Molecular packing in elastoidin spicules

J. Mol. Biol. (1978)

118. 567478

Molecular

Packing in Elastoidin *JOHN WOODHEAD-GALLOWAY

Spicules f

Department of Rheumatology, University of Manchester Stopford Buildi~ng, Manchester M 13 9PT. Englad DAVID

Department of Medical Stopjord Building.

W. I,. HUKIKS$

Biophysics. Manchester

DAVID

C:niversity of Manchester Ml3 9PT. England

P. KNIGHY

Department of Biology, King Alfred’s Collolleg~ Win.chester SO2 2N R. England PENELOPE

A. MACHIS

Atlas Computing Division Rutherford Laboratory Chilton, Oxon OX17 OQY: England AND JAQUELINE

B. WEISS

Department of Rheumatology, University of Manchester Stopford Building. Manchester Ml3 9PT. England (Received 28 March, 1.977. ad

in, revised form, 72 September 7977)

Low-angle X-ray diffraction shows that, despite the well-defined regular axial11 projected structure, there is no long-range lateral order in the packing of molecules in native (undried) or dried elastoidin spicules from the fin rays of the Squulua acanthias. The equatorial intensity distribution of the X-ray spurhound diffraction pattern from native elastoidin indicates a molecular diameter of 1.1 nm and a packing fraction for the structure projected on to a plane perpendicular to the spicule (fibril) axis of 0.31 (the value for tendon is much higher at around O-6). Density measurements support this interpretation. When the spicule dries the packing fraction increases to 0.43 but there is still no long-range order in the structure. The X-ray diffraction patterns provide no convincing evidence for an) microfibrils or subfibrils in elastoidin. Gel electrophoresis shows that the three chains in the elastoidin molecule are identical. The low packing fraction for collagen molecules in elaatoidin explains the difference in appearance between electron micrographs of negatively stained elastoidin and tendon collagen. In elastoidin, but not in tendon collagen, an appreciable proportion of t,he stain is able t,o penetrate between the collagen molecules.

t Present address: Medical $ To whom correspondence

Research Council, 20 Park Crescent,, London should be addressed. 56;

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4AL, England.

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Introduction

Elastoidin spicules form needle .shapcd fish fin rays and appear to he very wide (up to ahout 0.5 mm in Squwlus ucanthias, single collagen fib& tht ’ source of our material) (MeGavin, 1962; Woodhead-Galloway & Knight, 1977). In contrast. collagen fihrils in. for example, tendon have diameters of the order of only lo2 nm (Fitton *Jackson. 1968). Woodhead-Galloway & Knight (1977) have confirmed that in axial projection the arrangement of molecules is the same as in rat, t,endon and (Aher collagen fihrils (Hodge & Petruska, 1963). Both low-angle X-ray diffraction patterns and electron micrographs of elastoidin have been published previously and have provided information on its axially projected structure (Bear. 1952; McGavin, 1962; Wrag, 1972; Woodhead-Galloway 8~ Knight, 1977). We now present the first detailed analysis of the lateral arrangement of molecules and discuss its implicat’ions for the three-dimensional packing of molecules in the elastoidin spicule. Our analysis is aided by density measurements and electron micrographs; gel electrophoresis provides information on the chain composition of the molecules. We also indicate the implications of our results for the structural analysis of other collagen fihrils.

2. Materials and Methods (a) Materials

The largest spicules (about recently killed or deep frozen

O-5 mm (-ZO’C!)

diameter) were removed from the pectoral fins of adult specimens of the spurhound S. acunthias

