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Arthropod cuticle: A natural composite shell system Article in Composites Part A Applied Science and Manufacturing · October 2002 DOI: 10.1016/S1359-835X(02)00167-7
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Arthropod Cuticle – a natural composite shell system Julian F V Vincent Dept of Mechanical Engineering, The University of Bath, BA2 7AY, UK Fax +44 (0) 1225 826928,
[email protected] Abstract The cuticle of arthropods (jointed-limb animals), and especially of insects is, by biological standards, a relatively simple composite. It is a single external layer of material forming the skeleton and many sense organs. The fibrous phase is crystalline chitin making nanofibrils of about 3 nm diameter, a few hundreds of nanometers long and a modulus probably in excess of 130 GPa. At least two surfaces of the nanofibril can have silk-like protein attached through specific H-bonds; the rest of the protein is globular. The protein matrix stiffens through dehydration controlled by the introduction of hydrophobic phenolics. Crustacea add up to 40% calcium salts. The stiffness of cuticle can range from tens of GPa to 1 kPa. It can be hardened by the addition of Zn or Mn. It can form springs and change its stiffness and plasticity under the control of the animal. Keywords: Fibres, Nano-structures, Fibre/matrix bond, Mechanical properties Introduction Most biological materials are composites, either containing ceramic and protein phases (teeth, bone, mollusc shell, sea-urchin shell) or two organic phases, commonly a protein and a polysaccharide. Either phase may be fibrous, the other a matrix. Many soft tissues are composed of a protein fibre (collagen) in a protein-polysaccharide matrix (soft skins and connective tissues of nearly all animals). Many stiff tissues are composed of a polysaccharide fibre (cellulose) in a matrix which may also be mostly polysaccharide (hemicellulose, pectin etc) with a little protein (in most plant cells walls, woody and nonwoody); or polysaccharide fibre (chitin) in an extensive stabilised protein matrix (the cuticle, or outer covering, of insects, Figure 1) which may have a ceramic phase as well (cuticle of crabs, lobsters and other crustacea; a very few insects). These last animals are called arthropods or “jointed-limbed” animals. A chitinous cuticle is diagnostic of the arthropods. It is secreted by a single layer of cells (the epidermis) and gains its stiffness and structural complexity mainly by being folded and curved in many complex ways. Basic components of arthropod cuticle Chitin is a fairly completely acetylated polysaccharide akin to cellulose. Like cellulose, the monosaccharide units are joined by -1,4 links which make chitin chains very straight and ribbon-like. The chains are arranged anti-parallel ( -chitin) and combine into a highly crystalline structure within which the sugar residues are heavily H-bonded making the chains very stiff and stable. The stiffness of chitin cannot be less than that of cellulose, for which the best estimate is currently about 130 GPa, still only half the theoretical stiffness of 250 GPa. Within the body of the cuticle the chitin is assembled 1
into nanofibrils about 3 nm in diameter, each containing 19 molecular chains and about 0.3 µm long. Chitin can easily be isolated from crab shell, when it is found to be highly thixotropic (Vincent unpublished) and liquid crystalline [1, 2]. Since, in most biochemical studies, chitin is solubilised by deacetylation and the destruction of the crystalline order, there is relatively little information on the properties of the undeacetylated chitin nanofibril. The number of chitin chains in the nanofibril is probably close to a minimum for stability; hence the chitin nanofibrils present the maximum surface area for interfacial interactions within the cuticle. However, there is evidence that larger nanofibrils exist in some cuticles where resistance to compression may be important [3], so the optimisation of the size of the fibres in cuticle is unexplored. The crystalline structure of -chitin (Figure 2) has a number of surprises. Along the 010 face (the c-axis) the hydrogen chains are laterally spaced by 0.475 nm, the same as the lateral distance between the adjacent protein chains of an antiparallel -sheet such as silk. Along the length of each chitin chain the space between adjacent residues is 1.032 nm. Twice this repeat (2.064 nm) is almost exactly the same as three times the repeat of a protein -sheet (0.69 nm) [4]. Thus the lattice which appears on the 010 face of the chitin nanofibril can be related to the spacings in a chain folded sheet of protein -sheet [5] and the chitin is bonded to half the protein groups (Figure 3). Protein is probably synthesised within and is certainly secreted from the epidermis. The protein has to produce a matrix of varying mechanical properties, which will also interact with and stabilise the chitin. It interacts with the chitin in a narrow zone (the Schmidt layer) with relatively large amounts of water (about 90%), presumably to drive the hydrophobic interactions between chitin and protein. Electrophoretic 2-dimensional separation of extracted cuticular proteins on gels shows that they are many and varied. In soft cuticles such as the highly stretchy and soft cuticle of the locust there may be only 20 or so [6] whereas in some of the stiffer cuticles in the same insect there may be 200 or more [7]. This complexity is probably more apparent than real, since the conditions of separation of the proteins on gels picks up slight difference in charge, which could be