Cuticle differentiation in the embryo of the amphipod crustacean Parhyale hawaiensis
- 850 Downloads
The arthropod cuticle is a multilayered extracellular matrix produced by the epidermis during embryogenesis and moulting. Molecularly and histologically, cuticle differentiation has been extensively investigated in the embryo of the insect Drosophila melanogaster. To learn about the evolution of cuticle differentiation, we have studied the histology of cuticle differentiation during embryogenesis of the amphipod crustacean Parhyale hawaiensis, which had a common ancestor with Drosophila about 510 million years ago. The establishment of the layers of the Parhyale juvenile cuticle is largely governed by mechanisms observed in Drosophila, e.g. as in Drosophila, the synthesis and arrangement of chitin in the inner procuticle are separate processes. A major difference between the cuticle of Parhyale and Drosophila concerns the restructuring of the Parhyale dorsal epicuticle after deposition. In contrast to the uniform cuticle of the Drosophila larva, the Parhyale cuticle is subdivided into two regions, the ventral and the dorsal cuticles. Remarkably, the boundary between the ventral and dorsal cuticles is sharp suggesting active extracellular regionalisation. The present analysis of Parhyale cuticle differentiation should allow the characterisation of the cuticle-producing and -organising factors of Parhyale (by comparison with the branchiopod crustacean Daphnia pulex) in order to contribute to the elucidation of fundamental questions relevant to extracellular matrix organisation and differentiation.
KeywordsCuticle Extracellular matrix Chitin Parhyale hawaiensis (Crustacea) Drosophila melanogaster (Insecta)
The cuticle of arthropods is an extracellular matrix (ECM) with multiple functions. It protects the animal against environmental harm and dehydration and serves as an exoskeleton both allowing locomotion and supporting body shape. The functions of the cuticle are conferred by its stratified architecture (Locke 2001). The outermost lipid- and protein-containing envelope is involved in the control of water balance. The middle epicuticle, which is composed of a protein-catecholamine network, and the inner proteinchitin matrix called the procuticle together constitute the stiff, but elastic, exoskeleton. Despite the diversity of arthropods, the cuticle architecture that has been extensively described in the literature seems to be largely conserved.
To promote comprehension of the processes involved in cuticle differentiation, several factors that are required for correct cuticle architecture have been isolated and genetically and molecularly characterised from Drosophila. The majority of factors, including the chitin synthase-1 Krotzkopf verkehrt (CS-1/Kkv), Knickkopf (Knk), Retroactive (Rtv), Piopio (Pio) and Papillote (Pot), are membrane-associated and function in the synthesis, arrangement and attachment of chitin to the cell during procuticle differentiation (Bökel et al. 2005; Moussian et al. 2005a, b, 2006b). In addition, some extracellular factors belonging to the TweedleD class of proteins are predicted to act within the epicuticle where they are needed for body-size regulation (Guan et al. 2006). Orthologues of these factors are encoded by all arthropod genomes sequenced to date, suggesting that basic processes of cuticle formation and function are evolutionary conserved. Indeed, we are beginning to understand the molecular and cellular mechanisms controlling cuticle differentiation in the Drosophila embryo.
In addition to following a genetic approach to extend our knowledge of cuticle differentiation, it is equally fruitful to study and compare its basic underlying mechanisms in distantly related organisms within the same taxon. Such a comparison is intended to uncover not only those characteristics that account for naturally occurring differences, but also those invariable factors ensuring features typical for all branches of the taxon. Within the arthropods, insects probably derive from crustaceans, together constituting the pancrustacea (Mallatt et al. 2004). Therefore, as a next logical step to learning about cuticle evolution, representatives of the insects and crustaceans seem to be predestined for comparative analyses of cuticle differentiation in arthropods, which in principle has been concisely described in several insects, but not in crustaceans.
