximal regions of the procuticle but a minor or negligible loss from the distal procuticle. The consequence of this action is to produce an enhanced contrast between the proximal and distal regions, the discontinuity occurring approximately 0.6-0.7 pm below the epicuticle. It is also noticeable that whole cuticles are thinner after incubation in formamide or
LARVAL
CUTICLE
CHIRONOMUS
OF
483
appear to have an electron-dense core. The majority of the microfibres run parallel with the cuticle surface and, when seen in transverse section, they appear to be associated into sheets by interposed proteins. There is also some evidence of vertical linking of the fibres in different planes. Each ‘C’ in the gross lamellate pattern appears to be a composite structure made up of sections through a number of microfibres when seen at high magnifications (Fig. 12).
acetic acid because of shrinkage, especially in the proximal procuticle. On a few occasions sections have been cut through sites of muscle attachment to the cuticle. In such regions the tonofibrillae or muscle attachment fibres follow an apparently helicoidal path through the cuticle. The proximal procuticle in these areas is much less affected by incubation in formamide or acetic acid and relatively little material is lost (Fig. 6). The distinction between the distal and proximal procuticle seen elsewhere is not apparent at these sites. The two reagents have rather different effects on exuvial cuticle. Both result in some indication of lamellae in the most distal zone of the procuticle although they are more distinct after incubation in 1 M acetic acid which does seem to cause the loss of a little electron-dense material from the layer (Figs. 8, 9). However, the central, fragmentary zone of the procuticle is little affected by acetic acid except for some swelling whereas the structure of this band is totally lost after incubation in formamide. Formamide has little effect on the most distal zone. Treatment with either reagent causes the almost total disruption of the innermost zone of the exuvium. Following immersion in formamide or acetic acid, the size and deployment of chitin microfibres are much more clearly visible (Figs. 10-13). Each microfibre has a diameter of about 3 nm when seen in the negative contrast resulting from its association with proteins (Fig. 13). Occasionally, some of these electron-lucent microfibres
Fig.
3. Cuticle
of an older
layers.
exocuticlc
(exe),
epidermis Fig. contrast
was removed
4. Cuticle between
Fig. 5. Cuticle before marked.
before
of the
fixztion.
same
laarva (exe)
The
contrast
larva
without
exposure
to protein
than proximal
solvents.
ones cndocuticle
The (endo).
distal The
x46,0(H). but
incubated
in I M
and endocuticle
to that shown
between
the
exe
acetic
(endo)
acid before
is greatly
fixation.
enhanced.
The
x46,ooO.
in Figs. 4 and 5, but incubated
in formamide
(NO)
is again
and
endocuticle
(endo)
very
x46.000.
Fig.
6. Cuticle
of a larva,
region
of muscle
attachment.
adjacent
prepared
electron-dense
the cxocuticle of a similar
fixation.
larva
are more
Discussion
The fundamental difference between exocuticle and endocuticle is that the proteins of the former have undergone the process known as sclerotization, which has conferred unique properties upon them and therefore the exocuticle itself. As there is no universally acceptable understanding of the chemical basis of sclerotization, the presence of particular chemical groups in cuticles, or extracts made from them, cannot be regarded as incontrovertible proof that sclerotization has occurred (Andersen, 1979; Hillerton and Vincent, 1979). Furthermore, such demonstations as the presence of ketocatechols in cuticle extracts do not indicate the location of these compounds or their precursors in the original cuticle (Andersen and Barrett, 1971). Analysis of amino acid extracts have also been used to try and provide a biochemical distinction between ‘solid’ and ‘soft’ cuticles (Andersen, 1977). It is argued that cuticles containing relatively more amino acids with polar side chains are
area.
Fragments
incubated
in I M acetic
Far less protein of muscle
(M)
acid before
has been removed
remain
internal
fixation, from
sectioned
this cuticle
to the cuticle.
~13,500.
(CM)
through
a
than the
.’