(Woodhead-Galloway & Knight, 1977). Spicules were scraped and thoroughly washed in distilled water. Most were colourless and transparent but some had a slight yellowishbrown colour. Experiments were either performed immediately or on material stored in distilled water in the cold (5°C). Tendons used for comparative studies were dissected from the tails of freshly killed, adult albino rats. Collagen tape, also used for comparative studies, was manufactured by Ethicon Inc. (Somerville, N.J., U.S.A.) and given to us by Dr J. A. Chapman. Where necessary olastoidin spicules were dried at’ room humidity (-5,59/o) and temperature (m2O’C) under slight tension (0.1 N). (b) Gel electrophoresis

and hydroqproline

content

Two spicules were frozen in liquid nitrogen and disintegrated in a stainless steel mill (Steven, 1967). The resulting powder was heated (in a boiling water bath for 20 min) in urea (2 M) and sodium dodecyl sulphate phosphate buffer (0.01 M; pH 7.2) containing (0.2%). Eleotrophoresis patterns were obtained by the method of Furthmayr & Timpl ( 1971) using phosphate buffer (0.1 M ; pH 7.2) containing urea (2 M) and sodium dodecyl sulphate (O*1o/o). The apparatus was run initially for 15 min at a current of 2 mA per tube and then for a further 3 h at 6 mA per tube. Gels were stained with both Coomassie blue and Periodic Acid Schiff reagents (Smith, 1976). The results were compared with those obtained from type I collagen extracted from calf skin by the method of Jackson & Cleary ( 1967). Hydroxyproline was estimated by a modification of the method of Woessner (1976) using a semi-automated analysis system (Hook and Tucker, Croydon, Surrey). Redistilled propan-2-01 was used as the solvent for p-dimethyl aminobenzaldehyde instead of ethanol. The concentration of this reagent was 10% instead of 20%. Nitrogen content was measured hy Kjeldahl digestion and titration (Cannon et al., 1974) as a means of assaying the rlastoidin by taking the nitrogen content of collagen to be lS*6o/o. Mercaptoethanol reduction was carried out by the met,hod of Chung & Miller (1974).

The density of dried immersed in a mixture

elastoidin was Ineasured by a flotation method. Specimens were* of x,vlene and rarhon tet.rarhloride until they neither sank nor

MOLECULAR

PACKING

IN

ELASTOIDIN

569

floated. The density of the liquid was measured in a specific gravity bottle whose volume (9.76810.004 cm3) was first calibrated by weighing the bottle both empty and filled with distilled water. We used a combination of volume and dry mass determinations to find the mass of protein per unit volume of native elastoidin. The volume of a portion of a spicule was measured by displacement of water in a precision bore (l.OO* 0.01 mm) capillary tube; bores were checked with a travelling microscope and always found to be 1.00 mm. Tubrs were cleaned (with concentrated H,SO, followed by distilled water) and siliconised (with BDH Itepelcote silicone fluid). The length of a thread of distilled water in a tube was measured with a travelling microscope. Surface water was removed from a length of spicule with filter paper and its tapering ends cut off to obtain a cylindrical portion. This portion was immediately introduced into water in a capillary tube and its volume calculated from the length of the thread of water and the tube bore. The specimen was then dried to constant mass in an oven (1OO’C) and the mass of protein per unit volume of nativr, elastoidin calculated. (d) X-ray

diffruction

and electron

microscopy

X-ray diffracticn patterns were obtained using an evacuated Searle camera (MarconiElliot Avionic Systems Ltd., Borehamwood, Herts) equipped with two Franks (1958) mirrors to bring the beam to a point focus. Wet spicules were enclosed in a Perspex cell with 12 pm Melinex windows containing air in contact with distilled water. Equatorial intensity distributions on the diffraction patterns were measured with a Joyce-Loebl recording microdensitometer Mk IIIC (Joyce-Loebl and Co. Ltd, Gateshead); care was taken to ensure that the optical density did not exceed 1, to ensure that the response of the (Kodirex) X-ray film was linear. The specimen-to-film distance (about 10 cm) was obtained from the sharp meridional diffraction pattern of the native, undried elastoidin which corresponds to orders of a 66.8 nm grating (Woodhead-GallowTay & Knight, 1977). Electron micrographs of dispersed and negatively stained elastoidin and tendon were obt,airrcd by the methods described previously (Woodhead-Galloway & Knight, 1977). (e) Calculation

of equatorial

X-ray

inten,sity

distribution

The diameter, 2R, of the collagen molecule is customarily taken to be between 1.0 and 1.5 nm which is of the order suggested by molecular models. As a first approximation we suppose that the projection of the molecular structure perpendicular to its axis is a disc of this diameter, whatever the 3-dimensional arrangement of molecules may be. The equatorial X-ray intensity distribution yields information about t’he arrangement of molecules in the spicule. Results which we shall discuss later suggest that collagen molecules are packed in the spicule with no long-range lateral order. The Fourirr t#ransform of a single projected molecule (a disc) is given by F(K)

