ElectronMicrosc.Rev., Vol. 4, pp. 47~3, 1991. Printed in Great Britain. All rights reserved.
0892-0354/91 $0,00+ 0.50 © 1991PergamonPress plc.
STRUCTURE A N D BIOCHEMISTRY OF M A M M A L I A N H A R D KERATIN R. C. MARSHALL,*
D. F. G. ORWINt@
J. M . G I L L E S P I E *
*CSIRO Division o f Wool Technology, 343 Royal Parade, Parkville, Victoria, Australia 3052 t Wool Research Organisation of New Zealand, Private Bag, Christchurch, New Zealand Abstract--In this review, the structure and biological formation of hard ~t-keratins are drawn together. The hard keratins comprising wool, hairs, quills, hooves, horns, nails and baleen contain partly ~t-helical polypeptides which show homology with epidermal polypeptides only in the helical regions. These polypeptides (about 32 chains) are organized into intermediate filaments (IFs) of 7.5 n m diameter which are embedded in variable a m o u n t s of a matrix of non-helical cystine-rich proteins and glycine-tyrosine-rich proteins. The total number of proteins m a y exceed 100. In addition keratins contain a variety of lipid components. Wool and hair are produced in follicles in a multistep procedure. In the lower levels of the follicle, IFs without associated matrix are found. Subsequently matrix proteins are laid down between the IFs and further synthesis takes place concurrently. Finally the proteins are insolubilized by the oxidative formation of disulphide bonds. Keratinized fibres shows considerable complexity and diversity in the structural arrangement of IFs and matrix within cortical cells. Typically the IFs show hexagonal packing or give a whorl-like appearance in cross-section.
CONTENTS I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Morphology of hard keratin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Fibre characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Cell types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Subcellular components . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Structure and biochemistry of hard keratins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Solubility of hard keratins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Cellular and subcellular components . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Morphological location of the proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Disulphide bonds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Relationship to physical properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Development o f hard keratin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Follicle structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Ultrastructural development of hard keratin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biochemistry of hard keratin development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Variations in keratin structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cortical cell types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Cortical cell type distribution in wool . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Cortical cell types in other animal fibres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Structural differences between hard keratins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. A b n o r m a l fibre structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Structure and variability of keratin proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Constituent polypeptides of intermediate filaments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Matrix proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Variability in keratin composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Concluding statements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
48 49 49 49 51 52 52 53 53 57 57 58 58 58 60 65 67 67 68 69 69 71 71 71 72 74 76 77 77
R.C. Marshall et al. I. INTRODUCTION
Mammalian keratins such as epidermis and hair perform important functions in mammals. These are varied and include the retention of body fluids by the water impermeable epidermis, the prevention of heat loss by hair, the protection of digit ends by nails and claws, and the use of baleen in food gathering. Two distinct types of a-keratins have been recognized: (i) The soft a-keratin of epidermis and other epithelial tissues; (ii) The hard ~-keratins which, while they are derived embryologically from epithelial cells, are a group of derivatives in which hardness and durability are usually a feature. The hard a-keratins, the subject of this review, comprise wool, hair, nail, claw, hoof, the horns of cattle, goats, sheep and rhinoceros, the quills of porcupine, echidna and hedgehog, and the baleen of whale. While wool, hair and quill are fibrous in form and are produced by follicles, the other hard keratin derivatives have a variety of forms which reflect the organization of the tissues producing them. In spite of the large number of different forms of hard a-keratin, they show many similarities in chemical and structural features: (i) They have a unique arrangement of structural elements, comprising aligned intermediate filaments (IFs) traditionally termed microfibrils, about 7.5 nm in diameter, surrounded by a variable amount of non-filamentous protein matrix (Fraser et al., 1972). (ii) The IFs contain proteins descriptively termed low-sulphur proteins since they have less half-cystine than the original keratin. In contrast, the matrix always contains proteins (highsulphur proteins) richer in half-cystine than the keratin, and frequently another group which is rich in tyrosine and glycine and termed the high-tyrosine or high-glycine-tyrosine proteins (Gillespie, 1983).
(iii) Hard keratins are insoluble in conventional protein solvents except at extremes of pH or in the presence of oxidizing or reducing agents, due primarily to an extensive network of disulphide bonding (Fraser et al., 1972). The older terminology of the components of hard keratin, which arose from ultrastructural studies, is now at variance with the generally accepted nomenclature for epidermal cellular constituents, and increasingly microfibrils are referred to as intermediate filaments (IFs). Hard keratin IFs are composed of two families of lowsulphur polypeptides (Powell and Rogers, 1986) which are homologous with Type I (acidic) and Type II (neutral-basic) IF polypeptides of epidermis (Conway et al., 1989). IF polypeptides contain both helical and non-helical segments, and it is important to recognize that it is only in the helical segments that corresponding soft and hard keratin IF polypeptides are homologous (Conway et al., 1989; Sparrow et al., 1989). The matrix proteins (high-sulphur and high-tyrosine) are increasingly equated (Kuppers and Hocker, 1990) with intermediate filament associated proteins (IFAPs) of the epidermis (Dale et al., 1989), but as the two groups may well have different functions and in any case contain quite unrelated proteins, we have retained the older terminology. More detailed discussions of the IF polypeptides and matrix proteins are given in Sections VI. A and B. The major advances in the knowledge of keratin structure have come largely from research on wool, for apart from some investigations of human hair and mohair, extensive studies of other keratins have not been carried out. Thus wool, and to a lesser extent human hair, will be used as models for hard keratins. Reference to studies on other keratins will be made where appropriate. Information gained on one keratin is generally assumed to be applicable to other keratins since keratins form a group of homologous tissues. Knowledge obtained from biochemical and electron microscopical studies on wool, combined with X-ray diffraction data on porcupine quill, has allowed the spectacular recent advances in the knowledge of the structure of IFs of hard keratin (Conway et al., 1989).
Mammalian Hard Keratin II. M O R P H O L O G Y OF H A R D KERATIN
A. Fibre Characteristics Keratin fibres can differ markedly in diameter (10-250#m) (Bradbury, 1973; Kaplin and Whiteley, 1985) and in length. These physical characteristics depend primarily on the species of animal, and its nutritional and physiological state, but length is also determined by the timing of the anagen phase (Reis, 1979, 1989; Woods and Orwin, 1988; Hynd, 1989). Very few fibres are perfectly round, most show some ellipticity which is seldom perfect. Many vary in their cross-section along their length and this is very marked in 'doggy' wool (James and Ward, 1965), in some human genetic abnormalities (Price, 1979) and in wool following dietary and physiological manipulation of sheep (Chapman, 1989; Reis, 1989). B. CeH Types A schematic diagram of a keratin fibre showing the different constituent cell types is given in Fig. 1. The cortical cell, the major cell type in wool and hair, is spindle shaped (length 85 120F~m, width 4.5-6ktm: Bradbury, 1973; Hynd, 1989)
and is aligned in the growth direction. Up to about 90% of the cortical cell is made up of longitudinally-aligned IFs with accompanying matrix, the remainder being membranes and remnants from the nucleus and cytoplasm (Bradbury, 1973). The structural arrangements in other keratins can be different, for example the filaments in human nail are aligned perpendicular to the growth axis (see Section V.D). Most keratins contain a number of cortical cell types distinguished primarily by the arrangement of filaments. In the case of wool, ortho-, mesoand para-cortical cells (Fig. 2) have been defined. Various cell types are discussed in more detail in Sections III.C.1, V.A, B and C. Wool and hair fibres have a sheath of overlapping cuticle cells which surrounds the cortex and forms the external cell layer of the fibre (Fig. 3a). The cuticle is composed of roughly rectangular (30-50#m) flattened cells (0.2~0.7#m thick) whose edges overlap adjoining cuticle cells forming a characteristic scale pattern (Wildman, 1954; Appleyard, 1960). The number of overlapping cells making up the cuticle layer may vary from one (wool) to 35 (pig bristle) (Swift, 1977). However, in wool fibres with bilateral distribution of orthoand para-cortical cells, multiple layering and/or epicuticle ±ex°cuticle
• nuclear remnant
low-S high-S proteins proteins
I righthanded =:-helix
left-' handed coiled-coil rope
I L. . I matrix intermediate filament
[I aq I endocuticle . . . . ~ 1 I meoutta
membrane c o m p ex macrofibril
para cell [
J I 70000nm
Fig. 1. Schematicdiagramof a humanhair fibre showingthe major structural featuresexcept pigmentgranuleswhichare normally found in the cortical cells. (Adapted from Marshall, 1985.)
Fig. 2. Transmission electron micrographs at different magnifications of transverse sections of a Romney wool fibre stained by reduction and osmication. Ortho-cortical (0), meso-cortical (M) and para-cortical (P) cells are indicated. (a) Differentiation is shown at low magnification. Cytoplasmic and nuclear remnants are shown by arrows. Cu, cuticle. Scale marker indicates 2/tm. (b) Parts of ortho- meso- and para-cortical cells are shown. The characteristic whorl-like arrangement of ortho-cortical IFs, and the random packing of para-cortical IFs are evident. Areas of hexagonally-packed IFs (arrows) are apparent in the meso-corticat cells, cmc, cell membrane complex. Scale marker indicates 0.1/~m. 50
Mammalian Hard Keratin
Fig. 3. (a) Scanning electron micrograph of a Romney wool fibre showing the cuticle scale pattern. Scale marker indicates 10/~m. (b) Transmission electron micrograph of a longitudinal section of a wool fibre cuticle cell showing the a-layer (arrows), exocuticle (exo) and endocuticle (endo). Co, cortex. Scale marker indicates 0.5 #m. (Reproduced from Woods and Orwin (1980), with permission of Academic Press.) more extensive overlapping of fibre cuticle cells occurs on the para-cortical side of the fibre (Makinson, 1978). The main morphological features of the cuticle (Fig. 3b) are the sulphur-rich exocuticle and the endocuticle. Cuticle cells are restricted to wool and hairs. Echidna quills, porcupine quills and most coarse fibres are composed of elongated cortical cells with a central core of block shaped medullary cells. For fibres from different animal species, the columns of medullary cells show a wide
variety of forms and frequencies (Wildman, 1954; Appleyard, 1960; Brunner and Coman, 1974; Teasdale, 1988). The highly vacuolated cytoplasm of the medulla gives rise to reflective properties in the dry fibre, which differ from those of an unmedullated fibre.