To compensate for this discrepancy, we have studied the histology of cuticle differentiation in the embryo of the amphipod crustacean Parhyale hawaiensis by electron microscopy. We have chosen Parhyale as a model crustacean, as its entire development from zygote to the juvenile animal takes place within the eggcase, which is small enough to be immobilised by the high pressure freezing method prior to fixation. The cuticle in Parhyale is produced during the second half of embryogenesis (Browne et al. 2005). Soon after the formation of the layers has been initiated sequentially, as in Drosophila, the pro- and epicuticle differentiate simultaneously. Interestingly, unlike the overall uniform cuticle in Drosophila, the pro- and epicuticles at the ventral and the dorsal sides of Parhyale are dissimilar. The ventral epicuticle forms an even layer, whereas the ventral procuticle is eventually subdivided into two layers, the upper exo- and the lower endocuticle. By contrast, the dorsal epicuticle is interrupted and encloses electron-dense chambers that are coated by the envelope. Occasionally, similar but electron-lucid chambers are found within the dorsal procuticle. As in Drosophila, chitin microfibrils and laminae become visible long after chitin synthesis has been initiated suggesting that chitin synthesis and chitin arrangement may be separate processes. Based on the present framework of cuticle differentiation in Parhyale, we plan to investigate the function of its cuticle differentiation factors, which we are currently isolating by using the sequence information of the genome of the branchiopod crustacean Daphnia pulex (Colbourne et al. 2005, 2007).
Materials and methods
Animal maintenance and staging
Parhyale hawaiensis is a marine amphipod that is easy to maintain and breed in the laboratory. Its embryonic development is direct and takes approximately 10.5 days at 26°C.
Laboratory breeding cultures of Parhyale were maintained in shallow covered plastic trays on a day/night cycle. Water was circulated within trays by commercially available aquarium pumps. Phosphate-absorbing resin was used to control the accumulation of free phosphates and thus algal growth. Artificial seawater was prepared from commercial salt (Tropic Marin) to mimic natural seawater with a gravity of 1.018–1.022. About 50% of the water content per tray was changed every week. The animals received commercially available fish food (TetraRubin) every other day and dried yeast extract 3 times per week as a diet. For the methods described below, embryos were carefully taken from the ventral brood pouch of the mother manually. The developmental stages of the embryos were determined according to Browne et al. (2005).
Specimens for transmission electron microscopy (TEM) were prepared by high-pressure freezing followed by freeze-substitution and embedding in Epon following the protocol documented in Moussian et al. (2006a). In brief, embryos were immobilised within the eggcase in a high-pressure freezer (Bal-Tec HPM 010, Balzers, Liechtenstein) and freeze-substituted in 2% osmium tetroxide, 0.5% uranyl acetate and 0.5% glutaraldehyde in 97.5% acetone and 2.5% methanol at -90°C for 32 h, warmed up within 3 h to -60°C, kept at -60°C for 6 h, warmed up to -40°C within 2 h and kept for an additional 4 h at -40°C. After being washed with acetone, the samples were transferred into an acetone-Epon mixture at -30°C (1:1 for 4 h, 2:1 for 12 h), warmed up to room temperature, infiltrated in Epon (three changes within 30 h) and polymerised at 60°C for 48 h. Ultra-thin sections (50–70 nm) stained with 2% uranyl acetate in 70% methanol for 10 min and in 0.4% lead citrate in 0.1 N NaOH for 2 min were viewed in a Philips CM10 electron microscope at 60 kV.
For wheat germ agglutinin (WGA)-labelling, unstained Epon sections were incubated with biotinylated WGA (10 mg/ml; Vector Labs, Burlingame, USA) followed by rabbit anti-biotin antibodies (ENZO Life Sciences, Farmingdale, USA) and protein A conjugated to 10-nm gold (gift from Dr. York Stierhof, ZMBP Tübingen). Labelled sections were then stained with 1% aqueous uranyl acetate for 3 min and lead citrate.
For scanning electron microscopy (SEM), embryos were fixed for 5 h in 4% formaldehyde and 0.5% glutaraldehyde at 4°C. The chorion and vitelline membrane were dissected off the embryos manually with tungsten-wire needles placed in a syringe. They were osmium-treated (1% osmium tetroxide in 100 mM phosphate buffer, pH 7.2), dehydrated through an ethanol series, subjected to critical-point drying in CO2 and sputter-coated with 10-nm Au-Pd. Critical-point drying may have affected the appearance of the specimen’s surface. Nevertheless, the differences in the surface appearance of embryos at the different stages reflected changes in ECM composition during maturation. A Hitachi S100 field-emission scanning electron microscope was used to examine the samples.