= 2nR2 J,(KR)/KR,

(1)

where K = 2~1. Here 5 is the cylindrical polar radius in reciprocal space (i.e. corresponds to the distance along the equator and is equal to the reciprocal of a spacing measured in the object) ; throughout t’his section J, will denote a pth order Bessel function of the first kind. If the e-dimensional packing fraction, 7, of discs is greater than about 0.22, interference effects will occur between scattered X-rays (Woodhead-Galloway & Machin, 1976a). The intensity of X-rays scattered by an irregular arrangement of discs is then given hy I(K)

= NI,X(K)F2(K),

(2)

where I, is the intensity incident on the specimen, N is the total number of discs and S(K) is an interference function. Since neither I, nor N can he measured simply we obtain I on an arbitrary scale. S(K) may be calculated from S(K)

= 1 + n j{g(r)

-

1 } J,(iKr)dr,

(3)

570

J

W 0 0 D HE A 1) G A L L 0 \f. A Y E 7’ rl I,

where 12 is the number of projected molecules (discs) per unit area (~~:oodtlead-U~~ll~~~~~,~ & Machin, 1976a,b) and i is d-1. The radial distribution function, or pair correlation function, g(r) is the probability of finding some other disc per unit r7rt.a at a dixt,ancts 1’ from an arbitrarily chosen reference disc. \~roodllead-Galloway & Machirl ( 1!)76a) 11a~cs developed a theory for calculating g(r) from R and n analogous to that for t,he liqllitl state> (Woodhead-Galloway et aZ., 1968). 1%’a note that it and 11are related b> 7 = xR%.

(4)

The assumptions we have made are (1) that molecules may be treated as cylinders and (2) that we may neglect scattering from wat,er in the structure. \%‘c shall generally kc forced to make simplifying assumptions about molecular st,ructurr when calculating ttlr: intensity distribution to be expected from models of fibrous protein structure. because thra molecular structures are usually not known in detail and h-cause of t,tle magnitltde of the computing which would arise in taking account of sequence information and variable side-chain conformation (H&ins, 1975). 7’11~ approximation t,hat moltcnl~ are cylinders has been used before (Fraser et al., 1964) and would appear to be reasonable bccaus~ \VV are using low-resolution data, which are insensitive t,o details of molccldar strnctllrc~, arltl because diffraction data from fibres are cylindrically averaged. In any case collagen molrtcules are expected, because of thoir structure (Rich & Crick, 1961), t,o l~evc a reasonably uniform electron density distribution in projection. Trial calculations (based ot, the co-ordinates of the keratan sulphate molecule, taken from Arnott et al., 1974) show t,hat# this approximation is reasonable for predict’ing equatorial pcnk positions, cvc’n for a molecule which does not resemble a cylinder. The effect of water on fibre diffraction patterns has been considered by Langridge et al. (1960), Fraser et nl. (1965) and Arnott PL Hnkins (1973). Although Fraser et al. found quantitative cllangcs in peak posit,iona whrsn water was taken into account, water did not IlaT-e sucll a marked qualitative ef+ct~ as the kind we wish to explain when comparing diffraction pat,terns frorn nati\.c and dried elastoidin. The smallness of the changes just’ifies thr> approach used by l’ras~~r Kr Macltac(1958) and discussed by them subsequently (Fraser 8: MacRae. 1973), whi& amolmts to Ilrglect of any rnedium surrounding thr protein wlrc~l considering tlc(ll;rtorial diff’ract,iorl at low angles from large (at the atomic level) cylinders. We accept that we have adopted a simplified model in order to calculate the c quatorial X-ray intensity distribution but we have at,tempted to justify the simplifications mad(L. Some simplification is inevitable when performing calculations for sue11 a complex system as a liquid-like array of real protein moleculrs. A rigorous analysis of X-ray fibro diffraction patterns is probably only possible for rplativclp simple systems (Hnkins. 1975).