C. Subcellular Components In cortical cells, nuclear remnants and the typical IF/matrix complex are seen (Fig. 2b). The
R.C. Marshall et al.
IFs appear as lightly stained circular areas surrounded by a heavily stained featureless matrix. Some workers have observed a partially resolved internal structure within the IF giving the appearance of a ring and core (Fraser et al., 1972). The inter-IF spacing, and hence the relative proportion of matrix (volume fraction 28-56%), varies between keratins, but this distance and the IF diameter (7.2-7.8 nm) can only be accurately measured by X-ray diffraction of unstained specimens (Fraser et al., 1973). Both the cuticle and cortical cells are bounded by membranes which together with the intercellular material are known as the cell membrane complex. In the micrograph in Fig. 2b, the stained region of the cell membrane complex is termed the b-layer (approximately 15 nm wide) while the lightly stained or inert regions are termed the fl-layers. The cuticle and cell membrane complex play a more important role in determining certain fibre properties than would be expected from their proportions (Marshall, 1990). The cell membrane complex of wool fibres affects, for example, mechanical properties such as abrasion resistance, wrinkle recovery and resistance to torsional fatigue, and chemical properties such as resistance to attack by acids, proteolytic enzymes and diffusion and location of reagents within the fibre (Leeder, 1986; Marshall, 1990). Within the exocuticle, there is a layer of protein of very high sulphur content, about 40 nm thick, which is termed the a-layer (Figs 1 and 3b) (Lagermalm, 1954; Bradbury, 1973; Swift, 1977; Orwin, 1979; Chapman, 1986). The loss of the a-layer of the cuticle is probably the main cause of the weathering of trichothiodystrophic hair (Gummer et al., 1984).
IlL STRUCTURE AND BIOCHEMISTRY OF HARD KERATINS A. Composition 1. Protein
The hard ~-keratinous tissues are mainly protein but cover a very large range in compositions with
amino acid Lysine Histidine Arginine Aspar tic Acid Threonine Serine Glutamic Acid Proline
Glycine Alanine Half-Cystine Valine Methionine Isoleucine
Leucine Tyrosine Phenylalanine 0
residues % l
~ ] ~ Horn Range
Fig. 4. A schematicdiagram showingthe range in amino acid compositionfor a largenumber of hair and horn-likekeratins. significant variability in the proportion of nearly every amino acid. This is primarily due to their construction from variable proportions of cuticle, IFs, matrix and medulla. Analyses of a large number of different keratins indicate that they are relatively poor in methionine and histidine but rich in serine, glutamic acid, glycine, half-cystine and leucine (Fig. 4). The range in composition for each amino acid is frequently larger for horn-like keratins than for hair keratins. 2. Lipid
Wool contains about 1% lipid comprising mainly fatty acids, sterols, ceramides, triglycerides and glycolipids (for reviews, see Leeder, 1986; Marshall, 1990). The fatty acids are generally saturated C~4 C24 (mainly with an even number of carbon atoms and predominantly palmitic acid and stearic acid), with double and triple bonds being occasionally present. Cholesterol is the main component of the sterol fraction, and its precursor, desmosterol, is located in the keratinized wool fibre mainly in the cortex. The polar lipid cholesteryl sulphate has also been identified. It has been calculated that phospholipids constitute less than 1% of the total lipid (for reviews, see Leeder, 1986; Marshall, 1990), a result consistent with
Mammalian Hard Keratin the appreciable decrease in phospholipid content during keratinization (Braun-Falco, 1958). B. Solubility of Hard Keratins
Hard keratins are highly crosslinked polymers which can only be appreciably solubilized if covalent bonds, either peptide or disulphide, are broken. The most specific way to achieve solubilization is to break the disulphide bonds by reduction with reagents such as mercaptoethanol, thioglycollic acid or dithiothreitol. Complete reduction usually requires an excess of reducing reagent, a mildly alkaline pH and the presence of reagents such as concentrated urea, guanidine hydrochloride or lithium bromide to swell the keratin and render the disulphides accessible to the reagent. These dispersing reagents are also needed to dissolve the proteins. Since keratin proteins in the reduced state are readily reoxidized, the sulphydryl groups are generally blocked, usually with charged reagents such as iodoacetate which give the most soluble products (Gillespie, 1983). With Merino wool, the solubility frequently reaches 80%, while human hair is less soluble usually in the range 50-75% (Gillespie, 1983). Most of the soluble proteins originate from the IF/matrix complex as minor components are much less soluble. Solubilities as low as 5% for human hair have been reported, and in some cases low solubility appears to be an inherited trait (Gillespie, 1983). Solubility may be greatly reduced by mild alkali treatment, heating, cosmetic or textile treatments, or short exposure to sunlight (Kearns and Maclaren, 1979; Marshall et al., 1983; Leaver et al., 1985). However, severe treatments such as long exposure to sunlight (Leaver et al., 1985) or temperatures above 180°C (Marshall et al., 1983) actually increase the solubility probably due to peptide bond cleavage. C. Cellular and Subcellular Components 1. Cortex
Many hairs contain two or more types of cortical cells which, in the case of wool, are referred to
as ortho-, meso- and para-cortical cells (Fig. 2b). In fine Merino wool, ortho- and para-cortical cells are arranged bilaterally, but hair and the coarser wools have other arrangements (Fraser and Rogers, 1955; Rogers, 1959a; Kaplin and Whiteley, 1978; Orwin et al., 1984). Apart from their ease of staining with dyes such as methylene blue, the distingishing features of the ortho-cortical cells are macrofibrils (IF/matrix bundles) (Fraser et al., 1972) which are discrete and twisted along their long axis, and IFs which produce a whorl pattern because of twisting of the peripheral IFs around the central core (Fraser et al., 1972). In the more poorly-stained para-cortical cells, the macrofibrils appear to be fused and poorly defined especially around the cell periphery, with IFs in a pseudohexagonal array (Fraser et al., 1972). In meso-cortical cells, the IFs are hexagonally arranged (Kaplin and Whiteley, 1978, 1985). There is some evidence that para-cortical cells contain the most matrix, are the richest in high-sulphur proteins and are the repository of the ultra-high-sulphur (UHS) proteins (Fraser et al., 1972; Marshall and Gillespie, 1989; Dowling et al., 1990). The ortho-cortical cells, on the other hand, appear to contain all of the high-tyrosine proteins (Fraser et al., 1972; Kaplin and Whiteley, 1978; Dowling et al., 1990). There are many proteins present in extracts of keratin. The exact number is uncertain, and estimates of more than 100 have been made (Powell and Rogers, 1986). The proteins may be separated by chemical fractionation (Fig. 5) or by gel electrophoresis (Fig. 6) into multi-membered protein classes (IF, high-sulphur and high-tyrosine), each distinguished by a characteristic and unique amino acid composition (Table 1). The IF polypeptides alone contain a-helices and are richest in those amino acids favouring c~-helix formation, namely lysine, aspartic acid, glutamic acid and leucine, and are comparatively poor in half-cystine and proline. The high-sulphur proteins contain by far the most half-cystine and proline, and these amino acids together with threonine and serine constitute about two-thirds of the amino acid residues. Methionine is usually absent, while only small amounts of lysine, histidine and the aromatic amino acids are usually
lg * lOOml 0.2M mercaptoethanol in 8M urea, pH 10.6 2h, 40 C
0.4g i o d o a c e t i c acid (keep pH at 8-9 with tris), * zinc acetate to O.02M to dialysed protein (pH 5.8-6.0)
dissolve in O.02M sodium citrate
HT t y p e I proteins to 1% protein, 4 2 vol acetone, 0.3 vol sat ammonium sulfate
HT t y p e II proteins
IF p o l y p e p t i d e s
Fig. 5. A schematic diagram showing the isolation of the main classes of wool proteins.
found (Table 1). A wide range of half-cystine content is found in the high-sulphur proteins, and the proteins with half-cystine contents towards the upper limit are termed the UHS proteins (30-35 residues % of half-cystine). The two types of high-tyrosine protein differ in halfcystine content but both are very rich in glycine and tyrosine with 60-70% of the composition being contributed by these two amino acids and serine. The protein families may also be distinguished by size. The IF subunit polypeptides are the
largest with a molecular weight range from 45,000 to 58,000 (Woods, 1979). The high-tyrosine proteins are the smallest, all less than 10,000 (Gillespie, 1983) while the molecular weights of the high-sulphur proteins are between 10,000 and 30,000 (Gillespie, 1983). Estimates for the molecular weights of the UHS proteins range from 16,000 to 28,000 (Marshall and Gillespie, 1976b; MacKinnon, 1989). Further characteristics of the cortical proteins as well as factors affecting their variation in keratins, are given in Section VI.
Mammalian Hard Keratin
keratin (e,g. wool)
reduction and solubilization - S S - - ~
-SH-P - SCH2COO-
no protein fractionation
l IItl I SDS lowsulfur
& o oo
Fig. 6. Electrophoretic patterns of the constituent IF polypeptides, high-sulphur and high-tyrosine wool proteins, as well as a two-dimensional electrophoretic pattern of the unfractionated extract.