To identify the protein sequences of CS-1/Krotzkopf verkehrt Knickkopf and Retroactive in Daphnia pulex (DpCs-1 to 3 DpKnk and DpRtv), the respective Drosophila sequences were “blasted” against the translated genome of Daphnia at wFleaBase http://wfleabase.org/blast/). The domains of the identified proteins DpKnk and DpRtv were predicted by the SOSUI, PSORT, GPI-SOM and big-PI predictor on-line programs listed at the Expert Protein Analysis System (Expasy) site (http://www.expasy.org/tools/).
Course of cuticle differentiation during Parhyale embryogenesis
Parhyale embryogenesis is completed after 240 h post-fertilisation (hpf); this period has been subdivided into 30 stages (S1–S30; Browne et al. 2005). Cuticle differentiation has been observed to start around stage 26 (S26) at 180 hpf. To trace the cellular mechanisms of cuticle differentiation in the Parhyale embryo, we have analysed the ultrastructure of the cuticle of staged embryos from 120 hpf (S21) to 240 hpf (S30) by SEM and TEM.
Stages 22 and 23
At S22 (132 hpf), the apical surface of the epidermal cell starts to protrude microvillus-like structures (Fig. 2d). At the tips of these protrusions, fragments of a tripartite layer are formed that histologically resembles the insect envelope, which is also produced at microvillus-like structures. By S23 (144 hpf), the epidermal cells are unchanged compared with S22 but are now covered by a continuous envelope (Fig. 2e). Chitin has not yet been synthesised (Fig. 2f).
Stage 24 to early stage 26
Late stage 26 to early stage 27
Late stage 27
Stages 28 to 30
To gain wide-ranging information about basic mechanisms of cuticle differentiation by comparative morphology, a thorough analysis of the process in divergent, yet reasonably related, model animals is necessary. The evolutionary distance between insects and crustaceans promises a fruitful comparison of cuticle differentiation in these two major clades of arthropods. Cuticle differentiation has been well studied in the embryo of the insect Drosophila melanogaster (Hillman and Lesnik 1970; Moussian et al. 2006a). Here, we have presented our work on the course of cuticle differentiation in the embryo of the crustacean Parhyale hawaiensis in order to allow us to discuss some principles of cuticle differentiation.
The Parhyale embryo produces two cuticles
Chitin synthesis and arrangement
The chitin arrangements in the eggshell and the two cuticles produced by the Parhyale embryo are different. In the eggshell and the embryonic cuticle, chitin does not adopt the stereotypic organisation that, according to Bouligand (1965), is characteristic for the second juvenile cuticle. In several instances, the topology of the apical plasma membrane has been proposed to play an important role in chitin orientation. Indeed, microtubule-stabilised corrugations reminiscent of the apical undulae in Drosophila are present during the differentiation of the juvenile procuticle and absent when the embryonic procuticle differentiates. In addition, the more complex arrangement of chitin in the juvenile cuticle might require factors that are not expressed or functioning in the follicle cells and during chitin assembly within the simpler embryonic cuticle. Candidates for these factors are Retroactive (Rtv) and Knickkopf (Knk), which have been hypothesised to be involved in chitin microfibril orientation in Drosophila (Moussian et al. 2005b, 2006b). Both factors are indeed present in the genome of Daphnia (Fig. 8b,c) stressing that crustaceans share these sequences with insects. The alternative view that chitin orientation may be a property of the chitin synthase itself is less likely, since chitin synthesis and chitin lamina rotation seem to be separate processes, also during procuticle differentiation in the juvenile cuticle. Thus, it will be exciting to unravel the expression pattern of Parhyale chitin synthases, knk and rtv during embryogenesis and to analyse the phenotype of Parhyale embryos lacking the function of the chitin synthases, Knk or Rtv as generated by RNA silencing or morpholinos.