,-
LARVAL
CUTICLE
OF CHIRONOMUS
‘soft’ and those with fewer are ‘solid’. Unfortunately, it is not always clear whether the designation ‘soft’ or ‘solid’ is made before or after the analysis is undertaken. Therefore, biochemical analyses of cuticles cannot yet provide conclusive evidence of the extent of cuticle sclerotization or a ready distinction between solid and soft cuticles. Examination of the cuticle’s appearance would therefore appear to be the most appropriate means of determining the extent of its stabilization. The use of Mallory’s triple stain suggests that the bulk of the post-cephalic procuticle of C. riparius is not sclerotized. The red staining or refractory nature of the distal margin is, however, suggestive of its impregnation or sclerotization (Andersen, 1977). The extreme narrowness of this zone inhibits its clear resolution with the light microscope and resort to the electron microscope is required for further information. Nevertheless, the contrast between the chemical
nature of the head capsule and post-cephalic cuticle is abundantly clear and there is no doubt that the former is subject to far more extensive stabilization; it is shed with minimal resorption of endocuticle at the moult (Credland, unpublished observations). Ultrastructural examination of the postcephalic cuticle has revealed that a similar thickness of procuticle (about 04-1.0 pm) is secreted pre-ecdysially, is apparently slightly more electron-dense than postecdysial procuticle, and remains in the exuvium. Using the conventional criteria, such cuticle can appropriately be termed exocuticle. Whilst such observations reveal nothing about the chemical nature of the exocuticle, they are strongly suggestive of the fact that the layer is far more stable than the underlying endocuticle. The only explanations for the presence of procuticle in the exuvium depend either on the different chemical natures of the exo- and
Fig. 7. Exuvium of a post-cephalic segment shed at pupation. The procuticular element consists of a distal layer(h) which appears homogeneous at low magnifications, an intermediate fragmentary zone (f) and an internal continuous layer (arrowed). ~65,000. Fig. 8. Similar piece of exuvium but incubated in formamide before fixation. The fragmentary zone (f) and inner continuous layer (arrowed) show signs of further disruption. ~65,000. Fig. 9. An adjacent piece of exuvium to that shown in Fig. 8 but incubated in 1 M acetic acid and not formamide before fixation. Some swelling has occurred, especially in the fragmentary aone (f). rendering the exuvium thicker. ~65,ooO. Figs. 10-13. before fixation,
Areas of post-cephalic cuticlc from larvae, all incubated in protein to show the organization of chitin microfibres in the procuticle.
Fig. 10. Incubated in 1 M acetic acid and snowing the conventional cuticle with the ‘C’ pattern of microfibres. ~44,000. Fig. 11. Incubated in formamide and showing the characteristic of resolving individual microfibres. ~40,000.
lamellate
solvents
appearance
of
pattern but also the difficulty
Fig. 12. A small area of Fig. 11 much enlarged to show the discontinuities within each element of the pattern. Each apparently single microtibre is seen at this magnification to consist of a number of components whose plane varies, probably in accord with a model of sequential laminae laid down in a helicoidal manner. ~132,Otx). Fig. 13. Incubated in 1 M acetic acid and showing microfibres in different planes, each surrounded by a thin protein sheath. Unlike the protein, chitin is electron-htcent as it does not accept a heavy metal stain. When seen in transverse section, the microfibre appears as a lucent disc surrounded by a dense ring (arrowed). ~255,000.
32
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endocuticles or on the failure of enzymes in moulting fluid to digest the cuticle. The latter could occur because the supply of enzymes is exhausted, they are inactivated or because they simply do not have time to complete the dissolution of the entire procuticle. A simple and rather crude way of assessing which of these explanations may be correct is provided by the use of formamide or 1 M acetic acid. These two reagents are believed to remove from cuticles those proteins which are not sclerotized by the formation of covalent links between themselves or with chitin (Andersen and Barrett, 1971; Andersen, 1973, 1976; Thompson and Hepburn, 1978). Therefore one might anticipate that if a cuticle or exuvium contains many more loosely bound proteins which are electron-dense after heavy metal staining, it will appear significantly more electronlucent if treated with either of the reagents prior to fixation and preparation for electron microscopy. Chitin, because it does not stain with heavy metals, will not markedly affect the images seen although it is normally removed by enzymes in the moulting fluid (Jeuniaux, 1971). Actually, some proteins are inextricably bound to the chitin (Hillerton, 1980; Brine and Austin, 1981) and can be expecLed to remain even after incubation in formamide or acetic acid. Therefore, the essential difference between an unsclerotized cuticle from which certain proteins have been removed with these reagents and an exuvium will lie in the presence of chitin and its protein coat in the former. This is precisely what is seen in the endocuticle of the post-cephalic segments of C. ripari~~ which was rendered substantially electronlucent, in contrast to the exocuticle, after incubation in either formamide or 1 M acetic acid. Thus, the results substantiate the assertion that the majority of the proteins of the endocuticle are not covalently bound to each other, but those in the exocuticle, or at least a large proportion of them, are indeed sclerotized, being bound to each other and/ or the chitin. It is in cuticles from which loosely bound proteins have been extracted that the chitin microfibres can most easily be resolved. Seen effectively in negative contrast, surrounded by their protein coat, their diameter of about 3 nm is typical of the value observed in other insects (Filshie, 1980).