given by a single amino acid substitution in the sequence, which may not be so important in a structural protein. Also the proteins are probably not different in the different areas, as can be shown by immunological comparisons. The proteins can interact with each other giving well-defined complexes in the absence of H-bond breakers, suggesting that hydrophobicity is an important factor in their stabilisation [8, 9]. The proteins vary widely in hydrophobicity. In general proteins from more highly evolved insects tend to be less hydrophobic, even though they will form stiff cuticles in the same way. Much of the protein, especially in softer, more hydrated, cuticles can be extracted with mild solvents such as simple buffers; yield is often improved by going to low temperatures, a characteristic behaviour of hydrophobic proteins. In some cuticles up to 70% of the total protein can be removed in this way. The interaction of the remaining proteins with the chitin seems to be fairly consistent in that even in the softest of cuticles (in which the interactions are least developed) a strong solvent (e.g. 5% NaOH at 100C) is required to remove the protein from the chitin [10]. X-ray diffraction of the ovipositor (egg-laying tube) of the wood wasp Megarhyssa 2
suggested that the proteins surround the chitin in a regular manner [11]. Later work on the same system, allied with more careful molecular modelling and the analysis of the crystalline structure of the chitin nanofibril suggested that the protein is attached only to the 010 faces [5] (pers. comm.) and that the other faces of the nanofibril are essentially bare of bound protein. In recent years the increased amount of information on the amino acid sequence of these proteins allows much more detailed and interesting conclusions to be drawn about the nature of the matrix and its interactions both internally and with the fibrous phase. A socalled “conserved” sequence was identified [12] and it was tentatively suggested that this sequence could be important in interactions with the chitin. As noted above, this would necessitate its producing a -sheet in the protein. This has recently been confirmed, and it has been shown, experimentally, to bind to chitin [13]. Even so, the detailed nature of the chitin-protein bond in the cuticle is subject to speculation. The 3:2 relationship between the spacings of the chitin and protein residues along their respective chains accounts for only half of the bonds which the protein could make. Ted Atkins (who has worked most recently with the chitin system) is of the opinion that one, at least, of these groups could react covalently with the few unacetylated side chains of the chitin. This is still speculation, so while we cannot calculate the strength of the interfacial interaction between the chitin and the protein (fibre-matrix), we can at least put a lower bound on it. In the early stages of the development of the cuticle, before the insect has shed the cuticle of its previous stage, the amount of protein per unit chitin is much less than in the mature developed cuticle [14] and the protein which is present shows much more -sheet and so may be deduced to be interacting preferentially with the chitin [13]. Therefore the addition of matrix material to chitin is at least a 2-stage process; the chitin nanofibrils are first coated with a specific protein, then the rest of the protein is added, interacting with the proteins on the chitin. Conformations other than -sheets exist in cuticular proteins, although they have not been much explored. They have been mapped in some proteins [15]. In particular the cuticle protein(s) called resilin which are highly rubbery now appear to contain -turn structures similar to those found in elastin, a rubbery protein typical of vertebrate animals [16]. The -turn is present in many of the other cuticular proteins [7] and so may form an important component of hydrated cuticles. When dehydrated, such structures revert to the glassy state, since water plasticises proteins (but see below). Resilin is always highly hydrated. Calcium salts are present in many cuticles and are typical of crustacea, both aquatic and terrestrial. The salts are mostly carbonate, present as calcite and they dominate the mechanical properties. Extraction of the calcium with dilute acid leaves a tough leathery material. Their chemistry and interactions with the organic components and the control of calcification seem to be little understood. There are hardly any mechanical data, though the hardness of some of these materials has been shown to be outstanding [17]. Control of stiffness The stiffness of insect cuticle is due to the extent of interaction of the protein with the chitin, with other proteins and internally with itself. Stiffness can vary from a kPa in the highly hydrated extensible intersegmental membrane of the locust, via about 1 MPa for the rubbery protein, resilin, to a few GPa in well-tanned cuticles. The most widely 3