Dorso-ventral differences of cuticle architecture
The juvenile dorsal and ventral cuticles of Parhyale are dramatically different. The dorsal epicuticle and procuticle are characterised by inclusions that are missing in the respective layers at ventral positions. Moreover, pore canals, which have been proposed to be mineralised or to be routes for mineralisation, are only present in the dorsal cuticle. Overall, the ventral cuticle resembles the cuticle described in other crustaceans including copepods (Anomalocera patersoni, Cletocamptus retrogressus and Porocellidium viride), branchiopods (Daphnia magna, Triops cancriformis and Leptestheria dahalacensis) and decapods (Homarus americanus; Gharagozlou-van Ginneken and Bouligand 1973, 1975; Halcrow 1976; Arsenault et al. 1984; Bresciani 1986; Freeman 1989), whereas the dorsal cuticle seems to be a specific trait of amphipods and isopods (see below). By contrast, in Drosophila, the architectures of the cuticle at dorsal and ventral positions are indistinguishable. Two conclusions can be drawn from these observations. First, differentiation of the cuticle in Parhyale responds to patterning information established early during embryogenesis, whereas in Drosophila, cuticle differentiation seems to occur independently from pattern formation. Second, the exact boundary between the different types of cuticle strongly indicates that, at this position, cuticle differentiation is a cell-autonomous process. Indeed, the boundary separating the dorsal and ventral cuticles corresponds to the sites of cell-cell contacts. Conceivably, the establishment of the boundary is driven by dorsal- and ventral-specific sets of regulating and differentiation factors that respect positional information. For the further elucidation of these crucial aspects of cuticle differentiation, we will attempt to identify the factors that are responsible for the organisation of the dorso-ventral cuticle boundary.
Modelling of the dorsal epicuticle
The dorsal epicuticle of Parhyale harbours inclusions surrounded by the envelope. A similar epicuticular architecture has not been described for other crustaceans or arthropods including insects, except for another amphipod, Hyale nilssoni, and, with some modifications, for Idotea baltica, which belongs to the isopods, the closest relatives of the amphipods (Halcrow 1985; Powell and Halcrow 1985). Intriguingly, the formation of these inclusions through the invagination of the envelope at S27 in Parhyale occurs in the ECM with no physical contact to the apical plasma membrane. Neither in Hyale nilssoni nor in Idotea baltica has the formation of these inclusions been described in detail. In Drosophila and in insects in general, the maturation of the epicuticle involves extracellular (i.e. cell-independent) cross-linking of cuticle proteins with catecholamines in a process called sclerotisation but no dramatic structural configuration beyond smooth layering. Hence, the modelling of the Parhyale dorsal epicuticle is a striking example for self-organisation during cuticle differentiation in arthropods. It will be a great challenge to analyse the molecular mechanisms of this fascinating process.
The comparison of cuticle differentiation in Parhyale (presented here) and in Drosophila (published recently) allows two major conclusions. First, cuticle differentiation naturally integrates information from the epidermal cell, especially that defined by its apical plasma membrane and intrinsic properties of the cuticular components themselves, which, for instance, direct laminae rotation within the procuticle. Second, the molecular mechanisms controlling envelope and procuticle differentiation are probably more conserved across arthropods than those governing epicuticle construction. Indeed, an obvious difference in cuticle structure between Parhyale and Drosophila is observed within the epicuticle, whereas their envelope and the procuticle are similar. A review of the literature concerning cuticle structure largely supports the conclusion of a variable epicuticle compared with the stereotypic envelope and procuticle. In other words, the epicuticle has been more sensitive to selective forces during evolution than have the envelope and the procuticle.
- Bouligand Y (1965) On a twisted fibrillar arrangement common to several biologic structures. C R Acad Sci D 261:4864–4867Google Scholar
- Freeman JA (1989) The integument of Artemia during early development. In: MacRae TH, Bagshaw JC (eds) Biochemistry and cell biology of Artemia. CRC Press, Boca Raton, pp 233–256Google Scholar
- Merzendorfer H (2005) Insect chitin synthases: a review. J Comp Physiol [B] 176:1–15Google Scholar
- Moussian B, Tang E, Tonning A, Helms S, Schwarz H, Nüsslein-Volhard C, Uv AE (2006b) Drosophila Knickkopf and Retroactive are needed for epithelial tube growth and cuticle differentiation through their specific requirement for chitin filament organization. Development 133:163–171PubMedCrossRefGoogle Scholar