CREDLAND
Their arrangement, particularly in the ‘C’ pattern, conforms with the model proposed by Bouligand (1965) and indicates a structure of helicoidally arranged laminae (Figs. 11, 12). Their size and spacing is such that artefacts due to wetting or preparative techniques pointed out by Rudall and Kenchington (1973) appear to have been avoided although some shrinkage in cuticle thickness (Figs. 4, 5) after protein removal does seem to have occurred as expected (Neville, 1965). The possible argument that could be advanced that the exocuticle is relatively unaffected by formamide or acetic acid because it is masked or protected by the epicuticle and endocuticle is refuted by the exposure of exuvia to the same reagents. Similarly, the questions about the timing and availability of enzymes for dissolution of the exocuticle under normal moulting conditions are also answered. As has been demonstrated, very little electron-dense material is removed by either reagent from exuvia and one may conclude that the protein present in the exocuticle is therefore firmly bound or sclerotized. The correspondence between the thickness of the pre-ecdysial cuticle, the thickness of the procuticle left in the exuvium, and the extractability of the proteins in this layer, as compared with those in the more proximal part of the procuticle, would appear to provide ample justification for the assertion that a sclerotized exocuticle is present in the post-cephalic larval sclerites of Chironomus riparius. Furthermore, it represents about 25% of the maximum thickness of the procuticle except at sites of muscle attachment, which were previously reported to avoid digestion by moulting fluid in larvae of Cuffiphoru (Wolfe, 1954) on the basis of light microscope observations. At such sites, the exocuticle may constitute almost the entire thickness of the cuticle (Fig. 6). It should be noted, however, that no inference can be drawn either about the chemical nature of the sclerotization or the timing of its occurrence from the results presented. One feature which emerges clearly from these results is that the cuticle of C. riparius is not fundamentally different from the solid, tanned cuticle (Neville, 1975) despite it being very thin and ‘soft’ in the sense that it is deformed by slight forces, as used by
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OF CHIRONOMUS
Andersen (1977). The thicker cuticles of larval Luciliu (Filshie, 1970), Sarcophaga (Dennell, 1946), and pupal Tenebrio (Locke, 1961) are each reported to lack an exocuticle. However, in at least the first of these cases, Fig. 29 of Filshie’s paper (1970) appears to show a part of the procuticle of the second instar about to be shed in the exuvium. Similarly, illustrations of the Drosophila larva1 cuticle (Mitchell et al., 1971) suggest that about one half of the procuticle of the second instar is lost in the exuvium, a similar proportion to that secreted preecdysially. In itself, such evidence is not enough to prove the existence of an exocuticle but, conversely, if Locke’s (1974) definitions of exo- and endocuticles are accepted and it is sclerotization which prevents dissolution of endocuticle, then it is sufficient to question a simple statement that the cuticle of each larval instar is ‘soft and untanned’ as Filshie (1970) said of Luciliu. Whilst it would be most unwise to argue that all cuticles may have an exocuticle because ‘there is an inherent danger in attempting to formulate generalizations based largely on circumstantial evidence’ (Filshie, 1980), one might question whether any cuticle has ever been irrefutably shown to lack one. It should be appreciated that all the
conclusions reached as a result of this work are based on the defnitions and descriptive statements concerning cuticles cited previously. Evidence for them may therefore be regarded as circumstantial. However, until the chemical basis of sclerotization has been explained satisfactorily and a means of identifying the sclerotized proteins in situ has been produced, any distinction between exo- and endocuticle can only be made by deduction of the properties which sclerotization is presumed to confer. On a functional basis (Neville, 1975), there can be no doubt that the extremely thin cuticle of C. ripurius larvae does have an exocuticle whose behaviour precisely corresponds with that in other, larger insects with thicker cuticles. In itself, such a fact exposes the real problems of considering ‘soft’ cuticles as uniquely different from ‘solid’ ones, and highlights the dangers of thinking of thin larval cuticles as being distinct from thicker ones without a critical experimental approach being allied with straightforward observations. Acknowledgements I am grateful to Graham Lawes, Tony King
and Kevin Jennings for their technical assistance.