3. Chain Composition Figure The

former

1 compares migrates

gel electrophoresis

patterns

of elastoidin

as a single

with

R, (ratio

E band

an

and

type

of distance

I collagen. travelled

by

material in the band to distance travelled by solvent front) value of around 0.35 to 0.36. The latter migrates as two a bands corresponding to the al and a2 chains. Because type I collagen has twice as many al chains this band is more intense than the cc2band; the R, values are 0.33 to 0.36 for the al chain and 0.36 to 0.38 for the ~2. The presence of only one u band for elastoidin shows that its chains are identical. No difference was observed in the pattern after merca,ptoethanol reduction, indicating the absence of sulphydryl bonds. Hydroxyproline was calculated to be llyb of the total protein. This finding is in good agreement with the hydroxyproline content of collagens generally. No band could be seen after staining with Periodic Acid Schiff reagent indicating that the collagen was not highly glycosylated. This result is consistent with the large-diameter fibril (Grant et al., 1969). Our gel electrophoresis results show that the three chains in t’he collagen molecules of elastoidin are identical. Kulonen & Pikkarainen (1970) have previously reported

MOLECULAR

PACKING

IN

ELARTOII)TS

571

B

al a2

(a) F’rc:. 1. It, can be collagen. ident ical

(b)

Gel electrophoresis patterns of (a) spurhound elastoidin and (b) calf skin type I collagen. seen that alastoidin runs as a single a band in contrast to the ctl and a2 bands in type I Elastoidin also shows only a single fi band as would be expected from a collagen with a chains.

that collagen from the body wall of dogfishes consists of only one electrophoretically distinguishable component. Previous investigators have concentrated on tendon as a model system for understanding the structure of collagen fibrils. Tendon collagen is type I, i.e. it has two identical al chains and a third similar. yet distinct, a2 chain; collagens from most other tissues contain other types with three identical chains (Miller, 1973). Concentration on tendon as a model system for understanding the structure of collagen fibrils causes a bias towards type I collagen. Use of elastoidin, with three identical chains, as a model system tends t,o counter this bias.

4. Density Measurements content of native elastoidin of We obtained a value for the protein 0.458&0.009 g cme3. If we assume a value of 3 x lo5 for the molecular weight of collagen (Harrington & von Hippel, 1961), the number of molecules per unit volume is (0.458 x 6.025 x 10z3)/(3 x 105) == 9.2 >( 1017 cmm3. According to the accepted axially projected structure of the collagen fibril (Hodge C Petruska, 1963; see also Electron Microscopy, below) the axially-projected number density is 2.99 >
57.2

J. WOODHEAD-GALLOWAY

h’?’ A I,.

& standard deviation) which is in excellent agreement with t’he values obtained by us for dried tendon (1.34kO.01 g cm-3) and collagen tape (I.34 ) 0.01 g cm s), Using the value of the molecular weight given above. calculation of the expect~ed density of the protein produces results in the range 1.1 to 1.5 g cm m~3depending on the way the volume is estimated. Thus estimation of molecular volume could lead to errors of up to about 20% in density calculations. Decreasing the molecular weight used in our calculations to a value of 2.8x lo5 would yield an increased value for n of 0.33 nmm2. If 20% of the solid material in the spicule were not collagen (see Piez & Gross, 1959) then the calculations would be consistent, with an r~ value of 0.25 nmm2 (for a molecular weight of 3 x 105) or 0.27 nm m2 (for a molecular weight of 2.8 x 105). Conversion between n and 7 depends on the square of R, the molecular radius (eqn (5)), yet R is not known with any great, accuracy (see X-ray Diffraction below) and in any case is an inexact concept for a real molecule (Materials and Methods, section (e)). Thus calculations derived from density measurements are only approximate and are intended to check whether the interpretation of the X-ray diffraction data appears reasonable. Therefore small differences in, for example, our measured density of 1.34 g crnm3 for collagens and the oft-quoted value of 1.41 g cmm3 (which represents a 5% difference) are insignificant. We note that our value of 1.34 g cme3 is identical to that given by Rougvie & Bear (1953).