Cuticle cells have a laminated structure comprising the epicuticle, exocuticle and endocuticle (Figs 1 and 3b). The epicuticle (about 3 nm thick and not visible in standard electron microscopical preparations) is the outermost layer of the cuticle and is presumably derived from the plasma membranes of immature cuticle cells. It plays a dominant role in determining the surface chemistry of the fibre (Leeder, 1986). An analysis of the layer indicates that it contains about 80% protein, 5% lipid and 4% inorganic material (Bradbury, 1973). Apart from containing a high amount of cystine, the protein component of epicuticle contains high levels of serine, glycine and glutamic acid. Studies show that it is a hydrophobic, chemically resistant semi-permeable membrane. It envelops individual
cuticle cells rather than covering the whole fibre, and this was evident from experiments in which sacs were formed on the surface of wool fibres and isolated intact cuticle cells when they are placed in chlorine water (Allworden reaction) (for reviews, see Bradbury, 1973; Leeder, 1986). The amino acid composition of cuticle isolated from wool is given in Table 2. Compared with Merino wool and its cortex, the cuticle samples contain considerably more half-cystine, serine and proline, less aspartic acid and phenylalanine, and a small amount of citrulline. About one-third of the cuticle can be obtained in a' soluble and stable form by alkaline reduction in urea followed by alkylation with iodoacetate (Ley and Crewther, 1980). These soluble proteins are similar in composition to the high-sulphur proteins of the cortex but electrophoresis shows
R.C. Marshall et al. Table 1. Amino acid compositions (as residues%) of wool and its constituent proteins Amino acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine
Wool I 2.7 0.8 6.2 5.9 6.5 10.8 11.1 6.6 8.6 5.2 13.1 5.7 0.5 3.0 7.2 3.8 2.5
0.6 0.7 6.2 2.3 10.2 13.2 7.9 12.6 6.2 2.9 22.14 5.3 0.0 2.6 3.4 2.1 1.6
0.9 1.3 6.9 0.6 11.1 12.7 7.9 12.8 4.2 2.0 29.94 4.3 0.0 1.7 1.3 1.9 0.5
0.4 1.1 5.4 3.3 3.3 11.8 0.6 5.3 27.6 1.5 6.04 2.1 0.0 0.2 5.5 15.0 10.3
0.4 0.1 4.7 1.8 1.7 10.9 0.7 3.0 33.6 1.1 9.84 1.4 0.0 0.2 5.3 20.3 4.5
4.1 0.6 7.9 9.6 4.8 8. I 16.9 3.3 5.2 7.7 6.0 4
6.4 0.6 3.8 10.2 2.7 2.0
IGillespie and Marshall (1980). 2Gillespie (1983). 3Marshall et al. (1980). 4Measured as S-carboxymethylcysteine. that they share no c o m m o n c o m p o n e n t s (Ley et al., 1985; M a r s h a l l a n d Ley, 1986). The soluble cuticle proteins are extremely heterogeneous a n d cover a wide range in m o l e c u l a r weights. 3. M e d u l l a
The m e d u l l a (illustrated by guinea pig hair m e d u l l a in T a b l e 2) differs strikingly from the cortex by the virtual absence of half-cystine, the extreme richness in glutamic acid (41 residues%), a n d the presence of citrulline, a n d by its insolubility in disulphide b o n d - b r e a k i n g reagents. The insolubility is due to cross-linking of polypeptide chains with 7-glutamyl-c-lysyl isopeptide linkages. T h e proteins o f the i n n e r root sheath also c o n t a i n citrulline a n d isopeptide b o n d s (Rogers, 1983; Powell a n d Rogers, 1986). The presence of citrulline a n d its d e g r a d a t i o n p r o d u c t o r n i t h i n e in hydrolysates of keratins provides a useful i n d i c a t i o n o f the relative a m o u n t of m e d u l l a present. 4. Cell m e m b r a n e c o m p l e x
Observations made during development (for reviews, see B r a d b u r y 1973; Leeder, 1986) have s h o w n that the /~-layers of the cell m e m b r a n e
complex in the fibre are formed from the p l a s m a m e m b r a n e s of a d j a c e n t living cells in the follicle. As the m e m b r a n e s b e c o m e 'modified', the f - l a y e r material is laid d o w n between the cells a n d provides adhesion between them. Virtually n o t h i n g is k n o w n a b o u t the origin of this layer or its relationship to the cell prior to keratinization. The g-layer Table 2. Amino acid compositions (as residues%) of wool cortex and cuticle prepared after agitation of wool in formic acid for 1 min, and guinea pig hair medulla Amino acid
2.7 0.6 6.8 6.8 6.0 10.1 11.9 trace 6.4 9.7 5.3 9.7 5.2 0.4 3.0 7.9 4.4 3. I
2.6 0.8 4.6 3.0 4.6 15.1 8.3 0.2 10.9 8.8 5.6 17.4 6.7 0.3 2.0 5.4 2.7 1.3
6.1 0.9 3.5 4.5 1.6 2.3 41.0 23.6 0.0 2.9 2.1 0.0 1.6 0.2 1.0 6.7 0.8 1.6
Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Citrulline Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine ILey et al. (1988). 2Rogers (1983).
Mammalian Hard Keratin between cuticle ceils differs in composition from that between cortical cells (see Leeder, 1986, for a review) and there may even exist differences between the 6-layers found in the ortho- and para-cortex. After solubilizing keratin as completely as possible with a reducing agent in an alkaline solution of a disaggregating agent, a resistant membrane is obtained which contains approximately the same proportion of half-cystine as whole wool. This residue originates from the /~-layer and is very resistant to chemical and biological attack and to reaction with electron microscopical stains. The nature of this resistance is unknown but could be due to a particular arrangement of disulphide bonds or to an unidentified crosslink (Leeder, 1986). The amino acid composition of the cell membrane complex has been reported by a number of workers (for reviews, see Bradbury, 1973; Leeder, 1986). Differences exist between the results, which may be due to the method of preparation involving different extraction procedures and degrees of contamination. D. Morphological Location of the Proteins
The exact location of constituent proteins within keratins was for some time a matter of controversy, largely because no pure morphological component had been isolated and analyzed. A large body of electron microscopical and X-ray diffraction evidence linked the low-sulphur proteins with IFs and the high-sulphur proteins with the matrix. In the mid-1970s, IFs were isolated and the composition of the components confirmed by amino acid analysis and electrophoresis (Jones, 1975, 1976). Reassembly of soft keratin IFs from constituent polypeptides was successful (Steinert and Cantieri, 1983), but despite considerable efforts on a number of occasions, reassembly of hard keratin IFs proved difficult. Recently two groups have reported the formation of filaments (Heid et al., 1986; Thomas et al., 1986), but so far there has been no assessment of protein composition or comparison with native filaments. The location of the high-tyrosine proteins has not been determined directly. These proteins have
not been found in the cuticle (Bradbury, 1973; Marshall and Ley, 1986) and their proportion in many keratins (for example, 14% in some wool samples, 30% or more in echidna quill) is too large to be accommodated other than in the cortex (Gillespie, 1972a). Their absence in isolated IFs (Jones, 1975, 1976) and cell membrane complex (Bradbury, 1973), and the high correlation between the inter-IF volume fraction (matrix) of a range of keratins and the mass fraction of the constituent high-sulphur and high-tyrosine proteins (Fraser et al., 1973) makes it certain that they are only located in the matrix. E. Disulphide Bonds
In normal keratin, most of the half-cystine residues (> 95%) in IFs and matrix are involved in disulphide linkages. Mechanical strength, elastic properties, and chemical and biological resistance of keratins stem directly from the three dimensional structure stabilized by the disulphide bonds. In view of their importance, it is surprising that so little is known about their formation and overall disposition. The involvement of copper in the catalytic oxidation of sulphydryl to disulphide in wool and hair has long been known from observations such as the adverse effect on fibre properties from copper deficiency. However, the nature of the catalytic molecule is still unknown (Danks, 1983; Gillespie, 1983). In a severely copper-deficient subject, 50-70% of disulphides are still formed, suggesting that some disulphides, perhaps because of their molecular environment, do not require the intervention of a copper-based catalyst for their formation (Gillespie, 1990). For the high-sulphur proteins containing the pentapeptide repeating unit cys~-cys2-x-pro-y (Section VI.B. 1), it has been suggested on theoretical grounds that an intra-chain disulphide bond linking cysteine-2 of the first pentapeptide unit to cysteine-I of the second unit, is the preferred configuration. As a result, the peptide backbone of the protein is stabilized into a convoluted pattern (Parry et al., 1979; Fraser and MacRae, 1980). Fraser et al. (1988) have calculated that about 80% of the disulphides in human hair are
R.C. Marshall et al.
accommodated within the matrix, the majority of which are probably intra-chain. The locations of disulphides within the IFs have been investigated by Fraser et al. (1988). On the basis of data for globular proteins and on stereochemical constraints derived from the study of model compounds, they postulated that intramonomer and intra-dimer disulphides are unlikely but inter-dimer disulphides, presumably stabilizing the tetrameric structural unit, are likely. Nothing is yet known of IF-IF or IF-matrix types of disulphide linkages.
There is evidence for the mechanical effects of the matrix in that the transverse compressional inflexion modulus in water at 20°C is a function of the total content of high-sulphur and high-tyrosine proteins. The transverse swelling of keratins at 20°C from the dry to the fully swollen state in water or formic acid, is inversely related to the matrix content. These relations are generally regarded as due to the solvent-excluding properties of the matrix proteins in the IF/matrix lattice (Bendit and Gillespie, 1978; Bendit, 1980; Fraser and MacRae, 1980).
F. Relationship to Physical Properties IV. D E V E L O P M E N T OF H A R D KERATIN
Hard keratins are described mechanically as filament/matrix composites whose general mechanical properties arise from the properties of their component parts. These are listed as due to the mechanical properties of the filaments and matrix, filament length, filament orientation and packing, proportion and type of matrix and adhesion of matrix to filaments (Fraser and MacRae, 1980). This composite is not continuous but interrupted at cell boundaries. Attempts have been made over many years to develop a unified theory linking keratin structure and mechanical properties, but none has been very successful. Hearle and Susutoglu (1985) summarized the best information available to them, stating that longitudinal mechanical properties of the wool fibre, in extension and recovery, can be explained by a model involving fibrillar units (presumably IFs) joined at intervals along their length (presumably by disulphide bonds) to the matrix. It would be expected that changing the ratio of filament to matrix with a concurrent change in the extent of IF/matrix bonding would alter these mechanical properties. So far no experimental test of this hypothesis has been made. However there is good evidence that after removal of the effects due to fibre diameter, wool shows a comparatively wide range in fibre strength (reviewed by Orwin et al., 1985). One cause of such variability which might well reflect changes in the IF/matrix ratio, stems from variation in the ratio of ortho- to para-cortex (Orwin et al., 1985).