accepted version of stiffening (tanning, sclerotisation) of cuticle was suggested by Pryor [18], but the fact that 60 years later we are still uncertain of its mechanism makes one suspicious that the right trails are being followed. Immediately after the old cuticle has been shed, the epidermal cells, which secreted the cuticle, further secret into it a variety of ortho-dihydroxyphenolic compounds. They are converted into the more reactive quinone form and, supposedly, cross-link the proteins making the matrix stiff, hydrophobic, insoluble and chemically inert. At the same time much water is lost from the cuticle [19; 20] which can, anyway, be stiffened just as much simply by the removal of water [21]. Indeed, the naturally sclerotised and stiffened material can be reversibly softened and made rubbery by soaking it in formic acid [20]. This indicates that the secondary reactions made possible by the removal of water from the protein are much more important than the phenols in stiffening the cuticle, and that the control of stiffness is in general a matter of manipulating the water content. Ironically enough this experiment was performed by one of the arch proponents of the theory of phenolic tanning (as the only means of stabilisation of cuticular proteins) and he still hasn’t realised (pers. obs.) that this one experiment completely negates the theory!! Nowhere in the literature has any proponent of the phenolic theory tried to disprove the idea, thus establishing its credibility on a firmer footing. When the water has been removed the protein can form large amounts of -sheets [22, 23]. Under experimental conditions the cuticle can be shown to be extremely sensitive to water content, suggesting that an important source of stabilisation is H-bonding [24]. In the soft cuticle of maggots, which is later tanned with great loss of water [19], gradual reduction of water in the absence of tanning takes the cuticle through a transition of up to 10-fold increase in stiffness with a change in water content of only a few percent (Vincent, unpublished). In some cuticles the insect can increase the water content so that the modulus decreases. This happens in the blood-sucking bug Rhodnius, for instance, which can change the pH of the cuticle from about 7 to below 6, thereby increasing the charge density of the cuticular protein and increasing the cuticular water content from about 26% to 31%, dropping its stiffness from 250 MPa to 10 MPa and increasing its extensibility from 10% to more than 100%. Obviously the orientation of the chitin has a controlling influence on the mechanical anisotropy of a piece of cuticle, but surprisingly although most soft cuticles have equal amounts of chitin and protein, the stiffer cuticles have only about 30% chitin and 70% protein. This varies according to the particular type of cuticle [25, 14]. It seems that stiffness is mostly controlled by the secondary-bond cross-linking of the matrix.
Other functions Since the cuticle has to perform all the functions of skin and skeleton, it is praeternaturally multifunctional [26]. It not only supports the insects, it gives it its shape, means of locomotion (together with mechanisms for strain amplification for jumping), waterproofing and a range of mechanical tricks. For instance the cuticle can be locally hardened on the surface by the addition of up to 10% of Mn or Zn [27, 28, 29]. This enables locusts and other insects to eat plants, some of which contain large amounts of silica [30]. The cuticle also represents a sensory barrier, in that the insect has to see the outer world through it. Strains in the cuticle are detected by the deformation of holes
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whose size and shape vary depending on the type and direction of the expected strains [31, 32]. Learning from cuticle Many of the principles of the design and construction of composites are known and nature is as aware of them as we are. The most obvious advantage nature has is the assembly of materials from the molecular level upwards. Comparing the surface area available for bonding between matrix and protein; if the minimum practical size for a carbon fibre is a diameter of 5 m, but only half of the surface area of the chitin nanofibril is available for matrix bonding, then the surface area of chitin per unit volume of cuticle is 106 greater than the carbon fibre material. Although the interfacial shear strength of the protein-chitin interface is unknown, guessing that the shear strength of a single bond is of the order of 30 pN, and calculating that the area around one bond is about 10-18 square metres, then the shear strength will be about 30 MPa, or about half that measured for carbon fibres. But with the huge surface area difference it is fairly obvious that the total fibre-matrix interaction per unit volume in insect cuticle is still 5 x 105 greater than carbon fibre composites. Nature’s implementation of some of the desirable traits may yield other insights. The means of handling such fine fibres; the use of techniques of self-assembly; the coproduction of the components; the initial surface treatment of the fibres; the use of transitions in the control of stiffness. Perhaps these are uniquely biological and are optimisations and fudges in a system where control is difficult. But perhaps also these methods could suggest more efficient routes for our own technology. Insect cuticle remains the second most widely distributed material (the first is plant cell wall and wood) and the simplest high-performance composite. Its secrets are still untold. References 1. Murray SB, Neville AC. The role of the electrostatic coat in the formation of cholesteric liquid crystal spherulites from alpha-chitin. Int J Biol Macromol 1997;20:123-130. 2. Murray SB, Neville AC. The role of pH, temperature and nucleation in the formation of cholesteric liquid crystal spherulites from chitin and chitosan. Int J Biol Macromol 1998;22:137-144. 3. Gardiner BG, Khan MF. A new form of insect cuticle. Zool J Linn Soc 1979;66:91-94. 4. Fraenkel G, Rudall KM. The structure of insect cuticles. Proc R Soc Lond B 1947;134:111-143. 5. Atkins EDT. Conformations in polysaccharides and complex carbohydrates. Proc Int Symp Biomol Struct Interactions, Suppl J Biosci 1985;8:375-387.