5. X-ray Diffraction In this section we confine our attention to the equatorial intensity distribution of the low-angle X-ray diffraction patterns given by native and dried elastoidin spicules. These data provide information on the arrangement of rod-shaped molecules projected onto a plane perpendicular to the axis of the spicule. In this projection we view the lateral arrangement of molecules as an arrangement of discs. It has been noted previously (McGavin, 1962; Wray, 1972; Woodhead-Galloway & Knight, 1977) that the native elastoidin spicule gives a diffuse equatorial intensity distribution which falls to zero with no discernible peaks ; signs of a broad maximum have been noted in some patterns by A. Miller (personal communication) but this does not affect our argument since, as we shall see, it simply implies a somewhat increased packing fraction for the laterally disordered array. When the spicule dries the distribution changes to a single diffuse peak centred at about 0.8 nm- l. In Figure 2(a) and (b) we show microdensitometer traces across the equators of typical diffraction patterns from native and dried elastoidin. We can immediately make a deduction from the experimental data presented in Figure 2(a) and (b). In accordance with earlier observations no discrete Bragg reflections are observed so that the arrangements of molecules cannot involve any long-range order. Our initial deduction was confirmed more quantitatively by calculating the equatorial intensity distribution expected from an arrangement of parallel rodshaped molecules with no lateral long-range order (see Materials and Methods, section (e)) and then comparing the results with the experimental data. We obtained the best agreement between observed and calculated curves for native elastoidin when 2R = 1.1 nm and 7 = O-31 as shown in Figure 2(a). This value of 2 R, the diameter

MOLECULAR

0.5

IN

4573

ELrlSTOIDIN

-

I 0.0

PACKING

1

I

I

I

o-5

I

I

I

I

I I.0

( nm-’

2. Comparison of observed (--------) and calculated (-----) equatorial intensity disFIG. tributions of X-ray diffraction patterns from (a) native (undried) and (b) dried elastoidin spicules. Observed distributions are microdensitometer traces. Numerical values of the intensities are in arbitrary units, calculated values are scaled to give the best fit and both are plotted against I, the reciprocal spacing defined in eqn (1). The original X-ray diffraction patterns have been published previously (Hukins et aZ., 1976). The poor fit at low 8 values is in part a consequence of the hard disc approximation (Woodhead-Galloway & Machin, 19766); a similarly poor fit is obtained for liquids when a hard sphere approximation is used (Mikaloj & Pings, 1967).

of the rod, is in reasonable agreement with values for other collagen molecules ob& von Hippel, 1961), modeltained from light-scattering (1.5 nm; Harrington building (1.3 nm ; twice the sum of the maximum j3 carbon atom radius and the van der Waals’ radius of carbon; co-ordinates from Ramachandran & Sasisekharan. 1965) and analysis of diffuse equatorial X-ray scatter from rat tail tendon (I.2 nm: Woodhead-Galloway & Machin, 19763). The value of 7 is in good agreement with that obtained from density measurements (0.29). Figure 2(b) shows that for dried elastoidin there is good agreement between observed and calculated intensity distribution if 2R is kept at I *l nm and 77is increased to 0.43. We conclude that removal of excess water between molecules by drying the

574

I.