A. Follicle Structure
Hard keratin development of fibres takes place within wool and hair follicles. The general morphology and cytology of the follicle have been reviewed by Swift (1977), Chapman and Ward (1979), Orwin (1979, 1989) and Chapman (1986). Follicles are tubular downgrowths of the epithelium into the dermis (Fig. 7). The dermal components of the follicle are the connective tissue sheath (CTS) surrounding this downgrowth and the dermal papilla which is continuous with the CTS but enclosed within the follicle bulb. A basement membrane separates the epithelial-developed cells from the dermal components. Blood vessels, fibroblasts and, on occasion, mast cells are found in the CTS. These may also be present in the dermal papilla although blood vessels are generally restricted to larger papillae. Vibrissae, when present, have a thickened collagenous capsule containing blood sinuses as a surrounding extension of the CTS and a complex innervation. Accessory structures, i.e. sebaceous glands, sudoriferous glands and arrector pili muscles, may be present on some follicles. Fibre formation is the primary function of the epithelium-derived cells of the follicle although this is also dependent on the presence of the specialized population of fibroblasts of the dermal papilla (Oliver and Jahoda, 1989). The process is a dynamic one requiring cell proliferation, relative
Mammalian Hard Keratin
"'~-o~Zs- ........ •~ - O R ~ I F I C E CELLS
f C~TTTcXULTRA,STRUCTURALLYKERATINIZSO NOT ALL CORTICAL CELLS SHOWN APPRC0(. DIMENSIONS:
m m wide
2 . 3'7~IOP41 O-17mm wide
',',,_I / F SLOUGHING
U ~ l n SCOTSHEATH:
Fig. 7. Diagrammatic representations of non-medullated and medullated wool follicles showing the follicle zones referred to in the text. (Reproduced from Orwin (1979), with permission of Academic Press.)
R.C. Marshall et al.
cell movement, cell migration, differentiation of several cell lines and types and degradation of unwanted cells when fibre formation is complete (Orwin, 1989). The general sequence of events in fibre formation is similar in all follicles. Cells derived from a population of dividing cells in the bulb (zone A) move up the follicle towards the skin surface (Fig. 7). Above a 'critical level' (Auber, 1952; Orwin, 1971; Woods and Orwin, 1982) the potential for cell division is lost and further stages in differentiation become apparent. These are characterized by cell growth and shape changes in zones B and C; keratin synthesis in zones B, C and D; keratinization, stabilization and dehydration of the fibre in zone E; cornification and degradation of some non-fibre cell lines in zone F (Gemmell and Chapman, 1971; Pinkus et al., 1981; Orwin and Woods, 1985) to set the fibre free in the follicle neck (zone G) prior to its emergence above the skin surface. Within this general sequence, differentiation of each cell line or type is highly specific and may not follow all steps of this sequence. Differentiation of up to ten cell lines may take place from the population of dividing cells in the bulb. The outermost layer forms the outer root sheath (ORS) (Fig. 7); the next layer forms the
companion cell layer (Orwin, 1971; Ito et al., 1986a); then follows the inner root sheath (IRS) which is made up of Henle's, Huxley's and the IRS cuticle layers; the fibre cuticle; the fibre cortex which may be made up of ortho-, meso- and para-cortical cells and finally the central medulla. Consequently, only a relatively small proportion of germinative cells actually form the fibre cortex. Follicles lie at an angle to the skin and may vary in shape from straight to curved (Wildman, 1932; Nay, 1966), and it has been suggested that this may be a factor controlling crimp formation. Wool follicles in which the bulbs are deflected at an angle to the main axis of the follicle (Fig. 8) have a multilayered ORS and Henle's and Huxley's cells are larger on the side of the deflection (Auber, 1952). Hard keratin is the major component of cortical cells (Fraser et al., 1972), and the remainder of this section is primarily concerned with keratin development in these cells. B. Ultrastructural Development o f H a r d Keratin 1. Initial stages
The cells which differentiate into cortical cells derive from a cone-shaped population of mitotically
g l a n o
@ Fig. 8. Diagrammatic representationof skin showingwool folliclegroups. Scalemarker indicates 1 mm. (Reproduced from Orwin (1989) with permission of Chapman and Hall.)
Mammalian Hard Keratin active cells in the bulb (zone A) (Auber, 1952; Fraser, 1965). In follicles producing non-medullated fibres, the cortex is formed from bulb cells located around the dermal papilla in the upper region of the bulb and, in medullated fibres, from those adjacent to the middle, i.e. widest region, of the dermal papilla (Auber, 1952; Epstein and Maibach, 1969) (Fig. 7). While presumptive cortical cells in the bulb show no ultrastructural evidence of hard keratin formation, other forms of differentiation have been reported including alkaline phosphatase (Lyne and Hollis, 1967) and adenosine triphosphatase activities (Chapman and Gemmell, 1971b). In terms of ultrastructure, presumptive cortical cells in the bulb are characterized by large nuclei, numerous ribosomes and mitochondria, relatively small amounts of endoplasmic reticulum, vacuoles, lysosomes, Golgi complexes and coated vesicles (Orwin, 1979). Intercellular spaces are numerous and plasma membrane differentiations include desmosomes and gap junctions (Happey and Johnson, 1962; Roth and Helwig, 1964; Orwin et al., 1973a, b). Generally, presumptive cortical cells are ellipsoid in shape, while those adjoining the dermal papilla are columnar (Birbeck and Mercer, 1957). The proportion of sheep follicle stem cells which differentiate to cortical cells, one factor determining the efficiency of fibre formation, has been variously estimated to be in the range 10-40%. Many factors appear to be involved in controlling this proportion including breed of sheep, follicle geometry, time of day, and perhaps level of nutrition although this is subject to controversy (Schinckel, 1962; Short et al., 1965; Wilson and Short, 1979; Weinstein and Mooney, 1980; Hynd et al., 1986; Williams and Winston, 1987; Hynd, 1989). 2. Development in the cortex
Keratin development takes place in zones B, C and D (Fig. 7). The early stages of development in zone B occur when the cells are increasing in both volume and length (Auber, 1952). This is associated with the presence of microtubules, many of which are preferentially aligned parallel to the axis
of the follicle (Orwin, 1969; Orwin and Thomson, 1973). By mid zone C, the basic elongated shape of these cells has been established. Interdigitations with neighbouring cells at their ends but not their sides are a characteristic of cortical cells (Rogers, 1959a). The general ultrastructural development of cortical keratin has been described in several studies (see Orwin, 1979, for a review). IFs about 7.5nm in diameter are first seen in the cytoplasm at the start of zone B (Fig. 9a) as bundles. Increases in length and diameter of these bundles result from the addition of IFs. Initiation of IF bundles seems limited to zone B. Increases in keratin content in zones C and D result from increases in the diameter of existing bundles, now referred to as macrofibrils (Fig. 9b), rather than initiation of new ones (Forslind and Swanbeck, 1966). Microfibrils in the developing ortho-cortical cells appear to be twisted about their long axis (Birbeck and Mercer, 1957). Finally, macrofibrils form an interconnecting network oriented along the length of the cell. IF bundles may be initiated in association with desmosomes, presumably because of desmoplakin/ IF interactions (Skerrow, 1986; Franke and Heid, 1989), or in the cytoplasm (Roth and Helwig, 1964; Chapman and Gemmell, 1971a; Orwin et al., 1973b). The role of desmosomes in determining the general architecture of IF orientation and numbers is uncertain but may be important as desmosomes are at their maximum numbers, making up about 5% of the cortical cell surface, in zone B (Orwin et al., 1973b). The orientation of IFs is initially random but by upper zone B is parallel to the axis of the fibre (Birbeck and Mercer, 1957; Roth and Helwig, 1964; Chapman and Gemmell, 1971a). This occurs in conjunction with the cell shape changes described previously. Considerable differences exist between orthoand para-cortical cells in the way in which IF and matrix form. The presence of tactoid-like filament assemblies in human hair follicles (Jones and Pope, 1985) may indicate the mechanism by which these differences arise. In upper zone B of a wool follicle, the IF bundles in ortho-cortical cells are shorter and thinner than those in para-cortical cells, and
R . C . Marshall et al.
Fig. 9. Transmission electron micrographs of longitudinal sections of differentiating cortical cells. Scale markers indicate 2 t~m. (a) Low zone B. Groups of developing IFs (macrofibrils) are visible in the cytoplasm (arrowheads) or associated with desmosomes (arrows). (b) Low zone C. The macrofibrils (arrows) have increased in size and are oriented parallel to the fibre axis.