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6. Vincent JFV, Shawky NAF. The proteins of the urea-soluble fraction of locust intersegmental membrane. Insect Biochem. 1978;8:255-261. 7. Andersen SO, Hojrup P, Roepstorff P. Insect cuticular proteins. Insect Biochem molec Biol 1995;25:153-176. 8. Hillerton JE. Electron microscopy of fibril-matrix interactions in a natural composite, insect cuticle. J Mater Sci 1980;15:3109-3112. 9. Hillerton JE, Vincent JFV. Consideration of the importance of hydrophobic interactions in stabilising insect cuticle. Int J Biol Macromol 1983;5:163-166. 10. Hackman RH, Goldberg M. Comparative chemistry of arthropod cuticle proteins. Comp Biochem Physiol 1976;55B:201-206. 11. Blackwell J, Weih MA. Structure of chitin protein complexes: ovipositor of the ichneumon fly Megarhyssa. J mol Biol 1980;137:49-60. 12. Rebers JE, Riddiford LM. Structure and expression of a Manduca sexta larval cuticle gene homologous to Drosophila cuticle genes. J mol Biol 1988;203:411423. 13. Rebers JE, Willis JH. A conserved domain in arthropod cuticular proteins binds chitin Insect Biochem Mol Biol 2001;31:1083-1093. 14. Vincent JFV, Clarke L. Effects of diflubenzuron on the stabilisation of protein within the cuticular matrix of the locust . Entomol General 1985;11:15-24. 15. Iconomidou VA, Willis JH, Hamodrakas SJ. Is b-pleated sheet the molecular conformation which dictates formation of helicoidal cuticle? Insect Biochem Molec Biol 1999;29:285-292. 16. Urry DW. Elastic biomolecular machines. Sci Am 1995;January:44-49. 17. Currey JD, NashA, Bonfield W. Calcified cuticle in the stomatopod smashing limb. J Mater Sci 1982;17:1939-1944. 18. Pryor MGM. On the hardening of the ootheca of Blatta orientalis. Proc R Soc Lond B 1940;128:378-398. 19. Fraenkel G, Rudall KM. A study of the physical and chemical properties of insect cuticle. Proc R Soc Lond B 1940;129:1-35. 20. Andersen SO. The stabilization of locust cuticle. J Insect Physiol 1981;27:393-396. 21. Vincent JFV, Hillerton JE. The tanning of insect cuticle - a critical review and a revised mechanism. J Insect Physiol 1979;25:653-658. 22. Hillerton JE, Vincent JFV. The stabilisation of insect cuticles. J Insect Physiol 6
1979;25:957-963. 23. Hackman RH, Goldberg M. Some conformational studies of Calliphora vicina larval cuticular protein. Insect Biochem 1979;9:557-561. 24. Vincent JFV, Ablett S. Hydration and tanning in insect cuticle. J Insect Physiol 1988;33:973-979. 25. Vincent JFV. Insect cuticle: a paradigm for natural composites. In: JFV Vincent, JD Currey, editors. The Mechanical Properties of Biological Materials. Cambridge: The University Press, 1980; pp.183-210. 26. Vincent JFV. Ideas from Skins. Interdisciplinary Sci Revs 1999;24:52-57. 27. Hillerton JE, Vincent JFV. The specific location of zinc in insect mandibles. J exp Biol 1982;101:333-336. 28. Robertson B, Hillerton JE, Vincent JFV. The presence of zinc or manganese as the predominant metal in the mandibles of adult stored product beetles. J Stored Prod Res 1984;20:133-137. 29. Quicke D, Wyeth P, Fawke J, Basibuyuk H, Vincent JFV. Manganese and zinc in the ovipositors and mandibles of hymenopterous insects. Zool J Linn Soc 1999;124:387-396. 30. Lucas PW, Turner IM, Dominy NJ, Yamashita N. Mechanical defences to herbivory Ann Bot 2000;86:913-920. 31. Barth FG. Slit sense organs: "strain gauges" in the arachnid exoskeleton. Symp zool Soc Lond 1978. p. 439-448. 32.
Skordos A, Chan C, Jeronimidis G, Vincent JFV. A novel strain sensor based on the campaniform sensillum of insects. Phil Trans R Soc A 2001 (in press).
Rubrics Figure 1. Section through hard insect cuticle showing the layering arising from changes in the orientation of the chitin-protein fibres within the structure. The layers are parallel to the surface of the insect. Thickness of cuticle is about 10 m. Figure 2. View of chitin crystal structure looking along the c-axis (i.e. along the length of the nanofibril). The outline of a nanofibril is shown. From ref [5]. Figure 3. Interaction between chitin and protein. The plane of the paper is given by the c- and b-axes. 7