WOODHEAD-GALLOWAY

E 2' 1-lL.

elastoidin spicule increases the density of molecular packing so that, q increases from about 0.31 to about 0.43. Figure 3 presents this conclusion pict80rially by illustrating the liquid-like packing of molecules in the native and dried spiculcs projected on to a. plane perpendicular to the spicule axis. There is no need to postulate that the molecules aggregate into microfibrils or subfibrils in order to explain our data. The peak in the equatorial intensity distribution appears to arise from a fairly dense but, nevertheless disordered, lateral packing of single molecules.

(a)

lb)

FIG. 3. Molecular packing fractions in (a) native and (b) dried elastoidin represented as arrays of discs. In this Figure the arrays of discs have no long-range order and have the 2.dimensional packing fractions required to explain the observed curves in Fig. 2. This Figure provides a pictorial representation of the numerical data presented in X-ray Diffract,inn.

It has been suggested by a referee that our results could be explained by a random array of microfibrils. This suggestion requires the first peak in F2(K) to be absent. We have no reason to suppose that the first, and therefore most intense? peak in F2(K) is absent and find the suggestion contrived. Our argument is that the data from both native and dried elastoidin can be explained most convincingly without invoking a microfibril. No evidence has ever been presented to suggest that elastoidin consists of microfibrils.

6. Electron Microscopy We are now able to relate our X-ray diffraction and density measurements to the somewhat unusual appearance of dispersed and negatively stained elastoidin in the electron microscope. Negatively stained collagen fibrils usually appear at first sight to be alternating light and dark stained bands with a period of about 64 nm (Fig. 4(a) and (b)). (We note that X-ray diffraction studies of undried fibrils yield a period of 67 nm.) The conventional explanation for this observation is summarised in Figure 5(a). This Figure shows that the regular axially projected structure consists of “gap” and “overlap” regions. Gap regions can fill with electron-dense stain whereas overlap regions exclude it, so that there is high contrast between gap and overlap regions in the micrographs. The contrast leads to the alternating pattern of light and dark stained bands (Hodge & Petruska, 1963). This explanation has been confirmed by using a composite amino acid sequence derived from calf and rat skin collagen to explain the positively stained pattern of electron micrographs of calf skin collagen (Chapman & Hardcastle, 1974; Walton, 1974). Narrow white bands occur on micrographs of negatively stained specimens, presumably where bulky regions of the

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576

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collagen molecule (e.g. the terminal peptides) exclude almost all &win (Do,vlc P/ ~1.. 1974). We have used the white bands to align microglaphs of negativtaly stSainrd t,cndon collagen with those of elastoidin in Figure 4. It is clear from Figurtl l(c) and (d) that in elastoidin the stain penetrates, with apparently equal ease’. both gap and overlap regions. This observation may be explained by the elastoidin molecules being insufficiently densely packed to prevent stain from collecting between them. As shown in Figure 5(b) if sufficient stain can penetrate between molecules, there will be little

(b)

FIQ. 6. A 2-dimensional representation of the axially projected packing of molecules in the collagen fibril for (a) a relatively high and (b) a lower packing fraction. In (a) most of the electrondense stain will be in the holes of the gap region leading to high contrast between gap and overlap regions and, therefore, to micrographs like Fig. 4(a) and (b). In (b) much of the stain will penetrate between molecules leading to low contrast between gap and overlap regions and, therefore to micrographs like Fig. 4(c) and (d).

contrast between gap and overlap regions. Thus the low density of molecular packing in elastoidin spicules explains why, when dispersed and negatively stained, they do not have the familiar alternating light and dark banding pattern shown by other collagen fibrils.