Mammalian Hard Keratin are more numerous in the cytoplasm in contrast to those in para-cortical cells where they are preferentially aligned along plasma membranes (Orwin, 1979). Individual IFs are much easier to resolve in the bundles in para-cortical cells. In zone C, the matrix can only be seen between IFs of paracortical cells. By zone D, the two cell types have acquired the IF/matrix arrangement characteristic of mature cells of the fibre (Chapman and Gemmell, 1971a) (Section V.A). During differentiation and maturation, Short et al. (1965) estimated that cortical cell volume increases three-fold and dry mass by 13-fold. By zone E, keratin synthesis has ceased and a number of chemical and physical changes take place leading to a mature fibre. Most cystine residues are oxidized to give cystine cross-links both within and between protein chains (Gillespie, 1983). Thus disulphide cross-links lead to the characteristic insolubility and chemical inertness of the fibre. There is loss of osmiophilic character and most non-keratinous cellular constituents (for example nuclear remnants, lysosomes, mitochondria, ribosomes) are degraded to some extent (Birbeck and Mercer, 1957; Rogers, 1959a, 1964; Chapman and Gemmell, 1971a; Orwin, 1976). Finally the fibre is dehydrated leading to considerable shrinkage. Changes in zone E are often referred to as keratinization. No further changes have been reported above zone E. Although both DNA and RNA have been shown to be degraded during keratinization (Downes et al., 1966a), sufficient high molecular mass DNA (Higushi et al., 1988; Kalbe et al., 1988; Schreiber et al., 1988) remains in the keratinized hair fibre, to allow extraction and DNA typing. 3. Development of the fibre cuticle Development of the fibre cuticle takes place in two main stages. Between zones A and lower C, the fibre cuticle cell undergoes a series of shape changes which result in its flattened form (Fig. 10). This occurs in conjunction with shape changes in the IRS cuticle which impose the scale pattern shape on the fibre cuticle surface. Fibre cuticle surface ridges occur where two IRS cuticle cells are
apposed (Woods and Orwin, 1982). In the second stage, synthesis of the exocuticle proteins occurs in zones C and D, stabilizing the scale pattern (Woods and Orwin, 1982). Considerable variation in the scale pattern occurs particularly between hair samples from different species, although smaller variations can occur along the same fibre, between different fibres of the same sample, and between fibres of different samples (Wildman, 1954; Appleyard, 1960; Brunner and Coman, 1974; Woods and Orwin, 1982; Orwin and Woods, 1983). These variations reflect differences in the shape of the IRS cuticle cell surfaces apposing the fibre cuticle cell surface, and include surface roughness, and ridge height, slope and frequency. Some examples of the number of scale ridges per 100 #m are wool 4.5-10 (Orwin and Woods, 1983), cashmere 4-11 and yak 7-14 (Phan et al., 1988). By comparison with cortical keratin development, the formation of the exocuticle shows quite distinct differences. While a cytoskeletal/ filamentous system is present during shape changes and seems involved in the transport of exocuticle proteins to their final location in the cell, it does not play an integral role in the exocuticle structure itself (Happey and Johnson, 1962; Roth and Helwig, 1964; Bradbury, 1973; Swift, 1977; Woods and Orwin, 1982). Precursors of a-layer and exocuticle protein appear sequentially in the cytoplasm as globules of three distinct sizes. The first to appear are the smallest and come to lie against the peripheral plasma membrane where they presumably form the a-layer. The other two sizes of globules follow the same pathway (largest last) where they are transformed into a lamellar network on keratinization at the top of zone D. The remainder of the cytoplasm condenses at the same time to form the endocuticle (Woods and Orwin, 1980). 4. Development of the medulla Medulla cells originate from around the top of the dermal papilla but appear to have no mitotic cell region associated with their production (Auber, 1952). They form a central column in fibres with a wide variety of forms and frequencies
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Fig. 10. Transmission electron micrographs of longitudinal sections of differentiating fibre cuticle (Cu) and inner root sheath cuticle (IR) cells. Scale markers indicate 2 #m. (Reproduced from Woods and Orwin (1982), with permission of Academic Press.) (a) Mid zone B. The two cell types are quite regular in shape and lined up one for one. (b) Mid zone C. The cells are now elongated parallel to the fibre axis, and scale ridges are apparent on fibre cuticle cells adjacent to the apposed IRS cuticle plasma membranes (arrows). Co, cortex; Hu, Huxley's layer.
Mammalian Hard Keratin found within and between species (Wildman, 1954; Appleyard, 1960). Their development is less well characterized than other cell lines but is known to have distinctive features compared to cortical cell development. These are the formation of many vacuoles which may coalesce during differentiation and the production of medullary granules (Rogers, 1964; Roth and Helwig, 1964; Parakkal, 1969; see Orwin, 1979, for a review). These granules are amorphous and gradually increase in size until they transform into a hardened mass around the vacuoles and against the cell periphery in zone E. The vacuolar spaces are, therefore, preserved in the medulla cells of the keratinized fibre. Filaments are not associated with the transformation of these granules (Rogers, 1983) although they have been reported in human hair (Clement et al., 1980). 5. Development of the cell membrane complex The cytoplasmic components other than keratin, present in zone A cortical cells, are also present throughout zones B, C and D. The most relevant ultrastructural changes which occur in these zones appear to be modifications to the nucleus and plasma membranes (see Orwin, 1979, for a review). The plasma membranes of cortical and cuticle cells undergo differentiation as the cells pass through zones A to D. Ultrastructural changes beyond zone D/low zone E are minimal and it seems probable that by zone E the plasma membranes are in their final modified and stabilized form, and are then known as the cell membrane complex (Rogers, 1959a). These plasma membrane modifications clearly have a major function in enabling the cellular integrity of the fibre to be maintained in conditions of varying humidity. C. Biochemistry of Hard Keratin Development 1. Follicular site of synthesis of IF and matrix Several approaches have been used to establish the form in which IF polypeptides and matrix proteins are synthesized and the mode by which this occurs. One approach has been to compare the type of proteins found in follicles with those
extracted from the corresponding fibres. Roots obtained by plucking fibres from skin have been commonly used in these studies as the unkeratinized regions of the follicle left attached to the fibre usually include the differentiating cortical, cuticle and IRS cell lines down to, but usually excluding the mitotic zone (zone A). Examination of the urea-soluble proteins from plucked roots showed the presence of IF polypeptides similar to those of the mature fibre, and many corresponding high-sulphur proteins (Rogers, 1959b; Downes et al., 1966b; Clarke and Rogers, 1970; Steinert and Rogers, 1973a; Heid et al., 1986). Jones (1987) obtained similar results with dissected portions of developing human hair root tissue. The UHS and high-tyrosine proteins appear to be synchronized late in fibre development and are not ureaextractable. Autoradiographic studies of the incorporation of labelled amino acids over time have confirmed and expanded these findings. [35S]-cystine, for instance, is taken up within minutes in zones A and B following intravenous or subcutaneous injection. Subsequently, the label is incorporated into fibrils in the cytoplasm of the cortical cells in zones C and D reaching maximum uptake in 4--8hr post-injection (Bern et al., 1955; Ryder, 1958; Downes et al., 1962; Nakai, 1964; Forslind, 1971; Chapman and Gemmell, 1973). Isolation of labelled IF polypeptides and high-sulphur proteins from wool roots following injection of labelled [35S]-cystine has conclusively demonstrated that the labelled amino acid has been incorporated into the keratin proteins in these regions (Downes et al., 1963) (see also Section IV.C.2). At longer time intervals, radioactive label can be detected in the keratinized fibre (Bern et al., 1955; Ryder, 1956; Downes et al., 1967). Similar findings have been reported for the uptake of other labelled amino acids such as tyrosine (Sims, 1964) and leucine (Downes and Wilson, 1971), and also methionine which is found in IF polypeptides only (Wilson et al., 1971). The relatively rapid passage of labelled amino acids into cortical cells in regions ahead of cell movement may be facilitated by the extensive areas of gap junctions in the plasma membranes of differentiating cortical cells (Orwin et al., 1973a).
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Fig. 11. Immunogold electron microscopyof wool folliclecortex treated with an antibody against Type II IF polypeptide. Longitudinal section of keratin IF bundles. Scale marker indicates 100 nm. (Reproduced from J. Cell Biol. 1986, 102, 1412-1418, by copyright permission of the RockefellerUniversity Press.) A second approach has used immunolocation techniques with antibodies prepared against hard keratin IF polypeptides and three families of matrix proteins. Figure 11 shows the binding of monoclonal antibodies against wool Type II IF polypeptides to IF bundles in the developing wool fibre cortex. With this technique, both Type I and Type II hard keratin IF polypeptides are first detected in the upper bulb region, one or two cell layers out from the top of the dermal papilla and at maximum intensity in zones C and D. The absence of reaction in sections at higher levels (zone E) is presumed to be due to antibody access being prevented by protein cross-linking in the keratinizing fibre (Kemp and Rogers, 1970; Baden et al., 1980; Baden and Kubilus, 1984; French and
Hewish, 1986; Heid et al., 1986, 1988a, b; Ito et al., 1986a, b; Lynch et al., 1986). In some studies, hard keratin IFs were apparently found in medulla, cuticle, IRS and ORS cell lines (French and Hewish, 1986; Heid et al., 1986; Ito et al., 1986a). This may reflect a lack of specificity or a generality of the epitopes recognized by the antibodies as there is no evidence for the presence of IFs in these tissues when mature. However, it is o f some significance that Powell et al. (1989) using highly specific in situ hybridization could find no evidence for the expression of hard keratin IF Type II genes in IRS or ORS. Double immunolabelling of follicle cells has shown that both hard keratin IF and other cytokeratins may be expressed in the same cell, for
Mammalian Hard Keratin example in some bulb cells around the dermal papilla (Heid et al., 1988a, b). As cytokeratins are not found in the mature fibre, their suppression must be one of the early events in hard keratin differentiation. High-sulphur proteins were detected in the more developed differentiating regions (zones C and D) of the follicle in the cortical and cuticle cell lines using antibodies raised against one family of high-sulphur proteins (SCMKB2) (Frater, 1976), mixtures of high-sulphur proteins (French and Hewish, 1987) or IF plus matrix (Lynch et aL, 1986). Monoclonal antibodies to an atypical hightyrosine protein bound specifically to similar regions of the follicle but only in ortho-cortical cells (Hewish and French, 1986). A third approach, which was initially directed at establishing whether synthesis of keratin proteins occurred via the classical RNA pathway, has been to characterize the products of in vitro translation of polysomes or mRNA isolated from wool roots. In studies using both homologous (Freedberg, 1970; Steinert and Rogers, 1971, 1973b; Wilkinson, 1971) and heterologous (Ward and Kasmarik, 1980; Rogers et al., 1981) translational systems, the in vitro synthesis of the component proteins of IF and matrix (high-sulphur and high-tyrosine) has been demonstrated. The location of mRNA coding for Type II wool IF polypeptides in cortical cells in zones B, C and D has also been demonstrated by in situ hybridization (Powell et al., 1989). The probe used was derived from a highly conserved region of Type II IF polypeptides and labelled both ortho- and para-cortical cells. Studies on wool follicles strongly indicate that the proteins are synthesized de novo by distinct mRNAs and that, apart from oxidative cross-linking extensive modification of the proteins, e.g. by proteolytic processing, does not occur. Two lines of evidence suggest that mRNA turnover is likely to be slow. Autoradiographic evidence indicates that the uptake of labelled nucleotides is low in the upper keratogenous zones (zone D) where protein synthesis is high (Sims, 1967; Fraser et al., 1972), while a study of polysome profiles has shown that inhibitors of RNA synthesis take a relatively long time to affect these profiles (Wilkinson, 1970).
2. Sequential synthesis o f I F and matrix proteins
In the lower levels of follicle, IFs without associated matrix have been detected using X-ray diffraction (R. D. B. Fraser, personal communication). Immunolocation and studies of radiolabelled cystine uptake in wool follicle and in the constituent proteins in the follicle also indicate a sequential synthesis in which IFs are deposited first; subsequently matrix proteins are laid down between them and further synthesis takes place concurrently (Rudall, 1956; Downes et al., 1963, 1966b; Lynch et al., 1986; French and Hewish, 1987; Jones, 1987). It is not possible to explain the contrary views of Chapman and Ward (1979). Recent work with a probe to a UHS protein has indicated that this protein is expressed at higher levels in the follicle than normal high-sulphur proteins (MacKinnon, 1989).