References Andersen, S. 0. 1971. R&in. Comprehensive Biochemkfry (eds. M. Florkin and E. H. Stotz), Vol. 26C, pp. 633-657. Elsevier, Amsterdam. Andersen. S. 0. 1973. Comparison between the sclerotization of adult and larval cuticle in Schtirocerca gregaria. J. Insect fhysiol., 19, 1603- 1614. Andersen, S. 0. 1974. Cuticular sclerotization in larval and adult locusts, Schistocerca gregaria. 1. Insect Physiol., 20, 1537-1552. Andersen. S. 0. 1976. Cuticular enzymes and sclerotization in insects. TheInsect Inregumenr (ed. H. R. Hepburn), pp. l21- 144. Elscvicr, Amsterdam. Andersen, S. 0. 1977. Arthropod cuticles: their composition, properties and functions. Symp. zooi. Sot. Land., 39, 7-32. Andersen, S. 0. 1979. Biochemistry of insect cuticle. A. Rev. Em., 24, 29-61. Andersen, S. 0. and Barrett, F. M. 1971. The isolation of ketocatechols from insect cuticle and their possible role in sclerotization. J. Insect fhysiol., 17, 69-83. Bouligand, Y. 1965. Sur une architecture torsadee rCpandue dans de nombreuses cuticles d’arthropodes. C. r. hebd. SPanc. Acud. Sci., Paris, 261, 3665-3668. Brine, C. J. and Austin, P. R. 1981. Chitin isolates: species variation in residual amino acids. Camp. Biochem. PhysioL, 708, 173-178. Credland, P. F. 1973. A new method for establishing a permanent laboratory culture of Chironomur riparius Meigen (Diptera: Chironomidae). Freshwat. Biol., 3, 45-51.
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Credland, P. F. 1978. An ultrastructural study of the larval integument of the midge, Chironomus ripariur Meigen (Diptera: Chironomidae). Cell Tiss. Res., 186, 327-335. Dennell, R. 1946. The study of an insect cuticle: the larval cuticle of Sarcophaga folculato Pand. (Diptera). Proc. R. SOL, 133B, 348-373. Filshie, B. K. 1970. The fine structure and deposition of the larval cuticle of the sheep blowfly (Lucilia cuprina). Tissue & Cell, 2, 478-498. Filshie, B. K. 1980. Insect cuticle through the electron microscope-distinguishing fact from artifact. Inrec! Biology in the Future, ‘VBW 80’ (eds. M. Locke and D. S. Smith), pp. 59-77. Academic Press. New York. Hackman, R. H. 1974. Chemistry of the insect cuticle. The Physio/ogy of fnrecta (ed. M. Rockstein), 2nd edn, Vol6, pp. 215-270. Academic Press, New York. Hillerton, J. E. 1980. Electron microscopy of fibril-matrix interactions in a natural composite, insect cuticle. J. mater. Sci., l&3109-3112. Hillerton, J. E. and Vincent, .I. F. V. 1979. The stabilization of insect cuticles. J. fnrect Physiol., 25, 957-963. Jeuniaux, C. 1971. Chitinous structures. Comprehemive Biochemistry (eds. M. FIorkin and E. H. Stotz). Vol. 26C, pp. 595-632. Elsevier. Amsterdam. Locke, M. 1961. Pore canals and related structures in insect cuticle. .I. biophys. biochem. Cytol.. 10, 589-618. Locke, M. 1974. The structure and formation of the integument in insects. The Physiology of Insecta (ed. M. Rockstein), 2nd edn, Vol. 6, pp. 123-213. Academic Press. New York. Locke, M. and Huie, P. 1979. Apolysis and the turnover of plasma membrane plaques during cuticle formation m an insect. Tissue & Cell, 11, 277-291. Mitchell, H. K., Weber-Tracy, U. M. and Schaar, G. 1971. Aspects of cuticle formation in Drosophila melanogaster. J. exp. Zool., 116, 429-444.
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