7. Discussion We have shown that elastoidin is a simpler system than the collagen fihrils of tendon in that there is no evidence for long-range lateral order. The absence of this lateral order simplifies the analysis of our X-ray diffraction patterns ; there is no convincing evidence to suggest that the molecules in elastoidin aggregate into any of the various kinds of microfibrils or suhfihrils which from time to time have been postulated to occur in collagen. Rather we believe that the lateral organisation of the elastoidin spicule is a liquid-like array of single molecules. Our analysis is consistent with density measurements and with results obtained by clect’ron microscopy. X-ray

MOLECULAR

PACKING

IN

ELASTOIT)IN

577

diffraction (Hukins, 1977) and 13C nuclear magnetic resonance studies (Torchia & VanderHart, 1976) indicate considerable lateral disorder in fibrils of some other collagens which are nevertheless highly oriented in the axial direction. An analysis of the diffuse equatorial scatter in the X-ray diffraction patterns from tendon shows that it arises from a liquid-like array of single molecules (Woodhead-Galloway & Machin, 1976h). Our results show that elastoidin spicules have no long-range lateral order and yet, have a precisely defined structure in axial projection. Some theoretical calculations have recently been performed to further our understanding of this system (WoodheadGalloway & Young, 1977). We note that these properties define the spicule to be a type A smectic liquid crystal-this has some interesting implications for the structure of collagen fib& (Hukins & Woodhead-Galloway:. 1977). REFERENCES Arnott, S. & Hukins, D. W. L. (1973). J. iUoZ. Biol. 81, 93- 105. Arnott, S., Guss, J. M., Hukins, D. W. I,., Des, I. C. M. di Rees. D. A. (1974). .I. Mol. Rd. 88, 175-184. Bear, R. S. (1952). Ad~un. Protein Chem. 7, 69-160. Cannon, D. C.. Olitzky, I. bi Inkpen, J. A. (1974). In Clinical Chemistry Principles and Techniques (Henry, R. .J., Cannon, D. C. & Winkelman, W. J ., eds), 2nd edn., pp. 409 4 11, Harp ‘r Row, New York. Chapman, J. A. & Hardcastle, R. A. (1974). Conn. Z’iss. Res. 2, 13-150. Chutlg, E. & Miller, E. J. (1974). Science, 183, 1200. 1201. Doyb, B. B., Hulmes, D. J. S.. Miller, A., Parry, 1). A. D., Piez, K. A. & Woodhead(Galloway, ,J. (1974). Proc. Roy. Sot. ser. B, 187, 37--46. Fitton ,Jacksorl. S. (1968). In Treatise on Collagen (Gould, B. S.. ed.), vol. 2, pp. l-66, Academic Press, London and New York. Franks. A. (1958). Brit. J. Appl. Phys. 9, 349-352. Fraser, K. D. B. & MacRae, T. P. (1!158). Biochim. Biophys. Acta, 29, 229-240. Frasrr, K. D. 1%. & MacRae, ‘I?. P. (1973). Conf ormation in Fibrous Proteins, p. 503, .Icademic Press, London and New York. Fraser, R. D. Is., MaeRae, T. I’. & Miller, A. (1964). J. &lol. Biol. 10, 147-156. FrastAr, K. D. L<., MacRae. T. I’. and Miller, A. (1965). J. Mol. Rd. 14, 432-442. Furt’tlmayr, H. 8: Timpl, H. (1971). Anal. Biochem. 41, 510-517. Grat& M. E.. Freeman, I. L.. Schofield, ,J. D. & Jackson. D. S. (1969). Biochim. Biophys. .3cta, 177, 682-685. Harringtorr, W. F. & van Hippel, P. H. (1961). il&an. Protein Chem. 16, 1-138. Hodg,rcx, A. .J. &, Petruska, J. A. (1963). In Aspects of Protein Structure (Ramachandran, (:. N.. ed.). pp. 289-300, Academic Press, London. Hukins, D. IV:. L. (1975). In Structure qf Fibrous Biopolymers (Atkins, E. D. T. & Keller, A., ctds), pp. 293-305, Butterworth, London. Biophys. Res. Commun,. 77, 3356339. Hukins, D. W. L. (1977). Biochem. Hukills, D. W. L. & Woodllratl-(:allouray, !J. (1977). fiZo1. Cry&. Lig. Cry&. (Letters), 41, 33-39. Biophys. Res. Hukins, D. W. I,., Woodhead-Galloway, .J. & Knight. D. P. (1976). Biochem. Commlrn.

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