V. VARIATIONS IN KERATIN STRUCTURE A. Cortical Cell Types
Keratinized fibres show considerable complexity and diversity in the structural arrangement of IF and matrix within cortical cells (Rogers, 1959a; Bones and Sikorski, 1967; Kaplin and Whiteley, 1978, 1985; Orwin et al., 1984). The classification of cell types, based on Merino wool structure, has relied heavily on transmission electron microscope studies of transversely sectioned fibres (Fig. 2). It may be summarized as follows: (i) In ortho-cortical cells, the orientation of IFs in the centre of macrofibrils is typically parallel with the fibre axis. Towards the edges of macrofibrils, the IFs are tilted giving rise to a whorl-like appearance in cross-section. The IF/matrix ratio is high. The macrofibrils have the lowest diameter of the three cell types and are generally discrete with some cytoplasmic remnants between them. Nuclear remnants are centrally located in the cell but the distribution of macrofibrils throughout the cell cytoplasm appears to be less defined than in the other two cell types.
R.C. Marshall et al.
(ii) Meso-cortical cells appear as an intermediate type of cell showing characteristics ranging from ortho-cortex-like to para-cortex-like. They are often characterized by a clearly resolved hexagonal or pseudo hexagonal arrangement of IFs in macrofibrils. In general, macrofibrils are more consistently located along the cell periphery with macrofibril junctions being delineated by differences in IF orientation rather than cytoplsmic remnants. However, many forms occur. The IF/matrix ratio is intermediate. (iii) Para-cortical cells have IFs oriented predominantly parallel to the axis of the fibre without showing high degrees of packing. The IF/matrix ratio is low. Macrofibrils (Chapman and Gemmell, 1971a) are generally large because of fusion during development, few in number and located around the cell periphery. The central regions of the cell contain both the nuclear and cytoplasmic remnants. The basis of these different arrangements of IFs and matrix is unknown but may involve differences in matrix type or amount rather than the structure of the IFs. Electron diffraction studies indicate that the volume of IF as a percentage of the total IF-matrix complex ranges from 33-48% in para-cortical cells to 67-70% in ortho-cortical cells (Dobb, 1970). Ortho-cortical cells have a larger crosssectional area than para-cortical cells (Kassenbeck and Leveaux, 1957; Bones and Sikorski, 1967) by about 30% (6.5+_0.21 vs 8 . 5 + 0 . 2 4 # m 2) (Orwin et al., 1984). Ultrastructural (Chapman and Gemmell, 1971a), autoradiographic (Chapman and Gemmell, 1973), X-ray microanalytical (Carr et al., 1986) and histochemical (Wagner et al., 1983) studies, and chemical analysis of separated cells (Chapman and Bradbury, 1968; Kulkarni et al., 1971; Ito et al., 1985; Dowling et al., 1990) indicate that para-cortical cells contain higher concentrations of high-sulphur proteins than ortho-cortical cells. B. Cortical Cell Type Distribution in Wool
In wool, the numbers of each cell type and their distribution within a fibre cross-section and along
the length of the fibre show marked variation. Ortho-cortical cells are, on average, the predominant cell type, making up >50% of the fibre cross-section and volume (Ahmad and Lang, 1957; Snyman, 1963; Chapman, 1965; Jones, 1966; Bones and Sikorski, 1967; Orwin et al., 1980, 1984, 1985; Tester, 1987; Hynd, 1989). Studies of the distribution of ortho- and paracortex in a fine Merino wool fibre show a bilateral arrangement with approximately half the fibre cortex being composed of each cell type (Horio and Kondo, 1953). A wide variety of cell proportions and arrangements have been found in other wool types (Fraser and Rogers, 1955; Orwin et al., 1984; Orwin and Bailey, 1988). Highly crimped wool fibres are usually associated with a well defined bilateral segmentation of the cortex with the ortho-cortex on the outside of the wave. In poorly crimped fibres, para-cortical cells are partly replaced by meso-cortical cells. It has been generally assumed that the bilateral segmentation is the cause of crimp, but there is evidence that this segmentation is not the sole cause of crimp and may not even be involved at all (Campbell et al., 1972, 1975; Kaplin and Whiteley, 1978). For fibres of any one diameter within a wool sample, a wide range of cell type distributions and proportions are found. In general, for wool samples from sheep of different breeds or sheep of the same breed, as the diameter increases so does the proportion of ortho-cortical cells (Fig. 12). This relationship occurs in two forms, linear and curvilinear, although the slope of the relationship can vary between individual sheep (Orwin et al., 1980, 1984, 1985). As the proportion of orthocortical cells increases, the proportion of mesoplus para-cortical cells decreases, with a partial substitution of para-cortex by meso-cortex in fibres of greater diameter (Kaplin and Whiteley, 1978; Orwin et al., 1984). The relationships described above have been found in sheep fed to minimize body weight change and, therefore, presumably represent genetic differences between sheep (Orwin et al., 1985). Other evidence has shown that nutrition may change these relationships. For instance, general nutritional stress seems to increase the overall
Mammalian Hard Keratin
o g++ ,1: 0
Cortex diameter (,urn)
Fig. 12. Relationshipsbetween the proportion of ortho-cortex and fibre diameter (from Orwin et al., 1985). (a) Linear relationships with different,slopes for wools from two Romneysheep. (b) Curvilinearrelationships with differentslopes for wool from two Romneysheep. proportion of ortho-cortex in fibres (Orwin et al., 1984; Hynd, 1989). Various studies have suggested that cortical segmentation is determined in the bulb (zone A) and depends in part on the degree of deflection of the bulb (Fraser, 1964; Chapman and Gemmell, 1971a, b). However, the means by which individual follicles can change the proportions and arrangements of cell types are unknown.
C. Cortical Cell Types & Other Animal Fibres
A wide variety of other animal fibres have been studied to determine whether differences in cortical structure occur. Examination of camel, yak, alpaca, vicuna, llama, human, raccoon and dog hair fibres, cashmere, mohair and pig bristle has shown some general features (Fraser and MacRae, 1956; Kaplin and Whiteley, 1985; Tester, 1987; Tucker et al., 1988). Aggregation of IF/matrix into macrofibrils is common to all, and cell types similar to the ortho-, meso- and para-cortical cells of wool have been consistently recognized. Some fibres may consist of only one or two cell types, for example human hair (Kaplin and Whiteley, 1985), while the bilateral arrangement may predominate in other samples, for example yak and camel hair (Tucker
et al., 1988). Measurements of several cashmere samples found that, like wool, ortho-cortex was predominant (Tester, 1987). Some IF/matrix arrangemnts differ to that found in wool. These variants include the presence, in Samoyed dog fur and pig bristle, of cells containing mainly smaller macrofibrils whose IF packing was regular rather than the expected whorl-like arrangement. In raccoon fibres, cells with fused macro fibrils showed regular IF packing rather than less ordered para-cortex arrangements found in wool (Kaplin and Whiteley, 1985). D. Structural Differences Between Hard Keratins
As discussed previously, hard keratins are found in a wide variety of mammalian epidermal appendages, which, in spite of their differences in gross anatomical structure, have some common fine structural and developmental features (Fraser et al., 1972; Fraser and MacRae, 1980; Chapman, 1986). These include: (i) Interaction with dermal components often as specialized regions of the derivative, e.g. dermal papillae. (ii) A basal region or .layer of undifferentiated stem cells capable of mitosis. Continuous growth
R.C. Marshall et al.
is often a feature of hard keratin forming derivatives. (iii) Cell shape changes. These are generally specific to a derivative and seem basic to the mechanical functioning of the derivative. (iv) IF orientation is often associated with cell shape and in conjunction with matrix, gives rise to the major mechanical properties of the derivative (Fraser and MacRae, 1980). In spite of these similarities, many keratins differ significantly from wool. For example, in quills and some medullated fibres such as the tactile whiskers of lions and tigers, filament orientation may deviate from being parallel to the direction of fibre growth especially in the cortical cells surrounding the medulla (Makinson, 1954; Earland et al., 1962a, b). Nails have a complex layered structure of a thin dorsal plate overlying a thicker intermediate plate. A thin ventral plate is added during nail formation from the digit tip (Baden and Fewkes, 1983; Chapman, 1986). From a population of stem cells, nail cells differentiate to become oriented parallel to the nail surface and elongated at right angles to the direction of growth (Forslind, 1970). The cell margins are relatively straight in the dorsal plate but show many interdigitations in the intermediate plate. Intercellular material is derived from membrane coating granules (Hashimoto, 1971), a major difference from wool and hair (see Section III.C.4). Filament orientation in human nails follows cell shape and is generally at right angles to the direction of growth. X-ray diffraction patterns indicate that IF orientation in the dorsal plate is less ordered than in the intermediate plate (Forslind, 1970). Mammalian nails, which are intermediate in shape between human nail and claws, show IF orientations which are also intermediate (Baden, 1970). The orientation of IFs in nails is considered less marked than in hair (Baden, 1970) and this may arise from the presence of 10-20% soft keratin within the nail plate (Lynch et al., 1986). Mammalian claws are more curved, longitudinally and laterally, than nails, and extend well beyond the end of the digit. They are also multi-
layered with a superficial and deep layer of hard keratin (Clark, 1936; Thorndike, 1968; see Chapman, 1986, for a review). Filament orientation ranges from being parallel to the direction of growth to intermediate angles in some primate claws (Baden, 1970). The hooves of the horse and ox are complex histological structures (Bertram and Gosline, 1986, 1987; Chapman, 1986). The hoof wall is multilayered with a thin external layer, a middle layer (stratum medium) and an internal lamellar layer. The stratum medium makes up the bulk of the hoof wall and is derived from mitotically active epithelial cells around a number of dermal papillae and also the epidermal-like areas between. Those cells associated with dermal papillae give rise to a tubule of hard keratin-containing cells, in the centre of which is a column of medulla-like cells. The cells of the tubule are flattened and arranged concentrically around the medulla-like core (Ryder, 1962; Bertram and Gosline, 1986). Between the tubules, the epidermal-like areas give rise to the intertubular regions which consist of plate-like flattened cells whose longitudinal axis is parallel to the direction of growth of tubules. The predominant filament orientation follows the longitudinal axis of the cells, being parallel to the direction of growth in the tubules and at right angles to the direction of growth in the intertubular regions (Bertram and Gosline, 1986). This arrangement affects fracture propagation, such that longitudinal cracks (parallel to the direction of growth) are diverted into the intertubular regions, i.e. at right angles to the direction of growth. Horns in mammals usually grow on a bony core, as in Bovidae, although this can be rudimentary, for example as with rhinoceros horn. Horn arises from dividing epithelial cells around dermal papillae. Some cells give rise to columns of medulla-like cells which extend above the top of dermal papillae into the horn material. The majority differentiate into hard keratin containing flattened cells arranged concentrically around the medulla-like cells (Lyne and Hollis, 1973; Chapman, 1986). The predominant direction of the filaments in these cells seems to be at an angle of about 20-30 ° to the growth direction (Makinson, 1954).
Mammalian Hard Keratin Rhinoceros horn shows similarities to hoof wall in that it is composed of columns of tubular, hard keratin with a small amount of intertubular keratin between them. The central region of each tubule contains a column of medulla-like cells extending from the top of dermal papillae (Ryder, 1962). The orientation of the flattened cells in the tubules and of the filaments they contain is similar to that of other horns (Makinson, 1954). Baleen has a complex layered structure with a poorly defined orientation of filaments within the cells (Makinson, 1954; Gillespie, unpublished observations). E. Abnormal Fibre Structure
Abnormal hairs can exhibit a wide variety of aberrations involving changes in texture (for example woolly hair), in physical structure (for example bamboo hair), and in strength and chemical composition (for example the cystine-deficient hair of trichothiodystrophy) (reviewed by Price, 1979). Three mutations which cause major changes to groups of constituent wool and hair proteins are discussed in Section VI.C.2. Most commonly, aberrant hairs are the result of a mutation but some follow administration of chemicals, for example epidermal growth factor (Hollis et aL, 1983) (Fig. 13). VI. STRUCTURE AND VARIABILITY OF KERATIN PROTEINS A. Constituent Polypeptides o f Intermediate Filaments
All hard keratin IFs studied so far appear to be assembled from two families of polypeptides (Type I and Type II) which are of comparable size, charge and amino acid composition (Gillespie, 1983; Marshall, 1986). Considerable homology between comparable polypeptides from widely disparate keratins has been shown (Hewish et al., 1984). In the case of wool and mouse hair, each family contains four polypeptides (Bertolino et al., 1988; Sparrow et al., 1989), while for other keratins, the number is not known with certainty and may be somewhat different. While Heid et al.
(1986) reported that the number of IF polypeptide species in human hair was the same as for wool, Marshall (1983) reported that different samples contained either seven or eight polypeptides, while corresponding nail samples from the same individuals had eight or nine polypeptides, respectively. An analysis of the amino acid sequences from four wool polypeptides, two each from the Type I and II classes, has shown that they are built to a common plan illustrated in Fig. 14. Each component has a blocked N-terminal amino acid, probably N-acetylalanine, followed by a nonhelical domain, a rod-like central domain interrupted at three positions by short segments of non-helical residues and finally a C-terminus non-helical domain. In the rod domain the sequences show a succession of similar heptapeptide (heptads) sequences in which the first and fourth residues are usually hydrophobic. The helix-rich segments of the two wool families show considerable homology between themselves and with the helical regions of cytokeratin IFs. It is not surprising therefore that Weber et al. (1980) found an antibody raised against wool merokeratin (the helical portion of the IF) cross-reacted with IFs of some cytokeratins. The N- and C- terminal nonhelical regions which contain most of the halfcystine and proline, do not show much homology between wool families and none with the equivalent regions of other cytokeratin IFs (Conway et al., 1989; Sparrow et al., 1989). The formation oflFs can be visualized as a three step process: (i) The interaction of one Type I and one Type II polypeptide to form an obligate heterodimer with the chains parallel and in axial register (Fraser and MacRae, 1985); (ii) The aggregation of two dimers in antiparallel direction and out of register to form a tetramer (Woods and Inglis, 1984; Sparrow et al., 1989); (iii) The formation of an annulus (the IF) by the helical assembly of about eight tetrameric units per turn with an axial translation of 19.8 nm between successive units (Fraser and MacRae, 1985; Conway et al., 1989).
R. C, Marshall et al.
Fig. 13. Scanning electron micrographs of wool prior to, and following EGF infusion of sheep. (Reproduced from Hollis (1983), with permission of the publishers.) (a) Normal cuticular pattern prior to infusion. (b) Abnormal cuticular pattern following infusion.
B. Matrix Proteins 1. High-sulphur proteins A heterogeneous cystine-rich protein fraction is found in all hard keratins in relative proportion ranging from below 10% to about 50%. The highsulphur proteins are an example of a group of proteins of apparently similar function that exhibit an extraordinary degree of heterogeneity in both molecular size and chemical composition. The proteins are variable in half-cystine content, with a range from 12.5 residues% for a wool component to 35 residues% for a mouse hair component (Marshall and Gillespie, 1976a, b; Gillespie, 1983). q
The major components fall within a molecular weight range of 10,000 to 30,000 (Gillespie, 1983). Wool and hair high-sulphur proteins typically contain significantly more half-cystine and proline but less aspartic acid, glycine, leucine, tyrosine and phenylalanine than the horny keratin(s) highsulphur proteins from the same animal species (Gillespie and Marshall, 1977; Marshall and Gillespie, 1977). In general, for animals producing two forms of horny keratin (for example hoof and horn), the compositions of the high-sulphur proteins from the horny keratins resemble each other more closely than they resemble the comparable hair fraction. Domain
L12 2A L2
Fig. 14. Schematic diagram of an IF polypeptide showing substructure. Hatching defines segments having ~t-helical structure. The attached data shows the number of amino acid residues in each region of two Type I polypeptides (8c-1 from wool and M59K from mouse epidermis), and two Type II polypeptides (7c from wool and M67K from mouse epidermis). (The diagram and data are adapted from Conway et al. (1989) and Sparrow et al. 1989.)
Mammalian Hard Keratin The heterogeneity of the high-sulphur protein fraction is readily illustrated by the twodimensional electrophoretic pattern of the wool fraction where 20 or more components are separated in addition to the unresolved ultra-highsulphur proteins (Fig. 6). The full extent of the heterogeneity is unknown but at least sixty well-defined fractions in Merino wool have been reported on the basis of combinations of chromatographic and electrophoretic analyses (Joubert and Burns, 1967; Gillespie, 1983). Although the number of individual high-sulphur components within a hard keratin is high, the system is not as complicated as this indicates. The components fall into a limited number of families, perhaps six, with considerable intra-family heterogeneity involving loss of segments and point mutations (Swart et aL, 1976). For example, the 6 members of the SCMKB2 family encompass chain lengths ranging from 151 to 171 residues with an additional 13 amino acid replacements (Lindley, 1977). Amino acid sequences are known for a number of high-sulphur components (Swart et al., 1976). Most of these are from sheep wool with a limited number from mohair showing close similarities. An examination of the sequences of the wool high-sulphur proteins shows that there are three main classes: (i) BIIIB family with a molecular weight of
11,000, (ii) BIIIA family, molecular weight 16,000, and (iii) B2 family, molecular weight 19,000. All the components have half-cystine as the carboxy-terminal residue, and, except for a few 16,000 molecular weight components, have a blocked amino-terminal residue of N-acetyl alanine. There is little sequence homology between the families, except that the 16,000 and 19,000 molecular weight families (but not the 11,000 family) contain a repetitive half-cystine-rich pentapeptide which takes the form cys-cys-x-pro--y. Glutamine, glutamic acid or arginine frequently fill the 'x' position, and 'y' is often serine or threonine. In the major wool protein SCMKB2A of the 19,000 molecular weight family, about 85% of the molecule is constructed of this repeating EMR 4/I--D
sequence (Swart et al., 1976; Lindley, 1977). The possible structural significance of the pentapeptide repeating unit was discussed in the section on disulphide bonds (Section III.E). 2. Ultra-high-sulphur proteins
In addition to the normal set of high-sulphur proteins, hard keratins usually contain a variable amount of a distinctive group of proteins very rich in half-cystine, the UHS proteins (Table 1). Changes in the amounts of these proteins are responsible for the variations in the half-cystine content of wool caused by dietary or physiological manipulation of the sheep (Marshall et al., 1980, 1985b). The UHS proteins were originally detected as a peak in moving boundary electrophoresis patterns (GiUespie and Reis, 1966), and more recently as a diagonal wedge of unresolved protein in twodimensional gels (Gillespie and Marshall, 1983). Extensive fractionation of the proteins from mouse hair and wool indicated extreme heterogeneity in the UHS proteins and an almost continuous distribution of components (Lindley et aL, 1967; Marshall and Gillespie, 1976a, b). This accounts for the lack of specific ultra-high-sulphur protein bands and smeared background in gel electrophoretic patterns (Marshall et al., 1985b). Although pure protein components have not been obtained, it has been possible (MacKinnon, 1989) to derive limited sequence information by nucleic acid procedures (Section III.C.I). 3. High-tyrosine proteins
Many hard keratins contain a group of small proteins (molecular weight less than 10,000) rich in glycine, tyrosine and serine. These proteins are usually referred to as high-tyrosine proteins, and may be divided into two classes (Type I and Type II) on the basis of solubility and of amino acid composition, especially in the contents of half-cystine and phenylalanine (Table 1) (Gillespie, 1972a; Marshall et al., 1980). Both classes are heterogeneous, but only the Type I class has been examined in detail. Within this class, at least 30 components have been identified with the content
R . C . Marshall et al.
of tyrosine ranging from 7 to 20 residues%. Wide differences in the contents of many other amino acids are also found (Gillespie, 1972b; Gillespie and Frenkel, 1974a; Marshall et al., 1980). The amino acid sequences of two Type I class proteins have been determined by both conventional (Dopheide, 1973; Marshall et al., 1980) and DNA sequencing (Kuczek and Rogers, 1985). The sequence homology between these two is low, with only two small blocks of homology being separated by a region of random sequence. The content of high-tyrosine proteins in different keratins varies considerably, and seems unrelated to function. Some hairs and rhinoceros horn contain little or no high-tyrosine protein, while mouse hair and echidna quill are rich in these proteins (20% and 40%, respectively). Within the same type of keratin, a wide range is found in the amount of high-tyrosine protein, for example virtually no high-tyrosine is present in wool from Lincoln sheep while some samples of wool from Merino sheep may contain about 14% of these proteins (Gillespie and Frenkel, 1974b; Gillespie, 1983). Although the mammalian high-tyrosine proteins are similar in composition to the glycinetyrosine-rich proteins of avian beak, claw and scale, they appear to have no sequence homology with them (Marshall et al., 1980; Gregg et al., 1984). C. Variability in Keratin Composition 1. Molecular basis of variability
Hard keratins are not materials of unique amino acid composition; this variability stems from a number of factors. Keratins of one structural type, for example hair, can differ in composition between species, different animals within a species, between sites on one animal, or at different sampling times with one animal. Such observed differences in composition are due to alterations in the ratio of IFs to matrix, changes in the relative proportions of the two matrix protein groups, and variations in the constituent components of the matrix proteins. Variations in keratin composition between animal species also stem from the expected species specificity of
proteins (Gillespie and Frenkel, 1974b; Marshall et al., 1985a). 2. Genetic control
Although the relative proportions of contituent proteins in individual keratins are variable, a specific keratin type from each species appears to have its own characteristic compositional range. For example, human hair always contains large amounts of high-sulphur protein but little or no high-tyrosine protein, whereas mouse hair is moderately rich in both protein types. In general, the proteins expressed in a hard keratin, apart from some polymorphism, appear to be virtually constant within animals from the same species. Thus electrophoretic analysis of the constituent proteins of hard keratins may be used to discriminate between different species. Moreover in the cases of the proteins of wool, goat hair and human hair, sufficient genetic polymorphism exists to allow limited discrimination between individuals (Marshall et al., 1985a, b). In studies of different keratin types from the one animal, a number of workers have observed that the same constituent proteins are present although the relative proportions of protein components can be quite different (Baden et al., 1973; Gillespie and Marshall, 1977; Marshall and Gillespie, 1977). A comparative study of human hair and nail using highly resolving electrophoresis confirmed this finding for the major high-sulphur proteins, but found that hair contained an additional unique IF polypeptide (Marshall, 1983). High-sulphur proteins appear to have had a complicated genetic history involving gene duplication and amino acid replacements and deletions (Lindley, 1977). In the last decade, considerable interest has been shown in the IF polypeptides of hard keratins, which also have had a long genetic history, especially in the relation between the a-helical segments of these polypeptides and the corresponding regions of cytokeratin IFs (Blumenberg, 1989). Certain rare abnormal hairs stemming from genetic defects, may present very different patterns
Mammalian Hard Keratin of proteins (Gillespie and Marshall, 1989). Of the many hairs showing abnormal fibre structure (Section V.E), only three examples have been recorded with major changes to the constituent protein components: (i) The felting lustre mutant of the Merino sheep demonstrates complete suppression of the hightyrosine proteins of wool (Gillespie and Marshall, 1989). (ii) Trichothiodystrophy in man results in the loss of many normal high-sulphur protein components with replacement by components of lower than normal half-cystine content (Gillespie and Marshall, 1983; Gillespie et al., 1988, 1989). (iii) In the heterozygous naked mouse mutant, the high-tyrosine proteins are reduced by 50%, but a major disruption to the pattern of synthesis of both the matrix and IF polypeptides is observed in the homozygote (Raphael et al., 1984). In the cases of felting lustre mutant and trichothiodystrophy, the mutations may be affecting genes which regulate stem cell differentiation in the follicle. 3. Dietary control
Within certain limits, the half-cystine content of wool, and hence the relative proportions of its high-sulphur proteins (in particular the UHS proteins), are susceptible to dietary manipulation. The half-cystine content of wool increases by as much as 45% when cysteine, methionine or proteins rich in these amino acids are infused into the abomasum of sheep to avoid degradation by rumen microorganisms (Gillespie and Reis, 1966). Similar but smaller changes occur with alterations in the level of normal nutrition (Reis, 1979). The process of sulphur-enrichment appears to involve a reversible, rapidly-acting biological control mechanism (Gillespie and Reis, 1966) which, the work of Orwin et al. (1984) suggests, may operate by controlling the differentiation of follicle stem cells. These workers observed that as the level of nutrition changed so did the relative proportion of ortho- and para-cortical cells. The accompanying changes in wool composition are consistent
with the respective differences in composition between these cells. Substantial changes in wool protein composition can also be induced by feeding sheep imbalanced mixtures of amino acids, the mechanism of this control being unknown (Reis, 1979). 4. Physiological Control
Chemical substances such as epidermal growth factor (EGF), cyclophosphamide, ethionine, methoxinine, N-[5-(4-aminophenoxy)pentyl]phthalimide temporarily inhibit follicle activity and can markedly affect the composition and physical properties ( for example, see Fig. 13) of the wool grown after the treatment. This wool has an increased content of high-sulphur proteins, largely UHS type, and an almost complete suppression of high-tyrosine proteins (Gillespie et al., 1980, 1982; Reis and Gillespie, 1985). For example, the administration of 27.5 mg mouse EGF to a Merino sheep (Fig. 15, Table 3) causes the temporary inhibition of the activity of wool follicles and the shedding of the wool. After a delay of a few weeks, wool regrew and its composition changed progressively with time, reaching a maximum divergence from the control in the 3-4 week period. The composition progressively returned to the pre-treatment composition by about 10 weeks. Over this period, the proportion of high-sulphur proteins increased from 19% to a maximum of 30%, while the high-tyrosine protein content decreased from 12% to less than 5% (Gillespie et al., 1982). At present, the complex changes in wool protein synthesis which follow the temporary cessation of follicle activity cannot be fully explained. However, it is interesting to note that follicles in sheep and mice regenerating after chemical treatments or plucking produce wool and hair which is similar in composition to that produced in newly operating follicles in young lambs and mice, that is, deficient in high-tyrosine proteins but enriched in UHS proteins (Marshall et al., 1980). Because the tip of lamb's wool, which is largely para-cortex, has a similar composition (Nott and Sikorski, 1965) it is possible that physiological controls operate at the cellular level in follicle differentiation.
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5 - 6 WEEKS
Fig. 15. Two-dimensional electrophoretic patterns of wool proteins showing the effect on wool composition of a single subdermal infusion into sheep of 27.5 mg of EGF. (a) Pretreatment wool. (b) Regrowth wool (3~4 weeks). (c) Regrowth wool (5~5 weeks). First dimension (horizontal) in 8 M urea at pH 8.9. Second dimension (vertical) in SDS at pH 8.9. (Reproduced from Gillespie et al. (1982), with permission of Williams and Wilkins.)
VII. C O N C L U D I N G S T A T E M E N T S
In spite of the tremendous advances made in keratin research over the last thirty years, there are still large gaps in our knowledge, particularly of the biochemistry of the follicle. It is clear that the exact sequence of events in the synthesis of keratin proteins, their assembly into
the IF/matrix complex, the interactions between various proteins and assemblies of proteins, and the crosslinking between them are still matters of conjecture. We do not know whether the polypeptides of the IF are cross-linked to each other, although this now seems somewhat unlikely (Fraser et al., 1988), or cross-linked to other proteins in the complex. Although we know
Table 3. Amino acid (as residues%) and protein (grams per 100 grams wool) compositions of wool from sheep before and after administration of epidermal growth factor (Gillespie et al., 1982) Amino acid
5 ~ weeks
Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine
3.0 0.8 7.1 7.0 5.7 9.7 11.7 6.1 9.4 5.6 9.2 5.6 0.5 3,0 8.3 4.3 3.0
2.7 0.8 7. I 6.1 6.7 10.8 11.6 7.3 7.2 5.3 12.8 5.6 0.4 3.0 7.3 3.1 2.4
3.0 0.8 7.2 6.8 5.9 10.3 11.7 6.2 8.7 5.5 10.0 5.5 0.5 3.1 8. I 4.0 2.9
High-sulphur protein High-tyrosine protein
Mammalian Hard Keratin that c o p p e r is needed for part of the o x i d a t i o n process, we do n o t yet u n d e r s t a n d the mechanism(s) i n v o l v e d (Gillespie, 1990). A d v a n c e s in these fields have been h i n d e r e d by the difficulties experienced in o r g a n or tissue cultures o f follicle a n d in the great difficulties experienced in satisfactorily r e c o n s t i t u t i n g h a r d keratin IFs. c~-Keratins are u n i q u e a m o n g s t tissues in exhibiting very large variabilities in p r o t e i n composition which p r e s u m a b l y do n o t interfere with function. W e do n o t k n o w o f a n y reason for this p h e n o m e n o n . A l t h o u g h it was first systematically studied 25 years ago, the m e c h a n i s m by which it operates is still n o t u n d e r s t o o d a l t h o u g h there is some evidence that it takes place at the level of stem cell differentiation. I n the case of wool it is assumed, w i t h o u t m u c h solid evidence, that the o r t h o / m e s o / p a r a types of cortical cells differ in protein c o m p o s i t i o n a n d that dietary, genetic or physiological m a n i p u l a t i o n of wool c o m p o s i t i o n operates by v a r y i n g the p r o p o r t i o n s o f these cells. There is a n u r g e n t need for reliable i n f o r m a t i o n o n the p r o t e i n c o m p o s i t i o n of various cortical cell types, n o t only from wool, b u t also from other hairs. The s t r u c t u r a l complexity of m a m m a l i a n k e r a t i n a n d the large store of genetic i n f o r m a t i o n needed in its synthesis is very i n t r i g u i n g when we consider that a bird c a n m a k e a f u n c t i o n a l claw from only one m o d e r a t e l y p o l y m o r p h i c protein, in c o n t r a s t to the h u n d r e d or m o r e proteins needed by a m a m m a l . W e can ask whether the m a m m a l i a n claw is f u n c t i o n a l l y superior as a result o f its complex structure and, if not, what has been its survival value.
Acknowledgements--We thank Ms J. L. Woods for her help in the preparation of the manuscript, especially in providing the electron micrograph figures. Our thanks are also due to Dr L. N. Jones and Mr K. F. Ley for their comments on the manuscript.
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